Laser Capture Microdissection of Glioma Subregions for Spatial and Molecular Characterization of Intratumoral Heterogeneity, Oncostreams, and Invasion

All methods described here that use experimental animals have been approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Michigan.

NOTE: Glioma neurospheres generated from a GEMM or stable lines can be used for intracranial tumor engraftment in mice¹⁰ and processed for LMD and RNA sequencing. These cells constitutively express firefly luciferase and GFP proteins, which will be further utilized for tumor growth analysis and localization.

1. Generation of intracranial mouse glioma model from neurospheres derived from genetically engineered glioma models

1. To generate a primary culture of neurosphere cells from a genetically engineered mouse (GEMM) model, use the protocol described previously^10,11.
2. Prepare neurosphere culture medium as described: 500 mL of Dulbecco's Modified Eagle Medium F-12 (DMEM/F12) supplemented with 10 mL of 50x B-27 and 5 mL of 100x N-2 neuronal culture supplements, 5 mL of 100x Antibiotic-Antimycotic, and 1 mL of Normocin. Before plating the neurospheres, add 20 ng/mL human recombinant EGF and FGF to the culture medium.
3. Culture glioma neurospheres in a tissue culture incubator at 37 °C and 5% CO₂ for 2-3 days prior to intracranial tumor engraftment.
4. On the day of the surgery, collect the glioma neurospheres and spin them at 550 x g for 5 min at room temperature. Carefully remove the supernatant without disturbing the cell pellet.
5. To dissociate the neurospheres, resuspend the cell pellet in 1 mL of cell detachment solution and incubate at 37 °C and 5% CO₂ for 2-4 min. Following the incubation period, pipette the neurospheres up and down with a 1 mL micropipette to ensure a single cell suspension.
6. Inactivate the cell detachment solution by diluting the neurosphere suspension with 10 mL of un-supplemented DMEM/F12 media. Spin the neurospheres at 550 x g for 5 min at room temperature. Carefully, remove the supernatant.
7. Resuspend the cell pellet in 100 µL of DMEM/F12 media with no supplements. Make a 1:50 dilution of the cell suspension, and then add 50 µL of Trypan Blue to count viable cells. Use the following formula to determine the concentration of cells:
   cells/mL = [((∑cells counted per square) / # of squares counted) x 50 x 10,000 cells/mL]
8. After determining the cell count, to achieve a target concentration of 30,000 cells/µL in 100 µL volume, spin the neurospheres down and resuspend them in the appropriate volume of DMEM/F12 containing no supplements.
9. Place neurospheres in a pre-labeled 0.6 mL tube on ice.
10. Prepare working solutions of anesthesia, analgesic, and anesthesia reversal prior to implantation: For ketamine/dexmedetomidine anesthesia, add 0.6 mL of 100 mg/mL ketamine hydrochloride and 0.8 mL of 0.5 mg/mL dexmedetomidine hydrochloride to sterile 8.6 mL of 0.9% NaCl containing vial. For buprenorphine analgesic, add 1 mL of 0.3 mg/mL buprenorphine to 9 mL of 0.9% NaCl containing vial. For atipamezole anesthesia reversal, add 1 mL of 5 mg/mL atipamezole to 9 mL of 0.9% NaCl containing vial.
11. Use 6-8-week-old C57BL/6J female mice for intracranial tumor engraftment.
12. In a room approved for rodent survival surgery, set up sterile supplies for intracranial tumor engraftment. Perform surgeries on a rodent stereotaxic frame equipped with a sterilized 10 µL Hamilton syringe with a removable 33G needle. Utilize a bead sterilizer to sterilize tools between surgeries.
13. Anesthetize mice with a single intraperitoneal (i.p.) injection of the anesthetic solution prepared in step 1.11 (ketamine:75.0 mg/kg and dexmedetomidine:0.5 mg/kg). Approximately, a 250 µL volume of the anesthetic solution will be delivered to a mouse weighing 20 g. Ensure that the mouse is unresponsive to pedal reflex prior to proceeding.
14. Once the mouse is in a deeply anesthetized state, apply sterile petrolatum ophthalmic lubricant to the eyes to prevent drying. Shave the fur on the mouse cranium. Apply 10% povidone-iodine topical solution to the shaved area in order to disinfect.
15. Secure the mouse's skull in a stereotactic frame. First, carefully open the mouth with forceps and gently pull the tongue and move it to one side of the mouth to prevent choking. Keep the mouth open with the forceps and place the top incisors into the keyhole of the tooth bar on the stereotactic frame.
16. Holding the mouse head by the ears, place the ear bars against the postorbital bones and secure them. Ensure that the mouse's cranium is level with the surgical tabletop. Secure the ear bars carefully not applying pressure against the skull. Next, carefully secure the nose bar.
17. Using a size 15 scalpel blade, make an incision along the mouse head, exposing the cranium. Retract the skin at the incision site using Colibri retractors. Use a sterile applicator to remove all pericranial tissue.
18. Identify the bregma and lower the needle of the Hamilton syringe directly over it. Using the frame, position the needle 1 mm anterior of the bregma and 1.5 mm lateral. Using a 26G needle, mark this spot by scoring the cranium.
19. Use a cordless power drill equipped with a 0.45 mm drill bit to create a burr hole at the target site. Drill until reaching the underlying dura mater. Carefully extract the remaining bone in the burr hole with a 26G needle.
20. Using a pipette, homogenize the cells thoroughly and draw up 7 µL of the neurosphere suspension into the syringe. To ensure that that syringe is working properly, dispense 1 µL of the suspension onto a 70% alcohol-soaked pad.
21. Lower the needle to the surface of the dura mater. Lower the syringe 3.5 mm ventral and retract 0.5 mm. This will create a 0.5 mm space in the brain for the neurospheres to deposit when injected.
22. Leave the needle in place for 2 min for pressure equilibration. Slowly and smoothly deliver 1 µL of the cells over the course of 1 min. Allow the cells to settle in for 6 min. Remove the syringe slowly and smoothly from the brain over the course of 2 min.
23. Using sterile saline solution, wash the surface of the cranium three times. Dispense the excess cells in the syringe onto a 70% alcohol pad, and wipe the needle clean with another 70% alcohol pad. Rinse the syringe with sterile PBS to avoid clogging of the needle.
24. Remove the retractors and carefully take the mouse out of the stereotactic frame. Close the incision using 3-0 nylon sutures, forceps, and a needle driver.
25. Administer atipamezole (1.0 mg/kg), approximately 100 µL for a 20 g mouse. Administer buprenorphine (0.1mg/kg subcutaneous) subcutaneously, approximately 70 µL for a 20 g mouse. Place post-surgical mice into a clean recovery cage and monitor them until alert and active.

2. Animal perfusion and brain preservation

1. To monitor tumor progression, determine in vivo bioluminescence using an in vivo imaging system until animals show signs of tumor burden. Euthanize mice when they reach a signal between 10⁶ to 10⁷ photon/s.
2. Anesthetize mice displaying signs of tumor burden with intraperitoneal (i.p.) injection of ketamine (75.0mg/kg) and dexmedetomidine (0.5 mg/kg) solution. Deliver approximately 250 µL to a mouse weighing 20 g. Then, ensure that the mouse in unresponsive to pedal reflex.
3. Using forceps, hold the skin above the peritoneal cavity, and using a large pair of dissection scissors make a "Y" incision by penetrating the peritoneal wall, puncture the diaphragm, and then cut the rib cage.
4. Insert a blunt 20G cannula into the left ventricle of the mouse's heart. Then snip the right atrium of the heart to allow for exsanguination.
5. Allow oxygenated Tyrode's solution (0.8% NaCl, 0.0264% CaCl₂, 0.005% NaH₂PO₄, 0.1% glucose, 0.1% NaHCO₃, 0.02% KCl) to flow through the mouse circulatory system until the liver and lungs have completely cleared due to the removal of blood (~ 5 min).
6. Continue perfusing the animal with 30% sucrose solution dissolved in Tyrode's solution for an additional 15 min. To evaluate the success of the perfusion, confirm that the neck, tail and legs are rigid post 30% sucrose circulation.
7. Using a small pair of dissection scissors, cut the scalp at the midline. Starting at the occipital bone and working forwards towards the snout. This will expose the cranium.
8. Using a pair of rongeurs, break through the cranium beginning at the occipital bone and continue forward to totally expose the surfaces of the brain. Then turn the head side up and dissect the nerves at the base of the brain to release it from the cranium.
9. Prepare 30% sucrose solution with RNase free water and filter it through a 40 µm nylon mesh filter to reduce RNA degradation.
10. To maximize sucrose solution infiltration, place the dissected brains into 30% sucrose solution and store them at 4 °C overnight. Before further processing ensure that the brain reaches the bottom of the tubes containing the sucrose solution.

3. Cryopreservation of brains harboring glioma tumors

1. Prior to cryopreserving the brains, prepare a jar filled with cold isopentane/2-methylbutane and place the jar into a container filled with liquid nitrogen. Let the solvent cool down.
2. Remove the brain from the 30% sucrose solution and blot it dry on a filter paper.
3. Label the cryomold with a permanent marker. Carefully, add approximately 5 mL of OCT (optimal cutting temperature compound) into the center of the cryomold avoiding air bubbles.
4. Place the brain into cryomold containing OCT in the desired orientation. Fill the mold with OCT until the brain is fully submerged. Using clean forceps, quickly place the cryomold with OCT and the brain into the cold isopentane/2-methylbutane.
5. Once the OCT solidifies (~30-40 s), remove the cryomold with the brain and place it in dry ice. Do not leave the mold containing the brain in 2-methylbutane past 2 min as this may cause cracks in solid OCT. Wrap the cryomold with the brain in aluminum foil and store it at -80 °C.

4. Sectioning frozen brain tumor tissues

1. Label 2 µm polyethylene naphthalate (PEN) slides with the sample information. Tissue sections will be placed directly on these slides following sectioning.
2. Set the temperature of the cryostat chamber between -20 to -24 °C. Before sectioning, place the sample block in the cryostat chamber and let it equilibrate to the temperature in the chamber for 30-60 min.
3. Clean the cryostat chamber and the knife holder with 100% ethanol and spray the brushes to be used with RNase cleaning solution. Working inside the cryostat chamber, remove the mold and attach the OCT block containing the brain to the cryostat specimen disk with OCT. Place the block in the disk holder and align the block with the knife blade.
4. Install a disposable blade into the sectioning holder.
5. Section the brain at 10 µm thickness. Make sure there are no streaks or scratch lines in the tissue. Using a paintbrush, cautiously flatten and uncurl the tissue onto the cutting surface.
6. Carefully mount the tissue containing the brain sections onto RNase free PEN glass slides. Flip the positive charged glass slides with the fingers in direction of the tissue and smoothly press the glass slide down towards the tissue section.
   NOTE: Temperature of hands will help the tissue attach to the glass.
7. After mounting the brain sections onto the slides, keep slides in a box inside the cryostat chamber and then store them at -80 °C. Never keep the slides at room temperature.
   NOTE: Folding of the tissue, and tearing are common. For accurate posterior analysis, it is important to minimize these artefacts.

5. Fixation and staining of cryopreserved brain tissue sections

1. To preserve RNA integrity, clean all instruments to be used with RNase cleaning solution. Proceed with the fixation and staining protocol inside a fume hood.
2. Prepare the described fixative solutions in clean RNase free 50 mL tubes. Make all solutions with RNase free water on the day of the staining.
3. Same day the laser microdissection will be performed, prepare 100%, 95%, 70% and 50% ethanol solutions. Keep the solutions in tightly closed tubes at room temperature.
4. Prepare 4% Cresyl violet and 0.5% eosin Y in 75% ethanol solution. Vortex the solution vigorously for 1 min and filter them through a 0.45 µm nylon filter to eliminate traces of undissolved powder.
5. Place the tissue slides into a container with 95% ethanol for 30 s. Transfer slides to the tube containing 75% ethanol; leave slides there for 30 s.
6. Transfer slides to 50% ethanol and leave them there for 25 s. At this point, the OCT will be dissolved. Transfer the slide to 4% Cresyl violet solution for 20 s, and then transfer to 0.5% eosin Y solution for 5 s.
7. Take the slide out of the dye solution and blot the slide dry with a filter paper. Then, place the slides in 50% ethanol for 25 s. Transfer the slides to 75% ethanol for 25 s. Transfer the slides to 95% ethanol for 30 s. Transfer the slides to 100% ethanol for 60 s.
8. Rinse the slide with xylene. Transfer them to a container with xylene and wait 3 min.
9. Prepare mounting medium (e.g., Pinpoint gum) in RNase-free water. To mount mouse brain sections, dilute the mounting medium in RNase-free water at a ratio of 1:10.
10. Dry the slides on a RNase-free surface at room temperature for 10 s. Before the xylene dries, proceed to mounting the slides with the tissue sections.
11. Gently disperse mounting solution on top of the tissue on the slide with a sterile and RNase-free thin paintbrush. Wait 10-20 s, and then immediately transfer the tissue slides to the microscope microdissection platform.
    NOTE: The ratio of mounting medium to RNase-free water used for mounting varies depending on the tissue of interest. The mounting medium/water ratio preserves glioma tissue morphology for laser microdissection without affecting the RNA integrity.

6. Laser capture microdissection

NOTE: A laser capture microdissection microscope needs to be utilized to laser microdissect specific areas of interest within the tumor tissue. To minimize the time for tissue laser microdissection, have the LMD microscope prepared before fixation and staining.

1. To start the system, first turn on the power strip, followed by turning on the laser. Then, turn on the microscope controller and the computer. Start the LMD software.
2. Under Microscope control, select 10x magnification. Under Laser control, set the laser parameters for tissue dissection. Set a laser frequency of 120 Hz for best cutting results. Always set the laser current to 100%.
3. For accurate laser microdissection, set the speed at 10 and an aperture setting at 2.0-10.0 µm. Set the power to 53. Having the laser power at a higher setting may cause glass etching.
4. Load the tissue collector that will capture the tissue following dissection. Click the second unload button. Remove the empty collector and place the DNase/RNase free 0.5 mL PCR flat head tubes containing 30 µL of lysis buffer into the collector. Place the collector back into the machine and click Devam et on the software to proceed.
5. Load the processed (fixed and stained) specimen onto the microscope. First, click unload on the LMD software. Next, mount the sample on the slide holder and place the slide holder onto the stage. Click Devam et on the software to proceed.
6. Under Cut Shapes windows, select Draw + Cut. Use the microscope controls to find the area of interest. Draw the region of interest (ROI) and select a destination collector tube. It is possible to draw multiple ROIs to micro dissect different areas from a single slide at the same time.
7. Click Start Cut to proceed to tissue micro dissection. After sectioning the areas of interest remove the collector tubes from the holder and place the tubes on dry ice. Transfer the RNA tissue samples collected to -80 °C for long term storage.

7. RNA isolation of micro-dissected glioma tissue

1. For RNA extraction from LMD use an RNA isolation kit optimized for small samples and low RNA yield (see Table of Materials). Follow the manufacture instructions. Carry out all the isolation steps at room temperature (25 °C). To maintain RNA quality, work rapidly. Prepare all the solutions as indicated by the manufacturer.
2. Adjust the sample volume to 350 µL with lysis buffer with 1% β-mercaptoethanol. Vortex the sample for 40 s in order to reduce sample viscosity and increase RNA elution spin column efficiency.
3. Transfer the entirety of the sample to a gDNA eliminator spin column placed in a 2 mL collection tube. Centrifuge the tube for 30 s at 8,000 x g. Save the flow-through and ensure that no liquid is left on the column following centrifugation.
4. Add 350 µL of 70% ethanol to the flow-through and mix well by pipetting up and down. Transfer the sample, to an RNA elution spin column placed in a 2 mL collection tube. Close the lid gently, and centrifuge for 15 s at 8,000 x g. Discard the flow-through, saving the column.
5. Add 700 µL of RNA washing buffer 1 (20% ethanol, 900 mM GITC, 10 mM Tris-HCl pH 7.5) to the RNA elution spin column. Close the lid gently, and centrifuge for 15 s at 8,000 x g to wash the spin column membrane. Discard the flow-through.
6. After centrifugation, carefully remove the RNA elution spin column from the collection tube so that the column does not contact the flow-through.
7. Add 500 µL of the second RNA washing buffer (ethanol 80%, NaCl 100 mM, Tris-HCl 10 mM pH 7.5) to the spin column. Close the lid of the column and spin 8,000 x g for 20 s to wash the column membrane. Dispose the flow-through.
8. To wash the RNA elution column, add 500 µL of 80% ethanol, close the lid of the column and centrifuge at 8000 x g for 2 min. Throw out the tubes with the elution solution.
9. Use a new collection tube, open the lid of the spin column and centrifuge at 8000 x g for 5 min to dry the column. Discard the elution tube.
10. Place the RNA elution spin column in a new 1.5 mL collection tube. Add 12 µL of RNase-free water warmed at 37 °C directly to the center of the spin column membrane. Wait for 4 min and centrifuge for 1 min at full speed to elute RNA.

8. RNA quality control, library preparation and RNA-Seq analysis

1. Following RNA extraction and purification, amplify RNA and create a cDNA library using a special kit suitable for RNA isolation at pico-molar concentrations following manufacturer instructions (see Table of Materials). Follow manufacturer's instructions. Clean workstation in order to avoid contamination with other PCR products and nuclease degradation of samples.
2. If RNA has a RIN value greater than 4, meaning the RNA is of good quality or only partially degraded, proceed with the fragmentation step. Prepare all items on ice.
3. Create a master mix as indicated (Table 1). Create reaction mixture in a nuclease-free thin-wall 0.2 mL PCR tube. Incubate the tubes at 94 °C in a hot-lid thermal cycler. Fragmentation incubation time depends on the quality of the RNA. RIN ≥ 7: 4 min, RIN 5-6: 3 min, RIN 4-5: 2 min.
4. Following incubation, place the tubes on a PCR chiller rack that was previously cooled at -20 °C and let it sit for 2 min.
5. For each tube of RNA sample, prepare first strand synthesis reaction master mix (Table 2). Incubate the tubes in a hot-lid thermal cycler under conditions described in Table 3. cDNA products can be frozen at -20 °C until for two weeks before proceeding to the next step.
6. Create the PCR master mix as indicated in Table 4. Place the tubes in a hot lid thermal cycler and run the PCR reaction under the settings indicated in Table 5.
7. Allow the beads, which will be used to purify the DNA, to warm to room temperature. Once warmed, add 40 µL of the beads to each sample. Vortex the tubes to mix and spin down briefly to collect liquid at bottom of the tube. Let the tubes incubate at room temperature for 8 min to allow the DNA to bind to the beads.
8. Place the tubes on a magnetic separation device and let sit for about 5 min, or until the solution becomes completely clear. Keeping the tubes on the separation device, use a pipette to remove the supernatant carefully without disturbing the beads.
9. Keeping the tubes on the separation device, add 200 µL of freshly made 80% ethanol to the beads to wash without disturbing the beads. Wait for 30 s before removing the 80% ethanol. Repeat this step.
10. Briefly spin down the tubes and place the tubes back on the separation device. Remove any residual 80% ethanol without disturbing the beads.
11. Let the tubes air dry with the caps open for 5 min. Do not let it sit longer as the beads will over dry.
12. Keeping the tubes on the separation device, add 52 µL of nuclease-free water to cover the beads. Remove the tubes from the separation device and pipette up and down until all the beads are resuspended. Incubate at 5 min at room temperature.
13. Place the tubes back on the separation device until solution becomes clear, about 1 min. Transfer 50 µL of the resulting supernatant to the wells of an 8-well strip. Add 40 µL of new beads to each sample. Vortex thoroughly to mix. Allow beads to bind to DNA by incubating at room temperature for 8 min.
14. During this incubation period, begin thawing the components to be used for any rRNA present in the samples. Once thawed, place them on ice. Also, pre-heat a thermal cycler to 72 °C.
15. Place the samples on the magnetic separation device until the solution clears (about 5 min). Aliquot 1.5 µL of probes per sample to a chilled PCR tube. Place the tube in the preheated thermal cycler under the settings of 72 °C for 2 min and 4 °C.
16. Keeping the sample tubes in the magnetic separation device, remove the supernatant with a pipet without disturbing the beads. Then add 200 µL of freshly made 80% ethanol to the beads to wash without disturbing the beads. Wait 30 s before removing the 80% ethanol. Repeat this step.
17. Briefly spin down the tubes and place the tubes back on the separation device. Remove any residual 80% ethanol without disturbing the beads. Let the tubes air dry with the caps open for 2 min. Do not let sit longer as the beads will over dry.
18. Prepare master mix for all samples by combining the following components in the order presented (Table 6).
19. Mix the master mix by vortexing and add 22 µL of the mix to the dried beads of each sample and mix thoroughly to resuspend. Let it incubate at room temperature for 5 min.
20. Spin down the tubes and place them on the magnetic separation device for 1 minute or until the samples become clear. Transfer 20 µL of the supernatant, without disturbing the beads to new PCR tubes. Place the tubes in a pre-heated thermal cycler under settings in Table 7.
21. Prepare a PCR master mix for enough for reactions for each sample. Add the following components to the master mix in the indicated Table 8. Add 80 µL of the PCR master mix to each of the sample tubes from step 8.20 and place in the thermal cycler at the following settings (Table 9).
22. Allow beads, which will be used to purify the DNA, to warm to room temperature. Once warmed, add 100 µL of the beads to each sample. Let the tubes incubate at room temperature for 8 min to allow the DNA to bind to the beads.
23. Place the tubes on a magnetic separation device and let sit for about 5 min, or until the solution becomes completely clear.
24. Keeping the tubes on the separation device, use a pipette to remove the supernatant carefully without disturbing the beads.
25. Keeping the tubes on the separation device, add 200 µL of freshly made 80% ethanol to the beads to wash without disturbing the beads. Wait for 30 s before removing the 80% ethanol. Repeat this step.
26. Briefly, spin down the tubes and place the tubes back on the separation device. Remove any residual 80% ethanol without disturbing the beads.
27. Let the tubes air dry with the caps open for 5 min. Do not let sit longer as the beads will overdry. Pipette 20 µL of Tris Buffer to the dried pellet. Remove the tube from the separation device and mix thoroughly with a pipet to resuspend the beads. Let it incubate at room temperature for 5 min.
28. Place the tubes on the separation device for 2 min, or until solution becomes clear. Acquire the supernatant and transfer it to new tubes. Then store the collected solution at -20 °C.
29. Check the final libraries need for quality and quantity control. Pool the samples, clustered on and sequence, as paired-end 50 nt reads, according to manufacturer's recommended protocols.