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Letters to the Editor
mising data quality, as minor protocol deviations were commonly observed at initial implementation.
Between 2016 and 2019, MFC MRD results from 20 MM patients were compared within the EMN MRD QA program among four EMN reference laboratories willing to commit to EuroFlow protocols in the context of the EMN02/HO95 MM trial: Aalborg University Hospital, Denmark (laboratory 1), University Hospital Brno, Czech Republic (laboratory 2), Erasmus MC Rotterdam, the Netherlands (laboratory 3, EuroFlow member) and University of Turin, Italy (laboratory 4).10,11 In total, four QA rounds were organized, each comprising five differ- ent fresh samples from MM patients with variable levels of disease burden and variable treatment histories (Figure 1A). Samples were collected at Erasmus MC Rotterdam, the Netherlands and Ospedale Molinette di Torino, Italy, on random days throughout the year. This study was approved by the Medical Ethical Committees of Erasmus MC Rotterdam, the Netherlands and A.O.U. Città della Salute e della Scienza di Torino, Italy. Written informed consent was obtained from all participating patients, in accordance with the Declaration of Helsinki. Immediately after collection, samples were divided equally and shipped by overnight express courier to the participating laboratories. Samples from distributing hos- pitals were kept at room temperature for 24 h to ensure similar sample processing dates between laboratories. Using standardized forms, MRD results were collected centrally by one person, who kept these confidential until the end of each QA round, after which results were shared and discussed.
Timely sample processing is an essential prerequisite for high validity of MFC MRD results, as MM cells have a limited capacity to survive outside of the bone marrow. Hence, the IMWG recommends processing MFC MRD samples within 24-48 h. Considering all 67 samples from QA rounds 1-4, our data show that two laboratories were able to process 20/20 (100%) received samples within this recommended timeframe. Laboratory 1 processed 6/7 (86%) and laboratory 4 18/20 (90%) sam- ples within 48 h after sampling (Figure 1B).
Throughout QA rounds 1-4, laboratory 3 adhered strictly to EuroFlow standardized operating procedures, which was considered the reference for all other partici- pating laboratories. In QA rounds 1-2, second-generation flow protocols from EuroFlow were applied. QA round 1 was followed by a workshop to further standardize pro- tocols and gating strategies, which resulted in the use of significantly more comparable standardized operating procedures between laboratories in QA round 2 (Online Supplementary Tables S1 and S2). A minimal number of 20 monoclonal plasma cells (mPC) was required for MRD positivity.12 Despite complete standardization of proto- cols not being possible in laboratories 2 and 4 because of ongoing consumable contracts and local unavailability of certain reagents, MFC MRD results were highly concor- dant in QA rounds 1-2 at every level of residual disease. All participating laboratories reported the same MRD result for 9/10 (90%) samples (Figure 2A).
The ability to uniformly quantify MRD irrespective of daratumumab treatment status was tested in seven bone marrow samples that were distributed in QA rounds 3-4. Here, the EuroFlow next-generation flow pipeline was implemented. This pipeline contains a multi-epitope anti- body against CD38 in its staining panel, which circum- vents epitope blocking by daratumumab.6 Of note, at this stage all participating reference laboratories had commit- ted to fully standardized protocols in terms of data collec- tion, instrument setup, performance checks, sample
preparation, sample staining, data acquisition and data analysis (Online Supplementary Tables S1 and S2), result- ing in a second series of highly concordant MFC MRD results and 10/10 (100%) samples with a uniformly clas- sified MRD result (Figure 2A).
To compare interlaboratory test sensitivities in MRD- negative samples, the formula for limit of detection (LOD) was used: 20/number of acquired leukocytes. This showed a median LOD of 5.4 x 10-6 in the 34 MRD-neg- ative samples from QA rounds 1-4. Laboratory 3 reached a LOD <0.001% in 10/10 (100%) MRD-negative assays, whereas the other laboratories achieved a LOD <0.001% in 50-80% of MRD-negative assays. Overall, in all except one assay a LOD <0.01% was reached.
Recent reports indicate that the majority of newly diag- nosed MM patients have detectable mPC in their peripheral blood (i.e., circulating tumor cells) when the highly sensitive next-generation flow protocols are used.13 Recent reports indicate that the majority of newly diagnosed MM patients have detectable mPCs in their peripheral blood (i.e., circulating tumor cells) when the highly sensitive next-generation flow protocols are used.13 As mPC infiltration is typically low in both newly diagnosed MM peripheral blood and MRD bone marrow samples and the collection of peripheral blood is substan- tially less invasive than that of bone marrow, it has been questioned whether newly diagnosed MM peripheral blood samples could also be used for MM MRD QA pur- poses. The feasibility of doing so was assessed in QA round 4. Circulating tumor cells were uniformly detected in 2/2 (100%) peripheral blood samples from newly diag- nosed MM patients, both at highly comparable levels between 0.001% and 0.01%. This indicates that periph- eral blood samples from newly diagnosed MM patients may indeed be used as an alternative to MRD bone mar- row samples to assess interlaboratory standardization of MM MRD protocols.
To test the interlaboratory concordance of the detected mPC immunophenotypes, laboratories were asked to report staining intensities as positive, dim or negative. 10/20 (50%) samples were classified as MRD-positive and generally showed strong similarity between labora- tories for markers that are essential for mPC gating: CD38, CD138, CD45, CD19, CD56, CyIgK and CyIgL (Figure 2B). The reported expression of other informative markers (i.e., CD27, CD81 and CD117) showed more variability. Even though this did not affect mPC quantifi- cation, it underscores the importance of using strict defi- nitions in terms of data analysis to ensure reproducibility of mPC immunophenotype data.
Finally, MFC MRD assessment has the advantage over molecular MRD techniques that it also generates infor- mation on the cellular composition of non-MM popula- tions, which could be used to infer the quality of bone marrow samples. To this end, the EMN suggested in its consensus from 2008 that the polyclonal plasma cell (pPC) levels should always be stated in the final MRD report.14 To test the concordance of this reference popula- tion between laboratories, information on pPC levels was collected from all samples in QA rounds 2-4 (Figure 2C). As expected, peripheral blood samples had a lower medi- an pPC level than bone marrow samples. The interlabo- ratory concordance of reported pPC levels was generally good, although it was inferior to that of reported MRD levels, suggesting that pPC levels are more susceptible than mPC levels to interlaboratory variations in sample processing and data analysis.
In conclusion, our data indicate that full standardiza- tion of interlaboratory MM MFC MRD assessment is fea-
haematologica | 2021; 106(5)