The overall goal of this procedure is to recover high quality RNA from spinal cord motor neurons without contamination by RNA from neighboring cells. This is accomplished by first recovering, dividing and embedding the cord in cryo embedding medium and freezing at negative 160 degrees Celsius. The second step is to cryo section the cord onto pen membrane slides at negative 20 degrees Celsius.
Next, the slides are washed in 70%ethanol and stained with Azure B in 70%ethanol. The final step is laser capture micro dissection of identified motor neuron cell bodies into guine thiocyanate lysis buffer. Ultimately, total RNA is prepared from the combined lysates and tested to uncover differences in general or specific RNA transcription in neurons of mutant versus wild type animals.
Laser capture microdissection allows recovery of motor neuron cell bodies from spinal cord sections without contamination by the more numerous Scalia and other cell types. Affording a chance to prepare RNA and compare transcription between neurons from an diseased animal and a normal one. The most difficult aspect in developing this procedure was to find a staining method that would allow us to detect motor neurons in spinal cord sections while maintaining high levels of RNA integrity and extract ability, both before and during laser capture.
We found that staining sections in Azure B and 70%ethanol met these criteria when combined with collection of the dissected cell bodies directly into Guine Thiocyanate lysis Buffer demonstrating the procedure is ine Van Padia, a postdoc in our lab For the best results. Make sure that all equipment is cleaned with RNAs decontaminating solution followed by RNAs free 70%ethanol after euthanasia and trans cardial perfusion Carefully isolate the spinal cord. Rinse the cord for 10 seconds in RNAs.
Free water to wash off any residual blood and lay it on an RNAs free glass slide very gently but quickly. Next, divide the spinal cord transversely using a clean razor blade into nine to 10 pieces. Position the pieces in a cryo mold filled with OCT and align them vertically using a clean needle.
Place the mold in a shallow tray containing two methyl butane pre-cool with liquid nitrogen. To avoid RNA degradation, place the tray into a liquid nitrogen bath. To fast freeze the spinal cord pieces.
Store the OCT embedded block at negative 80 degrees Celsius for up to six months before sectioning. When ready for sectioning position, the OCT embedded block into a chilled cryostat. Produce 20 micron slices each containing nine to 10 spinal cord cross sections.
Place the sections on RNAs free pen membrane. Two micron slides kept initially at room temperature to ensure that the sections adhere. Well immediately refreeze the section on a negative 20 degrees Celsius surface inside the cryostat.
Keep the slides at negative 80 degrees Celsius until used on a clean rack range. Six 50 milliliter conical tubes filled with RNAs free.70%ethanol. The depth should be sufficient to cover all the sections on a slide when the slide is immersed.
Next place 1%Azure B solution on the same rack. To minimize the time between changing solutions during washing or staining, keep three or four lab wipes in front of the rack to drain extra solution quickly. When ready, take the slides out of the negative 80 degrees Celsius freezer and place them onto dry ice thaw one slide by placing it on a gloved palm.
Remove most of the moisture and condensation on the slide by wiping it off with a lab wipe, being careful not to touch the sections. Put the slide on fresh silica gel desiccant in a Petri dish for 30 to 40 seconds to dry completely. When dry, dip the slide into the first tube of RNAs, free 70%ethanol.
Soak the slide for 30 seconds and then wash off the OCT by dipping it up and down for 45 seconds to one minute. Next, take the slide out of the solution and drain the excess liquid on lab wipes. Repeat this process by dipping the slide in the second tube of 70%ethanol.
If OCT remains around the spinal cord sections, repeat the washing in the third tube. After draining the excess liquid, submerge the slide in 1%Azure B solution in 70%RNAs, free ethanol for 30 to 45 seconds. Drain the excess Azure B on a lab wipe.
At this point, the whole slide will be blue. Submerge the slide in the next tube of 70%ethanol and dip up and down quickly, three to four times to remove excess dye and to make the specific motor neuron staining visible. If the sections look very darkly stained, repeat the des staining step in a fresh tube of 70%ethanol.
If the staining seems too light, return the slide to the Azure B solution for another 32nd and repeat the des staining. If the staining is satisfactory, allow the slide to air dry briefly and proceed to the next step. After DES staining and drying, the gray matter will be light blue and the deep blue stained motor neurons will be easily visible.
Start the dissection immediately after turning on the microscope. Unload the tube holder to attach the cap of a 0.6 millimeter micro fuge tube. For sample collection, put 30 microliters of guine and thiocyanate lysis buffer into the cap and cover the surface by spreading the solution.
With a pipette tip, align the cap with the hole and the objective. With the microscope software wear, keep the cap in the covered position until starting laser capture. Place the air dried slide upside down on the stage of the microscope so that the membrane side faces down and the section is aligned with the hole.
With the slide in place, focus on a spinal cord section with the five x objective and then move to the 20 x objective for dissection. First, mark a dummy region with the light pen and allow the laser to cut it out without collecting it. This relaxes the membrane and makes it possible to mark and cut multiple regions successively.
Next, refocus and identify motor neurons in the ventral horn region. These neurons are recognizable by their location parmal morphology, large size and dark blue staining with Azure B.Using the light pen, select the drawing tool and mark along the edge of the motor neurons making a complete outline. Try to make the outline as close as possible to the motor neuron to avoid contamination from other cells.
Multiple cells can be marked before cutting since the software will remember the positions when ready to cut, bring the cap underneath the cutting position and click the start button to initiate motor neuron dissection with the laser. Collect all the motor neurons from all of the sections on one slide into the cap of one micro fuge tube. After collection is complete, take the micro fuge tube out of the holder by unloading the tray and gently dislodging the tube completely dissolve the motor neurons and buffer by pipetting the solution up and down.
Move the solution into the body of the tube by centrifuging, freeze the sample immediately in dry ice and store it at negative 80 degrees Celsius after staining with Azure B.The motor neurons appear as large darkly stained cell bodies. They're confirmed by anti CHATT antibodies, staining and are easily differentiated from smaller neighboring cells. In this example, the motor neuron cell bodies have been outlined with the light pen and numbered by the software.
Here the cell bodies have been cut out with the laser and dropped into the collection tube with the outlines turned off. The extent of the laser damage to neighboring regions is apparent. Typical results of an electrophoretic analysis of the RNA integrity of a sample from approximately 4, 000 mouse motor neuron cell bodies collected by LMD are shown here.
Note the prominent ribosomal RNA peaks. This process from slide staining through motor neuron dissection should be completed within 30 minutes for all of the sections on one slide to minimize RNA degradation. When attempting this procedure, it's important to remember to work as quickly as possible, both in the initial dissection and embedding of the cord, as well as in the final steps of staining and laser capture microdissection of the motor neurons.
