A 63-year-old man was admitted to the Department of Hematology, People's Hospital of Longhua (Shenzhen, China) in August 2023, presenting with cough, dizziness, weakness, fatigue, blurred vision and severe anemia. The symptoms had worsened (dizziness and fatigue) with paroxysmal cough and expectoration following an upper respiratory tract infection 1 month prior. An ophthalmological consultation due to blurred vision confirmed bilateral retinal hemorrhage, bilateral macular edema and bilateral ametropia. After a series of examinations, the patient was diagnosed with WM. Owing to the patient's concerns about treatment efficacy, he was transferred to the Department of Hematology, Shenzhen People's Hospital (Shenzhen, China) for further treatment. The patient was managed with a stepwise strategy of symptomatic plasma exchange followed by bortezomib plus dexamethasone chemotherapy, resulting in satisfactory clinical symptom remission and resolution of cold agglutination.

Upon admission, a series of laboratory examinations were performed to make a definite diagnosis. Of note, initial hematology analysis triggered a clot detection alarm, with the EDTA-anticoagulated blood specimen showing fine sand-like erythrocyte agglutination adherent to the tube wall (Fig. 1A). Following centrifugation, the serum stored at 4˚C rapidly formed semisolid gel-like aggregates (Fig. 1B), which gradually liquefied at room temperature (~25˚C) (Fig. 1C), confirming severe cold agglutination.

To eliminate the interference of cold agglutination and obtain valid hematological parameters, the specimens were incubated at 37˚C for 30 min and analyzed immediately. Key findings included severe anemia, as evidenced by a decreased red blood cell count (1.73x10¹²/l; reference range: 4.1-5.8x10¹²/l), hemoglobin concentration (54 g/l; reference range: 125-174 g/l), hematocrit (17.4%; reference range: 40-50%), white blood cells (5.1x10⁹/l; reference range: 3.5-9.5) and platelets (109x10⁹/l; reference range: 125-350x10⁹/l). The biochemical analysis revealed markedly elevated total protein (131 g/l; reference range: 63-82 g/l), globulin (92 g/l; reference range: 24-35 g/l), hypercalcemia (serum Ca²⁺: 3.15 mmol/l; reference range: 2.1-2.69 mmol/l) and impaired renal function [creatinine (143 µmol/l; reference range: 58-110 µmol/l); creatinine clearance (45 ml/min; reference range: >80 ml/min)]. Additional abnormalities included elevated β2-microglobulin (6.16 mg/l; reference range: 1.3-3.0 mg/l); 24-h urinary total protein (24-UMTP) (1,018.8 mg/24 h; reference range: 50-80 mg/24 h), markedly elevated IgM (83.6 g/l; reference range, 0.3-2.2 g/l), decreased IgG (6.05 g/l; reference range, 8.6-17.4 g/l) and IgA (0.55 g/l; reference range, 1.0-4.2 g/l).

Given the markedly elevated serum IgM level, serum protein electrophoresis was performed via capillary electrophoresis using the HYDRAGEL PROTEIN(E) kit (cat. no. PN4140; Sebia) according to the manufacturer's protocol, which demonstrated an elevated γ-globulin fraction (61.3% of total protein) and a distinct monoclonal protein peak accounting for 53.2% of the γ-globulin fraction (Fig. 2A). The agarose gel immunofixation electrophoresis using the HYDRAGEL IF kit (cat. no. PN4309; Sebia) according to the manufacturer's protocol to further identified the monoclonal component as IgM-λ in the serum (Fig. 2B). In addition, urine protein components were analyzed by agarose gel immunofixation electrophoresis using the aforementioned HYDRAGEL IF kit. Distinct precipitation bands were detected in both the λ light chain and free λ light chain lanes (Fig. 3A), indicating positive urine Bence-Jones protein of the free λ light chain type. Urine immunofixation electrophoresis also revealed an abnormal monoclonal band in the λ lane, indicating that the monoclonal immunoglobulin was of the free λ light chain type (Fig. 3B). All electrophoresis experiments were performed on the Sebia HYDRASYS 2 SCAN FOCUSING automated electrophoresis system (Sebia) in accordance with the manufacturer's reagent protocols.

Morphological examination is a crucial approach for evaluating hematological disorders (8). In the present study, Wright-Giemsa staining (Solution A, cat. no. BA4017C; Solution B, cat. no. BA4017D; Baso Diagnostics Inc.) was performed to assess the morphology of peripheral blood and bone marrow specimens, according to the manufacturer's protocol. Briefly, smears were stained with solution A for 1 min, mixed with solution B and incubated at room temperature for 3-10 min. After rinsing and air-drying, the smears were examined under a light microscope with x100 oil immersion. Peripheral blood smear showed rouleaux formation and atypical lymphocytes (Fig. 4A). Bone marrow cytology also demonstrated rouleaux formation and an increased lymphocyte proportion with frequent plasmacytoid lymphocytes (Fig. 4B). Furthermore, hematoxylin and eosin (H&E) staining was used to evaluate the cellular morphology of bone marrow biopsy specimens. Briefly, specimens were fixed in 10% formalin for 12 h at room temperature, embedded in paraffin and cut into 5-µm section. The sections were then deparaffinized (xylene I: 5 min; xylene II: 5 min), hydrated through graded ethanol (100% for 3 min, 90% for 3 min, 80% for 2 min and 70% for 2 min), stained with hematoxylin for 5 min, differentiated in 1% hydrochloric alcohol for 3 sec, blued in tap water for 15 min, counterstained with 0.5% eosin for 10-15 sec, dehydrated through graded ethanol (80% for 5 sec, 90% for 15 sec, 95% for 30 sec and 100% for 5 min), cleared in xylene (xylene I: 5 min; xylene II: 5 min) and mounted with neutral balsam; all at room temperature. Morphological observation was carried out using an Olympus BX53 microscope (Olympus Corporation). It revealed focal collagenous fibrosis of the medullary stroma, a diffuse lymphocytic infiltrate exhibiting plasma cell differentiation and small clusters or focal aggregates of plasma cells. The bone marrow was extensively infiltrated by B-cell lymphomatous cells, which accounted for ~80% of nucleated cells, while abnormal plasma cells comprised 10%, with only 10% residual normal hematopoietic elements remaining (Fig. 4C). These morphological features are consistent with B-cell lymphoma, which is characterized by lymphoplasmacytic infiltration and suppression of normal hematopoiesis (1,8).

To identify the specific B-cell lymphomatous subtype, immunophenotyping by FCM was performed on bone marrow cells (13-15). Fresh bone marrow aspirate (anticoagulated with EDTA) was subjected to red blood cell lysis (BD Pharm Lyse^™; cat. no. 555899; BD Biosciences) and mononuclear cells were resuspended in PBS at 1x10⁶ cells/ml. Surface staining with fluorochrome-conjugated antibodies (cat. nos and dilutions in Table SI) was performed for 20 min on ice. Intracellular light chains were assessed after fixation/permeabilization (cat. no. 554722; BD Biosciences) using anti-cytoplasmic (c)κ-FITC and anti-cλ-PE. Internal negative controls were used for both fluorescence compensation and gating. T cells (CD3⁺CD19^-) and B cells (CD3^-CD19⁺) served as reciprocal negative controls, with compensation adjusted to achieve proper alignment of populations in all dot plots. Data acquisition was performed on a BD FACSCanto II (50,000-100,000 events/tube) and analyzed with FlowJo v10.8.1 (both from BD Biosciences). Gating employed CD45/SSC-A for initial lymphocyte discrimination, followed by CD19-positive B-cell enrichment for immunophenotypic characterization. The results revealed an increased lymphocyte proportion (60.4% of nucleated cells, P4), with CD19-positive cells (P9) accounting for ~72.2% of lymphocytes, suggesting the presence of abnormally mature B lymphocytes. Furthermore, the immunophenotypes of these abnormally mature B lymphocytes were analyzed. These cells exhibited strong positivity for cλ, CD19, CD20, CD79b and human leukocyte antigen-DR; partial positivity for CD23, CD25, CD27 and CD200; and negativity for cκ, CD5, CD10, CD22, CD38, CD103 and follicular mantle clone 7 (FMC7). Additionally, plasma cells exhibiting strong CD38 expression were identified, accounting for 0.30% of the cells (P10). Subsequent analysis of CD27 and CD38 expression levels in plasma cells revealed that the proportion of normal plasma cells was ~0.10% (P11), whereas abnormal plasma cells constituted ~0.20% (P12). Abnormal plasma cells showed strong expression of cλ, CD19, CD38 and CD138, partial expression of CD20 and CD27, and absence of cκ, CD5, CD10 or CD56 (Fig. 5).

Genetic testing plays a crucial role in accurately diagnosing WM and distinguishing it from other hematologic malignancies (2). IgM multiple myeloma (IgM-MM) often harbors the t(11;14) (q13; q32) translocation or other 14q32 [immunoglobulin heavy chain (IGH)] rearrangements, whereas the MYD88 L265P mutation is detected in >90% of WM cases (1,3,14,16). Accordingly, the MYD88 L265P mutation was detected by allele-specific polymerase chain reaction (AS-PCR) at Guangzhou Huayin Medical Laboratory Center (Guangzhou, China). The following primers were used: Forward 5'-CCTTGGCTTGCAGGTGC-3', reverse 5'-AGGATGCTGGGGAACTCTTT-3', and the following probes: Mutant (FAM-MGB) 5'-AAGCGACCGATCC-3', and wild-type (VIC-MGB) 5'-AAGCGACTGATCC-3'. Each 20-µl reaction contained 10 µl 2X qPCR Mix [with uracil-N-glycosylase (UNG)], 0.4 µl each primer (10 µM), 0.2 µl each probe (10 µM), 2-5 µl template DNA and nuclease-free water to a final volume of 20 µl. The thermal cycling conditions were as follows: 50˚C for 2 min, 95˚C for 10 min; 45 cycles of 95˚C for 15 sec and 60˚C for 60 sec. Fluorescence was acquired at FAM (mutant allele) and VIC/HEX (wild-type allele) channels at the end of each cycle. In this patient, genetic testing confirmed positivity for the MYD88 L265P mutation. AS-PCR showed a Cq value of 25.52 for the mutant allele and 23.03 for the wild-type allele (Fig. 6A). Subsequently, fluorescence in situ hybridization (FISH) was performed on bone marrow samples to detect the t(11;14) (q13;q32) translocation using the Vysis LSI IGH/cyclin D1 (CCND1) DF FISH Probe Kit (cat. no. 08L58-020; Abbott Laboratories, Inc.), following the manufacturer's instructions. FISH analysis, performed using an Olympus BX53 fluorescence microscope (Olympus Corporation), revealed a normal signal pattern in 199 out of 200 interphase nuclei (99.5%), with two copies each of CCND1 and IGH and no evidence of gene fusion, indicating the absence of the t(11;14) (q13;q32) translocation (Fig. 6B).

Due to the patient showing serum IgM >40 g/l, complicated by hyperviscosity syndrome and cold agglutination, a stepwise therapeutic approach was chosen in accordance with the latest WM clinical practice guidelines (12). This strategy prioritized symptomatic relief of hyperviscosity with TPE followed by etiological chemotherapy targeting the underlying malignant lymphoplasmacytic proliferation. Initially, one cycle of TPE comprising five sessions was performed to rapidly mitigate acute hyperviscosity-related manifestations-a key and potentially life-threatening complication of WM-with fresh frozen plasma (FFP) as replacement fluid. Specifically, 2,000 ml FFP was administered in the first session, followed by 2,500 ml per session for each of the subsequent four sessions. During the first TPE session, the patient developed urticarial rash after FFP infusion, which resolved with temporary infusion cessation and intravenous dexamethasone (5 mg). Subsequent sessions were premedicated with intravenous dexamethasone (5 mg) 30 min prior to TPE, with no further allergic reactions. Upon completion of TPE, systemic chemotherapy with a bortezomib-dexamethasone regimen was initiated, consistent with frontline therapeutic recommendations for WM (1,3,12), to further control the disease and reduce tumor burden. The regimen was administered as follows: 2 mg of bortezomib was subcutaneously injected twice weekly (on days 1 and 4) for 2 consecutive weeks, and 10 mg of dexamethasone sodium phosphate was intravenously administered immediately after each bortezomib dose to enhance therapeutic efficacy and minimize adverse events. Concurrently, targeted supportive therapy was provided to manage other accompanying symptoms and improve the patient's treatment tolerance.

During one month of treatment, serial monitoring of clinical symptoms and core laboratory parameters were conducted, as shown in Table I. Before treatment, the patient presented with severe hyperviscosity syndrome and multiple organ damage, characterized by a markedly elevated serum IgM level (83.6 g/l), total protein (131 g/l) and globulin (92 g/l). Concurrently, the patient exhibited renal impairment (creatinine: 143 µmol/l; creatinine clearance rate: 45 ml/min; 24-UMTP: 1,018.8 mg/24 h), hypercalcemia (calcium: 3.15 mmol/l), and high tumor burden (β2-microglobulin: 6.16 mg/l, reference range: 1.3-3.0 mg/l). Immediately after TPE, the serum IgM level rapidly decreased by 48.5% to 43.04 g/l, with a concurrent reduction in total protein (from 131 to 99.3 g/l) and globulin (from 92 to 61 g/l), indicating immediate relief of hyperviscosity. Renal function also showed rapid improvement, with creatinine decreasing to 96 µmol/l, and hypercalcemia was corrected to 2.47 mmol/l within the normal range. At 1 month after bortezomib-based chemotherapy, sustained therapeutic effects were observed. The serum IgM level remained lower than before treatment (from 83.6 to 59.9 g/l) and the β2-microglobulin level dropped to <0.191 mg/l, demonstrating profound suppression of the underlying WM clone. Renal function was further improved, with creatinine normalized to 94 µmol/l and creatinine clearance rate increased to 74.32 ml/min. Hypercalcemia was completely corrected, with the calcium level stabilizing at 2.22 mmol/l. The 24-UMTP decreased from 1,018.8 to 867 mg/24 h, indicating ongoing recovery of renal injury.

After discharge, a structured follow-up plan was established, consisting of regular monthly visits with hematology and ophthalmology specialists to monitor disease activity. Follow-up evaluations included key indicators such as physical examinations, complete blood counts, serum biochemical tests and immunoglobulin quantification. However, follow-up was prematurely terminated after the first month (November 2023) because the patient required re-admission to the hospital for treatment of a pulmonary infection. Notably, the patient ultimately developed severe pulmonary infection and was admitted to the intensive care unit for management, thereafter progressing to multi-organ failure in the setting of profound systemic immunocompromise. Regrettably, in January 2025, the patient's family requested discharge and chose to withdraw active treatment.