Download Neogenesis of Lymphoid Structures and

Published OnlineFirst July 31, 2012; DOI: 10.1158/0008-5472.CAN-12-1377
Cancer
Research
Microenvironment and Immunology
Neogenesis of Lymphoid Structures and Antibody
Responses Occur in Human Melanoma Metastases
ate2,
Arcadi Cipponi1, Marjorie Mercier1, Teofila Seremet1, Jean-François Baurain5, Ivan The
Joost van den Oord6, Marguerite Stas7, Thierry Boon3, Pierre G. Coulie1,4, and Nicolas van Baren1,5,3
Abstract
Lymphoid neogenesis, or the development of lymphoid structures in nonlymphoid organs, is frequently
observed in chronically inflamed tissues, during the course of autoimmune, infectious, and chronic graft rejection
diseases, in which a sustained lymphocyte activation occurs in the presence of persistent antigenic stimuli. The
presence of such ectopic lymphoid structures has also been reported in primary lung, breast, and germline
cancers, but not yet in melanoma. In this study, we observed ectopic lymphoid structures, defined as lymphoid
follicles comprising clusters of B lymphocytes and follicular dendritic cells (DC), associated with high endothelial
venules (HEV) and clusters of T cells and mature DCs, in 7 of 29 cutaneous metastases from melanoma patients.
Some follicles contained germinal centers. In contrast to metastatic lesions, primary melanomas did not host
follicles, but many contained HEVs, suggesting an incomplete lymphoid neogenesis. Analysis of the repertoire of
rearranged immunoglobulin genes in the B cells of microdissected follicles revealed clonal amplification, somatic
mutation and isotype switching, indicating a local antigen-driven B-cell response. Surprisingly, IgA responses
were observed despite the nonmucosal location of the follicles. Taken together, our findings show the existence of
lymphoid neogenesis in melanoma and suggest that the presence of functional ectopic lymphoid structures in
direct contact with the tumor makes the local development of antimelanoma B- and T-cell responses possible.
Cancer Res; 72(16); 3997–4007. 2012 AACR.
Introduction
Many if not all human tumors express antigens that can be
recognized by autologous T lymphocytes (1). Cancer patients
have circulating antitumoral antibodies and T lymphocytes
(2, 3). Tumors are often infiltrated by inflammatory cells,
principally lymphocytes and macrophages. Many studies support the notion that this inflammation accompanies an
immune reaction mounted by the host against its tumor. It
has been shown that tumor-infiltrating B and T lymphocytes
can be directed at tumor antigens (4–7). In several types of
tumors such as melanoma and ovarian, colorectal and breast
carcinomas, the presence of tumor-infiltrating T cells has
catholique de Louvain;
Authors' Affiliations: 1de Duve Institute, Universite
2
Service d'Anatomie Pathologique, Cliniques Universitaires Saint-Luc;
3
Ludwig Institute for Cancer Research Ltd, Brussels Branch, 4WELBIO,
catholique de Louvain, Brussels; 5Medical
de Duve Institute, Universite
Oncology Department, Centre du Cancer des Cliniques Universitaires
catholique de Louvain, Bruxelles; 6Laboratory for
Saint-Luc, Universite
Translational Cell & Tissue Research, Department of Imaging and Pathology, and 7Department of Surgical Oncology, University Hospitals, Katholieke Universiteit Leuven, Leuven, Belgium
Note: Supplementary data for this article are available at Cancer Research
Online (http://cancerres.aacrjournals.org/).
Corresponding Author: Nicolas van Baren, Ludwig Institute for Cancer
Research Ltd, Brussels Branch, Avenue Hippocrate 74, B1.74.03, B-1200
Brussels, Belgium. Phone: 32-2-764-75-08; Fax: 32-2-764-75-90; E-mail:
[email protected]
doi: 10.1158/0008-5472.CAN-12-1377
2012 American Association for Cancer Research.
been associated with a more favorable clinical evolution
(8–11). Altogether these observations indicate that cancer
patients can mount spontaneous adaptive immune responses
against their tumor. These responses have been extensively
studied mainly through the identification of their target
antigens. However, little is known about the mechanisms that
trigger off these responses, at which stage of the disease and
where they originate, and how they are maintained. It is
commonly thought that these events take place in the lymph
nodes that drain tumor sites.
The secondary lymphoid organs, which include the lymph
nodes, the spleen, and mucosal associated lymphoid tissues,
are the major sites of lymphocyte activation during adaptive
immune responses (12). In the absence of antigenic stimulation, resting B lymphocytes aggregate around a network of
follicular dendritic cells (FDC), forming clearly recognizable
structures called follicles. Outside of these follicles, the T-cell
zones contain mainly resting T cells, interdigitating dendritic
cells (DC), and fibroblastic reticular cells. To reach the lymph
nodes, resting lymphocytes circulating in the blood adhere to
specialized blood vessels located near the B- and T-cell zones,
the high endothelial venules (HEV). The specificity of this
binding is mainly mediated by L-selectin (CD62L) expressed
on naive and central memory lymphocytes, and their receptors,
the peripheral node addressins (PNAd), which are sulfated
glycoproteins expressed selectively on the endoluminal side of
HEV. Migrating B cells are attracted toward the follicles by the
CXCL13 chemokine, secreted mainly by FDC, whereas T lymphocyte migration toward the T-cell zones is mainly controlled
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Cipponi et al.
by CCL19 and CCL21, produced principally by fibroblastic
reticular cells.
Following antigenic stimulation, B cells proliferate vigorously in germinal centers within the follicles. Activated helper
T cells migrate within the follicle, and interaction with these
follicular helper T cells induces expression of activationinduced cytidine deaminase (AID) in the germinal center B
cells (13). This enzyme controls somatic hypermutation in the
variable region of immunoglobulin (Ig) genes, which introduces additional diversity in the antibody repertoire (14, 15).
Competitive interaction with FDC, which present the antigen
in limited amount, results in the clonal selection of the B cells
with the highest Ig affinity. AID also directs isotype switching,
resulting in the preferential production of either IgG, IgA, or
IgE (14, 15).
In most cases of infectious aggression, the coordinated
action of innate and adaptive immunity results in the elimination of the pathogen and restoration of tissue integrity.
Failure to do so leads to chronic inflammation, characterized
by lymphomonocytic infiltration and attempts to confine the
infection by encapsulation or granuloma formation. In addition, ectopic lymphoid structures, also called tertiary lymphoid
structures or organs, develop within the inflamed area, a
phenomenon called lymphoid neogenesis. These structures
resemble the B- and T-cell areas of lymph nodes, in that they
contain follicles comprising B cells and FDC, and distinct T-cell
areas comprising mature DC, T-cell clusters, and neighboring
HEV-like blood vessels. This phenomenon is thought to
improve adaptive responses against persisting antigens, by
bringing the whole immune response machinery to the site of
the aggression (16, 17). Ectopic lymphoid structures are found
in many chronic infectious diseases such as Helicobacterassociated gastritis, chronic HCV hepatitis, and Lyme arthritis.
Lymphoid neogenesis is also frequently observed in noninfectious chronic inflammatory diseases, including many autoimmune diseases such as thyroiditis, rheumatoid arthritis, and
multiple sclerosis, as well as in chronic allograft rejection (16).
It has been shown that these structures can host B- and T-cell
responses directed at antigens expressed in the diseased tissue
(18–20).
The molecular mechanisms that lead to lymphoid neogenesis are only partially understood. They seem to be similar to
those that lead to the formation of new secondary lymphoid
organs during fetal development. This development is orchestrated by cells of hematopoietic origin, the lymphoid tissue
inducer cells (LTi), which drive endothelial cells to express
PNAd and become HEV, and stromal cells to differentiate into
FDC and fibroblastic reticular cells. It is not known whether
LTi-like cells contribute to the formation of ectopic lymphoid
structures. An alternative mechanism has been proposed, by
which activated lymphocytes substitute for LTi (16, 21).
Malignant tumors resemble chronic infections, autoimmune diseases and chronic allograft rejection in that they host
a persistent immune infiltration that fails to clear the antigenic
insult. Consistent with this, the presence of ectopic lymphoid
structures has been reported in several neoplastic diseases.
Non–small cell lung cancers were found to host such structures, combining B-cell follicles, HEV associated with enriched
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Cancer Res; 72(16) August 15, 2012
naive T cells, and clusters of mature DC and T cells whose
density was associated with a better prognosis (22, 23). B-cell
follicles containing germinal centers were observed in primary
medullary and ductal breast cancer. In these tumors, molecular
characterization of Ig gene rearrangements showed that clonal
proliferation, somatic Ig mutation, and clonal selection all
occurred (24–26). The analysis of primary germ cell cancers
has revealed the presence of lymphoid follicles with germinal
centers, in which ongoing antigen-driven antibody responses
occurred (27). Finally, ectopic lymphoid structures have been
detected in colorectal cancer, with their presence being correlated to a better prognosis (28). In all these tumors, lymphoid
neogenesis seems to be associated with an inflammatory
infiltrate.
In this work, we report the presence of ectopic lymphoid
structures in metastatic lesions of melanoma, a tumor type
that is frequently considered as particularly immunogenic, and
characterize the B-cell responses that take place in these
structures.
Materials and Methods
Patients and tissue samples
Tumors and healthy tissues were obtained as surgical discard samples, or as research-aimed surgery or biopsy after
informed consent, under approval of the Commission d'Ethique Biomedicale Hospitalo-Facultaire, Cliniques universitaires
Saint-Luc, Brussels, Belgium. Fresh tissue samples were snap
frozen on dry ice or in isopentane shortly after surgical
resection, embedded in Tissue-Tek OCT Compound (Sakura)
and stored at 80 C.
Immunohistochemistry
Seven mm-thick cryosections were obtained from frozen
OCT-embedded tissue samples, air dried, stored at 80 C
until use, then thawed and fixed in 4% paraformaldehyde for
5 minutes. Endogenous peroxidase activity was blocked with
Peroxidase Blocking Reagent (Dako Cytomation). For PNAd
immunostaining, sections were treated with Biotin Blocking
System (Dako) to neutralize endogenous biotins, incubated for
30 minutes with the biotinylated MECA-79 antibody, washed,
and incubated for 30 minutes with streptavidin coupled to
horseradish peroxidase (Vector Laboratories). For all other
antigens, sections were incubated for 30 minutes with unlabeled primary antibody (see list in Supplementary Table S1),
washed, and incubated for 30 minutes with a secondary
polyclonal goat anti-mouse antibody coupled to horseradish
peroxidase. After washing, bound antibodies were detected
with 3-amino-9-ethylcarbazole and sections were counterstained with hematoxylin. All reactions were carried out at
room temperature. A reactive lymph node and a reactive tonsil
from noncancerous persons were used as positive controls for
all antigens except Melan-A, for which a lymph node melanoma metastasis with confirmed expression of the Melan-A gene
by reverse transcriptase PCR (RT-PCR) was used. Negative
controls included omission of primary antibody on the control
tissue sections. Stained slides were digitized by automated
whole-slide image capture, using a Mirax Midi scanner (Carl
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Published OnlineFirst July 31, 2012; DOI: 10.1158/0008-5472.CAN-12-1377
Melanoma Tumors Host Functional Ectopic Lymphoid Structures
Zeiss MicroImaging), equipped with a Zeiss Plan-Apochromat
20 NA 0.80 objective lens and a Hitachi HV-F22 acquisition
camera, providing an object pixel size of 0.23 mm. Image
acquisition was controlled with the Mirax Scan software
(Zeiss). Image files were compressed with 0.80 jpeg, and
analyzed with the Mirax Viewer software (Zeiss).
Laser capture microdissection
Additional cryosections (9–16 mm thick) were cut from the
remaining frozen tissues and were mounted onto ultraviolettreated membrane-coated slides (PALM Microlaser Technologies). Samples were fixed in 75% ethanol for 1 minute at
20 C, stained for 30 seconds with 1% cresyl violet in ethanol,
and dehydrated in consecutive washes of 75% and 95% ethanol
for 30 seconds each, and for 2 minutes in 100% ethanol. Slides
were allowed to air dry for 5 minutes. Follicular and extrafollicular areas were identified and selected based on their
morphology and on CD20 immunostaining of an adjacent
tissue section. The number of B lymphocytes in each isolated
follicle was estimated by manual counting of CD20þ cells in the
adjacent section. Selected groups of cells were microdissected
using a PALM MicroBeam laser capture workstation (Carl
Zeiss). Isolated fragments were catapulted on the cap of
AdhesiveCap 500 Opaque tubes (Carl Zeiss) and were snap
frozen in dry ice and stored at 80 C until further processing.
RNA extraction from tissue samples, cDNA synthesis,
and qPCR
RNA samples were prepared from frozen tissue sections or
tissue fragments using the guanidinium isothiocyanate/
cesium chloride procedure (29). RNA quality was verified with
an Agilent Bioanalyzer. Total RNA was treated with TURBO
DNAse (Turbo DNA-freeTM Kit; Ambion). The first-strand
cDNA was produced from 2 mg of total RNA with the M-MLV
reverse transcriptase (Invitrogen) and a random hexamer
primer (Fermentas), according to the manufacturer's protocol.
One-fortieth of the reaction volume was used for PCR. Quantitative RT-PCR amplifications were carried out as described
(25), using the PCR Core Kit (Eurogentec), in a final volume
of 25 mL with 0.625 U of HotGoldStar DNA polymerase (Eurogentec), 300 nmol/L of each primer, 100 nmol/L of probe,
200 mmol/L of each dNTP, and 5 mmol/L of MgCl2 in an ABI
7300 Real-Time PCR system (Applied Biosystems). The following PCR conditions were used: 94 C for 10 minutes, 45 cycles of
94 C for 15 seconds, and 60 C for 1 minute. The qPCR primers
and probes (Eurogentec) are reported in Supplementary
Table S1. Samples for quantitative PCR were assayed in duplicate. Levels of expression were normalized to b-actin.
Construction of Ig heavy chain cDNA libraries
RNA was purified from microdissected tissue sections using
the TriPure isolation reagent (Roche) according to the manufacturer's protocol, except for the addition of 20 ng tRNA at
the start, and 20 mg of glycogen before the precipitation step.
Recovered RNA was resuspended in 5 or 10 mL of RNase-free
water and stored at 80 C. The mRNA encoding Ig heavy chain
genes was selectively retrotranscribed using Superscript II
reverse transcriptase (Invitrogen) and a set of degenerate
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primers matching the DNA region encoding the transmembrane region of the IGHD, IGHM, IGHG1, IGHG2, IGHG3, IGHG4,
IGHA1, and IGHA2 genes (see Supplementary Table S1 for
primer sequences). After RNaseH treatment (Fermentas), the
resulting cDNA was divided in 4 equal parts, one for each
isotype family (except IgE). Each part was amplified by 2
rounds of nested PCR, according to a previously reported
method (30). The first PCR used a set of degenerate forward
primers matching the leader (L) sequence of the IgVH family
genes, and a reverse primer matching the respective Ig heavy
chain constant gene. Amplification was carried out with Taq
DNA polymerase (Takara) in 1x PCR Buffer n 1 from the
Expand Long Template PCR System Kit (Roche), with 5 mL of
cDNA in a 50 mL volume, and started with 4 minutes at 94 C,
followed by 30 cycles of 45 seconds at 95 C, 45 seconds at 50 C,
and 45 sec þ 2 sec/cycle at 72 C, followed by 10 minutes at
72 C. The second PCR used a set of degenerate forward primers
matching the Framework 1 region of the IgVH family genes and
a more proximal reverse primer matching the respective Ig
heavy chain constant gene. Amplification was carried out with
5 mL of a 1/100 dilution of the product of the first PCR in a 50-mL
volume, using the same enzyme and buffer, and started with 4
minutes at 94 C, followed by 30 cycles of 45 seconds at 94 C, 30
seconds at 55 C, and 45 seconds þ 2 sec/cycle at 72 C, followed
by 10 minutes at 72 C. The PCR products were separated by
electrophoresis on an agarose gel, and a gel slice containing
DNA in the desired range was excised. The DNA was purified
with the GeneJet Gel Extraction Kit (Fermentas) and cloned
into TOP10 bacteria using the TOPO TA Cloning Kit (Invitrogen). Sequencing of individual DNA clones was carried out
using the Big Dye Terminator Cycle Sequencing Kit (Applied
Biosystems). The sequences were compared with published
germline Ig genes using the international ImMunoGeneTics
information system (IMGT) database (website http://www.
imgt.org; ref. 31). The VH, DH, and JH segments in each
sequenced clone were assumed to be derived from the most
homologous corresponding germline genes. Comparison
between cloned and germline sequences allowed to identify
junctional diversities and somatic mutations. In addition, as
the PCR product included the 50 end of the Ig constant region,
the Ig isotype of each clone could be determined, including
within the IgG and IgA subfamilies.
Results
Sample collection and processing
A series of 29 tumor samples was obtained by surgical
excision of skin metastases from melanoma patients. Samples
were selected prospectively or retrospectively on the basis of
tissue integrity and RNA quality. The cutaneous or subcutaneous origin of the tumors was distinguished from possible
superficial lymph node metastases by anatomical location,
presence of skin structures, and absence of residual lymph
node tissue and capsule on histologic examination. Lymph
nodes and a tonsil from noncancerous patients were used as
positive controls. All the tissue samples were frozen shortly
after resection. Sequential cryosections were used either for
immunohistochemistry or for gene expression testing by RT-
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PCR or quantitative RT-PCR (qRT-PCR). Additional cryosections were obtained for laser capture microdissection. The
main characteristics of the patients and tumor samples are
reported in Supplementary Table S2.
Cutaneous melanoma metastases contain fully
developed ectopic lymphoid structures
We observed on conventionally stained sections of some
cutaneous melanoma metastases structures resembling lymphoid follicles. To confirm this, we used immunohistochemistry to study the type and distribution of immune cells in skin
metastases from melanoma patients. Using an anti-CD20
antibody specific for B lymphocytes, we detected B-cell aggregates in 14 of the 29 tumor samples. These aggregates often
formed dense structures resembling lymph node follicles.
Plasma cells stained for CD138 were distributed more diffusely
and irregularly outside the B-cell aggregates. We then searched
for the presence of FDCs, which in lymph nodes form an
interdigitating network in the center of B-cell follicles. In 10
of the 29 tumor samples, we observed that dense B-cell clusters
were associated with more centrally located cells expressing
CD21, a complement receptor expressed selectively by FDC.
These cells were also stained by anti-VCAM1 and anti-FDCag
antibodies, 2 other FDC markers (data not shown). We concluded that these structures correspond to lymphoid follicles.
An illustrative example of follicles in a melanoma metastasis is
shown in Fig. 1. Another example can be viewed in Supplementary Fig. S1 and control stainings in Supplementary Fig. S2.
To determine whether some follicles contained germinal
centers, we stained adjacent cryosections against AID, a protein that is specifically expressed in the nuclei of germinal
center B cells. Several follicles were found to host a small group
of cells with nuclear expression of AID (see Fig. 1). The largest
of these cell clusters was also stained for nuclear Ki-67,
indicating intense B-cell proliferation, as is frequently observed
in germinal centers from secondary lymphoid organs (data not
shown). Germinal centers were only observed in follicles that
were in close contact with melanoma cells.
We next looked whether HEV-like blood vessels were present in our tumor samples. Immunostaining with the MECA-79
antibody, which is specific for PNAds, revealed the presence of
small PNAdþ blood vessels in the neighborhood of most of the
melanoma associated follicles, but not in other parts of the
tumors. We observed that most follicles were also intimately
associated with neighboring concentrations of CD3þ T cells,
including CD8þ T cells often clustered around DC-LAMPþ
mature DCs and surrounded by PNAdþ HEVs, as occurs in Tcell zones in secondary lymphoid organs (see Fig. 1 and
Supplementary Fig. S1 and S2). DC/T-cell clusters were also
repeatedly observed in or outside most of the tumor samples,
including those in which B-cell follicles and HEV-like blood
vessels were absent (data not shown).
Altogether, among the 29 cutaneous melanoma metastases
analyzed, 7 contained complete ectopic lymphoid structures,
combining follicles, HEV-like blood vessels, and T-cell mature
DC clusters. Four of these tumors hosted at least one follicle
containing a germinal center. Of the remaining 22 tumors, 3
contained HEV associated with B-cell clusters without evi-
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Cancer Res; 72(16) August 15, 2012
dence of FDC, 3 others had complete follicles without HEV, and
the 16 remaining tumors had no evidence of lymphoid
neogenesis.
Confirmation of FDCs and germinal centers in follicles
We microdissected 7 individual follicles from cryosections
cut from 3 selected metastases with AIDþ cells and carried out
qRT-PCR amplification of the CD21L transcript, a long CD21
mRNA isoform that is selectively expressed by FDCs (32).
CD21L was detected in the follicles, but not in tumor areas
taken outside the follicles from the same cryosection (Fig. 2).
Similarly, we detected the expression of the AID gene in most
microdissected follicles, but not in the extrafollicular areas
(Fig. 2). Altogether, these data confirmed that cutaneous
melanoma metastases contain B-cell follicles and suggested
that some of them are reactive and host B-cell responses.
Lymphoid neogenesis is irregularly distributed among
different types of melanoma lesions
We assessed whether ectopic lymphoid structures were also
present in primary melanomas and in other types of metastases. We used qRT-PCR to quantify the CD21L transcript as a
molecular marker for the presence of FDCs, and thus also of
lymphoid follicles, in a large series of melanoma samples.
Lymph node metastases were not included, because they often
contain normal lymphoid tissue and thus would give false
positive results. Consistent with our previous observations,
CD21L transcripts were detected in 9 of 63 cutaneous metastases. In contrast, none of 25 primary cutaneous melanomas
were positive, neither was normal skin. In visceral metastases,
FDC were detected in some lung, muscle, and gut metastases,
but intriguingly in neither of 12 liver and 12 brain metastases
(Fig. 3).
Molecular characterization of the immunoglobulin gene
repertoire expressed in melanoma-associated ectopic
lymphoid structures
To assess whether active B-cell responses are taking place in
B-cell follicles present in melanoma metastases, we analyzed
the diversity of rearranged heavy chain immunoglobulin genes
expressed in 4 different follicles isolated from 3 selected
tumors, T3 (2 follicles), T4, and T9 (1 follicle each). Four
consecutive cryosections were obtained from each tumor, and
whole follicular areas were isolated from each section by laser
capture microdissection. We chose to isolate whole follicles
instead of germinal centers, because the latter were often
undetectable or limited to very few AIDþ cells, difficult to
identify in tissue sections for microdissection. The selected
follicles all contained AIDþ cells, as confirmed by immunostaining or qRT-PCR of close cryosections (data not shown). Ig
heavy chain cDNA libraries were constructed independently
from each follicle section as illustrated in Supplementary Fig.
S3. Briefly, RNA extracted from each follicle section was reverse
transcribed with a mix of primers specific for the transmembrane exons of all the constant region genes except IGHE. IgEencoding transcripts were excluded from the libraries because
they had not been detected by RT-PCR in our tumor samples,
as opposed to the other Ig isotypes (data not shown). By
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Melanoma Tumors Host Functional Ectopic Lymphoid Structures
B
CD20 (B cells)
CD138 (plasma cells)
C
CD21 (follicular dendritic cells)
D
AID (germinal center B cells)
G
DC-LAMP (mature dendritic cells)
H
PNAd (HEV-like blood vessels)
500 µm
50 µm
2 mm
A
Melan-A (melanoma cells)
F
CD8 (cytolytic T cells)
500 µm
2 mm
E
Figure 1. Immunohistologic detection of the indicated antigens in adjacent cryosections of cutaneous melanoma metastasis T3. Positively stained antigens
appear in red.
excluding secreted Ig, we restricted our analysis to naive,
memory, and germinal center B cells and largely excluded
plasma cells, as the latter can migrate from distant secondary
lymphoid organs via the blood to inflammatory sites, and thus
might have infiltrated the tumors independently of local B-cell
responses (33). The resulting cDNA was amplified separately
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for each IgD, IgM, IgG, and IgA isotype by nested PCR, based on
the approach described by Wang and colleagues (30). The
amplified products, which included the 50 extremity of the first
exon of the constant region genes (CH1) to allow isotype
identification, were cloned and sequenced. Thus, 64 independent VDJCH1 cDNA libraries were obtained, from each of 4
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Cipponi et al.
200
393
CD21L
% Expression relative to lymph node follicles
150
Figure 2. Expression of CD21L (top)
and AID (bottom) by qRT-PCR in 7
microdissected follicles (F1, F3
to F8) isolated from cutaneous
melanoma metastases T3, T4,
and T9. Extrafollicular areas
microdissected in the same tumor
sections were used as negative
controls (EF) and follicles
containing germinal centers
microdissected from a normal
reactive lymph node as positive
controls (gray boxes). Results are
numbers of cDNA copies
normalized to b-actin and with the
mean level of expression in normal
follicles set to 100. Error bars
indicate one SD. n, the
number of biologic replicates
tested. Replicates are the same
follicle in adjacent tumor sections
or different follicles isolated from
the same lymph node section.
No PCR product was amplified
from extrafollicular areas
microdissected from the same
tumor sections, indicating less
than 5 cDNA copies for CD21L
and less than 2 for AID.
100
50
0
0
0
0
100
80
AID
60
40
20
n=2
n=2
n=2
n=2
n=2
n=2
T9/F4
T9/F5
T9/F6
T9/F7
T9/F8
T9/EF
Follicles isolated from tumors
different follicles, 4 adjacent sections, and for each of 4 different
isotypes.
In total, we obtained 672 interpretable VDJCH1 sequences
corresponding to 304 individual B-cell clones defined by their
unique VDJ rearrangement, junctional diversity, and common
somatic mutations. None of these 304 clones was present in
more than one follicle. We first looked whether repeated
clones, that is, represented by more than one B cell in an
individual follicle, were present. These clones are likely to have
resulted from local clonal proliferation. A clone was considered
repeated if its VDJ sequence was amplified independently from
2 nonadjacent sections from the same follicle. This conservative definition excludes occurrences of single B cells divided
among 2 adjacent sections. It clearly underestimates the real
number of repeated clones in our samples. In total, 17 distinct
repeated clones were identified among the 4 follicles (Table 1).
We next looked at local occurrence of class switch recombi-
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Follicles from
lymph node
n=2
T4/EF
1
n=5
T4/F3
0
n=2
T3/EF
0
n=3
T3/F1
0
nation. We identified 8 clones in which a unique VDJ sequence
was coupled to either of 2 different constant regions. We
observed IgD to IgG1, IgG1 to IgG2, IgG1 to IgA1, and IgA1
to IgA2 transitions (Table 1). Clones with more than 2 different
isotypes were not detected. Finally, we also searched for clonal
variants within individual B-cell clones. Two or more VDJCH1
sequences were considered to be clonal variants derived from a
single B-cell precursor if they shared the same VDJ rearrangement and junctional diversity, and if they differed by more than
5 nucleotides. The latter definition was deduced from the error
rate of retrotranscription and PCR amplification in our experimental setting, according to the law of Poisson. This rate was
0.003, measured as the number of mismatches observed in a
large number of CH1 sequences. In total, 17 different B-cell
clones had evidence of clonal variation (Table 1). Altogether, 27
B-cell clones (8.9%) showed evidence of affinity maturation,
including 13 that had more than one of the 3 cardinal features
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Melanoma Tumors Host Functional Ectopic Lymphoid Structures
% Expression of CD21L
Figure 3. Expression of CD21L
measured by qRT-PCR in primary
tumors and different types of
metastases from melanoma
patients. The y-values represent
numbers of CD21L cDNA copies,
normalized to b-actin and with the
level of expression in normal lymph
node set to 100.
relative to normal lymph node
40
30
20
10
Muscle
metastasis
Lung
Gut
Skin
0
Primary
melanomas
n = 25
Skin
n = 63
Lung
n = 14
Liver
n = 12
Brain
n = 12
Gut
n=7
Other
n = 18
Normal
tissues
n=3
Melanoma metastases (n = 126)
of Ag-driven B-cell responses. The aligned VDJCH1 sequences
of B-cell clone B2 are shown in Fig. 4 as an illustrative example.
Among all the Ig sequences derived from the 27 B-cell clones
undergoing affinity maturation, about 10% seemed to be
nonfunctional because of frame shifts resulting either from
out-of-frame VDJ rearrangements or from base insertion or
deletion (data not shown). These features are characteristic of
ongoing germinal center B-cell responses, during which a
number of irrelevant Ig gene rearrangements occur before
being selected negatively (26).
We concluded that part of the B cells present in melanomaassociated ectopic lymphoid structures have undergone clonal
expansion and selection, somatic hypermutation, class switch
recombination, and abortive Ig rearrangements, displaying the
characteristics of an Ag-driven B-cell response that takes place
in secondary follicles. These Ig responses are oriented toward
the IgM, IgG1, IgG2, IgA1, and IgA2 isotypes.
Primary melanomas contain incomplete ectopic
lymphoid structures
Lymphoid follicles are usually not observed in conventionally stained histologic sections of primary melanomas, with the
exception of desmoplastic/neurotropic melanomas (34), suggesting that these tumors do not contain ectopic lymphoid
structures. Consistent with this, we failed to detect CD21L
expression in 25 primary melanomas. However, the presence of
tumor-associated HEV-like blood vessels expressing PNAd has
recently been reported in 11 of 18 primary melanoma samples
(35). We addressed this paradox by testing 10 primary melanomas with known lymphocyte infiltration for the presence of
lymphoid follicles, HEV, and clusters of mature DC and T cells
by immunohistochemistry. Nine tumors contained CD20þ B
cells forming a loose infiltrate rather than a dense follicle-like
structure. No CD21þ FDC were observed (Table 2 and Supplementary Fig. S4). All tumors contained a CD8þ T-cell
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infiltrate associated with mature DC. Six of 10 tumors contained PNAdþ blood vessels, closely associated with the T- and
B-cell tumor infiltrate. This suggested that primary melanomas
host HEV, but no FDC and no follicles, and thus that the
process that leads to full scale lymphoid neogenesis is incomplete in these tumors.
Discussion
We report here for the first time the presence of ectopic
lymphoid structures in human melanoma metastases. These
structures associate lymphoid follicles, comprising mainly B
lymphocytes clustered around FDC, with HEV and with clusters of DCs and T cells. The presence of germinal centers in
some follicles and the mature phenotype of neighboring DC
associated with T cells suggest that adaptive immune
responses take place in these structures. Here we confirm this
for B-cell responses, by showing that the Ig transcript repertoire in melanoma follicles is enriched for clones that display
the main characteristics of affinity maturation. Our observations do not allow to determine whether these local immune
responses are directed at melanoma antigens, but we feel that
this is a realistic possibility. Tumor-associated ectopic lymphoid structures are presumably not connected to the specialized networks of afferent lymph and blood vessels that
carry antigens to B- and T-cell zones in lymph nodes and spleen
and are therefore more likely to sense antigens present locally,
as in the case of mucosal associated lymphoid tissue. It must be
noted that all the follicles containing germinal centers that we
observed were invaded by or were in close contact with tumor
cells.
Primary melanomas sometimes host HEV and neighboring
clusters of DC and T cells, but in contrast to metastases they
never contain lymphoid follicles, which seem to be replaced by
less organized accumulations of B lymphocytes. The absence
of FDC is the likely explanation for this absence of follicles. The
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Cipponi et al.
Table 1. List of the 27 B cell clones isolated from 4 tumor-associated follicles and involved in an Ag-driven
Ig response
B cell clone no.
Rearranged VDJ
Tumor/follicle/sections
Isotype(s)
Clonal variants
B1
B2
B3
B4
B5
B6
B7
B8
B9
B10
B11
B12
B13
B14
B15
B16
B17
B18
B19
B20
B21
B22
B23
B24
B25
B26
B27
V3-9 D3-16 J4
V3-11 D3-22 J4
V3-23 D2-21 J6
V3-30 D2-15 J3
V3-48 D3-3 J6
V5-51 D6-13 J4
V3-74 D5-24 J4
V5-51 D3-3 J4
V5-51 D6-6 J5
V1-3 D6-6 J2
V3-11 D6-19 J6
V3-48 D5-24 J4
V5-51 D3-3 J4
V3-9 D1-26 J3
V3-15 D2-8 J6
V3-15 D4-17 J5
V3-23 D5-24 J4
V3-23 D3-9 J6
V3-33 D3-3 J4
V3-66 D4-17 J6
V3-66 D7-27 J6
V4-59 D5-5 J3
V5-51 D1-7 J4
V5-51 D5-5 J6
V5.a D3-22 J2
V3-30 D? J3
V3-66 D6-19 J6
T3/F1/S2,3,4
T3/F1/S1,2,3,4
T3/F1/S1,2
T3/F1/S2
T3/F1/S1,2,4
T3/F1/S1,2,4
T3/F2/S1,2
T3/F2/S1
T3/F2/S2
T4/F3/S1,2,3
T4/F3/S1,2,4
T4/F3/S1,2
T4/F3/S3
T9/F4/S2,3,4
T9/F4/S3,4
T9/F4/S1,2,3
T9/F4/S2,4
T9/F4/S1,2,3,4
T9/F4/S2,4
T9/F4/S1,4
T9/F4/S2,3,4
T9/F4/S2,3,4
T9/F4/S1,3
T9/F4/S1,2,4
T9/F4/S1,3,4
T9/F4/S1,2
T9/F4/S2,3,4
IgA1
IgG1, IgG2
IgA1
IgG1, IgG2
IgG1
IgM
IgG1, IgG2
IgM
IgA1
IgA1
IgG1
IgD, IgG1
IgA1, IgA2
IgG1
IgG1, IgA1
IgA1
IgA1, IgA2
IgA2
IgA2
IgG1
IgA1, IgA2
IgG1
IgA1
IgG1
IgA1
IgM
IgG1
No
Yes
Yes
No
No
No
No
Yes
Yes
No
Yes
Yes
Yes
No
No
Yes
Yes
Yes
Yes
Yes
Yes
Yes
No
Yes
No
Yes
Yes
NOTE: These clones display one or more of the following characteristics: Clonal amplification illustrated by repeated occurrences of
clonal B cells in the same follicle, isotype switch, and clonal variation. T3, T4, and T9 identify the tumor samples, F1 to F4 the follicles,
and S1 to 4 the tissue sections from which the sequences were isolated. Ig sequences from clone B2 are shown in Fig. 4 as a
representative example.
accumulated B cells could be naive or memory B cells that
migrated from the blood to the tumor via the neighboring
HEV, but even though some of them might have encountered
their cognate antigen, an appropriate B-cell response ought to
be impaired because of the absence of FDC. It will be important to understand why FDC are missing in primary
melanomas.
The antibody responses that take place in melanoma follicles are switched toward the IgG1, IgG2, IgA1, and IgA2
isotypes. To the best of our knowledge, this is the first time
that IgA responses are documented in ectopic lymphoid
structures, presumably because they were not specifically
searched for in previous studies, many of which point to IgG
as the major isotype produced. Could antimelanoma IgA
influence tumor development? As opposed to IgG, IgA do not
activate the complement and do not mediate antibody-dependent cellular cytotoxicity. It seems therefore unlikely that such
IgA responses could have an antitumoral effect. Why do
antibody responses in melanoma metastases switch toward
4004
Cancer Res; 72(16) August 15, 2012
IgA? Isotype switching is mainly driven by specific cytokines,
usually produced by follicular helper T cells (Tfh) in germinal
centers. One such cytokine, TGF-b, has been identified as the
key inducer of T-cell–dependent IgA class switching (36–38).
The detection of IgA responses in all of the 4 follicles that we
have analyzed suggests that TGF-b modulates the microenvironment of melanoma metastases. This cytokine plays a dual
role in cancer progression. In early stages, it acts negatively by
inhibiting cellular proliferation, whereas in more advanced
disease it stimulates cellular dedifferentiation and invasiveness
and acts as a potent inhibitor of inflammation. Its capacity to
inhibit T-cell activation makes it a frequently cited candidate
to explain the resistance that tumors display against immunemediated destruction (39).
Because lymphoid neogenesis develops as a consequence of
ongoing adaptive responses in contact with persistent antigens, its presence in melanomas indicates that these tumors
host an active immune reaction, and that melanoma-infiltrating lymphocytes are not completely inhibited by the tumor
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Melanoma Tumors Host Functional Ectopic Lymphoid Structures
Figure 4. Ig sequences derived
from B cell clone B2 isolated from
lymphoid follicle F1 present in
melanoma metastasis T3. DNA
sequences of the 12 bacterial
colonies (S01 to S12) with a
V3-11 01, D3-22 01, J4 02 gene
rearrangement are aligned to
the corresponding germline
immunoglobulin VDJ and IGHG1
genes (bottom, in black and bold
characters). Matched and
mismatched nucleotides are
displayed in gray and black,
respectively. All these sequences
originate from a single B cell
precursor, as they share the same
junctional diversities as well as many
somatic mutations. Sporadic
mismatches are assumed to result
from reverse transcription or PCR
errors. Underlined mismatches
represent somatic mutation variants
that identify B cell subclones. All
sequences have the IgG1 isotype,
except sequence S02, which has the
IgG2 isotype (IgG2 nucleotides
appear in small characters).
Table 2. Immunohistochemical analysis of primary melanomas with known lymphocyte infiltration
Tumor no.
HEV (PNAdþ)
B cells (CD20þ)
T30
T31
T32
T33
T34
T35
T36
T37
T38
T39
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
FDC (CD21þ)
NT
NT
CTL (CD8þ)
Mature DC (DC- LAMPþ)
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
þ
NOTE: The presence (þ) or absence (–) of the indicated cells is shown. An illustrative example (T33) is shown in Supplementary Fig. S4.
Abbreviation: NT, not tested.
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Cipponi et al.
environment. Does this presence counteract or favor tumor
survival and progression? Ectopic lymphoid structures may
play an active role in the immune control of tumors, by
allowing naive or memory lymphocytes to become activated
in situ. In lung and colorectal carcinomas, the presence of these
structures seems to be associated with a better prognosis
(22, 28). Alternatively, an opposite, protumoral effect can not
be excluded. Ectopic lymphoid structures might be involved in
maintaining peripheral immune tolerance, just like lymph
nodes are. In a mouse model of tumor-induced tolerance, the
forced expression of CCL21 by melanoma cells induces LTi
recruitment and lymphoid tissue development in grafted
tumors, followed by tumor growth (40). In humans, ectopic
lymphoid structures associated with renal allografts can be
associated with donor-specific tolerance rather than rejection
(41). Experimental models have shown that humoral immunity
can promote tumor survival and growth. Several molecular
mechanisms have been proposed to explain this effect, such as
the impairment of antitumoral T-cell responses by blocking
antibodies, and the modulation of the inflammatory response
by immune complexes (42, 43). On the basis of our findings, we
can also hypothesize that locally secreted IgA might compete
with IgG for binding to tumor cells, thereby inhibiting IgGmediated cytotoxicity. The detrimental effect of such an IgA–
IgG competition has been shown recently with HIV vaccines
(44). To assess whether lymphoid neogenesis helps or impairs
tumor progression, it will be important to evaluate its prognostic impact in a large series of melanoma patients. This study
does not allow to answer this question, as it was designed to
provide a proof-of-principle that lymphoid neogenesis is present in melanoma and that it hosts ongoing adaptive responses.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
Authors' Contributions
Conception and design: A. Cipponi, J.-F. Baurain, I. Theate, P.G. Coulie, N. van
Baren
Development of methodology: A. Cipponi, M. Mercier, T. Seremet, I. Theate,
N. van Baren
Acquisition of data (provided animals, acquired and managed patients,
provided facilities, etc.): M. Mercier, T. Seremet, J.-F. Baurain, I. Theate, J. van
den Oord, M. Stas, N. van Baren
Analysis and interpretation of data (e.g., statistical analysis, biostatistics,
computational analysis): T. Seremet, J.-F. Baurain, I. Theate, J. van den Oord, T.
Boon, N. van Baren
Writing, review, and/or revision of the manuscript: A. Cipponi, T. Seremet,
J.-F. Baurain, I. Theate, J. van den Oord, M. Stas, T. Boon, P.G. Coulie, N. van Baren
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): M. Mercier, M. Stas
Study supervision: J.-F. Baurain, P.G. Coulie, N. van Baren
Acknowledgments
The authors thank Madeleine Swinarska and Francis Brasseur for expert
technical assistance and Suzanne Depelchin for editorial help.
Grant Support
This work was supported by grants from the Belgian Programme on Interuniversity Poles of Attraction initiated by the Belgian State, Prime Minister's
Office, Science Policy Programming, by grants from the Belgian Cancer Plan
(Action 29_049), the Fonds National pour la Recherche Scientifique (Belgium),
the Fondation contre le Cancer (Belgium), the Fondation Salus Sanguinis
(Belgium), and the Fonds Maisin (Belgium).
The costs of publication of this article were defrayed in part by the payment of
page charges. This article must therefore be hereby marked advertisement in
accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received April 10, 2012; revised May 18, 2012; accepted May 19, 2012;
published OnlineFirst July 31, 2012.
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Neogenesis of Lymphoid Structures and Antibody Responses
Occur in Human Melanoma Metastases
Arcadi Cipponi, Marjorie Mercier, Teofila Seremet, et al.
Cancer Res 2012;72:3997-4007. Published OnlineFirst July 31, 2012.
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