Interlaboratory Analytical Validation of a Next-Generation Sequencing Strategy for Clonotypic Assessment and Minimal Residual Disease Monitoring in Multiple Myeloma

Bone marrow samples from 101 patients with MM were collected at diagnosis and early follow-up time points (after the end of induction or 3 months after transplant), irrespective of their response, as part of the GEM2012MENOS65 clinical trial (NCT01916252).¹¹  Sixty-nine had been included in a previous publication from our group, exploring the clinical significance of MRD evaluation by next-generation techniques.³²  Patient selection was based on the following criteria: (1) immunophenotypic studies performed with conventional 8-color MFC (diagnosis) or EuroFlow's NGF (follow-up), and (2) DNA availability for paired diagnostic and MRD studies. Responses to treatment were evaluated according to the IMWG response criteria.¹⁹ 

Samples were distributed and analyzed according to Figure 1. Thus, the Salamanca laboratory carried out the clonotypic characterization of diagnostic samples targeting IGH (FR1 and FR2 regions) using Sanger sequencing, while the San Diego laboratory used LymphoTrack (Invivoscribe Technologies, San Diego, California) targeting IGH (FR1, FR2, and FR3) and IGK rearrangements. For MRD studies both centers used NGS (FR1, or FR2 if necessary), allowing the measurement of interlaboratory precision. In addition, baseline and follow-up MFC studies had been previously carried out in Salamanca. However, bone marrow samples for MFC and NGS were extracted in different pulls.

The Ethical Committee of the University Hospital of Salamanca (Salamanca, Spain) gave approval for this study, in accordance with the Spanish law and the Declaration of Helsinki principles. Written informed consent was obtained from every patient before their inclusion.

Genomic DNA (gDNA) was isolated from bone marrow aspirates with the automated Maxwell DNA Purification Kit (Promega, Madison, Wisconsin). DNA quality and quantity were first assessed in the Salamanca laboratory using NanoDrop2000 (ThermoFisher, Waltham, Massachusetts) and then distributed. Samples were once more quantified by each laboratory, in this case using Qubit 2.0 and the dsDNA BR assay (ThermoFisher).

Follow-up samples with insufficient DNA concentration for MRD purposes (those with less than 100 ng/μL) were ethanol precipitated.³³  Sodium acetate 1/10 as well as 2-2, 5x volume ethanol (100% and stored at −20°C) of the total sample volume were added to the samples, then incubated overnight at −20°C. Afterwards, samples were centrifuged at 17 900g for 10 minutes at 4°C. The nucleic acid pellet was later washed with 500 μL ethanol (70%) and centrifuged again at 17 900g for 5 minutes at 4°C. Finally, the pellet was dried and rehydrated in ≈12 μL of water and quantified by using Qubit dsDNA BR assay.

Limits of detection (LOD) and quantification (LOQ) of the LymphoTrack assay were assessed with the following experiment: diagnostic bone marrow biopsies from 20 patients with myeloma were sorted to select CD138⁺ cells. Genomic DNA was obtained from all, and the LymphoTrack IGH panel (Invivoscribe Technologies) was used to determine the tumor-related clonotypic rearrangement. Afterwards, gDNA from all patients was pooled together at the same ratio (100 cells/patient/μL) based on the quantification made with Qubit 2.0.

Then, 5 serial dilutions at prespecified cell proportions (100, 30.3, 9.2, 2.8, and 0.85 cells/patient/μL) were made by using Tris-ethylenediaminetetraacetic acid buffer. Each was further diluted into buffy coat obtained from 7 healthy donors and tested with the LymphoTrack panel at 3 different total DNA inputs in 50 μL (DNA input: 500 ng) or 100 μL (DNA input: 1 μg or 2 μg) per reaction, so that 15 artificial MRD samples, each comprising 20 clonal rearrangements, were tested in the range of 1.3·10⁻³ to 2.7·10⁻⁶ (Supplemental Table 1, see Supplemental Digital Content 1, containing 7 tables and 1 figure at https://meridian.allenpress.com/aplm in the July 2022 table of contents). A constant proportion of spikein, consisting of a known quantity of DNA (equivalent to 100 cells) from a Waldenström macroglobulinemia cell line (MWCL1, kindly provided by Stephen Ansell, MD, PhD, Mayo Clinic, Rochester, Minnesota),³⁴  was added to each reaction. Sequencing was performed in the MiSeq platform (Illumina, San Diego, California) using v3 reagent kits and 2×251 cycles of sequencing.

For the amplification of complete VDJH rearrangements at diagnosis, the Salamanca laboratory used the BIOMED-2 (now Euroclonality) FR1 or FR2 primers in multiplexed PCR reactions.³⁵  All reactions were carried out in a 25-μL mixture containing 20 to 100 ng of baseline DNA and 10 μmol of forward and reverse primers. Monoclonal assessment of amplified products was carried out by GeneScanning using 1 μL of PCR reaction. PCR products were sequenced in an automated ABI3500 XL DNA sequencer using Big-Dye terminators v3.1 (Applied Biosystems, Foster City, California).

The LymphoTrack IGH panel was used for the analysis of diagnostic (San Diego laboratory) and MRD samples (both laboratories), aiming to detect tumor clonotypic rearrangements. Briefly, this commercial strategy uses primers targeting the immunoglobulin framework regions to amplify V(D)J rearrangements. In 1-step PCR (Figure 2, A through E) amplicons are generated and 1-side indexed, allowing the simultaneous sequencing of up to 24 samples in a single run.

All baseline samples were sequenced in San Diego, using 50 ng of DNA. Whenever it was possible, at least 650 ng of follow-up DNA was used in 1 or 2 PCR replicates to reach a sensitivity level of 10⁻⁵ (assuming 6.5 pg of DNA per cell, cell input for NGS would be at least 100 000). In addition, 1 μL of MWCL1, corresponding to 100 to 1000 cells, was added in each follow-up PCR reaction as a spike-in, to allow the absolute quantification of tumor plasma cells.

After a purification step using Agentcourt AMPure XP microbeads (Beckman Coulter Inc, Brea, California) and 70% ethanol, and once the purity and quantity of the amplicons were assessed with the TapeStation 4200 (Agilent, Santa Clara, California) and the KAPA Library Quantification Kit (KAPA Biosystems, Boston, Massachusetts) or Qubit 2.0, libraries of between 12 and 20 pM were prepared. These libraries were later sequenced in a MiSeq instrument using v3 reagent kits and 2×251 cycles of sequencing, aiming for 1 million reads per sample. To prevent cross-contamination during the sequencing step, samples from the same patient were always sequenced in different runs, using different indexes when possible. In addition, libraries were sequenced within 2 runs of each other, performing a sodium-hypochlorite post-run wash on the instrument to reduce carry-over contamination.

Sanger sequences were analyzed by using the IMGT/V-Quest database³⁶  taking into account the option for insertions/deletions. IGHV, IGHD, and IGHJ genes, as well as complementarity-determining region 3 (CDR3), were identified. Mutational status was assessed by using the closest germline gene as a reference.

FastQ files generated during NGS were processed with the LymphoTrack Software-MiSeq (Invivoscribe Technologies) to retrieve sequences from virtually every clonal B cell in the samples. Tumor clones at diagnosis were identified following 3 criteria: (1) 20 000 or more total reads; (2) at least 1 but not more than 2 merged top reads with 2.5% or greater of total reads; and (3) top first or second merged reads at least two times more abundant than the third most frequent read to be considered clonotypic. An NGS report, listing the top 5 rearrangements and sequencing parameters for all patients and primer sets, is included as Supplemental Digital Content 2.

Minimal residual disease was calculated with the LymphoTrack MRD Data Analysis tool 1.2.0 (Invivoscribe Technologies) considering the number of spike-in cells, the number of total cells that were used for each reaction, as well as tumor and spike-in read counts (Figure 2, E). Sequencing results were considered invalid when fewer than 20 000 total reads were retrieved. A sample was considered positive for MRD assessment when at least 2 identical tumor clonotypic reads were detected (allowing for up to 2-bp mismatches).

Samples were analyzed within 48 hours after the extraction, using a conventional 8-color multiparametric flow cytometry panel (CD38/CD138/CD45/CD19/CD56/CD117/CD27/CD81) or the 2-tube, 8-color EuroFlow NGF panel (tube 1: CD38/CD138/CD27/CD45/CD19/CD56/CD81/CD117; tube 2: CD38/CD138/CD45/CD19/CD56/CyIgK/CyIgλ) for diagnosis and follow-up time points, respectively, following EuroFlow's guidelines for each assay.^26,29  Commercial antibody panels for diagnostic and MRD evaluation were used (Cytognos S.L., Salamanca, Spain). At least 500 000 (diagnosis) or 10 000 000 (follow-up) bone marrow cells were acquired in a BD FACSCanto II (BD Biosciences, Franklin Lakes, New Jersey). Data analysis was performed by using the calculation function of the INFINICYT v2.0 software (Cytognos S.L.).

Patients' characteristics were tested with SPSS 20.0 software (IBM, Armonk, New York), using Fisher exact test for discrete variables and the Mann-Whitney test for continuous variables. Bland-Altman,³⁷  Cohen κ, and linear regression tests were used to assess potential agreement, concordance, and bias between methods.

LOD and LOQ were calculated as described by Ching et al.³⁸  The LOD, or sensitivity, was calculated by using a probit model and defined as the lowest number of input tumor cells at which the probit curve detected MRD with a type II error rate of 5%. Here, detection was considered positive when at least 2 identical clonal rearrangements were present, and negative otherwise. The LOQ was calculated as the lowest number of input tumor cells quantitated within a maximum total error of 70%. Total error was calculated as the root-mean-square error (RMSE) divided by the number of input tumor cells. RMSE was calculated as the square root of the squared bias plus the variance. Prespecified accepted values for the LOD and the lowest LOQ were both set at 2 ± 1 malignant cells per total number of input cells. All reported P values were obtained by a 2-sided exact method, at the conventional 5% significance level (P < .05). To compare the overall performance of MRD methods, NGS results from the 2 laboratories were merged, considering “NGS positive” those cases for which MRD was detected in at least 1 laboratory, or “NGS negative” otherwise.

One hundred one patients met the inclusion criteria. Follow-up samples were collected 3 months after autotransplant from 81 patients and following the end of induction from the remaining 20 patients. Median follow-up since MRD evaluation was 36.9 months (interquartile range [IQR], 27.27–44.32).

Clinical variables from our cohort are summarized in Supplemental Table 2. Male to female ratio was 62:38. Median age at diagnosis was 59 years. Patients with high-risk cytogenetics represented 21.7% (22 of 101) of the present series. The R-ISS³⁹  stages I, II, and III represented 21.8% (22 of 101), 62.1% (63 of 101), and 16.1% (16 of 101) of patients, respectively. At the corresponding MRD evaluation time point, the proportion of patients achieving sCR, CR, very good partial response, partial response, and stable/progressive disease was 32% (32 of 101), 20% (20 of 101), 30% (30 of 101), 14% (14 of 101), and 4% (4 of 101), respectively.

Based on the data from the 5 serial dilutions and the 3 DNA input levels (2 μg, 1 μg, and 500 ng), the probit model estimated the LOD to be 1.30 myeloma cells (95% CI, 0.89–1.72) for any used total gDNA input (Supplemental Table 3). Herein, the lowest amount of MRD that could be effectively detected in 95% of measurements using 500 ng, 1 μg, and 2 μg of gDNA would be 1.7·10⁻⁵, 8.2·10⁻⁶, and 4.2·10⁻⁶, respectively.

As expected, the number of input tumor cells greatly influenced precision in the quantification ability of the assay. The LOQ was set at 8 tumor cells, with total estimated errors ranging from 32.3% when using dilutions with 100 tumor cells/sample/μL to 1770% when using dilutions containing 0.85 tumor cells/sample/μL (Supplemental Table 3).

All samples were successfully characterized by using flow cytometry. Phenotypic features and clonal plasma cell infiltration in the bone marrow met the IMWG criteria to confirm symptomatic MM diagnosis in the entire population. Median plasma cell infiltration in the bone marrow was 6.85% (IQR, 2.69%–17.16%); the median proportion of clonal, pathologic plasma cells was 99.85% (IQR, 99.33%–100%).

All samples but 1 (100 of 101, 99%) were monoclonal after PCR amplification, GeneScanning, and Sanger sequencing. Ninety-nine of 101 (98%) were successfully characterized by using FR1 primers. One could not be identified through FR1 Sanger sequencing but was later detected by FR2. Additionally, in another sample, the clonal rearrangement was not identified by FR1 or FR2 sequencing, most probably owing to the high polyclonal background observed by flow.

Regarding NGS by LymphoTrack, 83.2% (84 of 101), 79.2% (80 of 101), 62.4% (63 of 101), and 86.1% (87 of 101) of cases were considered monoclonal for VDJH-FR1, VDJH-FR2, VDJH-FR3, and VJK sequencing, respectively (see also the supplemental files). Overall, when combining the 3 IGH regions and IGK, LymphoTrack showed 100% of clonality detection ability (meaning at least 1 valid result, as shown in Supplemental Table 4). Of note, LymphoTrack allowed the detection of 7 of 101 biallelic rearrangements (6.9% of the total), all undetected by Sanger. One patient with an unproductive rearrangement detected by Sanger was found to have a double-unproductive rearrangement when NGS by LymphoTrack was performed; this patient had nonsecretory myeloma. VDJH-FR3 sequencing using LymphoTrack showed poor performance results, with only 47 of 63 cases (74.6%) displaying the same CDR3 region that was detected by Sanger sequencing; in addition, IGHV genes of these sequences were misidentified in 29 of 47 cases (61.7%).

Therefore, we decided to consider only results obtained with FR1 and/or FR2: 95 of 101 cases (94%) fulfilled criteria to be considered clonal by LymphoTrack. Of these, 78 of 95 cases (82.1%) shared exactly the same CDR3 region when compared to Sanger sequencing. Fifteen of 95 (15.8%) had minor discordances in the nucleotide sequence, affecting the codification of 1 or 2 CDR3 amino acids; the last 2 of 95 cases (2.1%) were completely different in terms of clonal rearrangement and CDR3 amino acid composition (Supplemental Table 5). Thus, the LymphoTrack approach for FR1 plus FR2 was applicable in 95 of 101 cases (94%); of these, 93 of 95 (97.9%) were concordant with Sanger sequencing.

After considering the results obtained with baseline samples, MRD studies were performed only in the 95 cases for which LymphoTrack was previously applicable. NGS results were compared in 82 of 95 cases (86.3%) for which parallel sequencing in Salamanca and San Diego was possible. Additionally, 11 cases were sequenced only in 1 laboratory, 3 in Salamanca (2 MRD-positive and 1 MRD-negative), and 8 in San Diego (4 MRD-positive and 4 MRD-negative). Another 2 cases were sequenced in only 1 laboratory (one in Salamanca, another in San Diego) but they could not be further analyzed owing to low total read counts (<20,000 total reads).

Ethanol precipitation often led to the loss of DNA instead of increase in its concentration, thus affecting the applicability of LymphoTrack for MRD analyses. Because of this, the minimum number of cell equivalents to reach a virtual sensitivity of 10⁻⁵ could only be used in 87.9% of cases (89 of 101) by Salamanca and 60.4% of cases (61 of 101) by San Diego (Table). NGF was successfully used for the evaluation of our entire series, with a significantly higher number of cells required to perform the studies at the preestablished sensitivity threshold.

As shown in Figure 3, we found a good correlation for NGS between centers (r = 0.932; R² = 0.869, P < .001) with only 12 of 82 discordances (reproducibility: 85.4%) that could be due to several circumstances (Supplemental Table 6): most of the positive samples detected only in the Salamanca laboratory would be explained by a higher number of input cells than that used by the San Diego laboratory, while discordances in the other cases may be related to sample aliquoting, sample concentration, or suboptimal sequencing. High correlation was maintained between NGF and NGS performed in each center (NGS–San Diego versus NGF: R² = 0.856, P < .001; NGS–Salamanca versus NGF: R² = 0.746, P < .001) (Supplemental Figure 1, A and B). When NGS results from the 2 laboratories were merged, only 13 of 93 discordant cases (14%) between techniques were detected (4 NGF⁺/NGS⁻; 9 NGF⁻/NGS⁺). A total of 19 patients had relapse in our series, all preceded by a positive MRD result (mean, 11.6 months earlier) by NGS and NGF (Supplemental Table 7). When plotting progression-free survival curves considering only patients for whom a minimum sensitivity of 10⁻⁵ was achieved (Figure 4, A through C), no significant differences were observed between methods or laboratories. Thus, progression-free survival rates at 36 months for MRD-positive versus MRD-undetected were 62% versus 94% for NGS in Salamanca (P = .003), 59% versus 96% for NGS in San Diego (P = .002), and 59% versus 98% for NGF (P < .001), respectively.

Finally, Bland-Altman plots (Figure 5, A and B) showed a considerably high level of agreement between NGS and NGF. Overall, the mean difference in the quantification of MRD levels detected by NGF and NGS was lower than 2-fold (1.15-fold and 1.86-fold when comparing NGF with NGS in Salamanca and San Diego laboratories, respectively). This bias was significant only when comparing NGF versus NGS in San Diego (bias: −0.27; 95% CI, −0.55 to −6.2E-05; P = .049), showing a trend toward a slight overestimation made by NGS in this case, while it was not significant when comparing NGF versus NGS in Salamanca (bias: 0.06; 95% CI, −1.5E-05 to 0.12; P = .07). Differences between methods were homogenously distributed across the range of MRD levels, with more than 70% of cases showing differences within the maximum range of ±1 log.

Minimal residual disease represents one of the most relevant prognostic factors for patients with MM, showing a significant advantage over previous, well-established prognostic factors such as cytogenetics and conventional responses.^15–17  From a revision of the results reported on clinical trials implementing this method, the IMWG encouraged the use of 2 different tools to detect MRD: EuroFlow's NGF and Adaptive's ClonoSEQ (NGS) approaches.¹⁹  While equivalent validated methods are also accepted, there is scarce information about NGS alternatives; in fact, ClonoSEQ is the only sequencing-based method validated by the FDA to report MRD in myeloma to date.³¹  To investigate new sequencing tools, we had previously evaluated the LymphoTrack assay.³²  Although our results showed that this strategy could be considered as a feasible alternative to NGF, given that the prognostic value was not altered, a more detailed characterization of its performance in both baseline and MRD settings was still needed. Here, we have evaluated LymphoTrack as a suitable and reproducible strategy for diagnostic and follow-up characterization of 101 patients with myeloma, while assessing its technical and analytical characteristics.

The LymphoTrack assay showed a high sensitivity, with a LOD of 1.3 malignant cells per reaction tube. However, the LOQ was outside the prespecified accepted values, allowing to precisely quantitate up to 8 tumor cells. Ideally, LOD and LOQ should be close to provide accurate and unbiased detection of an analyte. This level of discordance between parameters could be due to several reasons: (1) the statistical models were underpowered owing to a limited population size; (2) other factors contributing to total error (such as reagent lots, sequencer, operators, etc) were not taken into account; and (3) using a single cell line as a spike-in for normalization is a main source of error. From our perspective, the last point may be the most important factor, since amplification is not always proportionally adjusted to the initial concentration of DNA templates.⁴⁰  Further experiments are needed to verify the LOD and LOQ of the LymphoTrack assay, and a robust quantitation method should be developed to make this tool useful for clinical purposes.

When LymphoTrack was evaluated for clonality assessment and compared with conventional Sanger sequencing in baseline samples, the applicability of both techniques was similar, taking into account FR1 and FR2 sequencing: 95 of 101 (94%) for NGS and 100 of 101 (99%) for Sanger sequencing. Moreover, combining FR1 and IGK could be even more effective, as in our study 98 of 101 cases (97%) were monoclonal. Hence, our results using NGS would be in line with previous Sanger sequencing reports for MM, where up to 10% of cases can be missed owing to somatic hypermutation hampering primer annealing or to a high polyclonal background.³⁵  In fact, the use of longer primers for NGS would explain why some additional cases are lost. NGS primers are much longer than conventional Sanger sequencing primers (90 bp versus 25 bp) and this could be one reason to explain a poorer performance. A previous study that used LymphoTrack⁴¹  showed even lower rates of clonality detection using only FR1 primers in plasma cell neoplasms (∼70%), but these rates were improved after combining 2 or 3 targets for clonality detection, implying that this strategy should be followed to bypass the effect of hypermutation. Additionally, in this study the authors observed that applicability rates of FR1 primers greatly differ between B-cell lymphoproliferative disorders, ranging from ∼60% in follicular lymphoma to more than 95% in mantle cell lymphoma, implying that other tumor characteristics play a role (eg, DNA integrity, tissue of origin, somatic hypermutation) and somehow have a greater impact on NGS.

Remarkably, the use of a lower cutoff to establish clonality by NGS (ie, 2.5% of total reads) allowed us to identify 5 tumor rearrangements that were concordant with Sanger and would not have met the threshold for clonal rearrangement if the 5% cutoff, instead of the 2.5% cutoff, had been used. In addition, 3 had detectable residual disease when baseline rearrangements detected by NGS were tracked in follow-up samples, which may suggest this threshold is optimal to confidently determine whether a rearrangement is truly tumor specific.

Nonetheless, NGS and Sanger sequencing results can be different in a small subset of cases: in our series, 17 of 95 cases (17.9%) had distinct VDJH usage and/or CDR3 sequences. When follow-up samples were evaluated, all the MRD-positive cases detected by NGS showed full clonotypic concordance with baseline NGS results, not with Sanger. This is crucial since it would indicate that Sanger sequencing is more error-prone and using NGS at baseline would be preferable when accurate detection of V(D)J rearrangements or somatic hypermutation is needed (eg, CDR3 identification as biomarker for MRD, or prognostic impact of somatic hypermutation in chronic lymphocytic leukemia). This is also a clear demonstration that VDJH rearrangements and CDR3 regions are stable over time⁴²  and are not subjected to further somatic hypermutation cycles once the myeloma clone leaves the germinal center. Compared to standard Sanger sequencing, NGS could also improve the analysis of samples with low plasma cell infiltration levels, or with a high polyclonal background, which are common findings in the bone marrow of patients with myeloma at diagnosis owing to the patchy nature of the disease and potential hemodilution.¹  In our series, 44 of 101 (43.6%) diagnostic samples had abnormal plasma cell infiltration rates below 5% of total cells by flow, and significant amplification of polyclonal B cells was observed in 31 of 101 cases (30.9%) by Sanger.

Despite being limited by the available gDNA amount in specific samples, NGS and NGF showed high concordance rates in terms of MRD detection, reproducibility of results, and quantification ability in a head-to-head comparison, even when samples were obtained from independent bone marrow pulls. The reported detection sensitivity at the level of 10⁻⁵ in this article may not reflect what can potentially be achievable in other laboratories or clinical settings, as it was severely biased by the performance of ethanol precipitation or the maximum amount of available gDNA, which is sometimes low in samples extracted from the hypocellular marrow of posttreatment patients. Sequencing samples in duplicate or triplicate, increasing the total number of DNA templates, could further improve the limits of detection and the accuracy of MRD evaluation by NGS, which is a limitation of this study. It must be noted that, as several reports have stated before,^17,19  MRD evaluation in the clinical setting should be performed whenever patients achieve CR, and it gains significance with lower sensitivity, that is, when achieving the LOD of 0.0001% or 10⁻⁶. While in clinical trials this helps in distinguishing truly negative, long-term surviving cases, this was not the scope of our study. Although we detected 13 discordant cases between NGS and NGF that could be explained by less restrictive experimental conditions on NGS, none of those patients have had relapse to date (median follow-up since MRD evaluation, 36.9 months; IQR, 27.27–44.32 months). By contrast, the 19 patients with relapse in our cohort (5 in CR/sCR at the time of MRD evaluation) had detectable MRD with both methodologies. Furthermore, survival plots showed an improved prognosis for those patients with undetectable MRD at a minimum sensitivity of 10⁻⁵, implying that in our series, the prognostic value of MRD remained significant irrespective of the selected method. However, future implementation of stringent rules in the collection, processing, and analysis of MRD samples should be applied in order to harmonize NGS and NGF reports and reduce potential sources of error.⁴³ 

Our results support the use of the LymphoTrack strategy to detect and evaluate clonality in patients with MM, with excellent applicability and comparable results to the current gold standard, namely Sanger sequencing. Although notable differences between protocols were present owing to suboptimal procedures and lack of standardization for this NGS strategy, the LymphoTrack-based NGS and NGF assays were also comparable in terms of MRD evaluation, indicating that both are suitable options for MRD studies in patients with myeloma, as our previous publication already pointed out.³²  A similar publication from the Memorial Sloan Kettering Cancer Center group using LymphoTrack has recently described similar findings, including a high clonality detection rate, and a good concordance with high-sensitivity flow cytometry techniques.⁴⁴  For any institution, the choice of methodology would depend on secondary factors, such as expertise or availability of equipment. Multiparametric flow cytometry has a shorter turnaround time and does not depend on the identification of clones at diagnosis, but samples must be rapidly processed to maintain cell viability, and a high level of expertise is required to deliver an accurate interpretation of the results. In contrast, NGS approaches can be successfully applied to formalin-fixed, paraffin-embedded or fresh tissues, as well as frozen DNA samples; both normal and tumor-related clones can be tracked over the course of the disease with NGS, allowing a dynamic analysis of the B-cell compartment.

Altogether, these findings support the usefulness of alternative NGS approaches in MM, demonstrating a statistically significant level of agreement with previously validated methods routinely used for clonality detection and MRD assessment, and underline the need for further standardization of quantitation procedures of the LymphoTrack assay for use as a suitable alternative to the ClonoSEQ assay.

The authors wish to thank Fuensanta Martinez-Ruicobo, PhD; Alicia Antón, AS; Rebeca Maldonado, AS; Montserrat Hernández-Ruano, AS; Ana Balanzategui, BPharm; Estrella Arnés, AS; Rocío Corral, MD; and Francisco Boix, PhD (University Hospital of Salamanca, Spain) for their technical support. The authors are also very grateful to the patients who participated in this study.