Hematopathology / B-CELL CLONALITY IN REACTIVE SETTINGS
Prominent Clonal B-Cell Populations Identified
by Flow Cytometry in Histologically Reactive Lymphoid
Proliferations
Steven J. Kussick, MD, PhD,1 Michael Kalnoski, MD,1 Rita M. Braziel, MD,2
and Brent L. Wood, MD, PhD1
Key Words: Flow cytometry; Follicular hyperplasia; Clonality; B cell; Polymerase chain reaction; PCR
DOI: 10.1309/4EJ8T3R2ERKQ61WH
Abstract
We describe 6 cases from the University of
Washington Hematopathology Laboratory (Seattle) in
which prominent, clonal, follicle center B-cell
populations were identified by flow cytometry and
confirmed by molecular methods, but in which the
histologic features showed reactive follicular
hyperplasia without evidence of bcl-2 overexpression or
the t(14;18). The 6 cases included 5 lymph node biopsy
specimens and 1 tonsillectomy specimen. Of the 6
cases, 5 occurred in young males (8-28 years) with no
known immunologic abnormality; the other case was a
32-year-old, HIV-positive woman. In all 6 cases, clonal
CD10+ B cells representing at least 20% of the total B
cells were identified. Available clinical follow-up
ranging from 13 to 56 months revealed no evidence of
lymphoma in any of the 6 patients. Our findings add
rare cases of follicular hyperplasia to the list of
histologically reactive settings in which clonal B-cell
populations might be present.
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Whether demonstrated directly by the Southern blot
technique or the polymerase chain reaction (PCR) or indirectly by the identification of light chain restriction by
immunophenotyping, the presence of clonality among B
lymphocytes historically has been viewed as a marker of
malignancy in lymphoid proliferations. However, several
studies have demonstrated that clonal B-cell populations
can be identified by molecular techniques in nonmalignant, antigen-driven proliferations of B cells such as
lymphoepithelial sialadenitis,1-3 chronic gastritis due to
Helicobacter pylori infection,4-7 lymphocytic thyroiditis
that has not evolved fully to lymphoma, 8 and orbital
lymphocytic proliferations.9 In addition, several studies
have demonstrated that clonal B-cell populations may be
identified by PCR in individual, microdissected tissue
fragments 10 or in individual follicles from paraffinembedded benign lymph nodes,11,12 presumably reflecting
the limited number of B-cell clones known to populate any
given germinal center.13 While there have been a number
of studies using molecular techniques to demonstrate Bcell clones in benign settings, only 1 study addressed the
possibility of light chain restriction, as determined by flow
cytometry, in such settings, and that study was limited to
lymphocytic thyroiditis.14
We describe 6 biopsy specimens of histologically reactive lymphoid tissue that contained prominent surface light
chain–restricted, follicle center–type B-cell populations by
4-color flow cytometry and clonal B-cell populations by
PCR or the Southern blot technique. Since none of the
cases showed bcl-2 overexpression by immunohistochemical analysis or the t(14;18) by PCR, and none of the
patients were reported to have developed lymphoma during
© American Society for Clinical Pathology
Hematopathology / ORIGINAL ARTICLE
follow-up periods ranging from 13 to 56 months, these
cases seem to represent benign clonal proliferations of
follicle center B cells.
Materials and Methods
The 6 cases in this series were selected to have both
flow cytometric and molecular evidence of clonality in the
setting of reactive-appearing morphologic features. All cases
were identified in the University of Washington
Hematopathology Laboratory from January 1998 through
December 2002. Two additional cases showing light
chain–restricted B-cell populations by flow cytometry but in
which clonal B-cell populations could not be identified by
PCR or the Southern blot technique were not included in the
series. The procedures used in the study were approved by
the University of Washington Human Subject Review
Committee (No. 02-3505-E 01) and were in accord with the
ethical standards established by the Helsinki Declaration of
1975.15
Flow Cytometric Analysis
In all cases, the fresh tissue was collected in RPMI
supplemented with 10% fetal calf serum. To increase the
number of cells for analysis, these specimens were finely
minced and strained through 40-µm cell strainers (Becton
Dickinson [BD], San Jose, CA) during processing. The cells
were washed once in a solution of phosphate-buffered
saline/0.3% bovine serum albumin/0.1% sodium azide,
resuspended in 100 µL of RPMI/10% fetal calf serum, incubated with the relevant antibodies for 15 minutes at room
temperature in the dark, washed again in phosphate-buffered
saline/bovine serum albumin/azide, and fixed in 1%
paraformaldehyde before flow cytometric analysis. All antibodies used in the flow cytometric analysis were conjugated
to fluorescein isothiocyanate (FITC), phycoerythrin (PE),
PE-Texas Red (ECD), or PE-Cy5. The specific fluorochromes and clones used in the flow cytometric analysis of
the B cells were as follows: CD10-FITC (clone D12, BD) or
CD10-PE (clone HI10a, BD), CD19-ECD (clone HD237,
Beckman Coulter [BC], Hialeah, FL), CD20-FITC (clone
H299, BC) or CD20-PE (clone L27, BD), CD45-ECD (clone
J.33, BC), CD5-PE-Cy5 (clone BL1a, BC), κ-FITC (clone
TB 28-2, BD), and λ-PE (clone 1-155-2, BD). In each case,
the amount of antibody used was based on the manufacturer’s suggestion or titration experiments to optimize the
signal/noise ratio. In all cases, the B cells were evaluated
using the following 4-color combinations: κ-FITC, λ-PE,
CD19-ECD, CD5-PE-Cy5 and CD20-FITC, CD10-PE,
CD19-ECD, CD45-PE-Cy5 or CD10-FITC, CD20-PE,
CD19-ECD, CD45-PE-Cy5.
To facilitate the demonstration of light chain restriction
in 3 of the 6 cases, additional 4-color antibody combinations
were used to separate the follicle center population from the
non–follicle center B cells based on the increased CD10 or
CD20 expression of the former: κ-FITC, λ-PE, CD19-ECD,
CD10-PE-Cy5 (1 case) or κ-FITC, λ-PE, CD19-ECD,
CD20-PE-Cy5 (2 cases).
In all cases, the flow cytometry was performed on
Coulter Epics XL instruments (BC). In all cases, more than
10,000 cellular events were analyzed per tube of antibodies,
and in several cases, 100,000 cellular events were analyzed
per tube. The flow cytometric data were compensated and
analyzed on Macintosh G3 or G4 computers (Apple, Cupertino, CA) using Winlist software (Verity House Software,
Topsham, ME) or software developed in our laboratory
(B.L.W.). In all cases, events with very low forward scatter,
which were likely to include degenerated cells, were
excluded from the analysis by forward vs side scatter gating.
CD19 vs side scatter gating was used to limit the flow cytometric analysis to the B cells.
Histologic and Immunohistochemical Analysis
All cases were evaluated by using H&E staining,
immunohistochemical analysis of bcl-2 expression, and in
situ hybridization to detect Epstein-Barr virus (EBV)
early RNA 1 (EBER1) messenger RNA. All immunohistochemical and in situ hybridization studies were
performed in the University of Washington Immunohistochemistry Laboratory. For the bcl-2 immunohistochemical
analysis, a 4-µm-thick section from 1 representative
paraffin block of each case was deparaffinized, rehydrated
in graded alcohol, blocked with 3% hydrogen peroxide,
and subjected to heat-induced epitope retrieval. After
cooling, the anti–bcl-2 monoclonal antibody 124 (DakoCytomation, Carpinteria, CA) was applied and incubated
for 40 minutes. Immunolocalization was performed via a
modified avidin-biotin immunoperoxidase technique with
a ferric chloride–enhanced 3,3'-diaminobenzidine chromogen. Finally, sections were counterstained with hematoxylin. Positive and negative control samples were used
for each immunohistochemical stain.
For the EBV in situ hybridization, a 4-µm-thick
section from 1 representative paraffin block of each case
was deparaffinized, rehydrated in graded alcohol, and
treated with 0.02 mg/mL of Proteinase K followed by
70% and 95% ethanol, before the application of a biotinylated probe to EBER1 messenger RNA (Operon Technologies, Alameda, CA). After a 40-minute incubation,
the slide was washed, blocked with 3% hydrogen
peroxide, and incubated with a mouse antibiotin antibody.
Immunolocalization was performed as described in the
preceding paragraph.
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Molecular Analysis
Fresh or frozen tissue samples were available for
analysis in all 6 cases. Genomic DNA was extracted using
the Puregene kit (Gentra Systems, Minneapolis, MN)
according to the method of the manufacturer, precipitated
with 70% ethanol, and resuspended in tris(hydroxymethyl)
aminomethane (Tris)–buffered EDTA. For the PCR analysis
of immunoglobulin heavy chain (IgH) gene rearrangements,
500 ng of DNA was incubated with 1 U of Taq polymerase
(Amplitaq, Applied Biosystems, Foster City, CA) and 200µmol/L concentrations of deoxynucleoside triphosphates, in
buffer containing a 12-mmol/L concentration of Tris (pH
8.3), a 1.2-mmol/L concentration of magnesium chloride, a
60-mmol/L concentration of potassium chloride, and 0.6
µg/µL of bovine serum albumin.
Consensus primers against complementarity determining
region 3 (5'-ACACGGC(C/T0GT(G/A)TATTACTGT-3') and
the JH region (5'-ACCTGAGGAGACGGTGACC-3') of the
IgH gene were used for the PCR amplifications. For the cases
analyzed since the year 2000, the primer sets included a fluorescent (Hex)-labeled primer, permitting the products to be
size-fractionated by capillary electrophoresis using an
Applied Biosystems ABI 310 Sequence Analyzer with
Genescan software. For cases before 2000, unlabeled PCR
primers were used, and the products were size-fractionated by
using agarose gel electrophoresis with ethidium bromide
staining. Negative and positive control samples for clonality
were used in all assays.
For Southern blot analysis, DNA was digested with
HindIII, BamHI, and EcoRI (all from Boehringer
Mannheim/Roche, Indianapolis, IN) for 12 to 16 hours
(overnight), size-fractionated on 0.7% agarose gels, and
transferred to nylon filters (Nytran Supercharge, Schleicher
& Schuell, Keene, NH) for 1 hour in a buffer of 5% standard
saline citrate. The blots were hybridized for 12 to 16 hours
with a phosphorus 32–labeled probe directed against the IgH
gene, in a sodium dodecyl sulfate hybridization buffer.
Autoradiograms typically were exposed to the blot for 24
and 72 hours. Positive and negative control samples were
used in all assays.
For the PCR reactions to detect the t(14;18), the unlabeled bcl-2 gene–derived primers MBR1 (5'-CGGGAATTCTTTGACCTTTAGAGAGTTGCTT-3'), MC7 (5'CGGGAATTCTCAGTCTCTGGGGAGGAGTGG-3'), and
MC8 (5'-CGGGAATTCGACTCCTTTACGTGCTGGTACC-3') were used in separate PCR reactions with the
JH primer 5'-CTCAAGCTTACCTGAGGAGACGGTGACC3'. The products were size-fractionated using agarose gel
electrophoresis with ethidium bromide staining. Positive
and negative control samples were used in all assays.
Results
The 6 cases in this series are summarized in ❚Table 1❚.
In all cases a prominent, surface light chain–restricted B-cell
population representing more than 20% of the total B cells
was identified at the first sampling of the lymphoid tissue. In
case 5, the initial sampling was a fine-needle aspiration
(FNA). Because of the identification of a B-cell clone in the
FNA sample, a second excisional biopsy was performed that
revealed the same abnormal B-cell population, although in a
lower percentage of the total B cells compared with the FNA
sample. In the remaining 5 cases, the first sampling of the
lymphoid tissue was an excisional biopsy.
Five of the 6 cases occurred in males younger than 30
years who had localized enlargement of lymphoid tissue,
predominantly in the head and neck region, and no apparent
history of immunologic problems. The other biopsy was
from a 32-year-old, HIV-positive woman with a peripheral
blood CD4+ cell count of 240/µL (240 × 106/L) at the time
of the biopsy.
❚Table 1❚
Cases With Light Chain–Restricted CD10+ B Cells and Reactive Histologic Findings
Case No./
Sex/Age (y)
1/M/8
2/F/32
3/M/14
4/M/17
5/M/28
6/M/10
Known Immune
Disorder
Biopsy Site
Histologic
Findings
No
HIV
No
No
No
No
Tonsil
Cervical LN
Cervical LN
Inguinal LN
Cervical LN
Parotid LN
FH
FH
FH
FH
FH
FH/PTGC
Follicle Center
λ Ratio
B-Cell κ/λ
0.35
4.3
10.2
0.13
9.0
12.5
Estimated Clonal
Molecular
Follicle Center B Cells* Results†
32
55
21
41
35 and 16§
27
–/+
+/+
+/ND
–/+
–/+
+/ND
Duration of
Follow-up (mo)‡
56
53
50
40
39
13
FH, follicular hyperplasia; LN, lymph node; ND, not done; PCR, polymerase chain reaction; PTGC, progressive transformation of germinal centers.
* Results are given as percentage of clonal CD10+ B cells among all B cells, based on flow cytometry. Note that all clonal follicle center B-cell populations expressed essentially
normal levels of CD19 (intermediate level), CD20 (intermediate to bright), and CD10 (low to intermediate), without CD3 or CD5. Owing to the straightforward follicle center
immunophenotype in these cells, we did not examine additional follicle center B cell–associated antigens such as CD23, CD38, or FMC7.
† Results are given for PCR/Southern blot analysis.
‡ Negative follow-up is based on lack of a subsequent biopsy of lymphoid tissue at the referring institution (for all but case 2) and lack of evidence of lymphoma in the available
clinical history. For case 2, a second lymph node biopsy was performed approximately 1 year after the initial biopsy; a clonal B cell population was not found.
§ Results are for samples obtained by fine-needle aspiration of the lesion and excisional biopsy, respectively.
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Hematopathology / ORIGINAL ARTICLE
In all cases, the light chain–restricted B-cell populations
identified by flow cytometry were of follicle center type,
showing the characteristic follicle center–associated expression of low- to intermediate-level CD10 and slightly brighter
CD20 and CD45 compared with the polyclonal non–follicle
center B cells in the specimen ❚Image 1❚ and ❚Image 2❚ (cases
3 and 2, respectively). Based on their forward and side lightscatter characteristics, the light chain–restricted B cells also
tended to be larger than the polyclonal B cells in the specimen. Four of the cases were κ light chain–restricted, while
the other 2 were λ-restricted. The levels of CD19 in the
clonal populations in this series were very similar to those of
the polyclonal B cells in the background. In addition, the
levels of expression of CD10, CD20, and CD45 were similar
to those typically seen by using flow cytometry in benign,
polyclonal follicle center populations. In case 6, cytoplasmic
Lymphocytes
FS
B cells
CD19 ECD
Follicle center B cells
λ PE
CD10 PE
CD45 PC5
SS LOG
Viable cells
bcl-2 protein expression was assessed by flow cytometry,
and there was no evidence of overexpression of bcl-2 in the
follicle center B cells (data not shown).
Molecular analysis of the IgH gene confirmed clonal Bcell populations in all 6 cases. PCR analysis identified clonal
B cell populations in a variable background of polyclonal B
cells in 3 of the 6 cases ❚Image 3❚ and ❚Image 4❚. In case 2,
Southern blot analysis confirmed the presence of a clonal Bcell population (Image 4), whereas in cases 3 (Image 3) and
6, the PCR positivity was confirmed by repeated PCR
assays. In the 3 PCR-negative cases, Southern blot analysis
demonstrated novel, rearranged bands that were not present
in the germline control (placental DNA), indicating the presence of a clonal B-cell population (data not shown).
❚Image 5❚ and ❚Image 6❚ summarize the histologic and
immunohistochemical findings in cases 3 and 2, respectively.
κ FITC
CD20 FITC
❚Image 1❚ (Case 3) Florid follicular hyperplasia in a cervical lymph node. Four-color flow cytometric evaluation demonstrates a
clonal follicle center B-cell population (blue) showing normal expression of CD45 and CD19 (second dot-plot from left), low-level
CD10, and increased CD20, compared with the non–follicle center B cells (second dot-plot from right), and κ light chain
restriction (rightmost dot-plot). ECD, PE-Texas Red; FITC, fluorescein isothiocyanate; FS, forward light scatter; PC5, PE-Cy5; PE,
phycoerythrin; SS, side light scatter.
Lymphocytes
FS
B cells
CD19 ECD
B cells
λ PE
CD10 PE
CD45 PC5
SS LOG
Viable cells
κ FITC
CD20 FITC
❚Image 2❚ (Case 2) Florid follicular hyperplasia in a cervical lymph node. Four-color flow cytometric evaluation demonstrates a
clonal follicle center B-cell population (blue) showing normal expression of CD45 and CD19 (second dot-plot from left), lowintermediate CD10, and increased CD20, compared with the non–follicle center B cells (second dot-plot from right), and κ light
chain restriction (rightmost dot-plot). ECD, PE-Texas Red; FITC, fluorescein isothiocyanate; FS, forward light scatter; PC5, PECy5; PE, phycoerythrin; SS, side light scatter.
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Case 3 demonstrated nodal follicular hyperplasia in a 14year-old boy without a history of an immunologic abnormality, while case 2 demonstrated florid follicular hyperplasia in a woman with HIV. Images 5A and 5B show a
pattern of typical follicular hyperplasia similar to those in 2
of the other nodal cases (cases 4 and 5) and the 1 tonsillar
case (case 1), all in male patients without a history of an
immunologic abnormality. Case 6 was slightly different; it
had focal progressive transformation of germinal centers in
addition to typical follicular hyperplasia.
Immunohistochemical evaluation of bcl-2 protein
showed no overexpression in the follicle center B cells
(Images 5C and 6C), consistent with the negative molecular
assay results for the t(14;18) in all cases. The large bcl2–negative area in Image 6C is a very large follicle center,
consistent with the floridly reactive appearance of the lymph
node. In addition, in situ hybridization studies revealed no
significant evidence of active EBV infection in any of the 6
cases, although there were slightly more EBV-positive cells
in the lymph node from the HIV-positive patient than in the
other tissue samples in the series (data not shown).
As shown in Table 1, during follow-up periods ranging
from 13 to 56 months there was no evidence to suggest the
development of lymphoma in any of these patients. In case 2,
a lymph node biopsy done approximately 1 year after the
biopsy specimen shown in Images 2, 4, and 6 was negative
and revealed no clonal B-cell population; to our knowledge,
none of the other patients underwent another biopsy during
the follow-up period.
Discussion
Our findings add rare cases of follicular hyperplasia to
the list of histologically reactive settings in which clonal Bcell populations might be present. There are 2 major differences between our findings and those of 2 earlier PCR-based
studies describing clonal B-cell populations in microdissected fragments of histologically reactive lymph nodes10,11:
(1) We detected clonality in large portions of tissue (excisional biopsy specimens). (2) We identified these populations by flow cytometry and used molecular methods only to
confirm the flow cytometric results. Moreover, Zhou and
colleagues10 found that whole tissue sections, unlike the
microdissected fragments, were “consistently polyclonal” by
IgH PCR. It is important to note that the cases we describe
are rare, representing only about 0.8% of all the reactive
lymph nodes that we evaluated by flow cytometry during the
Base Pairs
Fluorescence Intensity (Arbitrary Units)
60 80 100 120 140 160 180 200 220 240 260 280 300 320 340 360
7,600
7,200
6,800
6,400
6,000
5,600
5,200
4,800
4,400
4,000
3,600
3,200
2,800
2,400
2,000
1,600
1,200
800
400
0
❚Image 3❚ (Case 3) Florid follicular hyperplasia in a cervical
lymph node. Capillary electrophoresis shows the size
distribution of polymerase chain reaction (PCR) products
generated using consensus primers against the
complementarity determining region 3 and the JH region of
the immunoglobulin heavy chain gene (green),
demonstrating a prominent clonal B-cell population
corresponding to a PCR product of approximately 90 base
pairs. Red denotes size standards.
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Eco
C P
Bam
C P
Hind
C P
P
❚Image 4❚ (Case 2) Florid follicular hyperplasia in a cervical
lymph node. Agarose gel electrophoresis shows the size
distribution of polymerase chain reaction products generated
using consensus primers against the complementarity
determining region 3 and the JH region of the immunoglobulin
heavy chain gene, which demonstrates a small clonal B-cell
population in a polyclonal background in the lane designated
“P” (arrow). This finding is consistent with the Southern blot
results demonstrating faint rearranged bands in the EcoRI and
BamHI lanes, suggesting a small clonal population
(arrowheads). Lane C, control; lane P, patient.
© American Society for Clinical Pathology
Hematopathology / ORIGINAL ARTICLE
A
C
5-year period from January 1998 to December 2002. Therefore, our findings do not contradict those of a third study of
23 patients with reactive follicular hyperplasia,16 which
found no immunophenotypic evidence of light chain restriction in these cases but did find evidence of a clonal IgH
rearrangement by Southern blot analysis in 1 of the 23 cases.
In addition to the 6 cases presented herein, we also
identified prominent light chain–restricted, follicle center
B-cell populations by flow cytometry in excisional biopsy
specimens from 2 other patients during the study period,
but we omitted these 2 cases from this series because
clonal populations could not be documented by molecular
methods. If these 2 excluded cases are grouped with the
other 6, then “reactive” B-cell clones seem to be present
in 1.1% of the excisional biopsy specimens from reactive
lymphoid tissues evaluated during the 5-year period of
this study.
B
❚Image 5❚ (Case 3) Follicular hyperplasia in a cervical lymph
node. A and B, Sections show reactive-appearing morphologic
features (A, H&E, ×34; B, H&E, ×170). C,
Immunohistochemical analysis shows no evidence of bcl-2
overexpression in the follicles (bcl-2, ×34).
One of these excluded cases was morphologically
typical lymphocytic thyroiditis in a 32-year-old woman. In
this case, the initial sampling was by FNA and revealed an
apparent κ-restricted B-cell population representing about
35% of the B cells and 20% of all lymphocytes, while the
subsequent thyroidectomy specimen had a smaller abnormal
B-cell population representing about 9% of all lymphocytes.
Based on the histologic features, a large number of nonlymphoid cells, including many thyroid epithelial cells, were
present. Thus, the negative Southern blot result might have
reflected a false-negative finding owing to the relatively
small number of B cells compared with the other lymphocytes and nonlymphoid cells. The negative IgH gene PCR
result in this case and in all other PCR-negative cases in
this series likely reflected the phenomenon of somatic
hypermutation (reviewed by Spencer and Dunn-Walters17).
Note that there has been a single previous report of light
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A
B
❚Image 6❚ (Case 2) Florid follicular hyperplasia in a cervical
lymph node. A and B, Sections show some very large follicles
with reactive-appearing morphologic features (A, H&E, ×34;
B, H&E, ×170). The large area of bcl-2 negativity by
immunohistochemical analysis (C, bcl-2, ×34) and negative
polymerase chain reaction for the t(14;18) support the
impression that these represent floridly reactive follicles.
C
chain–restricted B-cell populations in benign lymphocytic
thyroiditis,14 although that report did not describe whether
the B cells were of follicle center origin.
The second excluded case was a tonsillectomy specimen
with reactive follicular hyperplasia and a λ-restricted follicle
center B-cell population from a 10-year-old boy. In this case,
only paraffin-embedded material was available for molecular
analysis, so only IgH PCR could be attempted. This assay
failed to reveal a clonal B-cell population, again presumably
because of somatic hypermutation.
The uniform lack of bcl-2 overexpression and the
t(14;18) in our cases argues strongly against the possibility
of “in situ” follicular lymphoma in these cases. As previously described by Cong and colleagues,18 cases of in situ
follicular lymphoma manifested at least focal bcl-2 overexpression in germinal centers, an identifiable t(14;18) in 43%
of evaluable cases, a much older age range (23-76 years),
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less of a male predominance (only 11 of 25 cases were
males), and the frequent identification of frank follicular
lymphoma either synchronously (5/18 evaluable cases) or
with follow-up (3/18 cases).
The specific cause of the lymphoid hyperplasia in our 6
cases remains uncertain, although the uniformly negative in
situ hybridization results essentially rule out EBV infection.
Infection by other viruses remains a distinct possibility,
although we do not have additional information about
specific viruses. Note that 1 previous report described a
clonal IgH rearrangement in a histologically reactive lymph
node in a 3.5-year-old patient with active human herpesvirus
6 infection,19 although immunophenotyping indicated polyclonality among the B cells, in contrast with our cases. It
seems likely that the emergence of the prominent clonal
populations in our cases reflects an extreme exaggeration of
the normal oligoclonal germinal center response to
© American Society for Clinical Pathology
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antigen.13,20 Among other examples, such skewing has been
described previously in surface light chain expression among
antibodies to gp120 and p24 in HIV-positive patients.21
Because prominent clonal B-cell populations may be
identified by flow cytometry in settings of reactive follicular hyperplasia, the flow cytometric identification of a
clonal follicle center B-cell population should not form the
sole basis for a new diagnosis of follicular lymphoma
when the follicle center B cells show normal levels of
expression of CD10, CD19, CD20, and CD45, particularly
in young patients or in patients with chronic immune
dysregulation such as HIV. In such settings, correlation
with the histologic features, as well as flow cytometric22 or
immunohistochemical evaluation of bcl-2 expression,
should be considered essential to make the final diagnosis.
In our experience, abnormally increased or decreased
levels of CD10, CD19, and/or CD20 in the setting of a
clonal follicle center B-cell population greatly increase the
likelihood of an underlying follicular lymphoma. This
impression is consistent with previously published reports
suggesting that abnormalities in CD10, CD19, and/or
CD20 expression are relatively common in follicular
lymphoma. 23,24 However, bcl-2 protein evaluation still
could be performed in such cases.
In FNA samples of lymphoid lesions in patients without
a previous diagnosis of lymphoma and in which a concurrent core needle biopsy specimen is not obtained for histologic evaluation, one needs to be especially careful about
making a new diagnosis of follicular lymphoma when a
clonal follicle center B-cell population with normal levels of
CD10, CD19, CD20, and CD45 is identified. Flow cytometric evaluation of bcl-2 expression might be particularly
helpful in such cases. In addition, it recently has been
reported that HLA-DO is expressed preferentially in follicular lymphoma and not in reactive follicular hyperplasia,25
so evaluation of this antigen might prove useful now that
there are commercially available antibodies.
The relationship, if any, of the biopsy specimens in the 4
youngest patients to previously reported cases of “pediatric
follicular lymphoma” is uncertain. A review of the photomicrographs in previous publications on this entity26-30 shows
apparent nodal effacement with cytologic features ranging
from grades 1 to 3, while none of the tissue samples in our
series were effaced. However, in all of the previous reports of
pediatric follicular lymphoma, at least a subset of the
lymphomas were negative for bcl-2 expression and the
t(14;18), and many of the patients entered a subsequent clinical remission, even without systemic chemotherapy. Hence,
our findings raise the possibility that some clonal B-cell proliferations diagnosed as pediatric follicular lymphoma might
have represented an aberrant immune response as opposed to a
neoplastic process. Therefore, we believe that cases similar to
the cases in our series should be treated conservatively, with
the initial clinical management being close follow-up.
From the 1Department of Laboratory Medicine, University of
Washington, Seattle; and 2Department of Pathology, Oregon
Health Sciences University, Portland.
Address reprint requests to Dr Kussick: Box 357110,
University of Washington Medical Center, 1959 NE Pacific St,
Seattle, WA 98195.
Acknowledgments: We thank the technical staff of the
University of Washington Hematopathology Laboratory for efforts
in the generation of the flow cytometric data and the University of
Washington Immunocytochemistry Laboratory for efforts in the
generation of the immunohistochemical data. We thank Dan
Sabath, MD, PhD, and the University of Washington Molecular
Hematology Laboratory for performing the PCR assays for B-cell
clonality and the t(14;18).
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