Variable CD52 expression in mature T cell and NK cell malignancies: implications for alemtuzumab therapy

Authors

  • Liuyan Jiang,

    1. Flow Cytometry Unit, Laboratory of Pathology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD
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  • Constance M. Yuan,

    1. Flow Cytometry Unit, Laboratory of Pathology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD
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  • Julia Hubacheck,

    1. Flow Cytometry Unit, Laboratory of Pathology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD
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  • John E. Janik,

    1. Metabolism Branch, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Mark O. Hatfield Clinical Research Center, Bethesda, MD, USA
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  • Wyndham Wilson,

    1. Metabolism Branch, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Mark O. Hatfield Clinical Research Center, Bethesda, MD, USA
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  • John C. Morris,

    1. Metabolism Branch, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Mark O. Hatfield Clinical Research Center, Bethesda, MD, USA
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  • Gregory A. Jasper,

    1. Flow Cytometry Unit, Laboratory of Pathology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD
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  • Maryalice Stetler-Stevenson

    1. Flow Cytometry Unit, Laboratory of Pathology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD
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Dr Maryalice Stetler-Stevenson, MD, PhD, Flow Cytometry Unit, Laboratory of Pathology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Building 10, Room 2A-33, 10 Center Drive, Bethesda, MD 20892, USA.
E-mail: stetler@mail.nih.gov

Summary

The anti-CD52 antibody alemtuzumab has been explored as a novel targeted therapy in T cell malignancies. To assess the suitability of alemtuzumab therapy, we carried out a comprehensive study of CD52 expression using flow cytometry (FC) in 78 untreated patients diagnosed with mature T/natural killer (NK) cell neoplasms, including 34 adult T cell leukaemia/lymphomas (ATLL), two anaplastic large cell lymphomas (ALCL), three angioimmunoblastic T cell lymphomas (AITL), 16 cutaneous T cell lymphomas (CTCL), four extra-nodal T/NK cell lymphomas (ENT/NKCL), four hepatosplenic T cell lymphomas (HSTCL), 13 peripheral T cell lymphomas, not otherwise specified (PTCL-NOS) and two T-prolymphocytic leukaemia (T-PLL). The level of CD52 expression was quantified using QuantiBRITE standard beads. The level of CD52 expression varied widely within each diagnostic category. All AITL, HSTCL and T-PLL cases were CD52-positive and the frequency of CD52 expression was high in PTCL-NOS (92·3%), ATLL (94·1%) and CTCL (87·5%), implying a rational role for alemtuzumab in the treatment of these diseases; however, CD52 expression was low in ALCL (50%) and ENT/NKCL (25%). FC testing for cell surface expression of CD52 is indicated in patients with T/NK cell malignancies being considered for alemtuzumab therapy. Further studies are necessary to determine if the level of CD52 expression correlates with response to therapy.

CD52 is a glycosylphosphatidylinositol (GPI) anchored low molecular weight glycoprotein (21–28 kDa) (Xia et al, 1993a) expressed on the surface of B and T lymphocytes, natural killer (NK) cells, monocytes, macrophages and some dendritic cells, but not on plasma cells, granulocytes, erythrocytes, platelets, or haematopoietic progenitor cells (Hale et al, 1990; Waldmann & Hale, 2005; Hernandez-Campo et al, 2007). CD52 exists in two forms, CD52-I and CD52-II. Both forms are recognized by alemtuzumab (Campath-1H), a ‘humanized’ rat IgG1 antibody developed by transferring the antigen-specific, complementary determining regions of the rat monoclonal antibody onto a human framework (Treumann et al, 1995; Waldmann & Hale, 2005). Although CD52 has little diagnostic value, the antigen has been shown to be a valuable target for antibody therapy in lymphoid neoplasia because of its abundant cell surface expression, close apposition to the cell membrane and lack of modulation after antibody binding (Bindon et al, 1988; Treumann et al, 1995). Upon binding to the cell surface CD52, alemtuzumab induces cell destruction via activation of complement-dependent cytotoxicity, antibody-dependent cellular cytotoxicity (ADCC) and induction of apoptosis (Xia et al, 1993b; The Non-Hodgkin’ Lymphoma Classification Project 1997; Nuckel et al, 2005). While all three activities have been demonstrated in vitro, the mechanism of in vivo cell killing remains unclear.

Mature T and natural killer (NK) cell neoplasms are uncommon and comprise <10% of non-Hodgkin lymphomas (The Non-Hodgkin’ Lymphoma Classification Project 1997). Except for anaplastic lymphoma kinase (ALK)-positive anaplastic large cell lymphoma (ALCL), which is curable in the majority of patients with standard chemotherapy, and T cell large granular lymphocytic leukaemia (T-LGL), which is commonly indolent, most T and NK cell neoplasms are clinically aggressive and show disappointingly short responses to conventional chemotherapy compared to their B cell counterparts (Jaffe et al, 2001).

Alemtuzumab has demonstrated significant activity against a number of B cell malignancies, particularly in refractory and relapsed chronic lymphocytic leukaemia, as well as other non-malignant haematopoietic disorders (Keating et al, 2002; Faulkner et al, 2004; Gupta et al, 2004). Several clinical trials have explored the role of alemtuzumab in the treatment of T cell disorders, including peripheral T cell lymphoma-not otherwise specified (PTCL-NOS), T cell prolymphocytic leukaemia (T-PLL), cutaneous T cell lymphoma (CTCL) and adult T cell lymphoma/leukaemia (ATLL) (Pawson et al, 1997; Dearden et al, 2002; Lundin et al, 2003; Zhang et al, 2003; Enblad et al, 2004; Gallamini et al, 2007; Kim et al, 2007). Most of these studies have demonstrated antitumour activity; however, in these trials CD52 expression by the malignant cells was not established prior to initiation of therapy. Alemtuzumab can also result in substantial toxicity due to attendant immunosuppression associated with its use, particularly increased risk of viral and other opportunistic infections (Enblad et al, 2004; Dearden & Matutes, 2006; Alinari et al, 2007). Pretreatment evaluation for expression of CD52 may aid in guiding patient management and limit unnecessary exposure to alemtuzumab’s potentially toxic effects.

Although several groups have investigated CD52 expression on malignant lymphocytes in selected mature T and NK cell lymphomas, these studies involved a limited variety and small number of cases. Moreover, most samples were evaluated using immunohistochemical methods on archived material (Rodig et al, 2006; Piccaluga et al, 2007). One retrospective study utilizing flow cytometry (FC) analyzed cryopreserved blood specimens (Ginaldi et al, 1998). Immunohistochemical studies (IHC) examining CD52 expression have limitations. Many cases of T and NK cell lymphomas demonstrate minimal to mild cytological atypia that is difficult to appreciate on IHC slides. These lymphomas are often associated with a prominent reactive background of normal B and T cells associated with fibrosis that may obscure visual identification of the neoplastic cells and limit interpretation. Given that CD52 is ubiquitously expressed by all mature lymphocytes, it is often difficult to distinguish expression by the small to medium sized malignant lymphoid cells from the surrounding reactive lymphocytes on IHC slides. In addition, a fibrotic background may interfere with an antibody’s ability to bind to its target antigen on the fixed embedded tumour cells. Additionally, there is the possibility of antigen loss through formalin fixation and paraffin embedding, which may diminish the sensitivity of the study. Furthermore, CD52 must be expressed on the cell surface for alemtuzumab to be effective and IHC cannot determine if a membrane-associated antigen is on the external or internal cell membrane. In addition, low-level CD52 expression may not be detected by IHC whereas flow cytometry can readily detect these cell populations. Therefore, we believe that flow cytometry analysis on fresh specimens offers clear advantages in evaluating CD52 expression for patients being considered for alemtuzumab therapy.

To our knowledge, a comprehensive study of the presence of cell surface CD52 expression by flow cytometry (FC) among a broad spectrum of T and NK cell neoplasms has not been reported. We therefore conducted a prospective FC study to determine cell surface CD52 expression in patients diagnosed with mature T cell and NK cell neoplasms prior to initiation of alemtuzumab therapy. The purpose of the study was to determine if alemtuzumab was a rational therapeutic choice in specific T cell and NK cell neoplasms, based on CD52 expression by the neoplastic clone.

Methods

Case selection

Specimens from 78 patients with a confirmed diagnosis of a mature T cell or NK cell lymphoma based upon the World Health Organization (WHO) classification (Jaffe et al, 2001) were submitted to the Flow Cytometry Unit, Laboratory of Pathology, National Cancer Institute (Bethesda, MD, USA) for evaluation of cell surface expression of CD52 by FC. Patients were undergoing eligibility evaluation for research protocols studying the efficacy of alemtuzumab alone or with chemotherapeutic regimens in various T cell lymphoproliferative disorders. All patients signed institutional review board-approved informed consent to be screened for eligibility. Clinical data were obtained through medical record review and by contacting the patients’ National Institutes of Health staff physicians. All patients had a confirmed diagnosis of T cell or NK cell neoplasia with a distinctively abnormal FC immunophenotype that could be employed to distinguish malignant cells from normal lymphoid cell populations. The cases included 34 patients with human T cell lymphotropic virus I (HTLV-I) associated adult T cell leukaemia/lymphoma (ATLL), two patients with anaplastic large cell lymphoma (ALCL), three angioimmunoblastic T cell lymphomas (AITL), 16 cutaneous T cell lymphomas (CTCL), four extra-nodal T/NK cell lymphomas (ENT/NKCL), four hepatosplenic T cell lymphomas (HSTCL) including three of γδ and one of αβ T cell origin, 13 peripheral T cell lymphomas, not otherwise specified (PTCL-NOS) and two T cell prolymphocytic leukaemia (T-PLL) patients. Two cases of acute myeloid leukaemia were selected as negative controls.

Tumour subclassification was based on the WHO criteria for haematological malignancies using a combination of morphological, immunohistochemical and flow cytometric immunophenotypic studies. When necessary, molecular studies for T cell receptor gamma gene rearrangement by polymerase chain reaction (PCR) and HTLV-1 serology (performed by enzyme-linked immunosorbent assay and confirmed by Western blot) and blood HTLV-1 viral load by real-time PCR were utilized. The diagnoses were confirmed by review of the original pathology reports, FC immunophenotype, IHC immunophenotype and cytology or histological evaluation by three haematopathologists (L. Jiang, C. Yuan and M. Stetler-Stevenson). All the specimens, including peripheral blood, bone marrow, body fluid and fine needle aspiration from lymph nodes, skin nodules and soft tissue mass, were collected and analyzed prior to initiation of alemtuzumab.

Immunophenotyping

All specimens were stained within 24 h of collection with a panel of antibodies. Specimens were washed with phosphate-buffered saline (PBS) to remove cytophilic antibodies before determining cell number. Cellularity was manually determined using a haemocytometer and viability was determined by trypan blue uptake. Specimens were stained for 30 min at room temperature with a cocktail of four antibodies (antibody concentration according to manufacturer’s recommendations). If red cells were present, erythrocytes were lysed by incubating with lysing solution (150 mmol/l NH4Cl, 10 mmol/l KHCO3, 0·1 mmol/l EDTA) for 10 min at room temperature at a ratio of 1:9 (sample volume:lysing solution volume. After incubation, cells were pelleted by centrifugation (500 g for 15 min at room temperature), the media was aspirated and the cells washed twice in a PBS solution containing 0·1% NaN3. The antibody panels were chosen based on the number of cells and diagnosis. The panel included antibodies against CD2, CD3, CD5, CD4, CD8, CD7, gamma/delta receptor, alpha/beta receptor, CD45 and CD52 in all the cases. Moreover, depending on individual cases, antibodies targeting specific antigens characteristically expressed in the patient’s lymphomas were also used, including CD10 for AITL, CD25 for ATLL and CD30 for ALCL. All cells were fixed in 1·0% paraformaldehyde and stored at 4°C for up to 12 h before FC acquisition. Normal lymphoid cells within specimens served as internal controls. Normal T cells were evaluated in 20 control specimens.

Four-colour cytometry was performed using a BD Biosciences FACSCalibur flow cytometer (BD Biosciences, San Jose, CA, USA). The sensitivity of fluorescent detectors was monitored using Calibrite beads (BD Biosciences) according to the manufacturer’s recommendations. Data, collected in list mode, were analyzed with CellQuest Pro software (BD Bioscience) and FCSExpress (De Novo Software, Los Angeles, CA, USA). At least 5000 lymphocytes were acquired per tube. For analysis, relevant cell populations were analyzed by gating on forward scatter (FSC), side scatter (SSC), CD45, CD3 and characteristic markers for each specific entity (i.e. CD25 for ATLL), and the cell population of interest subsequently examined for staining with anti-CD52 (Southern Biotech, Birmingham, AL, USA).

The antibody-binding capacity (ABC) per cell of the malignant lymphoid cells was determined for anti-CD52 clone CF1D12 (Southern Biotech) using the BD Biosciences QuantiBRITE system for fluorescence quantification. The CD52 ABC value is the measurement of the mean value of the maximum capacity of each cell to bind the anti-CD52 antibody. QuantiBRITE PE Beads (BD Biosciences) are precalibrated standard beads containing known levels of PE molecules. QuantiBRITE beads were acquired on a FACSCalibur flow cytometer on the same day at the same instrument settings as the individual patient specimens. A standard curve comparing the geometric mean of fluorescence to known PE content of the QuantiBRITE beads was constructed using QuantiCALC software (BD Biosciences). The regression analysis, slope, intercept and correlation coefficient were determined. Analysis gates were drawn based upon immunophenotype and cell/forward light scatter size to include only the malignant cells for determination of the geometric mean fluorescence of CD52 staining. The ABC values were generated from the measured geometric mean fluorescence of only the malignant cells using the QuantiBRITE standard curve. As a negative control, CD4 ABC values were determined in normal CD4-negative T cells.

Results

Seventy-eight cases of mature T cell and NK cell neoplasms, encompassing eight WHO classification diagnostic categories were included in the study. These specimens were obtained as part of protocol eligibility screenings or clinically indicated diagnostic procedures. The specimens included 52 peripheral blood samples, three bone marrow aspirates three body fluid samples, 18 fine needle aspirates and two tissue (nasal) biopsy specimens.

Cell surface expression of CD52 on the tumour cells was demonstrated in all (100%) cases of AITL, HSTCL and T-PLL studied (Table I). There was a high frequency of CD52 expression by neoplastic cells in the PTCL-NOS (92·3%) and ATLL (94·1%) cases studied (Table I). However, in one PTCL-NOS and two ATLL cases the neoplastic cells failed to express CD52 on the cell surface (Table I, Fig 1), indicating that alemtuzumab therapy would be unlikely to be active in these patients due to lack of expression of the antibody target.

Table I.   Summary of patients, specimens and frequency of CD52 expression.
WHO classificationNo. of cases (n = 78)CD52 expression (%)Specimen type
  1. PB, peripheral blood; LN, lymph node; FNA, fine needle aspirate; BM, bone marrow.

Angioimmunoblastic T cell lymphoma (AITL)33 (100)2 PB, 1 LN FNA
Hepatosplenic T cell lymphoma (HSTCL)44 (100)2 PB, 2 BM
T-Prolymphocytic leukaemia(T-PLL)22 (100)2 PB
Peripheral T cell lymphoma, not otherwise specified (PTCL-NOS)1312 (92·3)6 PB, 5 LN FNA, 1 BM, 1 CSF
Adult T cell leukaemia/lymphoma (ATLL)3432 (94·1)23 PB, 1 CSF, 1 Ascites, 9 FNA (7 LN, 1 liver, 1 subcutaneous mass)
Cutaneous T cell lymphoma (CTCL)1614 (87·5)16 PB
Anaplastic T cell lymphoma (ALCL)21 (50)2 FNA (1 LN, 1 abdominal mass)
Extranodal T/NK cell lymphoma (ENT/NKCL)41 (25)2 nasal BX, 1 PB, 1 FNA (subcutaneous mass)
Figure 1.

 CD52 Expression in T cell malignancies. CD52 expression as measured by anti-CD52-PE antibody. (A) High level CD52 expression (ABC-24 894). (B) Low level CD52 expression (ABC-1317). (C) Negative for CD52. X-axis-CD3, Y-axis CD52, oval indicates malignant cells (based upon complete immunophenotypic data).

All of the 16 CTCL cases studied were mycosis fungoides/Sézary syndrome. Fourteen of these cases (87·5%) demonstrated cell surface expression of CD52 (Table I). One of two ALCL (50%) showed expression of CD52 on the CD30+ tumour cells. Only one of four (25%) of the extranodal T/NK cell lymphomas was positive for CD52 (Table I, Fig 2). CD30 expression was studied on a limited number of cases. It is of interest to note that many of the cases of CD52-negative lymphomas expressed CD30. All of the CD52-negative ATLL and PTCL-NOS cases and one out of three CD52-negative extranodal T/NK cell lymphoma were CD30-positive (2/3 were CD30-negative). Of the ALCL cases, both were CD30-positive, with one CD52-positive and one negative case. CD30 expression was not studied in the CTCL cases as they were all demonstrated to be mycosis fungoides/Sezary syndrome.

Figure 2.

 CD52 expression in NK/T cell malignancies. CD52 expression as measured by anti-CD52-PE antibody. (A) Positive for CD52 expression (ABC-1336). (B) Negative for CD52. X-axis-CD56, Y-axis CD52, oval indicates malignant cells (based upon complete immunophenotypic data).

The level of CD52 expression varied widely (Table II, Figs 1 and 2). There was no significant difference in the mean anti-CD52 antibody-binding capacity (ABC) of the neoplastic T cells in ATLL AITL, HSTCL and PTCL-NOS. Anti-CD52 ABC values were lower in the single ALCL and ENT/NKCL cases positive for CD52 and in the two T-PLL cases in comparison to the mean ABC values in the other diagnostic categories. As normal T cells are CD52-positive, the mean negative ABC value was determined using anti-CD4 and examining the CD4-negative T cells. The mean ABC value in a negative population was 82·15 ± 13·7 (mean ± SD).

Table II.   CD52 cell surface antibody-binding capacity (ABC) by tumour cells in CD52-positive cases.
DiagnosisNo. of casesMean CD52 ABCSDRange
  1. CD52 ABC value is the mean capacity per malignant cell to bind anti-CD52 antibody.

  2. *ABC value was not available for one case of gamma delta HSTCL.

Angioimmunoblastic T cell lymphoma (AITL) 3622531653113–9440
Hepatosplenic T cell lymphoma (HSTCL) 3*564150442631–11 464
T-Prolymphocytic leukaemia(T-PLL)2827455505–1149
Peripheral T cell lymphoma, unspecified (PTCL-NOS)12459040751114–13 539
Adult T cell leukaemia/lymphoma (ATLL)3254274437314–18 450
Cutaneous T cell lymphoma (CTCL)14648469191276–24 894
Anaplastic T cell lymphoma (ALCL)1597597
Extranodal T/NK cell lymphoma (ENT/NKCL)113361336

Discussion

Mature T cell and NK cell leukaemias and lymphomas are uncommon, comprising a small minority of non-Hodgkin lymphomas. With the exception of ALK-positive anaplastic T cell lymphoma, early stage mycosis fungoides and T cell large granular lymphocyte leukaemia, the majority of T and NK cell neoplasms are clinically aggressive and resistant to conventional chemotherapy. Therefore the promising outcomes attained in several clinical trials including alemtuzumab as a single agent or in combination with other chemotherapeutic agents in treatment of T cell neoplasia have generated great interest (Dearden & Matutes, 2006). The general applicability of these studies, however, depends in part upon the frequency of CD52 expression across the T cell lymphoproliferative diagnostic categories. Unfortunately, as these studies failed to establish CD52 expression on the neoplastic cells prior to initiation of therapy, no inference could be drawn between the presence of the therapeutic target, CD52, on the neoplastic cells and the response to alemtuzumab.

Our study attempted to bridge this gap, by examining CD52 expression on patient tumour specimens from eight WHO diagnostic categories of mature T and NK cell neoplasms. We observed a high frequency of CD52 expression by neoplastic cells in AITCL, HSTCL, T-PLL, PTCL-NOS and ATLL. This is in contrast to previous smaller studies that reported a significantly lower frequency of CD52 expression among these entities (Rodig et al, 2006; Piccaluga et al, 2007). Based on our data, alemtuzumab therapy is likely to be more effective in these patients than previously suspected. The percentage of cases with expression of CD52 in AITL, HSTCL and PTCL-NOS in our study was much higher than that reported in other studies utilizing IHC techniques. Piccaluga et al (2007) reported that only 41% PTCL expressed CD52; Rodig et al (2006) detected expression of CD52 in only 40% of AITL, 33% of HSTCL and 35% of PTCL-NOS. The difference between our observations and previous reports may be explained by a higher sensitivity of detection of antigen expression by FC compared to IHC (Thakhi et al, 1996). We found that CTCL, ALCL and ENT/NKCL demonstrated a lower frequency of CD52 expression. This finding strongly implies that alemtuzumab therapy may be useful in a smaller percentage of patients with these neoplasms. We observed that CD30 expression was frequently associated with absence of expression of CD52; although this was not true in all cases it further highlights the importance of evaluating CD52 expression in all patients.

As the assessment of CD52 expression typically occurs when the patient undergoes initial diagnostic evaluation and fresh tissue from the original diagnostic biopsy is no longer available, one might argue that detection of CD52 by IHC on paraffin-embedded tissue blocks is more convenient, feasible and minimizes unnecessary duplication of procedures for the patient. However, in our study, the majority of cases had FC CD52 assessment performed on a fresh specimen obtained by non- or minimally-invasive techniques. Only two cases required surgical biopsy to obtain a specimen for flow cytometric immunophenotyping (a nasal biopsy in two patients with extra-nodal T/NK cell lymphoma). The other specimens included 52 peripheral bloods, three bone marrows, three body fluids and 18 fine needle aspirations, all obtained using minimally invasive techniques and with no significant patient morbidity. The majority of these specimens also yielded additional important staging and immunophenotypic information. In view of the known lower sensitivity of IHC on paraffin tissue compared to FC in fresh tissue (Thakhi et al, 1996) and the benefit of obtaining additional data concerning the extent of disease, we believe that re-sampling to obtain a fresh specimen for flow cytometric immunophenotyping for CD52 assessment is warranted and will be important in clarifying the level of antigen expression and response to therapy. The frequent observation of circulating malignant cells in peripheral blood of patients with T and NK cell neoplasms highlights the importance of flow cytometry on peripheral blood as a simple non-invasive site for detecting T cell neoplasms.

A number of clinical trials have shown promising overall response rates with alemtuzumab in T-PLL, CTCL and PTCL-NOS; however, the response rate reported among these studies varied significantly (Dearden et al, 2001, 2002; Dearden & Matutes, 2006). For example, the overall response rate varied from 55% to 100% in CTCL, 50% to 100% in T-PLL and 36% to 60% in PTCL-NOS in different clinical trials. In our study, we found a much higher rate of CD52 expression in these three lymphoma categories (see Table I). There is clearly a discrepancy between the clinical response rate and expression rate of CD52. The reason for this discrepancy is unclear. Ginaldi et al (1998) suggested that the level of CD52 expression on the surface of the tumour cells may determine the response rate to alemtuzumab therapy. By using quantitative flow cytometric analyses, they demonstrated that CLL and T-PLL patients who failed to respond to alemtuzumab treatment had lower levels of CD52 cell surface expression, whereas the highest levels were present in patients with major responses to therapy. We found that levels of CD52 expression varied widely, from 314 to 24 894 anti-CD52 antibody-binding capacity units. Thus, the variable response to alemtuzumab therapy of patients with mature T/NK cell lymphomas/leukaemias may be due to varying levels of CD52 expression in positive cases, in addition to a subset of patients lacking CD52 cell surface expression by the tumour cells. IHC cannot accurately quantify cell surface expression of antigens, further bolstering the need for flow cytometric immunophenotyping to provide objective and precise quantification of antigen expression.

In conclusion, the frequency of CD52 expression by mature T cell and NK cell neoplasms varies according to the specific diagnostic category. Our data indicates that alemtuzumab therapy may be ineffective in a significant number of cases of ALCL and ENT/NKCL, where the frequency of CD52 expression is low. Furthermore, even in diagnostic categories exhibiting a high frequency of CD52 expression, individual cases where the neoplastic cells fail to express CD52 are encountered. This may significantly impact an individual’s response to alemtuzumab treatment. Flow cytometry is more sensitive than IHC in detecting CD52 expression, and can specifically detect expression on the cell surface, whereas IHC cannot. In addition, flow cytometry can accurately quantify levels of CD52 expression. We therefore strongly recommend flow cytometric assessment of CD52 cell surface expression prior to the initiation of CD52-targeted therapy.

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