Comprehensive Protocol to Sample and Process Bone Marrow for Measuring Measurable Residual Disease and Leukemic Stem Cells in Acute Myeloid Leukemia

The overall goal of this protocol is to perform an accurate testing of bone marrow samples from acute myeloid leukemia patients for measurable residual disease including the quantification of leukemia stem cells. This method can provide an accurate assessment of measurable residual disease in bone marrow samples of acute myeloid leukemia patients. The main advantage of this approach is that a close following of the protocol yields highly-reproducible, high-quality results.

The implications of this technique extends toward clinical decision-making as it can be used as a read-out of patients'response to treatment. Demonstrating the procedures will be Jeroen Janssen, hematologist, Gerrit Sijm, technician of the diagnostic hematology laboratory and Jennifer Scheick and Sander Snel, technicians of the MRD team. With the patient in the lateral decubitus position, mark the superior posterior iliac spine with a pen and disinfect the skin of the intended biopsy area with 0.5 to 1%chlorhexidine in ethanol in an outward circular motion.

After administering a local anesthetic, pick up a 15-gauge 2.8-inch needle with the proximal end in the palm and the index finger against the side of the metal shaft of the needle near the tip for control. Using gentle but firm pressure, introduce the needle with a quick alternating pronating supinating movement through the anesthetized skin toward the iliac spine. When the needle comes into contact with the posterior iliac spine, remove the stilette and attach an empty 10-milliliter syringe to the needle.

Gently withdraw the plunger to create negative pressure within the syringe until up to 10 milliliters of bone marrow spicules have been harvested. Do not take more than 10 mils of bone marrow per aspirate to avoid hemodilution. Remove the syringe and place the stilette back onto the needle.

Then, eject about two milliliters of the marrow into a watch glass and ensure that enough sample is drawn to be injected into a heparin-coated eight-milliliter tube and immediately invert the tube. To prevent clotting, it is important to invest the anticoagulant-containing tube as soon as the bone marrow is added. For optimal morphological assessment, use a plastic spatula to transfer spicules from the aspirate in the watch glass onto a glass slide, and carefully glide another glass slide on top of the marrow.

After drying, stain the spicules with May-Grunwald Giemsa for analysis by light microscopy. Place the bone marrow tube in a plastic holder and place the holder into a plastic container with an ambient gel pack and a completed request form. Before analyzing the bone marrow by flow cytometry, start up the flow cytometer and open the flow cytometry analysis software to create a new experiment with a forward scatter versus side scatter dot plot.

For assessment of the size and granularity of the cells, load unlabeled lysed peripheral blood cells from a healthy donor and select the Acquire Cells function. Gate the lymphocytes and adjust the forward and side scatter voltages to reach the appropriate mean target values for the gated cell population. Then, acquire data for at least 10, 000 events.

To set up the target florescence channel photo multiplier tube voltages, first use eight-peak rainbow bead calibration particles according to the manufacturer's instructions to acquire 10, 000 events at a low flow rate. Gate the singlet beads in the forward versus side scatter dot plot followed by gating the 7th peak in the FITC-PE dot plot. Gate the resulting bead populations for each fluorophore.

Continue the acquisition of the eight-peak rainbow bead suspension and adjust and fine-tune PMT voltages in all fluorescence channels to reach target MFI values according to the manufacturer's instructions. Use Record to collect data of about 2, 000 events and check the MFI for the 7th peak beads. Next, add a sufficient volume of each experimental flourochrome conjugated antibody to stain the beads in the corresponding florescence tubes minus one control and vortex the tubes thoroughly.

Create a new blank experiment with the same compensation parameters taking care that the forward scatter threshold is set to 5, 000 and the compensation values of all the flourochromes are zero. To create the compensation controls, select the Create Compensation Control function and load the unstained control tube. Adjust the P1 gate around the singlet bead population and confirm that the P1 gate contains only singlet beads.

Right-click the P1 gate and select Apply to All Compensation Controls. Then, record data for all of the single labeled florescence tubes confirming that the P2 gate encompasses the positive population on each florescence histogram. To stain the cells for flow cytometric analysis, transport the bone marrow to the laboratory, check the request form to verify patient study number and date of birth and determine what needs to be stained.

For MRD, we use a staining panel consisting of four tubes. Here we demonstrate the staining of the LSC panel which consists of one tube. After cell count, transfer the appropriate volume of bone marrow into a new 15-milliliter tube.

Add lysis buffer at 10 times the experimental cell volume to lyse the red blood cells. Invert the tube to mix and incubate the cells for 10 minutes at room temperature. At the end of the lysis period, pellet the cells by centrifugation.

Resuspend the cells in 15 milliliters of wash buffer at room temperature and harvest the cell pellet again by centrifugation. In the meantime, pipette the appropriate amount of antibody cocktail solution in five-milliliter FACS tubes. Shown here is the panel for the stem cell tube.

Resuspend the pellet in wash buffer and add the cell's suspension to the FACS tube containing the monoclonal antibody cocktail solution for a 15-minute incubation in the dark. Then, wash the cells in wash buffer and pellet the cells by centrifugation. After centrifugation, resuspend the pellet in 400 microliters of fresh wash buffer.

To analyze the leukemic stem cells, add four microliters of blank beads to the cell suspension that is stained with the stem cell antibody cocktail. Then, read the samples using the previously set parameters acquiring at least four times 10 to the 6th events from the experimental patient samples. To identify the leukemia-associated immunophenotype observed in approximately 90%of acute myeloid leukemia patients, it is crucial that the blast gates are set appropriately as illustrated.

Here, examples of data from individual patients harboring different types of aberrancy on the CD34-positive primitive cells are shown. In these graphs, a patient who remains in complete remission and a patient who relapsed after having measurable residual disease positive results after the second cycle of induction therapy can be observed underscoring the need for accurate gate setting prior to sample analysis. The identification and quantification of LSC requires additional analysis of aberrancies specifically on CD34-positive CD38-negative blast cells as shown in these graphs.

While performing this protocol, it's very important to have specialized and backup personnel for the bone marrow sampling, the transport, the experimental work and analysis steps of this procedure. This procedure can also be used to perform cytogenetic and molecular analysis to answer additional questions about the correlation between specific cell populations and molecular aberrancies of interest, or to refine the risk group of a patient for predicting clinical outcome. After its development, this technique paved the way for researchers in the field of hematology to explore residual disease in AML patients after treatment.

After watching this video, you should have a good understanding how we accurately identify and quantify residual disease by using flow cytometry including the collection and transportation of bone marrow samples of AML patients.