Biochemistry
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Optimal Preparation of Formalin Fixed Samples for Peptide Based Matrix Assisted Laser Desorption/Ionization Mass Spectrometry Imaging Workflows
Chapters
Summary January 16th, 2018
This protocol describes a reproducible and reliable method for the sublimation-based preparation of formalin fixed tissue destined for imaging mass spectrometry.
Transcript
The overall goal of this procedure is to reproducibly prepare a tissue sample for peptide mass spectrometry imaging. This method allows novice mass spectrometry users to develop their own imaging mass spectrometry workflows without going through lengthy empirical determinations. The main advantage of this technique is that it is highly reproducible, simple to implement, and uses cost-effective methodologies.
To prepare the nitrocellulose coated slides, pipette 40 microliters of liquid nitrocellulose onto one edge of the conductive indium tin oxide microscope slide. Using a regular glass microscope slide, drag the nitrocellulose under surface tension across the slide to create an even thin-film coating. Allow the nitrocellulose to dry at room temperature for 20 seconds, and then store until needed at room temperature.
Next, create custom vapor chambers by cutting a piece of thick blotting paper big enough to fit the bottom half of a standard plastic petri dish. Cut the paper to leave a rectangular strip in the center the same size as a microscope slide. Leave enough excess so that the paper will maintain its position in the petri dish.
For FFPE tissue, float mount the sections onto the nitrocellulose pre-coated ITO slides as described in the text protocol. Submerge the slides in fresh xylene to remove residual paraffin for two minutes. Wash the deparaffized samples by submerging the slides in a graded solvent series consisting of a 70%volume for volume ethanol water solution, followed by 100%ethanol, for 30 seconds.
Then, submerge the slides into Carnoy's fluid for two minutes. Finally, submerge the slides into a series of 100%ethanol, ultrapure water, and 100%ethanol again, for 30 seconds. Load the sample slides into a plastic slide box that is filled to the top with 20 millimolar tris HCO.
Seal the box, and place it in a water bath containing 500 milliliters of water, allowing the box to touch the bottom of the bath. Then heat the box for 15 minutes at 120 degrees celsius in a pressure cooker, which can achieve an operating pressure of 70 kilopascals. Remove the slides, allow them to cool, and let them dry at ambient temperature for 15 minutes.
Once hydrolyzed, coat the samples with 10 microliters of trypsin solution in ultrapure water by first pipetting 10 microliters of the solution onto the edge of the tissue section. Then, using the same pipette tip, drag the droplet across the whole surface of the tissue under surface tension. After allowing the samples to dry at ambient temperature, mount them inside the top of the previously constructed vapor chamber with autoclave tape on either edge of the slide.
Carefully pipette 600 microliters of a solution containing a one-to-one volume for volume mix of 100%acetonitrile and 50 millimolar ammonium bicarbonate onto the blotting paper finger in the bottom part of the petri dish, until the finger appears to be evenly wet. Pipetting is critically important because incorrect pipetting will result in the delocalization of your matrix and surface analoids which will destroy the spatial information contained within your sample. Place the top half of the vapor chamber on the bottom half, ensuring the paper finger and sample slide align perfectly, and seal the chamber around its equator with paraffin film.
Leave the sample over night in a 37 degree celsius incubator to allow complete digestion. Once digested, leave the sample slide on a five-digit microanalytical balance. Mount the slide onto the cooling finger of the sublimation apparatus, and secure it with copper tape in the same way as described for the vapor chamber.
Place 300 milligrams of CHCA matrix into a glass petri dish at the bottom of the chamber and spread it evenly to create a thin layer of matrix crystals. Assemble the sublimator. And secure the two halves with the horseshoe clamp.
Suspend the assembled unit 15 to 20 centimeters above a sand bath, preheated to 220 degrees celsius, by placing it in a metal ring connected to a retort stand. Connect the chamber to the vacuum source, engage the vacuum, and allow it to stabilize down to approximately 25 millitorr for five minutes. Pack the cooling finger to the top with ice, and add 50 milliliters of water.
Allow the apparatus to settle for a further five minutes before proceeding. Lower the chamber onto the surface of the stand, ensuring the sand completely contacts the bottom of the chamber. Leave it for 45 minutes to create an ideal coating of 0.22 milligrams per square centimeter.
After 45 minutes, remove the chamber from the sand bath by raising the metal ring, and vent the chamber. Once sublimated, mount the sample slide inside the top of the previously constructed vapor chamber, as before. Carefully pipette 600 microliters of a solution containing a one-to-one volume for volume mix of acetonitrile and trifluoroacetic acid in water onto the blotting paper in the bottom part of the petri dish to ensure an even coating.
Next, assemble the chamber, ensuring the paper tab and microscope slide align perfectly. Leave the chamber in a 37 degree celsius incubator for one hour. Scan the slide in a flat-bed scanner to create a digital image.
Load the sample into the mass spectrometer, and then analyze using the appropriate software platform. Finally, analyze the samples as described in the text protocol. Shown here is a correctly processed tissue specimen that has been imaged at 50 microns and shows good macro structure and clear difference between white matter and gray matter.
Successfully compared samples will show clear differentiation at different tissue locations. Here, there's a clear region of upregulation of the peptide represented by a mass-to-charge ratio of 1, 085 in the white matter region, as represented by white regions. By contrast, this incorrectly prepared sample shows a large-scale level of delocalization.
The patterns of the molecules present across the three panels clearly demonstrate that there is no clear definition between biological regions or differences in the abundance of the molecules displayed. After watching this video, you should have a really good idea as to how prepare your own samples for peptide imaging analysis. With proper procedure, this method can be performed within eight hours over two days, allowing for an overnight digestion step.
While preparing this methodology, it's important to take care during each step. Any physical damage that happens to the tissue will destroy the spatial information contained within your sample. We had the idea for this methodology when we first started our own IMS space investigations and realized that there were no other definitive methodologies that were published.
With small modifications, our methodology can be applied to the analysis of proteins, peptides, lipids, and other small molecules. Following this procedure, other histological techniques such as immunofluorescence and hematoxylin and eosin staining can then be applied to your sample in order to get a reference point for your imaging mass spectrometry data. When working with potentially explosive chemicals, such as liquid nitrocellulose, it's important to maintain the correct safety procedures.
Minimizing working volumes and disposing your waste immediately is critically important.
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