Spatial Molecular Imaging of the Glycome using Mass Spectrometry

Abstract

The spatial organization of the glycome within tissues is key to the molecular basis for physiological function. The diverse and dynamic glycome is critical for fundamental cellular processes including metabolism, signaling, and adhesion. Innovations in spatial biology have ushered in new avenues for spatial molecular imaging of diverse glycome classes. Here, we describe an optimized protocol for spatial biomolecular imaging of the glycome in fresh-frozen mouse liver. The workflow comprises (1) rapid harvesting and freezing, (2) cryostat sectioning at optimal thickness and position, (3) tissue preparation and on-tissue enzyme digestion using carbohydrate-active enzymes, (4) matrix application and data acquisition by matrix-assisted laser desorption/ionization mass spectrometry imaging (MALDI-MSI), and (5) data processing and visualization to place the findings in biological context. Use of this approach allows acquisition of detailed spatial maps of N-linked glycans and glycogen, revealing key physiological and cellular features. These data allow definition of key spatial glycomic heterogeneity associated with liver function and dysfunction. This workflow enables highly reproducible and sensitive spatial glycomics of mouse liver. Additionally, it is readily adaptable to other tissues or species, facilitating novel spatial insights into glycome biology in health and disease.

SUMMARY:

This protocol details the key steps to enable rigorous and reproducible measurement of both glycogen and N-linked glycans using mass spectrometry imaging.

INTRODUCTION:

The field of spatial biology is rapidly expanding, providing critical insights into the dynamic spatial biomolecular complexity of living organisms. Multiple approaches are being employed to study distinct biomolecules in a biological context. While some are highly specialized techniques, mass spectrometry (MS) has emerged as an important general technique to quantify a wide range of biomolecules in the context of cells, tissues, or organisms.

Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry Imaging (MALDI-MSI) is one of the emerging general techniques for multiomic spatial biomolecule imaging. It was initially established for visualization of proteins and peptides in tissue slices ¹. Since then, it has been extended to many diverse biological samples and molecules ². The technique can be applied to cellular samples ³, organoids ⁴, model animals ⁵, and patient samples ^6,7. Some molecular classes can be directly measured using MSI-based approaches, including many metabolites and lipids, while other types of more complex biomolecules such as proteins and glycans require enzymatic or chemical treatment for them to be suitable for MSI-based detection ^2,8.

MALDI-MSI has also been applied to the spatial analysis of N-linked glycans and glycogen utilizing workflows that require enzymatic processing to produce glycan fragments suitable for MSI detection ^9-15. N-glycans are cleaved from the asparagine residue of proteins with PNGase F and glucose chains of glycogen are cleaved with isoamylase. Key advantages of the MALDI glycogen analysis is that it provides spatial definition of glycogen distribution in the biological sample, relative quantification, and information about the branching of the glucose chains. Because both N-glycan and glycogen workflows require enzymatic processing of the sample for MSI analysis, they have been combined into an integrated workflow for glycome analysis (Fig. 1A). This workflow requires careful preparation of biological samples and specific processing to enable MS analysis of the glycome, followed by data acquisition and processing. While the workflow requires a series of specialized steps, it produces rich high-quality spatial glycome data that is highly reproducible and can be adapted to many different types of samples ¹⁶. Additionally, the workflow is modular, such that the sample processing steps can be performed by independent research laboratories before submission of the samples to specialized facilities that perform the later steps of MALDI-MSI data acquisition and analysis.

Figure 1:

MALDI-MSI glycome analysis of mouse liver. A) Schematic for critical methods in sample preparation, data acquisition, and analysis. B) Sample mass spectra derived from the approach. Arrows in red demonstrate the regularly spaced oligosaccharides derived from glycogen, differing by one glucose monosacchride. Arrows in blue denote N-glycans, which are found distributed throughout the spectra. In liver, glycogen peaks are generally more prominent, especially at higher m/z. C) Spatial distribution of maltohexaose (DP6) derived from glycogen in the liver. D) Distribution of glycogen chain lengths in the liver. E-G) Spatial distribution of different N-linked glycan species in the liver. H) Hematoxylin and Eosin (H&E) staining for a focused central region, along with anatomical annotations. I) Corresponding focused inset showing the spatial localization of additional N-linked glycan (Hex5dHex1HexNAc5) in portal tract elements.

PROTOCOL:

This research was performed in compliance with institutional animal care and use committee (IACUC) guidelines at the University of Florida.

1. Harvesting liver

1.1 Animal euthanasia and dissection

1.1.1 Perform live cervical dislocation and decapitation to euthanize the mouse.

NOTE: For this example, a 6-month old wild-type C57BL/6J mouse was utilized. At this age, mice are ~25-33g. A wide range of ages, sizes, and strains are well suited for this analysis.

1.1.2 Immediately place the animal on an absorbent underpad and expose the abdominal cavity using a sterile scalpel.

1.1.3 Using sterile forceps and scissors, isolate the liver with minimal manipulation and tissue damage.

NOTE: Metabolic changes occur rapidly. Avoid perfusion or sedation if possible, and ensure dissection steps are performed as rapidly as possible ¹⁷.

1.2 Rapidly wash and clean tissue

1.2.1 Gently dip the liver in 1x phosphate-buffered saline (PBS) to remove any blood.

1.2.2 Dip the tissue in deionized (DI) water twice.

1.2.3 Blot excess water by gently touching the tissue onto a clean task wiper.

NOTE: Samples larger than what can fit on a standard microscope slide should be cut into pieces prior to freezing.

1.3 Sample freezing

1.3.1. Place the cleaned liver directly into a pre-chilled disposable polystyrene weigh boat floating on liquid nitrogen.

1.3.2 Once tissue changes color to a shiny opaque, approximately 3 min, then place the tissue into premade and labeled aluminum foil packets on dry ice.

1.3.3 Place the aluminum foil packets with the samples into a −80 °C freezer for storage. The samples can be stored long term at −80 °C.

2: Preparation of Biological Sample

2.1 Preparation for cryosectioning

2.1.1 Ensure the cryostat is turned on and equilibrated to a temperature appropriate for the tissue, between −16 °C and −20 °C. The exact temperature will depend on the type of tissue. Liver should be sectioned at a slightly warmer temperature, approximately −16 °C to −17 °C, to prevent tears in the tissue while other tissues should be sectioned at −20 °C.

2.1.2 Ensure there are at least two chucks, two thin paintbrushes, a pair of thin tweezers, and a 70mm glass insert at temperature in the cryostat. Additionally, label slides in pencil prior to placing them in the cryostat to acclimate.

2.1.3 Transfer the tissue from the −80 °C freezer to the cryostat on dry ice.

2.1.4 Allow tissues to acclimate to the temperature in the cryostat for at least 30 min.

2.1.5 Ensure there is no damage to the edge of the glass insert and that it is smooth where the tissue will meet the anti-roll guide by running a gloved finger along the edge.

2.1.6 Place the glass insert into the anti-roll guide.

2.1.7 Take a low-profile blade and slide it into the open blade holder carefully and close the blade holder. Place the red blade guard down over the blade until sectioning.

2.1.8 Mount the tissue onto the chuck using M1 mounting media, which is compatible with MS. Place room temperature M1 mounting media onto the chuck inside the cryostat, covering at least two rings on the chuck so the tissue is secured in place.

Note: Optimal cutting temperature (OCT) compound is often utilized in cryosectioning, however it causes ion suppression in MS applications and must be avoided. If OCT must be used then an additional multi-step protocol should be utilized to minimize or remove the OCT. Additionally, some MS resource centers will not analyze samples that were embedded in OCT.

2.1.9 Take a chilled pair of tweezers and remove the tissue from the aluminum foil. Use the tweezers to position the tissue onto the chuck in the desired orientation.

2.1.10 Place the chuck down in the cryostat to allow the mounting media to freeze, approximately 2 min. Once it is frozen, it will be hardened and appear opaque.

2.2 Cryosectioning liver tissue

2.2.1 Place the chuck with the mounted tissue into the chuck holder. Position the chuck to where the tissue will cut at a 90 ° angle to the blade.

2.2.2 Retract the chuck holder until the tissue does not have contact with the blade. Slowly bring in the chuck until the tissue is immediately adjacent to the blade to begin sectioning.

2.2.3 Begin sectioning with a thickness slice of 30 μm to remove unwanted tissue.

2.2.4 Ensure the roll plate depth creates a smooth section and the tissue does not roll on top of the roll plate. If this is the case, then loosen the roll plate slightly. If the tissue moves the roll plate and hits it a second time on the reset of the rotation, then tighten the roll plate slightly.

2.2.5 As the desired section is approaching, decrease the intended thickness slide to 10 μm, or another desired thickness suitable to the specific application.

2.2.6 Complete 10 rotations so the chuck is at the correct thickness to prevent lagging and uneven sections.

2.2.7 Use a chilled paintbrush to clear the stage in preparation for sectioning. Close the roll plate over the blade and slowly take a section of the tissue.

2.2.8 Lift the roll plate and use the two chilled thin paintbrushes to gently tease the section away from the blade. Pull the section to the side of the stage for ease of mounting on the slide.

2.2.9 Take a chilled slide from a slide carrier in the cryostat and place an index finger on the opposite side of the slide from where the tissue will be positioned for 10 seconds.

2.2.10 Gently place the slide down onto the tissue and lift it back up. Do not push down hard, as this will tear the tissue. The tissue should melt onto the slide and become translucent.

2.2.11 Rapidly transfer the slides into a desiccator for drying. Dry the slide in the desiccator for 1 hr.

2.2.12 After drying, slides can be directly processed for MS or vacuum sealed and stored at −80 °C.

3. Slide preparation for MSI of glycomics.

3.1 Preparation of enzymes

3.1.1 MS-ready low salt lyophilized PNGase F can be purchased and directly utilized.

3.1.2 Isoamylase requires dialysis to remove excess salt for MS applications. Prepare one or more dialysis cups by placing in HPLC water for 15 min.

3.1.3 Dialyze 200 uL of stock isoamylase using a 500 uL dialysis cup in 14.3 mL of HPLC water for 2 h at 4 °C, without stirring. Replace the 14.3mL of HPLC water with fresh HPLC water and dialyze for an additional 16 h (overnight) at 4 °C. Replace the 14.3 mL of HPLC water with fresh HPLC water and dialyze for an additional 8 h at 4 °C.

3.1.4 Gently remove the isoamylase from the cup with a pipette and place into a 1.5 microcentrifuge tube. Measure the total volume of isoamylase solution after dialysis (in μL). From the starting activity of the stock, typically 200 Units/mL and final volume calculate the total volume required for 3Units of isoamylase.

3.1.5 Pipette aliquots of 3 Units isoamylase into thin-walled PCR tubes. Snap freeze tubes in liquid nitrogen. Store at −80 °C. One tube of dialyzed isoamylase solution can be used for each spraying protocol, which is sufficient for four slides. Avoid freeze/thaw cycles to maintain enzyme activity.

3.2. Fixation and washing of tissue

3.2.1 Place slides in a slide holder and submerge in a glass container filled with 10% Neutral Buffered Formalin (NBF) for 1 h.

3.2.2 Remove the slides from the NBF. Submerge the slides in 70% ethanol for 2 h.

3.2.3 Prepare four Coplin jars containing 70%, 80%, 90%, and 100% ethanol. Label each jar clearly to avoid confusion during sequential washing.

3.2.4 Successively transfer slides as follows: 70% ethanol for 5 min, 80% ethanol for 5 min, 90% ethanol for 5 min, 100% ethanol for 5 min.

3.2.5 Transfer the slides to a glass petri dish filled with fresh 100% xylene. Ensure the slides are fully submerged. Incubate the slides on a shaker at 70 rpm for 1 hr, depending on the lipid content of the tissue. Repeat this step two additional times using fresh xylene for each wash.

NOTE: This protocol is designed to allow maximal delipidation, since lipids are known to result in ion suppression, and thus maximum reproducibility. However, this does result in delocalization of signals thereby decreasing the resolution that can be obtained. Decreased washing steps, limiting processing time, and careful control of humidity may decrease delocalization and thereby result in improved data resolution.

3.2.6 Place slides in a desiccator overnight to dry.

3.2.7 Place slides in 70% ethanol for 1 minute to rehydrate. Place slides in DI water for 3 min. Repeat the DI water wash with fresh DI water.

3.2.8 Dry the slides in a desiccator for 10 min, or until completely dry.

3.3 Preparation of slides for enzyme application

3.3.1 Prepare an HTX humidity chamber for incubation by placing a paper towel soaked with DI water below the metal rack. Place the chamber in a 37 °C incubator.

3.3.2 Prepare a food steamer by filling the water reservoir to the top with tap water. Set the timer to 60 min to turn the unit on.

3.3.3 Prepare citraconic buffer by adding 50 mL of HPLC-grade water, 25 μL of citraconic anhydride, and 2 μL of 12 M HCl to a 50 mL Falcon tube. Vortex the solution for 10 seconds to mix thoroughly. Each 50 mL falcon tube of citraconic buffer is enough for four slides. Verify the pH of the solution is 3.0 (acceptable range: 2.5–3.5).

3.3.4 Fill each mailer with the prepared citraconic buffer. Place slide with the tissue facing inward, to prevent the tissue from touching the sides and allow for effective antigen retrieval. Two slides can be used per mailer, using the two edge positions with both slides having tissues facing inward.

3.3.5 Verify that steam is being emitted by the food steamer. Place the mailers in the food steamer and incubate for 30 min.

3.3.6 Transfer mailers to a DI water bath for 5 min. Replace half of the citraconic buffer in the mailer with DI water and let stand for another 5 min. Repeat this dilution two more times.

3.3.7 Remove all solvent and wash the slides by adding 100% DI water to the slide mailer and then remove the DI water. Using a lab wipe, spot dry around any areas in which tissue is not found.

3.3.8 Draw a circle and triangle on the back of each slide to mark internal standard locations. Apply 1 μL of 1 mg/mL Horseradish Peroxidase (HRP; circle) and 1 μL of 10 mg/mL rabbit liver glycogen (triangle) to the front of the slide.

3.3.10 Place slides in the desiccator for 15 min to dry completely.

3.4 Enzyme spraying and digestion

3.4.1 Remove one tube of 100 μg lyophilized PNGase F (500 units/μL) and one tube of 3 Units isoamylase and allow to thaw for 10 min. Spin tubes at ~2000 X g for 5 seconds in a microfuge. This is sufficient enzyme to spray 4 slides.

3.4.2 Pipette 50 μL DI water to each lyophilized PNGase F tube, vortex gently for 10 seconds, and spin in a microfuge at ~2000 X g for 5 seconds. Pipette the solution of 50 μL PNGase F directly into the isoamylase tube. Add HPLC water to a final volume of 1 mL.

3.4.3 Vortex the dual enzyme microcentrifuge tube 10 seconds and spin down at ~2000 X g for 5 seconds.

3.4.4 Turn on the HTX M5 sprayer and open the HTX M5 software. Ensure the tray type is set to “Ambient” under the “System” tab.

3.4.5 Turn off the Knauer pump by pressing the stop button on the front right side. Power on the external syringe pump using the switch on its back panel.

3.4.6 In the HTX software, under the “Methods” tab, select the appropriate method for dual enzyme spraying. Parameters must be set to nozzle temperature of 45 °C, 15 passes, flow rate of 0.025 mL/min, velocity of 1200 mm/min, track spacing of 3 mm, CC pattern, pressure set to 10 psi, gas flow rate of 3 L/min, and nozzle height of 40 mm.

3.4.7 Navigate to the “Temp” tab and set the spray nozzle temperature to 45 °C. Use a wrench to move the sprayer line to the top connector.

3.4.8 Open the ultra high purity nitrogen gas valve and ensure the sprayer pressure is 10 psi. On the syringe pump, set the flow rate to 95 μL/min.

3.4.9 Using a fresh syringe and needle, fill with 4 mL HPLC-grade water. Remove all bubbles. Run 2 mL of water through a ~6-inch section of the sprayer line into waste beaker.

3.4.10 Attach the syringe to the sprayer line and run water through the syringe pump by starting and running at 95 μL/min for 5 min.

3.4.11 While flushing the syringe pump, tape the slides to the bottom-left corner of the heated tray using a metal alignment guide to ensure the tissue lies within the spray area. Under the “Sample” tab, define the X and Y parameters for the spray region.

3.4.12 After 5 min, pause the pump, disconnect the syringe, draw air into it, and then push air through the sprayer line to purge any residual water. Repeat as needed.

3.4.13 Load the dual enzyme solution into a new syringe and connect to pump, ensuring no bubbles remain.

3.4.14 Attach the syringe to the sprayer line. On the syringe pump, adjust the flow rate to 25 μL/min.

3.4.15 Once the spray nozzle reaches 45 °C, press “Start” under the “Cycle” tab. Click “No” when prompted to turn on the Knauer pump. Press “Start” on the syringe pump to begin the spray.

3.4.16 Place a blank slide under the spray nozzle to confirm visible enzyme spray deposition. Once observed, click “Continue” on the software, place the glass cover on the sprayer, and monitor the spray for an even application, as indicated by uniform wetting.

3.4.17 When the spray cycle is complete, press stop on the pump, turn off the nitrogen gas, and click “Valve load confirm.” Clean the line and stage of the sprayer and shut off the sprayer, ensuring the Knauer pump is back on.

3.4.18 Incubate the slides in the HTX humidity chamber at 37 °C for 2 h, placing the slides facing upwards. Once complete place slides in the desiccator for 15 min. Slides can remain in the desiccator overnight or the protocol can be continued.

3.5 MALDI matrix preparation and application

3.5.1 Prior to matrix application, capture an optical image of the slide using a scanner.

3.5.2 Weigh 40 mg of solid CHCA into a 15 mL conical tube. Dissolve the CHCA powder in a solution of 0.1% trifluoroacetic acid and 50% acetonitrile (ACN) at a concentration of 7mg/mL

3.5.3 Sonicate the matrix solution for 10 min using cycles of 90 sec on and 30 sec off. Repeat these steps 5 times to ensure full dissolution and homogeneity.

3.5.4 Turn on the HTX M5 sprayer and open the HTX M5 software. Open the nitrogen gas valve and set the system pressure to 10 psi.

3.5.5 In the HTX software, under the “Methods” tab, select the appropriate method for CHCA matrix spraying. Parameters set to nozzle temperature of 79 °C, 10 passes, flow rate of 0.1 mL/min, velocity of 1300 mm/min, track spacing of 3 mm, CC pattern, pressure set to 10 psi, gas flow rate of 3 L/min, and nozzle height of 40 mm.

3.5.6 Navigate to the “Temp” tab, set the spray nozzle temperature to 79 °C. Under the “Pump” tab, set the flow rate to 100 μL/min.

3.5.7 Tape the slides to the bottom-left corner of the heated tray using a metal guide to ensure proper alignment in the spray area. Under the “Sample” tab, define the X and Y parameters for the spray region.

3.5.8 Use a disposable syringe and needle to draw 5.5 mL of 50% ACN (remove bubbles).

3.5.9 Set the sprayer to “LOAD” mode. Insert the ACN syringe and inject the ACN. Remove the ACN syringe and immediately replace it with a new syringe containing 5.5 mL of filtered CHCA solution, being sure to avoid bubbles. Set the sprayer to “SPRAY” mode.

3.5.10 Once the spray nozzle reaches 79 °C, press “Start” under the “Cycle” tab. Click “Yes” when prompted.

3.5.11 Use a blank test slide and place under the sprayer nozzle. To confirm visible matrix spray deposition a yellow matrix deposition will be found on the slide. Once observed, click “Continue”, place the glass cover on the instrument, and monitor the spray for consistency. Consistent spray will produce a slide with uniform opaqueness.

3.5.12 When the spray cycle is complete, set the sprayer to “LOAD” mode and click “Valve load confirm.”

3.5.13 Clean sprayer by setting pump to 300 psi and putting temperature down to 30 °C. Uptake and push through the line 5mL of 50% acetonitrile twice and 5mL of 50% methanol twice. Wipe the stage clean with 100% methanol. Once temperature reaches 30 °C, switch to “LOAD,” turn off gas, shut off sprayer, and ensure the Knauer pump is back on.

3.5.14 Place the slides in the desiccator for 15 min until the matrix is completely dry. The slides should be run as soon as possible after spraying, within 14 days at maximum, to ensure reliable data collection with appropriate signal to noise.

4. MALDI-MSI glycomics data collection

4.1 Preparing for data collection

4.1.1 Secure the slide into the MALDI instrument’s sample holder with slide adapter. Insert the sample holder into the MS and ensure it is properly seated for imaging.

4.1.2 Open the flexImaging software and import the optical slide image previously recorded, see 3.4.1. If required, calibrate the image alignment with the instrument stage so that the tissue sections and standard spots are correctly positioned for region selection.

4.1.3 Define the imaging regions for the tissue sections, the HRP standard, the glycogen standard, and control matrix square in the software. Outline each as a separate region.

4.1.4 Set the raster step size to 50 μm, yielding a pixel resolution of 50 × 50 μm.

4.1.5 In the instrument control software, select positive-ion MS mode and set the scan range to m/z 500–4000 with a low-mass cutoff of 700.00 m/z.

4.1.6 Load the pre-optimized MALDI imaging tuning parameter set in the instrument control software (or enter the values manually if needed). Verify that the source and ion optic voltages are set to the specified values: MALDI Plate Offset 50.0 V; Deflection 1 Delta 70.0 V; Funnel 1 RF 500.0 Vpp; Funnel 2 RF 500.0 Vpp; Multipole RF 500.0 Vpp; Ion Energy 5.0 eV. Confirm the collision cell and TOF timing settings: Collision Energy 10.0 eV; Collision RF 4000.0 Vpp; Transfer Time 180.0 μs; Pre Pulse Storage 25.0 μs.

4.1.7 Confirm laser settings under the Laser tab. Select "Imaging 50 μm" as the application default.

4.1.8 Calibrate the mass spectrometer using an external calibrant mixture such as tune mix. Introduce the calibrant and run the instrument’s calibration routine. Adjust the calibration so that the mass accuracy error is within 0.001% and the calibration accuracy score is 100%.

4.1.9 Set the acquisition to perform a burst of 320 laser shots at 10,000 Hz frequency. Laser attenuation should be adjusted to 37% to maintain signal intensity within the optimal dynamic range.

4.1.10 Use the instrument’s automated beam alignment function, target profile adjustment, and focus adjustment to ensure the laser is properly focused on the sample. On a region off the tissue, execute the “Auto Beam Adjust” under the Laser tab in the control software to align the laser with the ion optics. This will ensure that the laser and stage are adjusted for optimal data collection. Finally, execute target profile generation and focus tuning adjustments in the sample carrier tab.

4.1.11 Confirm that the internal standard spots produce the expected ion peaks, indicating readiness for data collection. HRP should have a prominent glycan peak at 1211.42. Glycogen should have regularly spaced prominent peaks, with the highest intensity peaks at 1013.32, 1175.37, and 1337.43.

NOTE: If there are issues identified with intensity, revisit laser configuration. If there are issues identified with mass accuracy, revisit calibration.

4.2 MALDI-MSI data collection

4.2.1 Begin the MALDI-MS imaging acquisition for the defined regions by clicking the “Start” button in the imaging software. The instrument will automatically raster the laser over each defined region at 50 μm intervals, acquiring a positive-ion MS1 spectrum (m/z 500–4000) at each pixel. Typical data collection time for a liver section at this resolution is 135 min.

4.2.2 After the run, the instrument will automatically save the spectra for every pixel acquired creating a .d folder with the raw data.

5. MALDI-MSI data analysis

5.1 Data import and export

5.1.1 Import the dataset into SCiLS Lab, or equivalent MSI analysis software, to visualize ion intensity maps of the glycome across the tissue sections.

5.1.2 Open SCiLS Lab. Select “New.” Select the .mis file from the MALDI run.

5.1.3 Once the .mis file opens in SCiLS, click “Next.” Select Automatic range for m/z Range. Select automatic axis for Axis Parameters. Select absolute bin size. Click ‘Next” and then “Import.” Select the folder where the file will be saved.

5.1.4 After the import is finished, open the new .slx file.

5.1.5 Click file, Import, and select feature list from file option. Import glycome annotations and create ion images by going to feature list pane, selecting correct list, and click the create ion images button.

NOTE: Robust feature lists are available for N-glycans ^16,18,19 and glycogen ¹³.

5.1.6 Normalize the data, as appropriate. Total Ion Count (TIC) is the most commonly utilized normalization.

5.1.7 Once the ion images have loaded go through the feature list and ensure correct peak integration, manually adjusting for those features in which the peak is not fully integrated. To adjust left click on the peak while holding shift. Once peak is integrated hit control and space bar on the keyboard. This will generate a new ion image for the adjusted peak.

5.1.8 After repeating for the rest of your feature list. Within the ion image panel, select save feature list as, name the list, and click save.

5.1.9 Go to file, feature table, and open the feature list table that was just saved. Sort by m/z. The adjusted features will appear with an empty cell in the name column. Rename the adjusted peaks by copying and pasting the name and removing the original peak by selecting the remove selected row from feature list button located towards the top middle of the feature table.

5.1.10 After adjusting all names, a final annotated peak list is obtained. Click the add intensity button located near the delete button. From here select the samples of interest to export the average intensity for the annotated features for each region imaged.

5.1.11 Ensure to select the Peak Area interval processing mode and Total Ion Count for the normalization. Click ‘OK’. New columns will appear with the corresponding data. To export the data, select the outward pointing arrow and save as .csv.

REPRESENTATIVE RESULTS:

Careful processing of a liver from a wild-type mouse, including cutting a full transverse section (Fig. 1A), allowed definition of the glycome by MALDI-MSI, with diverse glycogen-dervied oligosaccharides and N-linked glycans over a range of m/z (Fig. 1B). Glycogen was broadly localized throughout the liver hepatocytes and lacking in vessels and connective tissue in the portal tracts, as expected (Fig. 1C-D). Importantly the MALDI-MSI data allowed measurement of both the spatial distribution of glycogen as well as chain-length distribution, a key feature of glycogen in normal metabolism and disease states (Fig. 1D). Additionally, diverse N-linked glycans exhibit diverse organizations. The observed spatial distribution ranges from widely distributed in hepatocytes (Fig. 1E) to regional foci, consistent with the sinusoidal organization of the liver (Fig. 1F), to specific labeling of portal tract elements (Fig. 1G-I).

DISCUSSION:

MALDI-MSI is an emerging tool for glycome spatial biology. Importantly, while this protocol focuses on data collection using specific hardware, it can be readily adapted to MALDI-MSI platforms from multiple vendors ¹³. Sample processing is identical up until method 4, although some vendors may require use of specialized slides. Modifications of data collection, using parameters specific for the MALDI-MSI platform, enable rapid adaptation of this protocol. Vendor-specific software for data processing can be utilized, as detailed in method 5, or, alternatively, platforms for rigorous and reproducible MALDI-MSI data processing can be employed ¹⁹.

MALDI-MSI is a particularly useful tool for understanding biology since different organs, tissues, and cells have different patterns for their glycome. Liver is well suited for glycome analysis in that it has high levels of glycogen and diverse N-linked glycans ¹³. Other organs have higher levels and diversity of N-linked glycans and lower levels of glycogen ¹³. Importantly, the glycome varies in different organs depending on the metabolic and disease state of the organism and so the glycome provides a rich source of biological information ^5,14,15,20. Continuing development of unique carbohydrate-active enzymes for MALDI-MSI analysis promises to extend glycome analysis to additional orthogonal biomolecules in the glycome.

While having many advantages, MALDI-MSI is currently limited in resolution compared to other tools utilized in spatial biology. This protocol utilized 50 μm resolution data collection. While excellent for physiological analysis, data at single-cell resolution would significantly advance the field ²¹. In particular, delocalization can be a significant barrier to increased resolution in methodologies that utilize enzyme spraying and subsequent incubation in humid conditions. Recent advances in sample processing hardware, sample handling methodologies, data acquisition hardware, use of selective labeling, and data processing are allowing MALDI-MSI data collection to approach cellular resolution ^22,23. However, there are tradeoffs between resolution and signal to noise and inherent limitations based on current sample preparation methods for MALDI that need to be considered. Very specific applications are already achieving the goal of single cell-resolved MALDI-MSI using innovations in both sample processing ²⁴ and novel algorithms, suggesting general solutions to these problems are within reach ^25-27.

Following data collection, the slide can be stained via H&E for the purposes of histological annotation. However, cellular and tissue damage is associated with MALDI-MSI data collection, so this strategy limits the extent of cellular annotation. Alternatively, sequential tissue slices can be utilized for H&E. This strategy provides high quality data, but requires alignment and integration of discreet sample cuts.

Of note, formalin-fixed paraffin-embedded (FFPE) tissues are very well suited for glycome analysis by MALDI-MSI with small changes in the protocol for sample preparation ¹³. However, FFPE tissue cannot be effectively utilized for metabolomics and other similar analyses, so fresh-frozen samples, as detailed here, are preferred for multiomics approaches ²⁸. Sample preparation is a critical component of any MSI pipeline for labile biomolecules, which requires special approaches for harvesting and handling of samples ^17,29,30.

Methodologies to expand the generality and wide-spread utilization of MALDI-MSI-based approaches are rapidly progressing with dramatic improvements in hardware and software ¹⁹. Additionally, recent cutting-edge multiomic ²⁸ and multimodal ²¹ approaches combined with machine learning and artificial intelligence are particularly promising for handling these highly complex datasets ³¹.