Standardized Multicolor Flow Cytometry Protocol for Measurable Residual Disease Assessment in Acute Leukemia

Summary

This protocol describes a standardized multicolor flow cytometry workflow for measurable residual disease assessment in acute leukemia, integrating controlled specimen processing, harmonized acquisition parameters, sequential gating strategies, and denominator-adjusted interpretation to improve analytical reproducibility in routine clinical laboratories.

Abstract

Measurable residual disease (MRD) is an important prognostic indicator in the management of acute leukemia. However, routine multicolor flow cytometric (MFC) assessment remains vulnerable to pre-analytical and analytical variability, including hemodilution, inconsistent acquisition depth, and subjective interpretation. This article presents a standardized operational workflow for routine MRD evaluation in B-cell acute lymphoblastic leukemia (B-ALL) and acute myeloid leukemia (AML). The protocol incorporates strict specimen acceptance criteria, with integrated hemodilution assessment, and uses a bulk red blood cell lysis method for bone marrow and peripheral blood specimens. Analytical standardization is achieved through disease-specific two-tube, eight-color antibody panels, harmonized acquisition targets requiring at least 500,000 CD45-positive events, and fixed sequential gating strategies. In addition, the workflow applies denominator-adjusted limits of detection (LOD) and quantification (LOQ) to support an objective three-tiered reporting system. Validation across 197 clinical specimens demonstrated strong repeatability, robust dilutional linearity, and high concordance with paired orthogonal molecular assays. Collectively, this protocol provides a practical framework for standardized MRD monitoring. for standardized MRD monitoring in routine clinical laboratories by integrating controlled pre-analytical handling and harmonized analytical parameters.

Introduction

Even after achieving morphological remission, patients with acute leukemia require ongoing monitoring for measurable residual disease (MRD) to assess relapse risk accurately. Conventional morphologic and cytopathologic examinations lack the sensitivity required to detect low-level residual leukemic populations. Consequently, MRD assessment has become an essential component of disease monitoring in both acute myeloid leukemia (AML) and B-cell acute lymphoblastic leukemia (B-ALL). Recent recommendations from the European LeukemiaNet emphasize that reliable MRD assessment in AML depends on technical standardization and interobserver reproducibility rather than simple assay availability^1,2. In B-ALL, multicolor flow cytometry (MFC) remains particularly important for patients without stable molecular targets. Previous studies have demonstrated that standardized multicolor flow cytometric approaches using structured antibody panel designs can provide reproducible and sensitive residual disease detection³. Flow cytometry is widely used because of its broad applicability, rapid turnaround time, and cost-effectiveness. Assay performance remains highly dependent on standardized specimen preparation, acquisition parameters, and data interpretation⁴.

One of the major technical challenges in flow cytometric MRD assessment is rare-event detection, which is strongly influenced by acquisition depth and total event counts. Identical antibody panels may produce different analytical sensitivities if specimens are acquired with inconsistent denominator sizes⁵. Standardization, therefore, extends beyond antibody selection and requires harmonized specimen processing, bulk red blood cell lysis, instrument compensation, acquisition targets, and gating strategies. In bone marrow specimens, hemodilution also represents a significant pre-analytical variable because peripheral blood contamination may artificially reduce the apparent residual leukemic burden^6,7. In addition, interpretation of low-frequency events requires clearly defined reporting thresholds based on specimen-specific acquisition depth. Recent studies have emphasized the importance of denominator-adjusted limits of detection (LOD) and quantification (LOQ) for improving the consistency of flow cytometric MRD interpretation⁸.

The overall goal of this protocol is to provide a standardized workflow for routine MFC-based MRD assessment in B-ALL and AML. The protocol integrates specimen acceptance criteria, hemodilution assessment, disease-specific antibody panel design, bulk-lysis sample preparation, harmonized acquisition targets, fixed sequential gating strategies, and denominator-adjusted reporting criteria into a single operational framework^9,10. By combining standardized pre-analytical and analytical procedures, this method provides a practical and reproducible approach for routine clinical laboratory implementation of MRD monitoring. Figure 1 illustrates the overall standardized pre-analytical and analytical workflow implemented in this study. Table 1 summarizes the cohort and specimen characteristics of the patients with B-cell acute lymphoblastic leukemia (B-ALL) and acute myeloid leukemia (AML) enrolled during the validation period across predefined monitoring windows (End of Induction, Consolidation, and Pre-HSCT).

Figure 1: Standardized workflow for routine multicolor flow cytometric MRD assessment. (A) Pre-analytical workflow showing specimen screening and quality assessment across predefined monitoring intervals. (B) Analytical workflow from bulk red blood cell lysis and antibody staining through sequential gating, denominator-adjusted interpretation, and final MRD reporting. Please click here to view a larger version of this figure.

Table 1: Consolidated patient demographics, clinical specimen characteristics, and routine acquisition metrics. This single revised table summarizes the baseline clinical features of the enrolled cohort comprising B-cell acute lymphoblastic leukemia (B-ALL) and acute myeloid leukemia (AML) cases and combines patient-level demographics with specimen-level characteristics, treatment monitoring intervals, viability, hemodilution scores, acquisition depths, and denominator-adjusted measurable residual disease classifications. Please click here to download this Table.

Protocol

All procedures involving human specimens were performed in accordance with institutional guidelines and approved by the Ethics Committee of The Second People’s Hospital of Neijiang (Approval No. PXTS887192). The study utilized de-identified residual clinical specimens obtained after routine diagnostic testing. Informed consent was waived because the study did not affect patient care, treatment allocation, or specimen collection procedures. The chemicals, reagents, and equipment used in the protocol are listed in the Table of Materials.

1. Specimen collection and pre-analytical assessment

1. Collect bone marrow aspirates as the preferred specimen type for measurable residual disease (MRD) assessment. Use peripheral blood specimens only when bone marrow aspiration is clinically contraindicated or during selected surveillance and comparator analyses.
2. Collect the bone marrow specimen during the first aspirate pull whenever possible. Avoid subsequent aspirate pulls to minimize peripheral blood contamination and underestimation of low-level residual disease.
3. Transfer specimens into dipotassium ethylenediaminetetraacetic acid (K2-EDTA) or sodium-heparin anticoagulant tubes immediately after collection. Store and transport the specimens at room temperature (18–25 °C) and protect them from direct sunlight.
   CAUTION: Do not refrigerate the specimens.
4. Process all specimens within 24 h of collection. For stability studies, retain aliquots for repeat analysis at 24 h and 48 h while maintaining room-temperature storage conditions.
5. Verify the patient identifier, anticoagulant type, specimen integrity, and collection time upon specimen receipt. Reject specimens demonstrating extensive clotting, severe hemolysis, or marked cellular degeneration
6. Assess bone marrow specimens for hemodilution before final interpretation. Calculate a semi-quantitative hemodilution score ranging from 0.00 to 1.00 to estimate peripheral blood contamination.
7. Determine this value by dividing the proportion of mature lymphocytes by the sum of CD34-positive progenitors, immature granulocytes, and mature lymphocytes, where a higher quotient indicates greater dilutional effect.
   NOTE: Compute the score using the following formula:
    
   where a higher score indicates greater peripheral blood contamination.
8. Classify hemodilution scores as follows: <0.20, acceptable; 0.20–0.39, mildly hemodiluted; and ≥0.40, substantially hemodiluted.
   NOTE: Do not assign definitive MRD-negative interpretations to markedly hemodiluted specimens unless adequate denominator size and preserved marrow precursor architecture are confirmed.

2. Disease-specific antibody panel preparation

1. Prepare standardized disease-specific two-tube, eight-color antibody panels for B-ALL and AML analysis. Achieve panel standardization by strictly adhering to fixed backbone markers across all tubes for stable population gating.
2. Utilize consistent fluorochrome conjugates for corresponding lineage antigens and strictly follow manufacturer-recommended titration volumes to minimize lot-to-lot fluorescence variability.
3. Refer to Supplementary Table 1 for marker combinations, fluorochrome assignments, antibody clones, staining volumes, and acquisition parameters.
4. For B-ALL assessment, use CD19, CD45, CD34, CD10, and CD20 as backbone markers in both tubes to define precursor populations and maturation patterns. Add CD38, CD58, and CD81 to Tube 1.
5. Add CD66c, CD123, and either CD73 or CD304 to Tube 2 according to the diagnostic leukemia-associated immunophenotype established at baseline evaluation.
6. For AML assessment, use CD45, CD34, CD117, CD13, CD33, and HLA-DR as backbone markers in both tubes to define blast and immature myeloid populations.
7. Add CD7 and CD56 to Tube 1 to evaluate cross-lineage antigen expression. Add CD11b and CD15 to Tube 2 to evaluate myeloid maturation abnormalities.

3. Bulk-lysis preparation and staining

1. Transfer 2.0 mL of well-mixed bone marrow or peripheral blood specimen into a 15 mL polypropylene tube. If specimen volume is limited, use a minimum input volume of 1.0 mL.
2. Add 10 mL of prewarmed ammonium chloride lysis buffer to the specimen. Incubate the suspension for 10 min at room temperature to lyse red blood cells.
3. Centrifuge the specimen at 500 × g for 5 min. Discard the supernatant carefully without disturbing the nucleated cell pellet.
4. Resuspend the pellet in 2 mL of phosphate-buffered saline (PBS) containing 0.5% bovine serum albumin (BSA) and 2 mM ethylenediaminetetraacetic acid (EDTA). Repeat the wash step once if residual hemoglobin contamination remains visible.
5. Determine nucleated cell concentration using an automated hematology analyzer or manual hemocytometer count with trypan blue exclusion. Consider a minimum nucleated cell concentration of 5 × 10⁶ cells/mL as optimal for standard acquisition.
   NOTE: If the cellular yield falls below this threshold, concentrate the specimen by an additional centrifugation at 500 × g for 5 min, and carefully reduce the resuspension buffer volume to maximize cellular density.
6. Aliquot 5 × 10⁶ nucleated cells per tube for routine staining. Increase the input to a maximum of 1 × 10⁷ cells per tube for high-sensitivity analysis when specimen recovery permits^6,7.
7. Add the individually titrated antibody cocktail and the fixable viability dye specified in the Table of Materials to each tube according to the validated staining panel configuration. The routine validated panel uses 5 µL per fluorochrome-conjugated antibody unless manufacturer-recommended titration indicates a different antibody-specific volume.
8. Ensure the final staining reaction volume is strictly maintained at 100 µL per test to preserve optimal antibody-to-antigen stoichiometry. Incubate the specimens for 15 min at room temperature in the dark.
9. Wash the stained cells once using wash buffer and resuspend the final pellet in 500 µL of acquisition buffer.
10. Prepare single-stained compensation controls using compensation beads on the same day as specimen acquisition.

4. Instrument setup and data acquisition

1. Perform daily start-up quality control on the flow cytometer before acquiring patient specimens. Verify fluidics, laser alignment, detector performance, and fluorescence target values using calibration beads, following the manufacturer’s instructions.
2. Prepare single-stained compensation controls using the same fluorochromes included in the antibody panels. Adjust the compensation matrix before patient acquisition and confirm that the backbone markers remain within the predefined fluorescence target ranges.
3. Use the same acquisition template, detector settings, compensation strategy, and gating hierarchy for all specimens analyzed during the validation period.
   NOTE: Ensure proper harmonization across instruments by establishing standardized target median fluorescence intensity values for all detectors using specialized calibration beads, and confirm that identical samples yield comparable fluorescence profiles across platforms before clinical data acquisition.
4. Acquire data at a low-to-moderate flow rate to reduce coincident events and preserve rare-event detection accuracy. Monitor the event rate continuously during acquisition.
5. Acquire at least 500,000 CD45-positive nucleated events for each reportable MRD result. When specimen quality and event stability permit, extend acquisition to 1.0–1.5 million total events, especially when low-level residual disease is suspected.
6. Pause or stop acquisition if fluidic instability, sample clogs, abrupt event-rate changes, or visible deterioration in scatter quality occur. Refilter or remix the specimen when appropriate, and reacquire the sample only after stable flow is restored.

5. Sequential gating and denominator-adjusted interpretation

1. Analyze the flow cytometry data using validated flow cytometry analysis software. Use the same sequential gating strategy for all specimens to reduce inter-operator variability.
2. Establish normal and regenerating maturation patterns by independently acquiring and analyzing a reference bank of at least 20 normal or regenerating bone marrow specimens to serve as baseline expression templates.
3. Exclude debris using forward scatter and side scatter parameters. Isolate single cells by applying forward scatter-area versus forward scatter-height gating.
4. Exclude non-viable cells using the viability dye gate listed in the Materials table. Eliminate these non-viable cells routinely because they exhibit non-specific antibody binding and can artificially elevate background noise or mimic rare aberrant populations.
   NOTE: Require a minimum viability threshold of 75% for optimal downstream analysis, noting that specimens falling below this value warrant cautious interpretation regarding potential rare-event artifacts. Define the leukocyte denominator by gating on viable CD45-positive events with appropriate side scatter characteristics.
5. For B-ALL specimens, restrict analysis to the CD19-positive precursor compartment. Implement an alternative gating framework for patients receiving anti-CD19 targeted therapies.
6. Shift the primary evaluation to CD22, CD24, or CD34-positive compartments to capture the leukemic population independent of CD19 expression. Evaluate CD34, CD10, and CD20 maturation patterns and identify aberrant populations using CD38, CD58, CD81, CD66c, CD123, and CD73 or CD304 expression patterns.
7. For AML specimens, define the blast or precursor region using CD45, side scatter, CD34, and CD117 expression.
8. Evaluate aberrant or asynchronous expression of myeloid and cross-lineage markers, including CD13, CD33, HLA-DR, CD7, CD56, CD11b, and CD15.
9. Calculate the specimen-specific limit of detection (LOD) and limit of quantification (LOQ) using the total number of acquired viable CD45-positive events as the denominator: 
    
    
10. Classify the final MRD interpretation into one of three categories: MRD-positive, below LOD, or analytically limited.
    1. Assign the result as MRD-positive when a clear, contiguous cluster of aberrant events exceeds the specimen-specific LOD and demonstrates an internally consistent immunophenotype distinct from normal or regenerating hematopoietic populations.
       NOTE: Do not classify scattered, non-clustered events as MRD-positive. Utilize an integrated analysis approach combining both the leukemia-associated immunophenotype (LAIP) tracked from the baseline diagnostic sample and different-from-normal (DfN) pattern recognition strategies.
    2. Assign the result as below LOD when rare suspicious events are present but do not reach the specimen-specific reportable threshold.
    3. Assign the result as analytically limited when the denominator size is insufficient, hemodilution is substantial, specimen quality is poor, or pre-analytical or analytical limitations prevent a confident negative interpretation.
11. Review all MRD-positive and borderline low-level cases independently by a second qualified analyst.
12. Finalize the flow cytometry interpretation before reviewing paired molecular comparator data, including quantitative polymerase chain reaction or next-generation sequencing results.  

6. Analytical validation methodology

1. Assess analytical repeatability by preparing duplicate aliquots from the same post-lysis cell suspension. Acquire and analyze at least 30 paired specimens independently.
2. Evaluate dilutional linearity by preparing serial dilutions across six levels within the expected low-burden range. Compare observed MRD values with expected MRD values across the dilution series.
3. Assess inter-operator and inter-instrument reproducibility by analyzing reference specimens across multiple qualified operators and harmonized flow cytometers.
4. Transform flow cytometric and paired molecular MRD percentages using log10 transformation before statistical comparison. Evaluate quantitative agreement using correlation analysis and Bland-Altman analysis.

Results

Characteristics of the routine implementation dataset

During the study period, 197 consecutive specimens passed eligibility screening and pre-analytical quality assessment and entered the reportable workflow. The study cohort included 98 patients, 62 (63.3%) with B-cell acute lymphoblastic leukemia (B-ALL) and 36 (36.7%) with acute myeloid leukemia (AML). The cohort consisted of 64 males (65.3%) and 34 females (34.7%), with a median age of 32 years (interquartile range [IQR], 13–49 years; range, 2–73 years). The median baseline blast percentage was 67.2% (IQR, 56.9% – 80.2%).

Most analyzed specimens were bone marrow aspirates (174/197, 88.3%), whereas peripheral blood specimens accounted for 23/197 (11.7%). According to the predefined monitoring intervals, 86 specimens (43.7%) were collected at the end of induction therapy, 65 (33.0%) during consolidation therapy, and 46 (23.4%) before hematopoietic stem cell transplantation (HSCT) (Table 1).

The median specimen viability was 88.3% (IQR, 84.4% – 92.6%). The acquisition workflow yielded a median of 1,116,901 total events (IQR, 908,521–1,258,946) prior to data cleaning. Following the sequential analytical exclusion of debris, coincident doublets, and non-viable cells, the gating strategy achieved a median of 907,325 viable single CD45-positive target events (IQR, 690,112–1,095,836), with a median acquisition time of 11.9 min. The median hemodilution score was 0.20 (IQR, 0.14–0.28). Following the denominator-adjusted interpretation, 41 specimens (20.8%) were classified as MRD-positive, and 156 specimens (79.2%) were classified as below the specimen-specific limit of detection (LOD) (Table 1).

Analytical validation: repeatability, linearity, and stability

Analytical performance of the standardized workflow was evaluated through repeatability, dilutional linearity, and short-term stability studies (Figure 2). Duplicate aliquots prepared from the same bulk-lysed specimens demonstrated strong repeatability, with a median duplicate coefficient of variation (CV) of 0.101% and a 95th percentile CV of 0.222%.

Figure 2: Analytical validation of the standardized multicolor flow cytometry MRD workflow. (A) Distribution of specimen viability across analyzed samples. (B) Correlation between expected and observed MRD values across serial dilution experiments on logarithmic analysis. (C) Agreement between independent replicate analyses during repeatability assessment. (D) Stability of MRD measurements after room-temperature storage at 0 h, 24 h, and 48 h. Please click here to view a larger version of this figure.

To evaluate low-level detection behavior, 12 serial dilution experiments comprising 72 total observations were analyzed across the expected lower MRD range. Observed MRD values showed strong agreement with expected values in a logarithmic analysis (r = 0.999, adjusted R² = 0.998, slope = 1.009, intercept = 0.018), indicating preserved linearity across low-frequency event ranges.

Short-term stability testing demonstrated progressive reductions in specimen viability over time, with median viabilities decreasing from 89.1% at 0 h to 84.9% at 24 h and 80.6% at 48 h. In contrast, the median absolute relative change in MRD burden remained limited at 24 h (6.2%) and 48 h (12.0%), supporting routine processing within a 24 h post-collection interval.

Inter-operator and Inter-instrument reproducibility

Inter-operator and inter-instrument reproducibility were evaluated using 24 reference specimens analyzed across multiple operators and harmonized flow cytometers, generating 96 total assessments. The median absolute relative deviation from the reference values was 13.7%. These findings demonstrate consistent analytical performance following standardization of specimen preparation, acquisition settings, and sequential gating strategies (Table 2).

Table 2: Analytical validation and method-comparison performance indicators for the standardized measurable residual disease workflow. This table is limited to validation outcomes, including intra-assay repeatability, dilutional linearity, short-term stability, inter-operator and inter-instrument reproducibility, and orthogonal molecular method comparison metrics; it does not duplicate patient demographics or specimen characteristics reported in Table 1. Please click here to download this Table.

Clinical application and orthogonal method comparison

An orthogonal comparison was performed using 117 paired specimens, including 75 analyzed by quantitative polymerase chain reaction (qPCR) and 42 by next-generation sequencing (NGS). The qPCR evaluations primarily monitored disease-specific fusion transcripts established at diagnosis, including BCR::ABL1, RUNX1::RUNX1T1, and PML::RARA. The NGS approaches assessed clonal immunoglobulin or T-cell receptor gene rearrangements for lymphoblastic leukemia and tracked recurrent somatic mutations using targeted myeloid panels for acute myeloid leukemia. Flow cytometric MRD values demonstrated a strong correlation with paired molecular results across measurable disease levels (r = 0.893 on logarithmic analysis).

Bland-Altman analysis demonstrated a mean log difference of −0.093 with limits of agreement ranging from −0.848 to 0.663 (Figure 3). Among discordant cases, one false-negative result was identified among 25 molecularly positive specimens. The workflow demonstrated a positive percent agreement of 78.1%, a negative percent agreement of 97.6%, and an overall concordance rate of 92.3%.

Figure 3: Clinical application and orthogonal validation of standardized MRD assessment. (A) Distribution of MRD-positive cases across disease groups and monitoring intervals. (B) Correlation between flow cytometric MRD values and paired molecular comparator results on logarithmic analysis. (C) Bland-Altman analysis comparing flow cytometric and paired molecular MRD measurements, with horizontal lines denoting the mean bias and the 95% limits of agreement across the analytical range. (D) Distribution of MRD-positive results across treatment monitoring intervals. Please click here to view a larger version of this figure.

These findings demonstrate that the standardized multicolor flow cytometry workflow achieved strong agreement with orthogonal molecular MRD assays across routine clinical specimens.

Supplementary Table 1: Comprehensive operational parameters for the standardized multicolor flow cytometry measurable residual disease workflow. This unified table delineates the essential methodological components divided into distinct sections. Section A details the disease-specific antibody panel configurations, in which CD73 or CD304 selection is based on the baseline diagnostic leukemia-associated immunophenotype. Section B outlines the standardized pre-analytical specimen processing and flow cytometric acquisition targets. Section C defines the denominator-adjusted reporting thresholds alongside final interpretation criteria.Please click here to download this file.

Discussion

The primary objective of this study was to establish a standardized workflow for multicolor flow cytometric assessment of measurable residual disease (MRD) in acute leukemia. Reliable MRD evaluation depends not only on antibody panel selection, but also on consistent specimen handling, acquisition parameters, and data interpretation¹¹. The results of this study demonstrate that harmonized specimen processing, controlled acquisition depth, and denominator-adjusted reporting criteria can improve analytical reproducibility in routine clinical laboratory settings. These findings are consistent with recent efforts to standardize flow cytometric MRD assessment across different institutions and operators¹².

An important component of this workflow is the ammonium chloride-based bulk red blood cell lysis procedure. Rare-event MRD detection requires acquisition of large cellular denominators to achieve adequate analytical sensitivity. Standardized bulk-lysis preparation enabled consistent recovery of sufficient nucleated cell numbers, frequently exceeding 1.0–1.5 million acquired events, while maintaining acceptable specimen viability. Previous studies from the EuroFlow Consortium similarly emphasized that high event acquisition is necessary to support sensitive and reproducible MRD detection¹³. In specimens with suboptimal analytical sensitivity, troubleshooting should include evaluation of specimen volume, cellular recovery, and acquisition depth.

This protocol also includes routine hemodilution assessment before interpreting low-level MRD-negative results. Hemodilution remains a major pre-analytical limitation in bone marrow-based MRD testing, as peripheral blood contamination can artificially reduce the apparent leukemic burden. Incorporating a semi-quantitative hemodilution score directly into the reporting workflow provides an additional safeguard against misinterpretation as a false negative. Previous studies have reported that hemodilution significantly affects the reliability of flow cytometric MRD analysis in acute myeloid leukemia⁷. Specimens demonstrating substantial hemodilution (≥0.40) should therefore be interpreted cautiously, particularly when acquisition depth or marrow precursor preservation is inadequate.

The orthogonal comparison performed in this study demonstrated strong overall agreement between flow cytometric and molecular MRD assessment methods. However, the discordant cases also underscore that flow cytometry and molecular assays assess distinct biological aspects of residual disease. Previous studies have shown that phenotypic shifts, clonal evolution, and regenerative hematopoiesis may contribute to discrepancies between analytical platforms¹⁴. In B-ALL, distinguishing residual leukemic precursors from normal hematogones remains particularly challenging at low disease burdens and requires integrated interpretation of multidimensional immunophenotypic patterns rather than isolated antigen abnormalities¹⁵. For this reason, all borderline and low-level MRD cases in this workflow were independently reviewed by a second analyst.

This study has several limitations. First, the validation was performed at a single institution using a limited number of harmonized instruments. Second, peripheral blood specimens were included only in selected clinical situations because they cannot replace bone marrow for routine post-treatment MRD assessment in most acute leukemia settings. In addition, some analytical parameters, including hemodilution scoring and low-level event interpretation, still require experienced operator review despite the standardized workflow. Nevertheless, the protocol provides a practical framework for routine clinical implementation of standardized multicolor flow cytometric MRD assessment^16,17. By integrating pre-analytical quality assessment, harmonized acquisition strategies, and denominator-adjusted interpretation criteria, this workflow supports more reproducible and clinically consistent MRD reporting across routine laboratory practice.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Acquisition buffer	Laboratory-prepared	PBS containing 0.5% BSA and 2 mM EDTA; final resuspension volume 500 µL	Final resuspension before flow cytometric acquisition
AML Tube 1 anti-CD117-APC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 104D2; fluorochrome: APC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Early myeloid/blast identification
AML Tube 1 anti-CD13-FITC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: L138; fluorochrome: FITC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Myeloid lineage anchor
AML Tube 1 anti-CD33-PE antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: P67.6; fluorochrome: PE; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Myeloid lineage anchor
AML Tube 1 anti-CD34-PerCP-Cy5.5 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 8G12; fluorochrome: PerCP-Cy5.5; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Immaturity marker
AML Tube 1 anti-CD45-APC-H7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 2D1; fluorochrome: APC-H7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Leukocyte backbone marker and denominator definition
AML Tube 1 anti-CD56-V500 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: NCAM16.2; fluorochrome: V500; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Aberrant blast-associated phenotyping
AML Tube 1 anti-CD7-V450 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: M-T701; fluorochrome: V450; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Cross-lineage aberrancy detection
AML Tube 1 anti-HLA-DR-PE-Cy7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: G46-6; fluorochrome: PE-Cy7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Maturation and subtype discrimination
AML Tube 2 anti-CD117-APC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 104D2; fluorochrome: APC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Early myeloid/blast identification
AML Tube 2 anti-CD11b-V450 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: ICRF44; fluorochrome: V450; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Maturation-discordance assessment
AML Tube 2 anti-CD13-FITC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: L138; fluorochrome: FITC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Myeloid lineage anchor
AML Tube 2 anti-CD15-V500 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: HI98; fluorochrome: V500; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Different-from-normal myeloid pattern assessment
AML Tube 2 anti-CD33-PE antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: P67.6; fluorochrome: PE; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Myeloid lineage anchor
AML Tube 2 anti-CD34-PerCP-Cy5.5 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 8G12; fluorochrome: PerCP-Cy5.5; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Immaturity marker
AML Tube 2 anti-CD45-APC-H7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 2D1; fluorochrome: APC-H7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Leukocyte backbone marker and denominator definition
AML Tube 2 anti-HLA-DR-PE-Cy7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: G46-6; fluorochrome: PE-Cy7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Maturation and subtype discrimination
Ammonium chloride-based red blood cell lysis buffer	Laboratory-prepared / commercial supplier	Ammonium chloride-based lysis buffer; 10 mL/test; 10 min at room temperature	Bulk red blood cell lysis
Analysis software	Beckman Coulter (Brea, CA, USA)	Kaluza 2.1	Data analysis and sequential gating
Automated hematology analyzer	Commercial supplier	Automated hematology analyzer used for nucleated cell concentration and viability assessment	Nucleated cell concentration and viability assessment
B-ALL Tube 1 anti-CD10-PE antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: HI10a; fluorochrome: PE; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Precursor B-cell definition
B-ALL Tube 1 anti-CD19-FITC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: SJ25C1; fluorochrome: FITC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	B-lineage identification and entry into precursor compartment
B-ALL Tube 1 anti-CD20-APC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: L27; fluorochrome: APC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Maturation pattern assessment
B-ALL Tube 1 anti-CD34-PerCP-Cy5.5 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 8G12; fluorochrome: PerCP-Cy5.5; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Immaturity marker
B-ALL Tube 1 anti-CD38-PE-Cy7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: HIT2; fluorochrome: PE-Cy7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Separation from regenerating normal precursors
B-ALL Tube 1 anti-CD45-APC-H7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 2D1; fluorochrome: APC-H7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Leukocyte backbone marker and denominator definition
B-ALL Tube 1 anti-CD58-V450 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: TS2/9; fluorochrome: V450; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Aberrancy detection in residual leukemic precursors
B-ALL Tube 1 anti-CD81-V500 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: JS-81; fluorochrome: V500; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Support for abnormal precursor recognition
B-ALL Tube 2 anti-CD10-PE antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: HI10a; fluorochrome: PE; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Precursor B-cell definition
B-ALL Tube 2 anti-CD123-V450 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 9F5; fluorochrome: V450; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Aberrant precursor phenotyping
B-ALL Tube 2 anti-CD19-FITC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: SJ25C1; fluorochrome: FITC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	B-lineage identification and entry into precursor compartment
B-ALL Tube 2 anti-CD20-APC antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: L27; fluorochrome: APC; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Maturation pattern assessment
B-ALL Tube 2 anti-CD34-PerCP-Cy5.5 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 8G12; fluorochrome: PerCP-Cy5.5; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Immaturity marker
B-ALL Tube 2 anti-CD45-APC-H7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: 2D1; fluorochrome: APC-H7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Leukocyte backbone marker and denominator definition
B-ALL Tube 2 anti-CD66c-PE-Cy7 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: KOR-SA3544; fluorochrome: PE-Cy7; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Leukemia-associated aberrancy support
B-ALL Tube 2 anti-CD73/CD304*-V500 antibody	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	Clone: AD2/AZ1; fluorochrome: V500; staining volume: 5 µL/test in the routine validated panel unless antibody-specific titration differs	Additional aberrancy confirmation according to diagnostic phenotype
Bone marrow / peripheral blood specimens	The Second People’s Hospital of Neijiang	Residual clinical specimens	MRD assessment
Compensation beads	Commercial supplier	Single-stained compensation beads used on the same day as specimen acquisition	Same-day compensation control preparation
Fixable viability dye	Commercial supplier	Fixable viability dye included in every staining tube	Exclusion of non-viable cells; viability gate referenced in Protocols 3.7 and 5.4
Flow cytometer	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	FACSCanto II / Navios EX	Multicolor flow cytometric acquisition
Manual hemocytometer	Commercial supplier	Manual hemocytometer count	Manual nucleated cell counting
PBS/BSA/EDTA wash buffer	Laboratory-prepared	Phosphate-buffered saline containing 0.5% bovine serum albumin and 2 mM EDTA	Cell washing and resuspension
Trypan blue solution	Commercial supplier	Trypan blue exclusion	Manual viability assessment by trypan blue exclusion
Validated antibody cocktails	BD Biosciences (Franklin Lakes, NJ, USA) / Beckman Coulter (Brea, CA, USA)	See antibody-specific rows below and Supplementary Table S1	MRD immunophenotypic detection