Minimal Residual Disease Detection by Flow Cytometry in Multiple Myeloma: Why and How?

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Minimal Residual Disease Detection by Flow Cytometry in Multiple Myeloma; Why and How? Mikhail Roshal

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To appear in: Seminars in Hematology Cite this article as: Mikhail Roshal, Minimal Residual Disease Detection by Flow Cytometry in Multiple Myeloma; Why and How?, Seminars in Hematology,doi:10.1053/j.seminhematol.2018.02.011 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Minimal Residual Disease Detection by Flow Cytometry in Multiple Myeloma; Why and How? Mikhail Roshal Hematopathology Service, Department of Pathology Memorial Sloan Kettering Cancer Center New York, New York, USA Ph: 212 639-6091 Fax: 929 321-1513 Email: [email protected] Conflict of interest: The author has received consulting fees from Celgene for a project that was not in connection with myeloma therapy or monitoring. Abstract The outlook for myeloma patients has steadily improved with the introduction of newer drug combinations in recent years. Unlike older therapies that largely achieved only modest levels of neoplastic clone reduction, the newer drug combinations have led to deeper suppression of myeloma clones in most patients. Frequently the neoplastic clones become undetectable with traditional disease evaluation approaches. Recent studies using ultrasensitive disease monitoring have demonstrated that patients with disease undetectable by traditional techniques show wide heterogeneity in disease levels varying by several orders of magnitude. Moreover, measurement of the depth of disease suppression even at very low level has emerged as the most powerful prognostication tool in myeloma. Minimal (or measurable) residual disease (MRD) evaluation has also been proposed as a relevant tool in assessment of drug efficacy and in selection of further therapeutic options. In the face of the robust MRD measurement utility data, it has become critical to develop widely applicable disease monitoring techniques that can be applied to more patients in a variety of clinical setting. Both DNA-based and flow cytometry-based approaches have been successfully developed for this purpose achieving sensitivity approaching 1 neoplastic cell in a million. This review article focuses on the theoretical and practical aspects and challenges of deep MRD monitoring in myeloma by flow cytometry. Challenges of flow cytometric disease monitoring in the era of antigen-directed therapy are also discussed. Evolving treatment for multiple myeloma results in deeper responses requiring more sensitive assays Until very recently multiple myeloma was considered an incurable disease with conventional therapeutic approaches. The assumption is based on the near uniform relapse of patients post treatment with older therapies [1, 2]. Relapses are almost certainly due to the residual neoplastic clones that constitute a reservoir of therapy-resistant disease. In the past, with less effective therapies the residual neoplastic cells could be readily demonstrated, even by relatively insensitive residual disease detection techniques such as fluorescence in-situ hybridization, immunohistochemical studies, and 4-color flow cytometric approaches with detection limits ranging from 0.01% to 5% [3-6]. Recent years have seen an explosion in availability of novel therapeutic options for treatment of the disease[2]. The advances in therapy are reviewed elsewhere in the issue. These newer therapies have resulted in longer progression free and overall survival improving the outlook of myeloma patients [2, 7, 8]. The improvements were also seen outside specific clinical trials. For instance, Fonesca et al. have shown a steady increase in overall survival in myeloma patients paralleling introduction of new therapies. Patients diagnosed in 2012 are 1.25 times more likely to survive the first two years compared to those diagnosed in 2006 according to US administrative claims [9]. Many patients are achieving durable remissions with newer therapies with residual neoplastic clones becoming undetectable by older methodologies. Fewer but nonetheless significant number of

patients becomes negative with ultrasensitive techniques with sensitivities of 5*10-5 to 10-6[10, 11]. For example 68% of patients showed MRD negativity at level of 2.5*10-5 with front-line transplantation program with Lenalidomide, Bortezomib, and Dexamethasone combination as induction and consolidation followed by Lenalidomide maintenance therapy[12], while only 42% of patients were MRD negative with a relatively insensitive (10-4) test when treated with 6 alternating cycles of vincristine, carmustine, melphalan, cyclophosphamide, prednisone (VBMCP) and vincristine, carmustine, doxorubicin, dexamethasone (VBAD), followed by high-dose melphalan and ASCT[13] While the data is not directly comparable, the studies illustrate a steady improvement in depth of therapy response with new drug combinations. Despite rapid progress with the modern drug combinations responses to therapies remain heterogeneous. The more durable clinical responses are strongly linked to deeper levels of suppression of the neoplastic clones. Two recent metanalysis studies confirm strong association between the depth of response and overall outcome across a number of recent clinical trials[14, 15]. Moreover the depth of therapy response as evidenced by the relative size of the remaining neoplastic clone is either the only or possibly one of the very few independent predictive factors for multiple myeloma patient outcomes regardless of a specific therapy[16-21]. Importantly, a retrospective pooled analysis of three large clinical trials demonstrated that patients who achieved complete remission without MRD negativity showed no significant increase in overall or progression free survival as compared to patients with at least partial response. In contrast, those who were MRD negative post treatment showed marked increase in progression free and overall survival[19]. Given the wealth of available highly effective treatment options, it is possible that measurements of residual disease might be used to effectively guide further therapeutic choices. This hypothesis is strengthened by a large clinical trial, which showed that thalidomide maintenance could convert some patients from MRD positive to MRD negative state [21]. For more extensive treatment of the topic readers are referred to an excellent prior review[22]. New sensitivity requirements for myeloma MRD evaluation As awareness of MRD measurement utility becomes more widespread, it becomes increasingly important to at least establish a common language of defining MRD positivity. Two consecutive surveys of large academic laboratories in the United States and a more general College of American Pathologists survey of labs claiming to offer myeloma MRD testing have demonstrated a wide heterogeneity in MRD cutoffs ranging from 10-3 (plurality in the CAP survey (50% of respondents)) to <10-5 [23-25]. Recent studies have shown that more sensitive assessment of myeloma MRD results in more robust longer-term prognostication. For instance using next generation sequencing approach MartinezLopez and colleagues showed that patients with at least very good partial response with myeloma clone suppression below 10-5 had median progression free survival (PFS) of 80 months, those with neoplastic clone in the 10-3 to 10-5 had PFS of 45 months, and those above 10-3 had PFS of 27 month[26]. It is therefore obvious that a negative test result with sensitivity of 10-3 has significantly different prognostic implications from negativity at 10-5. Moreover based on the distribution of results in the study a test with sensitivity of 10-3 would be considered a false negative in approximately 55% of patients compared to that with sensitivity of 10-5. It is in part based on the data that international myeloma working group (IMWG) has recommended MRD testing with sensitivity of at least 10-5 by either flow cytometric or next generation sequencing approaches [27]. The efforts to standardize or at least harmonize myeloma MRD testing are ongoing [28-31]. Next generation sequencing vs. flow cytometry for MRD evaluation Early molecular genetic approaches such as allele specific polymerase chain reaction have been proven successful in clinical trails, but are labor–intensive and suffer from relatively low applicability[27, 32]. These methods have now been largely subsumed by next generation sequencing. DNA-based approaches to myeloma MRD are covered elsewhere in the issue. IMWG did not express a preference for either testing modality and encouraged incorporation of both within future clinical trials to obtain further data on test performance[27]. Flow cytometry and NGS have unique advantages and disadvantages, these are summarized in table 1. Major advantages of NGS are higher theoretical analytical sensitivity and relative ease of standardization. In practice, sensitivity

gap between published validated flow cytometric and molecular genetic approaches is small (2*10-6 for flow and 10-6 for NGS) [28, 33-35]. The assessment sensitivity in practice may be further confounded by inability of DNA-based assays to distinguish between low quality hemodilute samples and those that are representative of marrow composition. Flow cytometry assays have the quality assessment built-in through analysis of cell composition of the sample. Unfortunately flow cytometry interpretation requires significant technical and analytical expertise making full standardization difficult. While EuroFlow assay utilizes automated analysis tool, expert review of the results may still be needed with the current implementation and challenging case analysis may carry an element of subjectivity[28]. On the other hand, flow cytometry is nearly universally applicable, while NGS has lower applicability [26, 36]. For instance Martinez-Lopez could only analyze 75% of patients by NGS due to inability to amplify high frequency diagnostic clones in presentation samples [26]. Similarly 98% of patient samples could be serially assessed by MFC, but only 77% could be studied by NGS in Korde et al. study of smoldering myeloma [37]. Efforts to improve amplification of clonal B cell receptor sequences in myeloma are ongoing. NGS assays with higher applicability are becoming available[27]. Nonetheless analysis of original diagnostic sample or labor-intensive flow cytometric cell sorting of an involved sample is expected to be required for NGS-based MRD for the foreseeable future. NGS may also be confounded by simultaneous presence of additional clonal B cell populations at diagnosis. Flow cytometry does not require a diagnostic sample prior to treatment, nor is confounded by presence of other neoplastic clonal B cell populations, which can be easily separated from abnormal myeloma clones. In our center this results in a significantly higher number of patient samples that can be analyzed for MRD by flow cytometry compared to NGS (internal unpublished data). The choice between the tests in the clinical practice will likely be driven by local availability, costs, and continuity of care considerations. Sampling consideration in myeloma MRD detection Multiple myeloma is largely a bone marrow based disease. MRD studies largely focus on bone marrow sampling. In addition to requiring sequential invasive procedures, this approach poses a few additional challenges. First, it will necessarily miss isolated extramedullary disease. Imaging studies such positron emission tomography and magnetic resonance imaging are complimentary to marrow-based evaluation [27]. Sequential imaging has now been incorporated as part of definition of MRD negative complete remission[27]. Second, great care must be taken in obtaining adequate samples. High quality minimally hemodiluted bone marrow aspirates are required for evaluation. While abnormal plasma cells may frequently be found in the blood of myeloma patients, there is at least 20-fold reduction in proportion of plasma cells in hemodiluted vs. non-hemodiluted bone marrow samples [38-41]. The differences in sampling can lead to major interinstitutional heterogeneity in plasma cell proportions, which can lead to differences in sensitivity despite identical analytical pipelines[42]. Third, bone marrow plasma cell disease may be patchy, suggesting that sampling at a single site may not be fully representative of disease burden. While this is of theoretical concern, bilateral bone marrow sampling showed no consistent advantage over unilateral sampling. Unilateral sampling is therefore sufficient for MRD evaluation[38] Finally it is clear that at least in situations when significant number of plasma cells are present for morphologic evaluation, they are often significantly underrepresented in flow cytometry samples as compared to bone marrow aspirate evaluations[43] [44] [45]. Moreover aspirate evaluations themselves often underestimate proportion of plasma cells as compared to core biopsies [46-48]. Some of the reported underrepresentation of plasma cell in the flow cytometry samples as compared to the aspirates is due to poor quality flow cytometry samples and may be corrected[44, 49]. However plasma cells are highly adherent to spicules and are frequently underrepresented in single cell suspension in flow samples [45]. It is therefore probable that even with high quality sampling, proportion of plasma cells will not always be fully reflective of the bone marrow aspirate cellularity. Moreover, if core biopsy is considered a gold standard of plasma cell evaluation, flow cytometry of even the high quality aspirate samples will frequently underestimate total disease burden. Therefore while flow cytometry-based quantitation of myeloma cells in the bone marrow is precise, it should not be interpreted as a fully accurate representation of disease burden. Despite the likely

underestimation of plasma cell proportion, flow cytometry can routinely detect residual abnormal clones in patients with no detectable morphologic disease because the analytical sensitivity of flow cytometry is at least 500-fold greater than that of morphologic assessment (5 in 100 vs. at least 1 in 100000) mitigating the effect of reduced plasma cell recovery. Of interest even when increased plasma cells are present by morphologic evaluation, plasma cell enumeration by flow cytometry has been proven to carry more prognostic information that morphology assessment and is better than morphologic assessment including immunohistochemstry at distinguishing polyclonal from neoplastic plasmacytosis at follow-up [43, 50]. In our experience the level of underrepresentation stays relatively stable for individual patients. Therefore, change in proportion of abnormal plasma cells in serial samples allows for accurate assessment of changes in neoplastic clone proportions. Considerations for bone marrow volume and cell numbers required for analysis Qualitative identification of minimal residual disease by flow cytometry requires a cluster of abnormal events. What constitutes a “cluster” is poorly defined and may vary from assay to assay and from laboratory to laboratory. In our experience 10 events constitute a reasonable target cutoff for a limit of detection (LOD) in a clinically validated standardized assay within a single laboratory. There may be higher minimal abnormal event requirement for assays run in multiple laboratories. The EuroFlow 2-tube 8-color assay was standardized between institutions and set LOD at 20 events for myeloma detection, while multi-institutional harmonization of a chronic lymphocytic leukemia assay required 50 events[28, 51]. The limits of detection must be validated for each individual test to assure that background noise due to non-specific changes such as sample degradation and minor variation in processing conditions as well as normal variation in phenotypes of non-neoplastic cells induced by therapy, inflammation and other changes in patient status do not interfere with identification of abnormal populations at the defined LOD threshold. In contrast to the LOD, which is empirical, Poisson counting statistics help estimate maximum obtainable precision of quantitation of abnormal events in the context of minimal residual disease detection. Coefficient of variation may be estimated as /N *100% where N is the number of events in the abnormal population (reviewed in [52]). Lower limit of quantitation (LLOQ) is defined as a minimum number of events at which precision remains acceptable for the purpose of the test. For instance at least 25 events would be required ( . Somewhat higher imprecision may be seen in multistep process such as flow cytomery testing. It is therefore critical to validate the LLOQ empirically under routine testing conditions. For MRD assays LLOQ with imprecision of 30% has been generally considered acceptable [53]. Guidelines for LOD and LLOQ validations for applicable to flow cytometry have been previously published [30, 53, 54] If a 20-cell cutoff is used for LLOQ and desired sensitivity is at least 10-5, greater than 2 million cells must be analyzed. This calculation is roughly in line with current recommendations from FDA-NCI round table (3 million), recently published consensus guidelines (3-5 million) and EuroFlow standardized approach (10 million)[28, 29, 31]. Acquisition of 10 million cells demonstrates effective sensitivity of 2*10-6[28]. Of note, due to the usual cell losses intrinsic to cell processing, the starting cell number should be at least in 2.5-fold excess of the cells required for final analysis. Traditional flow cytometry assays are generally performed in whole bone marrow using a fixed volume of sample between 50-200 microliters. However to reliably obtain such a high number of cells in nearly every sample, cells from greater volume of bone marrow must be analyzed. Figure 1 demonstrates the effect the number of cells can have on assay results. To reach the sensitivity of 10-5 -2*10-6 of bone marrow cells, first pull (most concentrated and least diluted sample) of 2-4 ml is required for adequate sampling[28, 55]. Samples must be collected in either EDTA or heparin and analysed as soon as possible, but no later than 48 hours to preclude false negative results due to sample degeneration or clotting[29]. In order to keep staining volumes fixed, bone marrow cells must first be concentrated prior to antibody staining. This is done by removing red cells from at least 2 and up to 4 ml of bone marrow by ammonium chloride lysis technique followed by collecting white cells by centrifugation[28, 34].

Antibody panel design considerations Flow cytometric identification of abnormal populations relies on recognizing antigenic differences between the neoplastic populations and their closest normal counterparts. This actually poses two related problems. First, plasma cells (both normal and abnormal) must be identified based on immunophenotypic parameters in a complex mixture of different cell types in a bone marrow sample. Second, a reliable myeloma MRD assay must be able to quantitatively distinguish neoplastic plasma cells from normal marrow resident plasma cells, even when normal plasma cells far outnumber the neoplastic clone. Even relatively infrequent normal plasma cell subsets should be excluded from abnormal cell quantitation. This places significant demands on number of markers required for sensitive and specific testing. Recognition of plasma cells requires at least three antigens. In the absence of CD38 targeting drugs, it is accomplished by combined gating using CD45, CD38, CD138 and light scatter parameters[30, 49, 56, 57]. Plasma cells usually express high density of CD38 and CD138 antigens with variable, but usually lower that mature lymphoid cells, expression of CD45. CD38 antigen density is frequently reduced in neoplastic plasma cells and CD138 is used as a “backup” antigen for sensitive plasma cell identification. On the other hand, CD138, while relatively specific is somewhat unstable and tends to be shed with sample aging. Separating normal from abnormal plasma cells poses a somewhat more difficult challenge. While one of the most common abnormal plasma cell immunophenotypes involves losses of CD19 and CD45 and expression of CD56, this phenotype is also seen in a subset of normal long-lived plasma cells and is therefore insufficient for the distinction [58-60]. Figure 2 demonstrates presence of the CD19+/CD56+/CD45 (dim) and other unusual immunophenotypes in the normal bone marrow. Using kappa to lambda light chain expression ratio in plasma cells in isolation is neither specific nor sensitive enough for disease follow-up and is a common source of interpretive errors[49]. Furthermore, expression of individual antigens known to be highly specific for abnormal plasma cells such as CD117 or CD20 occur only in minority of myeloma cases[59, 61]. Numerous additional informative markers have been reported, but no single target or a relatively small group evaluable antigenic targets could consistently separate normal from abnormal plasma cells in nearly all cases with high specificity and sensitivity [29, 60]. In an elegant experiment, investigators from EuroFlow consortium explored relative information value of many of the published antigenic targets. Using principal component analysis they confirmed that a combination of at least eight antigens (CD19, CD27, CD38, CD45, CD56, CD81, CD117 and CD138) could nearly always identify immunophenotypically aberrant plasma cells in the bone marrows of myeloma patients[28]. Furthermore, they confirmed that removing any of the markers from the panel resulted in significant test performance degradation when applied to a variety of abnormal cases. Testing the expression of these antigens within well-designed multicolor panels is now considered a requirement for clinically relevant myeloma MRD testing. Cytoplasmic kappa and lambda light chains are sometimes needed for confirmation of the clonal nature of the suspected neoplastic populations[29]. While this opinion is somewhat controversial, in our experience light chain staining enhances speed and accuracy of analysis when low-level MRD is present. It may also serve as additional lineage marker in difficult samples assisting in separating plasma cells from contaminating populations in a proper context. Moreover, because it is difficult to predict which samples will require confirmation with light chains prior to analysis, such staining may be done upfront for reasons of practicality and to prevent additional sample aging prior to second analysis. We have previously demonstrated that combination of surface and cytoplasmic staining in a single step/single tube analysis does not significantly degrade test performance compared to separate surface and cytoplasmic staining as implemented by EuroFlow [33, 34]. Additional highly informative targets such as CD200, CD307 and CD99 have been reported, but are yet to be incorporated into published assays with sensitivity of 10-5 and higher [62-66]. Result reporting Consensus reporting guidelines for myeloma MRD have been published[30]. In line with Bethesda international consensus recommendations, they call for descriptive reporting of immunophenotype of the abnormal population with each antigen staining intensity compared to reference normal

plasma cell obtained under the same condition and reported as normal, increased, decreased (absent) [67]. This report may assist in identifying the population in the future. Abnormal plasma cell population should be reported as a percent of all bone marrow nucleated cells (excluding debris) and as a proportion of total plasma cells. Reporting proportion of plasma cells among plasma cells may inform interpretation of morphologic plasma cell counts, if flow cytometry total plasma cell count proves to be an underestimate. At diagnosis, proportion of abnormal plasma cells among the total plasma cells has been shown to be of prognostic significance in precursor lesions (monoclonal gammopathy of unknown significance, smoldering myeloma) and may identify a subset of myeloma with better clinical outcomes[68, 69]. The report should also include validated LOD and LLOQ as well as number of plasma cells acquired Quality standards and annotation of problematic samples To assign clinical significance to an MRD result, stringent quality procedures and metrics must be implemented. Overall quality guidelines applicable to validation, performance and reporting of myeloma MRD assays are reviewed elsewhere [52, 53, 70-72]. Stringent instrument quality control including instrument standardization if the assay is run on more than one instrument and daily verification of instrument performance are required. In MRD assays, even a very small amount of sample-to-sample carryover has a potential to significantly affect the results. Procedures and practices to minimize and hopefully eliminate the possibility of carryover must be in place[52]. Implementation of the assays must undergo stringent validation of accuracy, reproducibility and claimed sensitivity[53]. If an assay is to be implemented in a multi-institutional setting, a fraction of samples and analysis files should be analyzed in parallel by all participating institutions to assure assay and interpretation standardization[28, 73, 74]. Once an MRD assay is validated and standardized, quality of each sample must be assessed to assure the validity of obtained results. This is particularly critical if a negative result is obtained. If the result is negative for MRD, but not enough cells were analyzed, the decreased sensitivity of detection should be properly annotated. While theoretically a new LOD and LLOQ could be calculated based on the number of cells actually analyzed, in our experience samples with low cell acquisition generally suffer from additional quality problems such as hemodilution that further reduce sensitivity. Hemodilution will affect the result even if sufficient number of cells is obtained. Assessment of hemodilution can be performed by analyzing additional cell populations with high degree of partition between marrow and blood compartment. These include nucleated red blood cells (cells negative for all antigens commonly tested in myeloma MRD tube), mast cells (exceptionally bright CD117 with somewhat increased side scatter), early maturing B cells (hematogones)(CD19 positive cells with bright CD81 and CD38 without cytoplasmic light chain expression with relatively dim CD45), and early myeloid and erythroid precursors (CD117 positive, CD27 negative cells). Evaluation of these additional populations was automated the EuroFlow assay and can also be easily performed manually outside of the platform. Example of such assessment is provided in Figure 3. Other issues such as sample aging, clotting and degeneration due to improper shipping and storage conditions (high or low temperature) may also significantly affect the results. Bone marrow aspirates are considered irreplaceable and should be analyzed whenever practical even if there are manageable problems. However, deviations from expected quality of the sample should be carefully annotated to avoid misinterpretation of a suboptimal result in clinical practice. Examples of high sensitivity flow cytometry assay implementations suitable for 6-10 color platforms The most well-known high sensitivity standardized flow MRD assay is the EuroFlow assay implemented by EuroFlow consortium[28]. This assay is best suited for 8-color instruments and has been validated on several instrument types[75]. The EuroFlow Consortium standardized EuroFlow assay uses 10 antigen testing split between two 8-color tubes: a surface only tube and a surface/cytoplasmic tube [28]. The assay has reported sensitivity of 2*10-6 bone marrow cells, requires acquisition of 10 million cells and shows very good correlation with NGS-based assessment. The most innovative aspect of the assay includes implementation of computer assisted population identification, which reduces interpretive variation, provides built in quality control, and paves the

way for enhanced standardization. The method supplanted prior 8-color approach used by PETHEMA/GEM group in clinical trials [76]. Although this method is clearly effective, the two-tube approach that shares 6 antibodies between the tubes has a drawback of increased costs resulting from multiple antibody duplication, increase in labor, and increased time of instrument acquisition and analysis. Full implementation also requires proprietary software. This may pose barriers for wide clinical adoption of the test outside dedicated centers and for applicability to patients treated outside major clinical trials. For example, in the United States, reimbursement is not provided for the increased cost and effort of implementing this resource-intensive method. To reduce additional costs and labor burden for the laboratories, Memorial Sloan Kettering group developed a streamlined approach suitable for 10-color instruments by combining surface and cytoplasmic staining in a single 10-color tube[34, 55]. Comparison of this approach with the EuroFlow 8-color 2-tube panel was performed in a series of 41 routinely obtained myeloma followup clinical samples covering the entire analytical range. The comparison showed similar test and interpretive performance while reducing costs and time of testing. Notably, quantitation of abnormal plasma cells as proportion of total marrow cells or as a proportion of plasma cells was not affected. The combined surface/cytoplasmic staining did result in a minor (approximately 15%) loss of total cells requiring a slightly higher initial cell input. This assay, however, has not been standardized across institutions and manual nature of analysis may make this standardization somewhat more challenging, though not unprecedented [51, 73, 74] A six color 8-antigen assay that excludes light chain staining has been proven to work well in the context of large clinical trials by United Kingdom’s Medical Research Council (MRC) Investigators. However, it has a target sensitivity of 0.01% and its performance below that limit has not yet been reported[18, 21]. The reported antibody combinations are provided in Table 2. Evaluation of MRD in the presence of surface antigen targeting agents Anti-CD38 directed monoclonal antibody (daratumumab) was recently approved for the treatment of relapsed multiple myeloma [77-79]. Presence of the drug saturates CD38 binding sites for several months following administration and results in marked degradation of performance of commercially available labeled monoclonal CD38 antibodies as gating reagents for identification of myeloma [28, 80]. This is illustrated in Figure 4. Lack of reliable CD38-based plasma cell identification results in significant overall test performance degradation. In an effort to overcome the competitive inhibition, EuroFlow group developed a multi-epitope mixture of anti-CD38 antibodies that bind to sites that are not occupied by the drug [28]. However, in our experience with many patients treated with daratumumab the commercially available multi-epitope reagent results in suboptimal separation due to relatively low staining intensity and partial interference by the drug (Roshal M, unpublished data). The observation has now been independently confirmed in a small study [80]. Further complicating future gating strategies CD138 was also successfully targeted, although no anti-CD138 drugs have been approved [81]. There is an urgent need to develop and validate alternative gating strategies and reagents. Substitution of current anti-CD38 reagents with either CD38 targeting nanobody JK36 or monoclonal HuMax-003, both of which have been reported to bind independently of daratumumab, offer one potential avenue; however, neither antibody is commercially available [80, 82]. Furthermore, daratumumab results in downregulation of CD38 on the surface of plasma cells, making long-term prospects of CD38 utility as a gating reagent in the presence of the drug questionable [82]. Several additional promising markers such as CD319, CD229 and CD54 were systematically evaluated. Of these only CD229 allowed consistent resolution between plasma cells and background cellularity [83]. Such gating would still require at least CD138 or another relatively specific marker. A monoclonal VS38c antibody directed against endoplasmic reticulum protein p63 has shown some promise and is under evaluation [84]. It is expected that CD38-independent robust gating strategies will soon emerge.

Future directions High sensitivity detection of multiple myeloma by flow cytometry serves as a paradigm for the future of MRD test development and utilization. There is robust clinical data in thousands of patients across multiple international clinical trials supporting prognostic utility of the test in this disease. There is keen interest from clinical and pharmaceutical groups in exploring MRD-directed therapeutic interventions. There is a consensus by experts on the optimal minimal sensitivity and test parameters required for successful testing. At least one standardized and partially automated version of the test is available for broad implementation. The practical sensitivity, speed, and broad applicability of flow cytometry testing makes the choice between NGS and flow cytometry a subject of intense research interest with no clear evidence of superiority of one platform over the other available at present. Of interest, EuroFlow investigators have now demonstrated a molecular geneticlike sensitivity of a new standardized test for B lymphoblastic leukemia/lymphoma suggesting that NGS-like sensitivity of flow cytometry in myeloma is not a singular exception [85]. Challenges in optimal sample procurement, defining most informative testing time points, test standardization, laboratory and practitioner education, as well as evolving surface antibody targeting drug landscape remain and will need to be addressed.

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Figure 1: Effect of cell number acquisition on the result of MRD test. Myeloma MRD test was run in parallel using whole marrow 8-color test (A) and bulk-lyse 10-color test (B). Kappa expressing plasma cells are shown in blue, lambda in red. The whole marrow staining procedure resulted in acquisition of 883 thousand cells and <10 abnormal plasma cell events (green, emphasized) corresponding to indeterminate/negative results. Bulk lyse procedure resulted in acquisition of 10 million cells with 84 abnormal plasma cell events leading to a clear positive MRD result. Figure 2: Diversity of normal plasma cell immunophenotypes: Plasma cells from a patient with no history of plasma cell neoplasm are shown in an MSKCC 10-color assay layout. Kappa light chain expressing plasma cells are shown in blue and lambda in red. Note that CD45-/CD19- and CD19-/CD45-/CD56+ (arrows) plasma cells are present and are polytipic. In addition small quantity of CD19+/CD56+ and CD81-/CD19+/- cells are also seen. Figure 3: Analysis of normal resident marrow cells in 10-color MSKCC layout: Early CD117 positive myeloid and erythroid precursors and show in yellow, CD117(bright) mast cells in purple and CD19+ early B cells hematogones lacking either kappa (blue) or lambda (red) cytoplasmic light chains are shown in light blue. Nucleated red blood cells can also be assessed (not shown) Figure 4: Potential alternatives to CD38-based gating in the presence of daratumumab. Sample from patient treated with daratumumab is shown. Plasma cells (blue) no longer separate from background cells based on CD38 alone (A). CD319 expression is seen on the plasma cells, but shows significant overlap with granulocytes (B). CD229 also shows overlap, but only with lymphoid cells which can be gated out based on scatter parameters and CD45 (not shown) (C). Cytoplasmic vs38c (cyPC) similarly shows overlap with myelomonocytic cells, but may be useful in combination (D).

Reported maximal sensitivity Data documenting clinical utility and test perfomance

Ease of standardization

Stabilty requirements Requirement for original diagnostic sample Proportion of patients who can be evaluated Interference by coexsiting clonal B cell prolifertions Sample quality assessment Costs Time to result

Table 1

NGS

MFC

1 cell/million

2 cells/million

Emerging (few large scale trials)

Easy Less rigid, DNA can be extracted for batch testing at later time

Extensive Difficult, but has been demonstared by Euroflow group Rigid (sample analysis within 24 hours preferred)

Required

Not required

75-95%

>98%

Yes

No

Limited Higher (but decreasing) Days to weeks

Built-in Lower Hours

Comparison of NGS and MFC methodologies

FITC MRC 1 CD27 MRC 2 CD81 EuroFl CD38 ow 1 (me) EuroFl ow 2 MSKC C

CD38 (me) cykapp a

PE CD56 CD11 7 CD56

CD56 cylamb da

Px5/ 5.5

PC7

CD19

CD38

CD19

CD38

CD45

CD45 CD11 7

CD19

APC CD13 8 CD13 8 CD11 7 cykapp a

CD19

CD13 8

CD19

APCR700

PB/B V421

BV BV 510 605

CD81

CD13 8

CD 27

cy-lambda

CD13 8

CD 27

CD81

CD 38

CD45 CD45

CD56

me=multiepito pe cy=cytoplasmi c Table 2:

APCH7/Cy7/C 750

Examples of 6, 8 and 10-color tubes used in myeloma MRD analysis

CD45

CD 27