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

Summary

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.

Abstract

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

Introduction

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). 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 synaptogenesis^3,4. Other types of lipids have shown promise in medical applications, including sphingolipids utilized in drug delivery systems and saccharolipids used to support the immune system^5,6. The numerous roles and potential therapeutic applications of lipids in the biomedical field have made lipidomics—the study of the pathways and interactions of cellular lipids—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 techniques^7,8. 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 compartments^9,10.

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 LDLR^15,16. 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^−/− mutant and wild-type Drosophila brains, as demonstrated in this study.

Despite the growing popularity of MALDI MSI, the instrument'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 example^11,17. 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.

Protocol

1. Fly head embedding

NOTE: The whole procedure takes ~45-60 min.

1. Prepare the optimal cutting temperature compound (OCT compound) stage with a flat surface.
   1. 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.
   2. 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 −80 °C.
2. Anesthetize adult flies using CO₂ (i.e., a CO₂ pad).
   1. 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.
   2. 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.
3. Transfer the OCT stage from dry ice to the dissection scope.
   1. 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.
   2. 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.
   3. 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.
4. After all the fly heads are aligned, let the OCT stage sit on the dry ice for another 5-10 min.
5. 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.
6. 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.
7. Store the frozen samples at −80 °C until ready for sectioning.

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.

1. 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.
2. Allow the tissues to equilibrate in the cryostat chamber for 30-45 min.
   1. 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.
3. 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.
4. Adjust the temperature of the cryostat chamber and specimen head according to the type of the tissue (e.g., −14 °C for liver, −20 °C for muscle, and −25 °C for skin¹⁰; −18 °C for fly heads in this protocol).
5. 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.
6. 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.
   1. 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.
7. Begin cutting in thick sections (50-100 µm) until the region of interest (e.g., the desired region of the brain) is found.
   1. Constantly brush off extra pieces with a precooled artist brush to keep the stage clean.
8. Change the thickness of the sections to 10-12 µm once the desired region is reached.
   1. 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 −18 °C.
9. Carefully collect the desired section and adhere it to the ITO slide. Perform this operation in the cryostat chamber.
   1. Take a room-temperature ITO slide and position it over the section, approach 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.
10. Place the slide aside in a rack or laboratory wipe outside the cryostat between the collections of multiple sections.
11. 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 variation. If necessary, separate to two slides, as the MALDI target holder can accommodate two slides in a single run.
12. 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 −20 °C throughout the process until storage at −80 °C within 24 h or shipping on dry ice to avoid the deterioration of lipids or metabolites.
13. Proceed to matrix deposition. Use 2,5-dihydroxybenzoic acid (DHB) in methanol/water (70/30, v/v) as the matrix.
14. If slides are not run immediately, either store the slides at −80 °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.
15. 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 °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.
16. 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.
    1. To calibrate the instrument, spot 0.5 µL 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.
    2. Set the laser spot diameters to Medium, as it is the modulated beam profile for 40 µm raster width, and gather imaging data by summing 500 shots at a laser repetition rate of 1,000 Hz per array position.
    3. 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 ±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&E) staining, as previously described¹¹.

Results

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^−/− mutant and wild-type Drosophila brains, which is elaborated on below.

The experiment was performed according to the workflow shown in

Discussion

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
2,5-Dihydroxybenzoic acid (DHB)	Millipore Sigma Aldrich	85707-1G-F
Andwin Scientific CRYOMOLD 15X15X5	Fisher Scientific	NC9464347
Andwin Scientific Tissue-Tek CRYO-OCT Compound	Fisher Scientific	14-373-65
Artist brush MSC #5 1/8 X 9/16 TRIM RED SABLE	Fisher Scientific	50-111-2302
autoflex speed MALDI-TOF MS system	Bruker Daltonics Inc		MALDI-TOF MS instrument
BD Syringe with Luer-Lok Tips	Fisher Scientific	14-823-16E
BD Vacutainer General Use Syringe Needles	Fisher Scientific	23-021-020
Bruker Daltonics GLASS SLIDES MALDI IMAGNG	Fisher Scientific	NC0380464
Drierite, with indicator, 8 mesh, ACROS Organics		AC219095000
Epson Perfection V600 Photo Scanner	Amazon	Perfection V600
Fisherbrand 5-Place Slide Mailer	Fisher Scientific	HS15986
Fisherbrand Digital Auto-Range Multimeter	Fisher Scientific	01-241-1
FlexImaging v3.0	Bruker Daltonics Inc		Bruker MS imaging analysis software
HPLC Grade Methanol	Fisher Scientific	MMX04751
HPLC Grade Water	Fisher Scientific	W5-1
HTX M5 Sprayer	HTX Technologies, LLC		Automatic heated matrix sprayer
Kimberly-Clark Professional Kimtech Science Kimwipes Delicate Task Wipers	Fisher Scientific	06-666A
MSC Ziploc Freezer Bag	Fisher Scientific	50-111-3769
SCiLS Lab (2015b)	SCiLS Lab		Advanced MALDI MSI data analysis software
Thermo Scientific CryoStar NX50 Cryostat	Fisher Thermo Scientific	95-713-0
Thermo Scientific Nalgene Transparent Polycarbonate Classic Design Desiccator	Fisher Scientific	08-642-7