With TRAP, we can investigate developmental processing at cellular resolution. We use cell-type-specific promoters to drive a tagged ribosomal sub-unit, and from there we can isolate polysome RNA. This protocol yields high quality RNA, but in very limited quantities, therefore, methods that are specialized for ultra-low input are needed to reliably produce high quality RNA libraries.
TRAP is an ideal tool for plant research, because many developmental processes rely on cell wall-related and mechanical signaling pathways. We use TRAP to study cell-cell communication in lateral root formation. As obtaining good quality RNA proved not as straight-forward as we anticipated, we hope that this step-by-step video instruction will increase TRAP applicability and help promote this powerful technique.
To sterilize the propagated homozygous lines, spread less than 300 microliters of seeds evenly on 12 by 12 centimeter squared Petri dishes, and stack the dishes on a 60 liter desiccator. After overnight gas sterilization, let the gas evaporate before collecting the seeds into a 50 milliliter storage tube. Before plating, dilute the sterilized seeds with 0.1%agar to obtain one milliliter of imbibed seed mix per plate.
To plate the seed mix, use square Petri dish lids to create a laminated template holder for planting three rows of seeds per plate, and place empty agar plates into the template holder. Distribute one milliliter of the imbibed seeds evenly onto three rows of plates. And place the processed plates in stacks in a laminar flow, until the seeds are dry.
When seeds have stuck to the agar's surface, close the lids and seal each plate with micropore tape. For exogenous treatment of Arabidopsis roots with an agent of interest, prepare 1.5 to two centimeter-wide, 10 centimeter-long, strips of tissue paper. Soak the tissue paper in 10 micromolar NAA, and use tweezers to apply a strip of tissue paper to each row of roots.
Then, gently press out any air bubbles, and empty the excess liquid from the plate, before labeling the plate with the time. At the appropriate experimental end point, use forceps to carefully remove the tissue paper strips from each plate without detaching the roots, and use a surgical blade to cut once per row along the shoot-root junction, in a single determined stroke. Use tweezers to swipe along the roots of each row, to collect the roots in three bundles.
And empty the the roots into a 50 milliliter tube filled with liquid nitrogen, for snap freezing. When all of the root samples per treatment have been collected, decant the excess liquid nitrogen using the tube lid to prevent the roots from spilling. And place the tube into a doer vessel, for a 80 degrees celsius storage.
For tissue grinding and homogenization, wear cotton gloves under standard lab gloves, and add PMSF to the polysome extraction buffer. Empty the tissue sample into a mortar containing liquid nitrogen, and use a cooled pestle to carefully grind the tissues until all the material appears as a white powder. When all of the tissue has been ground, add five milliliters of polysome extraction buffer to the sample, and quickly mix the frozen tissue fragments with the powder before the buffer freezes.
As soon as the mixture can be transferred, empty the slurry into a glass homogenizer on ice, and use an additional two milliliters of polysome extraction buffer to rinse the mortar and pestle. Manually grind the slurry until the extract is homogenous. Then, pour the crude root extract into a 50 milliliter centrifuge tube on ice, and process the next sample as demonstrated.
For total RNA collection, transfer 200 microliter aliquots of each crude sample into individual clean, pre-cooled, labeled microcentrifuge tubes. To remove cell walls, debris, and large organelles, centrifuge the samples and decant the supernatant into fresh, pre-cooled conical tubes for a second centrifugation. While the tubes are spinning, aliquot 60 microliters of magnetic GFP beads per sample, into a 1.5 milliliter tube.
And place the tube on a magnetic stand, to facilitate supernatant removal. Then, wash the beads two times in one milliliter of cold wash buffer per wash, and re-suspend the beads in 60 microliters of wash buffer per sample. Immediately after the centrifugation, decant the cleared supernatant into labeled 15 milliliter tubes, and add 60 microliters of washed beads to each sample.
Then, place all of the samples horizontally into an ice bucket, for a two hour incubation with rocking. At the end of the incubation, collect the beads on a magnetic stand on ice, and add PMSF to the remaining polysome extraction buffer. After discarding the supernatants, add approximately five milliliters of polysome extraction buffer to the samples, and re-suspend the beads within the samples with gentle tilting.
Rock the samples for 15 minutes on the shaker on ice, followed by two washes with fresh wash buffer on the magnet, as demonstrated. After the last wash, transfer the samples in one milliliter of wash buffer, into a 1.5 milliliter tube. And collect the beads one more time on the magnetic stand, to remove all of the supernatant.
Then, place the tubes on ice until all of the samples have been processed. For reconditioning of the lab supplies, first, rinse all of the equipment into the chemical waste. Hand wash the mortars, pestles, and homogenizers with soap, before rinsing each instrument thoroughly.
Next, place the equipment in a heat-safe container, and bake the materials overnight at greater than 220 degrees celsius. Brush all of the centrifuge tubes clean with detergent, before placing the tubes onto an autoclaveable tray with a rim and a fume hood. Pour a one milliliter of liquid diethyl pyrocarbonate and one liter of deionized water solution onto the tubes for three to 18 hours.
Then, collect the solution before sterilizing the tray of tubes in the autoclave. Here, representative plants with GFP signals are shown. Counter-staining with propidium iodide marks the cell wall outlines.
The lines correspond to the localization pattern of the endodermis, and the xylem-pole pericycle. In these representative measurements obtained from polysome RNA, most samples showed very little degradation, with RNA integrity numbers ranging from nine to 10. In these graphs, traces of successfully prepared libraries can be observed, highlighting the robustness of the procedure, despite scaled-down reaction volumes.
Before a genome-spanning dataset is produced, TRAP RNA from a pilot experiment can be probed by quantitative RTPCR to validate the treatment success and/or the experimental conditions, as well as the tissue specificity. In this analysis, the testing of three auxin-responsive genes confirmed a successful hormone induction after two hours of treatment. Tissue-specific marker gene profiles retrieved from the sequencing data confirmed the cell-type specificity.
Follow good practice advice when you handle RNA. Work at a sterile bench, clean your equipment with RNA-removing solution, wear gloves and immediately change them when they get contaminated. Be sure to appropriately handle and discard all toxic chemicals and gases that have been used during the procedure, especially phenol-DEPC chlorine gas and liquid nitrogen.
In the portfolio of omic studies, TRAP occupies an important niche, and has already been used to answer many biological questions, especially in developmental biology, TRAP has a huge potential.
