Flow Cytometric Patterns of CD200 and CD1d Expression Distinguish CD10-Negative, CD5-Negative Mature B-Cell Lymphoproliferative Disorders

Abstract

Objectives:

The importance of distinguishing mature B-cell lymphoproliferative disorders (B-LPDs) is highlighted by the distinct treatments used for and varying prognoses seen in association with these different diseases. Immunophenotyping allows for accurate and efficient differentiation of many B-LPDs. Recently, we showed that CD200 is highly expressed in hairy cell leukemia (HCL) but not in marginal zone lymphoma (MZL), lymphoplasmacytic lymphoma (LPL), or hairy cell leukemia-variant (HCL-v). Here, we assessed the usefulness of a flow cytometric panel combining CD200 and CD1d with CD25, CD103, and CD11c to distinguish CD10–, CD5– B-LPDs.

Methods:

We analyzed the expression of CD200 and CD1d by flow cytometric analysis in 79 cases of CD10–, CD5– mature B-LPDs.

Results:

Distinct patterns of CD200 and CD1d expression were seen in the examined B-LPDs. HCL showed bright positivity for CD200 along with positive staining for CD1d, whereas HCL-v showed low levels of expression for both markers. LPL demonstrated positive staining for CD200 in combination with dim to negative staining for CD1d. In contrast, MZL was commonly positive for CD1d and negative for CD200.

Conclusions:

Flow cytometric analysis of CD200 and CD1d, along with CD25, CD103, and CD11c, can aid in the diagnosis of CD10–, CD5– mature B-LPDs.

Materials and Methods

Immunophenotyping via flow cytometry allows for efficient and accurate differentiation of many mature B-cell lymphoproliferative disorders (B-LPDs), particularly in cases with positive expression of either CD10 or CD5, which help to identify follicular center–derived lymphomas or chronic lymphocytic leukemia (CLL) and mantle cell lymphoma (MCL), respectively. However, flow cytometric characterization of mature B-LPDs that lack expression of both CD10 and CD5 has traditionally been more challenging due to a lack of disease-specific markers, overlapping immunophenotypes, and antigenic variation that can be seen in such cases. Therefore, flow cytometric panels that include multiple immunophenotypic markers are often more effective in differentiating these B-LPDs than any single marker used alone. Recently, studies have suggested that flow cytometric analysis of CD200 and CD1d may be useful in distinguishing CD10–, CD5– B-LPDs.

CD200 (OX-2 antigen) is a type I immunoglobulin superfamily membrane glycoprotein that is normally expressed in multiple different cell types, including B lymphocytes, a subset of T lymphocytes, dendritic cells, endothelial cells, and neurons.¹ Among hematolymphoid neoplasms, CD200 is expressed in CLL,² multiple myeloma,³ acute myeloid leukemia,⁴ B-cell acute lymphoblastic leukemia,⁵ and classical Hodgkin lymphoma.⁶ CD200 expression has been identified as a negative prognostic indicator in multiple myeloma³ and acute myeloid leukemia,⁴ possibly due to direct suppression of the antitumor T-cell–mediated⁷ and natural killer cell–mediated⁸ immune responses by CD200. In addition, multiple recent studies have demonstrated that high CD200 expression can be a useful marker in distinguishing CLL from MCL and other mature B-LPDs.^5,9-13 Furthermore, we and others have shown that CD200 is highly expressed in hairy cell leukemia (HCL)^6,9-11,13,14 but not in hairy cell leukemia variant (HCL-v)⁹ or marginal zone lymphoma (MZL),^9,12,13 suggesting that this marker may be useful in the diagnosis of CD10–, CD5– B-LPDs.

CD1d is a highly conserved major histocompatibility complex class I–like cell surface glycoprotein that functions in presenting lipid and glycolipid antigens to T cells,¹⁵ including CD1d-restricted natural killer T cells.¹⁶ CD1d is normally expressed in a wide range of epithelial, mesenchymal, and hematopoietic cell types, including hematopoietic stem cells, thymocytes, monocytes, macrophages, and peripheral blood B cells.¹⁷ In B lymphocytes, CD1d expression is thought to be downregulated by B-cell activation, with resting, naive, and marginal zone B cells expressing higher levels of CD1d than activated and memory B cells.¹⁸ Recent work has shown that lower expression of CD1d is seen in CLL compared with MCL¹⁷ and that among cases of CLL, increased expression of CD1d is associated with poor prognosis.^19,20 In addition, CD1d expression has been demonstrated in splenic MZL but not in HCL-v or lymphoplasmacytic lymphoma (LPL).¹⁷

Results

Patient Samples

Given the difficulty in distinguishing CD10–, CD5– B-LPDs by flow cytometry, we examined the utility of a flow cytometric panel that included CD200 and CD1d as well as CD19, CD25, CD103, and CD11c in the evaluation HCL, HCL-v, MZL, and LPL.

Flow Cytometric Analysis

With approval from the Partners institutional review board, we analyzed samples received as part of routine clinical care from July 2013 to July 2016. Flow cytometric analysis was performed in the Brigham and Women’s Hospital Hematology Laboratory on 79 specimens from 71 patients with HCL, HCL-v, MZL, and LPL. Final pathologic diagnoses were determined using 2008 World Health Organization criteria and based on morphologic, immunohistochemical, flow cytometric, molecular, and clinical data, where applicable. Cases in which there was considerable diagnostic uncertainty or where a definitive lesional population was not identified in the sample examined were excluded from analysis. The cohort was comprised of 34 cases of HCL, three cases of HCL-v, 20 cases of LPL, and 22 cases of MZL. Molecular analysis confirmed the presence of a BRAF V600E mutation in all 22 tested cases of HCL and the presence of an MYD88 L265P mutation in 13 of 14 tested cases of LPL. The samples analyzed included 38 bone marrow aspirates, 28 peripheral blood samples, four spleen specimens, one lung tissue specimen, one pleural fluid sample, one parotid gland sample, and six lymph node specimens.

Antigen expression was measured using six-color flow cytometric analysis performed on a FACSCanto II (BD Biosciences, San Jose, CA) flow cytometer. All cases were initially characterized by flow cytometric analysis with a complete lymphoma panel. The immunophenotypic panel employed for the study included CD19–allophycocyanin (APC)–cyanine (Cy) 7, CD200–peridinin chlorophyll (PerCP)–Cy5.5, CD1d–R-phycoerythrin (PE), CD11c-APC, CD25-PE-Cy7, and CD103–fluorescein isothiocyanate (FITC) (all from BD Biosciences). In total, 20 μL antibody was added for FITC, PE, and PerCP-Cy5.5 reagents, and 5 μL antibody was added for APC, APC-Cy7, and PE-Cy7 reagents. Reagents were added to 100-μL aliquots of sample cells and incubated in the dark for 15 minutes. RBCs were lysed by the addition of 1 mL FACS lysing solution and sample incubation for 15 minutes in the dark, after which the cells were washed in isotonic solution, centrifuged, decanted, and resuspended in 0.5 mL deionized water. Analysis was performed using a BD FACSCanto II flow cytometer, and data were analyzed using FACSDiva software (BD Biosciences). Neoplastic cells were identified and distinguished from normal B cells based on CD19 expression, forward light-scatter location, and, where applicable, CD11c, CD25, and/or CD103 positivity. The mean fluo- rescence intensity (MFI) of CD200 and CD1d was determined for control and neoplastic populations using FACSDiva software, and expression levels of CD200 and CD1d were scored as negative, dim, positive, or bright positive. Normal B-cell staining for CD200 and CD1d was assessed in all cases where feasible, showed minimal variation between samples (see below), and constituted the longitudinal quality control.

Statistical Analysis

Discussion

In contrast to the findings for HCL cases, all three cases of HCL-v examined (including two peripheral blood specimens and one bone marrow specimen) were negative for both CD200 (mean MFI, 435) and CD1d (mean MFI, 532). These cases were negative for CD25 (3/3) but were positive (2/3) or dimly positive (1/3) for CD103 and positive for CD11c (3/3).

The 20 cases of LPL examined included 18 bone marrow specimens, one peripheral blood specimen, and one lymph node specimen. More than half of the LPL cases demonstrated a unique phenotype of CD200 positivity with dim to negative staining for CD1d (Figure 2). Positive staining for CD200 was seen in 13 (65%) of 20 cases (mean MFI, 1,640). The remaining cases showed dim staining (4/20; mean MFI, 781) or negative staining (3/20; mean MFI, 426) for CD200. CD1d expression was predominantly negative (15/20; mean MFI, 381) or dim (4/20; mean MFI, 853). One case of LPL showed positive staining for CD1d (MFI, 1,771). The results are summarized in Tables 1 and 2. There were no significant differences in CD200 or CD1d expression in bone marrow vs peripheral blood (data not shown) or in cases tested vs untested for the MYD88 mutation. Cases of LPL were also negative or dimly positive for CD25, CD103, and CD11c (6/20 dim and 14/20 negative for CD25; 1/20 dim and 19/20 negative for both CD103 and CD11c). The differences in CD200 and CD1d staining between HCL and LPL were statistically significant (unpaired t test, P < .0001 for each), as was the difference in pattern of CD200/CD1d staining in HCL vs LPL (Fisher exact test, P < .0001).

MZL was the only B-LPD for which a pattern of CD1d positivity and CD200 negativity was seen (Figure 2). The 22 cases of MZL included seven peripheral blood specimens, three bone marrow samples, four spleen samples, one pleural fluid specimen, one lung and one parotid gland samples, and five lymph node specimens. CD1d was positive in nine (41%) of 22 cases (mean MFI, 1,918), dim in seven (32%) cases (mean MFI, 782), and negative in six (27%) cases (mean MFI, 458) of MZL. Staining for CD200 was commonly negative in cases of MZL (13/22 [59%]; mean MFI, 316), with the level of staining being below that seen in reactive lymphocytes. Dim staining for CD200 was seen in eight cases of MZL (mean MFI, 808), and one case of MZL was positive for CD200 (mean MFI, 1,283). The results are summarized in Tables 1 and 2. There were no significant differences in CD200 or CD1d expression in bone marrow vs peripheral blood (data not shown). Cases of MZL were also predominantly negative for CD25 (15/22 negative, 4/22 dim, and 3/22 positive) and CD103 (20/22 negative and 2/22 positive) and showed variable staining for CD11c (8/22 negative, 9/22 dim, and 5/22 positive).

The differences in CD200 and CD1d staining between HCL and MZL and between LPL and MZL were statistically significant (unpaired t test, P < .0001, P = .0021, P = .0002, and P = .0030, respectively), as was the difference in pattern of CD200/CD1d staining in HCL vs MZL and in LPL vs MZL (Fisher exact test, both P < .0001). In addition, the CD200+/CD1d+ pattern, when present, was 94% sensitive and 98% specific for HCL; the CD200+/CD1d– pattern, when present, was 60% sensitive and 97% specific for LPL; and the CD200–/CD1d+ pattern, when present, was 41% sensitive and 100% specific for MZL.

While multiple modalities are now routinely used in the diagnostic workup of lymphoproliferative disorders, flow cytometry often provides immunophenotypic information at least 24 hours prior to morphologic and immunohistochemical results and often days to weeks prior to the availability of molecular data. Therefore, flow cytometric panels that can differentiate various B-LPDs are routinely used in clinical practice to provide rapid diagnostic information. In addition, not all cases of mature B-LPDs are submitted for molecular characterization. For example, in our study, clinicians at our tertiary care center did not order mutational analysis for the BRAF and MYD88 mutations in a subset of cases of HCL and LPL, respectively. In addition, many flow cytometry laboratories perform testing for multiple hospitals that may not perform a complete morphologic, immunophenotypic, and cytogenetic characterization of cases, or diagnostic specimens may be insufficient for complete and definitive characterization. For these reasons as well as speed, it is desirable for diagnostic cytometry laboratories to render as complete a characterization of mature B-LPDs as possible. Here, we present data that support the use of CD200 and CD1d in flow cytometric panels used to evaluate CD10–, CD5– B-LPDs, which can often be difficult to distinguish due to overlapping clinical and immunophenotypic characteristics. We find that most cases of HCL are brightly positive for CD200 and positive for CD1d, whereas HCL-v is often negative for these markers. LPL shows positive staining for CD200 with dim or negative staining for CD1d in more than half of cases, while MZL shows positive staining for CD1d in just under half of cases, commonly with negative staining for CD200. The levels of staining for CD200 and CD1d seen in HCL compared with that seen for LPL and MZL were statistically significantly different, as were the differences in the patterns of CD200/CD1d staining in the B-LPDs analyzed. Although the sensitivities of the various CD200/CD1d patterns for the diagnoses of HCL, LPL, or MZL vary from 41% to 94%, the patterns, when present, are highly specific for HCL, LPL, or MZL.

We previously examined expression of CD200 in a wide variety of B-LPDs.⁹ In the prior study, in which an MFI of 1,000 or less was considered negative and MFIs above 1,000 were considered positive, HCL was found to show the brightest expression of CD200, with a mean MFI of 15,411. LPL also showed positive staining for CD200 (mean MFI, 2,131), whereas HCL-v and MZL were negative for CD200 (mean MFIs, 742 and 913, respectively). Therefore, the patterns of CD200 seen in the current study match those seen previously.

The group of lymphoproliferative disorders examined here commonly shows antigenic variation among markers used to help distinguish these diseases, and we found that CD200 and CD1d were no exception. Although bright CD200 expression and positive expression of CD1d seem to be present in most HCL cases, the patterns of expression in other entities were more variable, with approximately half of LPL and MZL cases showing positivity for CD200 or CD1d, respectively. Given the potential for variable antigen expression, the interpretation of patterns of expression of multiple markers is likely to be most informative. This may be particularly useful in the context of CD103 positivity, which is most commonly seen in HCL and HCL-v but can also occur in MZL Figure 4.²¹ We found that HCL, typically positive for CD11c and CD25, showed bright positivity for CD200 and positive expression of CD1d. HCL-v and MZL are commonly negative for CD25 but can both be positive for CD11c and CD103. Both would be expected to show dim to negative staining for CD200; however, positive expression of CD1d would favor a diagnosis of MZL. In this case, the patient had a known history of splenic MZL that had previously been CD103 negative and had acquired CD103 expression at relapse. MZL can be particularly challenging to diagnose by flow cytometry due to the lack of diagnostic marker expression in many cases. Although CD1d positivity can be helpful, approximately 50% of MZL cases showed dim or negative expression for CD1d. In this context, we found that decreased expression of CD200, below the level seen in reactive B cells, in combination with dim expression of CD1d, is also suggestive of a diagnosis of MZL Figure 5.

With the increasing awareness of genetic alterations that underlie various malignancies, molecular and cytogenetic data will likely come to play a larger role in differentiating CD10–, CD5– B-cell lymphomas. Most cases of HCL contain BRAF V600E mutations,²² while the presence of an MYD88 mutation strongly supports a diagnosis of LPL.²³ At least a subset of cases of HCL-v and BRAF V600E negative HCL has been shown to harbor activating mutations in MAP2K1.^24,25 Finally, multiple recurrent genetic lesions have been shown to be characteristic of MZL. In addition to deletion of 7q in splenic MZL, both splenic MZL and nodal MZL are now known to show recurrent mutations in multiple genes, including NOTCH2, MLL2 (KMT2D), and KLF2, whereas nodal MZL appears to uniquely also harbor recurrent mutations in the PTPRD gene.^26-29 While knowledge of these genetic lesions will certainly augment the diagnosis of these neoplasms, genetic data are often not available for an extended period after specimens are collected, or genetic analysis may not be performed in all cases. In contrast, flow cytometric data are often available within 24 hours of specimen collection and can therefore provide rapid and valuable diagnostic information.

The importance of distinguishing mature B-LPDs is highlighted by the distinct treatments used for and varying prognoses seen in association with these different diseases. For example, HCL is often effectively treated with purine analogues, including cladribine and pentostatin, which can induce long-term remission in most patients³⁰; in contrast, HCL-v typically responds poorly to these medications, tends to pursue a more aggressive course, and may be treated with rituximab or anti-CD22 immunotoxin. We have found that HCL uniquely shows strong expression of both CD200 and CD1d, which distinguishes this disease from other CD10–, CD5– B-cell lymphomas.

The identification of bright CD200 expression on the surface of HCL cells may hold therapeutic implications. Studies have suggested that anti-CD200 therapy may be effective as a therapeutic target, possibly by blocking CD200-mediated immunosuppressive effects, thereby enhancing the antitumoral T-cell response.^31-34 The immunosuppressive effect of CD200 may in part be mediated through upregulation of the immune checkpoint protein PD-1 on T cells and inhibition of antitumoral T-cell activation.³⁵ These findings suggest that anti-CD200 therapy, such as the humanized, murine-derived antihuman CD200 antibody ALXN6000, which is currently under investigation, may be useful in the treatment of HCL, particularly in the context of relapsed/refractory disease.

In summary, we have found that flow cytometric analysis for CD200 and CD1d, along with CD25, CD103, and CD11c, is useful for the characterization of mature CD10–, CD5– B-LPDs. In our current practice, a lymphoma panel, which includes T-cell markers and B-cell markers, is employed for the characterization of possible lymphoproliferative disorders and includes CD19, CD20, CD5, CD10, CD23, and CD11c. A second tier panel, which includes CD25 and CD103, is employed for the analysis of possible cases of HCL and HCL-v. Based on the current study, we plan to expand our second tier panel to include CD200 and CD1d for the characterization of CD10–, CD5– B-LPDs.

References

Author notes