Hemolytic Indicators, Special Stains, Other Studies, and Flow Cytometry

• Bilirubin. Hemolysis usually raises the unconjugated (indirect) fraction. The magnitude depends on the rate of red-cell destruction and hepatic clearance, so liver disease or Gilbert syndrome can alter the pattern. Albumin-bound unconjugated bilirubin is not filtered by the glomerulus and does not cause bilirubinuria. Urine urobilinogen may increase.
• Plasma and urine hemoglobin, urine hemosiderin. Intravascular lysis releases free hemoglobin. Once plasma-binding and tubular-reabsorption capacity is exceeded, plasma may become pink to red-brown and hemoglobin appears in urine. A reagent strip positive for heme with few or no intact erythrocytes in the sediment indicates hemoglobin or myoglobin. Correlate it with the plasma and clinical findings. Renal tubular cells that reabsorb filtered hemoglobin later slough into the urine and stain with Prussian blue. Hemosiderinuria usually appears several days after intravascular hemolysis and can persist after plasma and urine free hemoglobin clear.
• Haptoglobin. Haptoglobin binds free hemoglobin, and the complex is rapidly cleared. A low result supports hemolysis and is most striking with intravascular lysis, although it can also fall in predominantly extravascular disease. Hepatocellular disease can lower haptoglobin. An acute-phase response can raise it. A normal value does not exclude mild or compensated hemolysis.
• Lactate dehydrogenase. LDH rises with red-cell destruction but is nonspecific. Liver disease, tissue injury, ineffective erythropoiesis, and in vitro specimen hemolysis can also raise it. A hemolyzed collection cannot establish that hemolysis occurred in the patient.

• Reticulocyte count, absolute count, and production index. The response usually rises within several days when marrow function and iron, vitamin B12, and folate availability are adequate. An elevated count in anemia supports blood loss or hemolysis. An aplastic crisis, marrow disease, nutrient deficiency, renal insufficiency, or immune destruction of erythroid precursors can blunt the expected response.
• Complete count and film. The film shows polychromasia and nucleated red cells. A rising MCV reflects the influx of larger shift reticulocytes. Compare it with the patient's own baseline because the MCV may still fall within the population reference interval. A rising RDW reflects the added variance from larger cells. Reactive leukocytosis and thrombocytosis are possible. Spherocytes occur with macrophage-mediated hemolysis, and schistocytes occur with fragmentation hemolysis. When schistocytes appear with thrombocytopenia, communicate the pattern promptly so thrombotic microangiopathy, disseminated intravascular coagulation, or another fragmentation process can be evaluated. Morphology alone does not distinguish these causes.
• Bone marrow, rarely needed, shows erythroid hyperplasia with a decreased myeloid-to-erythroid ratio. Core biopsy assesses cellularity more accurately than an aspirate.

Choose additional tests based on the pattern. Urinalysis can document hemoglobinuria, urobilinogen, or hemosiderin. The direct antiglobulin test detects immunoglobulin or complement bound to red cells and supports immune hemolysis when the rest of the findings agree. A positive result can occur without hemolysis, and low-level, low-affinity, IgA-mediated, or uncommon IgM-mediated immune hemolysis can be missed by a routine test. Add iron, vitamin B12, and folate studies when the indices, reticulocyte response, or film suggests a concurrent production problem.

Cytochemical stains demonstrate enzyme activity or cellular material on blood and marrow films. They support lineage assignment when morphology is ambiguous. Current acute-leukemia classification integrates morphology, immunophenotype, genetics, and clinical findings, so a stain pattern cannot establish a disease entity by itself. Enzyme activity can deteriorate in aged or improperly stored films, and fixation requirements differ by stain. Run an appropriate positive control each time and, when the method requires it, a negative or inhibited control.

• Myeloperoxidase (MPO) is present in primary granules of myeloid cells. Granulocytes and Auer rods stain strongly, monocytes are weaker, and lymphoid cells are negative. MPO-positive blasts provide strong evidence of myeloid differentiation. Maturing granulocytes serve as an internal positive population.^{11 Keohane EM, et al. Rodak's Hematology. 2025.}
• Sudan black B (SBB) stains phospholipids and other cellular lipids. Its pattern generally parallels MPO, with increasing granulocytic staining during maturation, weak or variable monocytic staining, and negative lymphoid cells. SBB is less lineage-specific than MPO.
• Esterase stains distinguish granulocytic from monocytic lineage. Naphthol AS-D chloroacetate esterase, the specific esterase, is positive in granulocytic cells including Auer rods and negative to weak in monocytic cells. α-Naphthyl acetate and α-naphthyl butyrate esterases, the nonspecific esterases, show strong diffuse positivity in monocytic cells, confirmed by inhibition with added sodium fluoride. Granulocytic and lymphoid cells are generally negative, with occasional dot positivity in lymphocytes.

Erythrocytes carry a net negative surface charge and normally repel one another. Elevated fibrinogen and immunoglobulins reduce that zeta potential and promote rouleaux, which increases the effective settling mass. The erythrocyte sedimentation rate (ESR) is the distance in millimeters that erythrocytes in anticoagulated blood settle in one hour while the tube stands undisturbed. Anemia can raise the result without a change in inflammation, and red-cell shape or plasma-protein abnormalities can dominate the measurement. The ESR measures a physical process and does not quantify any single analyte. It is used clinically to support the evaluation and follow-up of selected inflammatory conditions with the same method over time. The result does not identify the cause of inflammation.^{22 Jou JM, et al. ICSH review of the measurement of the erythrocyte sedimentation… 2011.}

Modified Westergren method. The International Council for Standardization in Haematology (ICSH) reference method uses blood anticoagulated with EDTA and diluted 4:1 with sodium citrate or saline before testing, or blood collected directly into sodium citrate at 4:1. Reject an ordinary 9:1 coagulation citrate tube, because it has the wrong ratio. Load the sample into a 200-mm Westergren pipette with a bore of at least 2.55 mm, and read it after 60 minutes standing vertically at 18 to 25 °C, protected from sunlight, drafts, and vibration. Mix a standard collection tube with at least eight complete inversions and remix it just before filling the pipette. Read from the bottom of the plasma meniscus to the top of the sedimented erythrocyte column, excluding the buffy coat.^{22 Jou JM, et al. ICSH review of the measurement of the erythrocyte sedimentation… 2011.} ICSH 2017 guidance addresses modified and alternate methods. Before clinical use, a laboratory compares at least 30 specimens spanning the instrument's range with its predicate or the Westergren method, verifies the reference interval, runs quality control each day the instrument is used, and participates in method-appropriate external quality assessment.^{33 Kratz A, et al. ICSH recommendations for modified and alternate methods… 2017.} CLSI H02-A5 is an archived standard that remains technically valid for United States practice. The historical Wintrobe method uses undiluted blood in a 100-mm tube. Its shorter column limits the reportable range, and its results are not interchangeable with Westergren values.

Sources of error. Correct fill volume preserves the required blood-to-anticoagulant ratio. Reject clotted, leaking, mislabeled, or improperly filled specimens. ICSH 2011 permits blood-citrate specimens to be stored for up to 2 hours at ambient temperature or 4 hours at 4 °C before testing.^{22 Jou JM, et al. ICSH review of the measurement of the erythrocyte sedimentation… 2011.} General EDTA guidance is testing within 4 hours at room temperature or 24 hours refrigerated. Exact conditions depend on the method.^{1313 Oregon Health & Science University. Erythrocyte sedimentation rate.} For example, the DIESSE MINI-CUBE manual specifies 18–25 °C for the 4-hour window and 2–8 °C for the 24-hour window, with the refrigerated specimen at room temperature for 15 minutes before mixing and testing.^{1414 DIESSE Diagnostica Senese. MINI-CUBE User Manual. 2018.} That rewarming period applies only to that method. Establish your laboratory's own validated storage, rewarming, and mixing policies and procedures, approved by the medical director and consistent with the applicable method instructions.

A slight tube tilt, vibration, or high temperature accelerates settling and raises the result. Delay and low temperature generally lower it. Severe anemia can produce a large, nonspecific increase, so interpret the result with the hematocrit and the method's limitations.^{44 Clinical and Laboratory Standards Institute. Procedures for the Erythrocyte Sedimentation Rate Test; Approved… 2011.}

Glucose-6-phosphate dehydrogenase catalyzes the first, rate-limiting step of the hexose monophosphate shunt, oxidizing glucose-6-phosphate to 6-phosphoglucono-δ-lactone, which hydrolyzes to 6-phosphogluconate, while reducing NADP⁺ to NADPH. Glutathione reductase then uses that NADPH to regenerate reduced glutathione, the erythrocyte's primary defense against peroxide-mediated oxidant damage to hemoglobin, membrane lipids, and enzymes. In the unstressed cell only 5 to 10% of glucose flux crosses this shunt. Under oxidant challenge, flux rises up to thirtyfold. G6PD deficiency reduces that reserve and increases susceptibility to oxidant-triggered hemolysis from drugs such as primaquine and tafenoquine, infection, or fava beans.^{55 Luzzatto L, et al. Glucose-6-phosphate dehydrogenase deficiency. 2020.}

• Quantitative spectrophotometric assay. The reference method measures the rate of NADPH generation from an erythrocyte hemolysate with substrate and cofactor, read as rising absorbance at 340 nm. Activity is normalized to hemoglobin concentration or red-cell count and interpreted with a method-specific reference interval. Marked leukocytosis can falsely increase activity unless white cells are removed according to the assay instructions.
• WHO variant classification. The 2022 WHO system classifies genetic variants from median steady-state activity in hemizygous males or homozygous females. Class A variants have median activity below 20% with chronic nonspherocytic hemolytic anemia. Class B variants have median activity below 45% with triggered acute hemolysis. Class C variants have median activity of 60% to 150% without hemolysis. Variants with uncertain clinical significance are class U. A newly found variant between 45% and 60% remains in that class until its hemolytic risk is established. Classification of a newly described variant requires steady-state results from at least three samples from unrelated males. The system does not classify an individual patient from one enzyme result.^{66 World Health Organization. Meeting report of the technical consultation to review the… 2022.}
• Qualitative and point-of-care tests. The fluorescent-spot test detects the fluorescence of NADPH at a visual endpoint. Many qualitative screens are designed to identify activity near or below 30% of normal. They detect most severely deficient hemizygous males and homozygous or compound-heterozygous females, but they can miss intermediate activity and many heterozygous females. Semiquantitative and quantitative point-of-care devices have different decision limits. Follow the manufacturer's instructions and verify performance for the intended population. Confirm a deficient or borderline screen with the quantitative assay.^{77 Roper D, et al. Laboratory diagnosis of G6PD deficiency. 2020.}
• Timing and interference. Reticulocytes have more G6PD activity than older erythrocytes. During or soon after acute hemolysis, selective loss of the most deficient cells plus reticulocytosis can produce a falsely normal or high activity. Recent red-cell transfusion can mask the patient's activity with donor cells. State these limitations in the report when they apply. If suspicion remains after a normal result, repeat the quantitative assay after the red-cell population has returned to steady state, commonly 2 to 3 months after a hemolytic episode. After transfusion, use the testing laboratory's interval because donor red cells can persist longer.^{77 Roper D, et al. Laboratory diagnosis of G6PD deficiency. 2020.}
• Molecular testing targets known variants. The great majority of G6PD variants are single-nucleotide substitutions. Sequencing can detect variants a targeted panel misses. A negative targeted result cannot exclude an untested variant. Genotyping can resolve an equivocal phenotype during reticulocytosis or after red-cell transfusion and can identify a heterozygous female. It does not measure the functional mosaic created by X-chromosome inactivation. Recent red-cell transfusion does not usually alter leukocyte genotyping, but allogeneic stem-cell transplantation or other donor leukocyte chimerism can make a blood DNA result reflect the donor genotype. Include the specimen history with the request.^{77 Roper D, et al. Laboratory diagnosis of G6PD deficiency. 2020.}

Heinz-body preparation. Heinz bodies are precipitates of denatured hemoglobin. Incubate fresh anticoagulated blood with a supravital stain such as new methylene blue or crystal violet, then look for dark inclusions attached to the inner red-cell membrane. They support oxidative injury, including G6PD deficiency or an unstable hemoglobin. They are transient in a patient with an intact spleen, because the spleen removes the inclusions and leaves bite or blister cells. Recent transfusion can dilute the affected population.^{11 Keohane EM, et al. Rodak's Hematology. 2025.}

Osmotic fragility and related membrane tests. The traditional osmotic-fragility test exposes red cells to progressively hypotonic saline and measures hemolysis against a concurrently tested normal control. Spherocytes have less membrane reserve and lyse at higher saline concentrations, producing increased fragility. Thalassemic and iron-deficient cells usually show decreased fragility. Incubation can increase sensitivity for mild hereditary spherocytosis. Immune spherocytes can produce the same pattern, and reticulocytosis or recent transfusion can mask an abnormal population.

Eosin-5-maleimide (EMA) binding is a common flow-cytometric screen for hereditary spherocytosis. Reduced fluorescence supports a membrane-protein defect. Several other membrane disorders can also reduce binding. Each laboratory establishes its own controls and decision limits because fluorescence values depend on the instrument and method. Osmotic-gradient ektacytometry adds a deformability profile that helps distinguish membrane hydration and structural disorders. Correlate borderline or discordant results with the blood film, indices, direct antiglobulin test, family pattern, and molecular or membrane-protein studies.^{88 King MJ, et al. ICSH guidelines for the laboratory diagnosis of nonimmune… 2015.}

Flow cytometry types individual cells in suspension by measuring light scatter and the binding of fluorochrome-conjugated monoclonal antibodies to surface and cytoplasmic antigens. A hydrodynamically focused single-cell stream crosses a laser: forward scatter approximates size, side scatter approximates internal complexity or granularity, and the fluorescence signal associated with each conjugated antibody reflects relative antigen expression after compensation or spectral unmixing. Multicolor panels combine several markers per cell and resolve populations that overlap on any single parameter. Normal hematopoietic maturation is continuous, so you recognize an abnormal population by its complete marker pattern, altered antigen intensity, maturation arrest, or asynchronous expression.

Specimens and processing. Peripheral blood and bone marrow aspirate are commonly collected in EDTA or heparin according to the assay's validated requirements. Anticoagulants are not freely interchangeable. Using an early marrow aspirate pull limits hemodilution. Place fresh tissue for lymphoma evaluation in the laboratory's validated isotonic transport medium, with a separate portion preserved for morphology and histology. Formalin-fixed tissue cannot supply the viable cell suspension used by routine leukemia and lymphoma flow cytometry. Process cerebrospinal fluid and other low-cellularity fluids immediately unless a validated stabilizing medium is used. Record the collection time, transport conditions, specimen age, clots, cell count, viability, and evidence of hemodilution because each can limit sensitivity.^{9,109 Clinical and Laboratory Standards Institute. Validation of Assays Performed by Flow Cytometry. 2021.10 Kroft SH, et al. Laboratory workup of lymphoma in adults. 2021.}

For leukocyte immunophenotyping, red cells are usually lysed before acquisition. The red-cell PNH aliquot remains unlysed. Surface staining is generally performed before fixation. Validated fixation and permeabilization steps are added for cytoplasmic or nuclear targets. Viability gating and doublet exclusion, when required by the assay, keep dead cells and cell aggregates out of the population being interpreted.

Analytical controls and gating. Daily instrument checks monitor fluidics, laser alignment, and detector response. Each panel also requires validated antibody clones and titers, single-color controls for fluorescence compensation or spectral unmixing, and appropriate internal or external positive and negative controls. Acquisition retains biologic events above the validated noise threshold. Analysis then removes unstable acquisition periods, debris, doublets, and nonviable cells before identifying populations. A CD45-versus-side-scatter plot is often the starting point for leukemia analysis. Lineage gates, sequential multiparameter gates, and back-gating are still needed to test the complete marker pattern. The final report describes the specimen, abnormal population, antigen pattern, proportion, and quality limitations. For rare-event assays, the laboratory's validated limit of detection and lower limit of quantitation determine whether a population can be called not detected, detected below the quantifiable range, or reported as a percentage. The report states the sensitivity when it affects interpretation.^{99 Clinical and Laboratory Standards Institute. Validation of Assays Performed by Flow Cytometry. 2021.}

Lineage and maturation markers.

CD45 density, displayed on a logarithmic scale, helps orient the analysis: lymphocytes are bright with low side scatter, monocytes are bright with intermediate scatter, granulocytes have high scatter with lower CD45, and many blasts are CD45 dim with low scatter. Erythroid precursors are usually CD45 negative. Abnormal populations can fall outside these usual regions, so the CD45 plot is a starting map and cannot serve as the only gate.

Immunophenotypes of major hematologic neoplasms. No single marker establishes a diagnosis. Interpret the panel as a pattern together with the morphology and genetics.^{11 Keohane EM, et al. Rodak's Hematology. 2025.}