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Extranodal lymphoid microstructures in inflamed muscle and disease severity of new-onset juvenile dermatomyositis.

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Vol. 60, No. 4, April 2009, pp 1160–1172
DOI 10.1002/art.24411
© 2009, American College of Rheumatology
Extranodal Lymphoid Microstructures in Inflamed Muscle and
Disease Severity of New-Onset Juvenile Dermatomyositis
Consuelo M. López de Padilla,1 Abbe N. Vallejo,2 David Lacomis,3
Kelly McNallan,1 and Ann M. Reed1
lar dendritic cells and high endothelial venules. They
also expressed high levels of CXCL13 and lymphotoxins
known to support lymphoid organogenesis. There were
also resident naive CD45RAⴙ T cells and maternally
derived B cells and PDCs. Patients with diffuse infiltrates or lymphocytic aggregates were responsive to
standard therapy with steroids and methotrexate, but
those with follicle-like structures tended to have severe
disease that required additional agents such as intravenous Ig or rituximab.
Conclusion. These data suggest that lymphoneogenesis is a component of the early disease process in
juvenile DM. Ectopic lymphoid structures could indicate a severe course of disease; their early detection
could be a tool for disease management.
Objective. Juvenile dermatomyositis (DM) is an
autoimmune disease of childhood characterized by lesions in skin and muscle that are populated by plasmacytoid dendritic cells (PDCs) and lymphocyte infiltrates.
We undertook this study to examine the cellular composition, organization, and molecular milieu of the
cellular infiltrates in muscle in juvenile DM and to
correlate the infiltrates with clinical disease manifestations.
Methods. Since PDCs and lymphocyte foci express CCL19 and CCL21, we investigated for in situ
formation of lymphoid microstructures that could be
sites of extranodal immune activation.
Results. Analyses of muscle biopsy samples from
children with new-onset juvenile DM showed 3 categories of lesions: diffuse infiltrates, lymphocytic aggregates lacking follicle-like organization, and follicle-like
structures. The last of these exhibited elements of
classic lymphoid follicles, including networks of follicu-
Juvenile dermatomyositis (DM) is a chronic, multisystem inflammatory disease involving small vessels of
skeletal muscle, skin, gastrointestinal tract, and other
organs (1). The clinical spectrum is very variable, from
mild disease that has minimal functional impact to a
chronic, severely disabling condition (1–4). Despite new
therapies, juvenile DM remains chronically active in a
large proportion of patients.
The initiating factors in muscle inflammation are
unknown, but immune-mediated processes have profound impact on the pathophysiology of juvenile DM.
The cellular infiltrate consists of T cells, B cells, and
plasma cells (1,5,6). We have recently reported that
many of the CD4⫹ cells in the inflamed muscle of
patients with new-onset juvenile DM are CD123⫹ plasmacytoid dendritic cells (PDCs) (7). Since PDCs play a
prominent role in adaptive immunity by directly activating naive T cells, their presence in inflammatory lesions
in juvenile DM muscle is consistent with an emerging
paradigm for a role of PDCs in the pathogenesis of
chronic inflammatory disease (8–10).
Presented in part at the 71st Annual Scientific Meeting of the
American College of Rheumatology, Boston, MA, November 2007.
Dr. López de Padilla’s work was supported by the Myositis
Association (grant MYOSITIS 1). Dr. Vallejo’s work was supported by
the NIH and the Arthritis Foundation. Dr. Reed’s work was supported
by the State of Minnesota Partnership, the NIH, the Mayo Foundation, and the Arthritis Foundation.
Consuelo M. López de Padilla, MD, Kelly McNallan, BS,
Ann M. Reed, MD: Mayo Clinic College of Medicine, Rochester,
Minnesota; 2Abbe N. Vallejo, PhD: Children’s Hospital of Pittsburgh,
Pittsburgh Cancer Institute, McGowan Institute for Regenerative
Medicine, University of Pittsburgh, Pittsburgh, Pennsylvania; 3David
Lacomis, MD: University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania.
Address correspondence and reprint requests to Ann M.
Reed, MD, Division of Rheumatology, Departments of Medicine and
Pediatrics, Mayo Clinic, 200 First Street SW, Rochester, MN 55905
(e-mail:; or to Abbe N. de Vallejo, PhD,
Department of Pediatrics, Children’s Hospital of Pittsburgh Rangos
Research Center, University of Pittsburgh, Pittsburgh, PA 15213
Submitted for publication July 14, 2008; accepted in revised
form December 30, 2008.
Cellular infiltration in the muscle in new-onset
juvenile DM appears to have varying degrees of organization (11). In some patients, PDCs, CD3⫹ T cells, and
CD20⫹ B cells are scattered throughout the affected
muscle. In other patients, the cells form discrete clusters
with discernible T cell and B cell zones. We recently
reported (7) that these PDC–lymphocytic aggregates
have high levels of CCL19 and CCL21, two chemokines
that guide normal lymphocyte traffic inside of secondary
lymphoid organs (12). These observations suggest that
the inflammatory microenvironment of the muscle in
juvenile DM could permit lymphoneogenesis.
Lymphoneogenesis is a process whereby cellular
infiltrates become organized in a manner similar to that
of lymphoid follicles, such that they can even form
germinal centers (GCs) within nonlymphoid tissues
(13,14). Lymphoid-like microstructures have been reported in some autoimmune conditions (15–18). If such
structures are true extranodal lymphoid tissues, then
they would express the same molecular effectors as
normal secondary lymphoid organs and display similar
tissue architecture (12,13,19–23).
In this context, we undertook studies to examine
the cellular composition, organization, and molecular
milieu of the cellular infiltrates in muscle in juvenile
DM and correlated the infiltrates with clinical disease
manifestations. We searched for networks of follicular
dendritic cells (FDCs) and high endothelial venules
(HEVs), and because juvenile DM is characterized by
microchimerism (24), we examined whether maternallyderived cells are components of lymphoid-like structures
in the muscle.
Patients and collection of muscle biopsy samples. This
study was approved by Institutional Review Boards of the
Mayo Clinic and the University of Pittsburgh, with signed
informed consent and/or assent. Muscle biopsy samples were
obtained from 23 children (ages 2–18 years) with new-onset
disease that fulfilled the Bohan and Peter criteria for the
diagnosis of juvenile DM (25). Control nonmyositic muscle
biopsy samples and waste tonsils were also obtained.
Disease activity was assessed using established disease
activity tools including muscle enzyme evaluation, manual
muscle testing, and the Myositis Disease Activity Assessment
Tool and Myositis Damage Index for myositis clinical trials
(26). Muscle samples were obtained as part of medical diagnoses. Of the 23 samples, 12 were reported previously (7) and
11 were new. All biopsy samples were examined, and the
histologic diagnosis of juvenile DM was confirmed.
Control muscle samples (n ⫽ 11) were biopsy samples
from patients without myositis, as reported previously (7). In
addition, tonsil tissue was obtained from patients who underwent routine tonsillectomy and was used as control tissue.
Tissues were embedded in TissueTek OCT compound
(Baxter Diagnostic, McGaw Park, IL) or gum tragacanth
(Fisher Scientific, Pittsburgh, PA) and stored at ⫺80°C (7).
Frozen sections were cut at 5–8-␮m thickness and used for
hematoxylin and eosin (H&E) staining, immunofluorescence,
immunoperoxidase staining, and laser capture microdissection.
Histopathologic analysis. Serial cryostat sections from
the entire muscle tissue (usually a 1 cm ⫻ 0.5–2–cm sample)
were stained with H&E. They were then examined by lowpower field screening to evaluate the patterns of distribution
and/or organization of inflammatory cells.
Immunofluorescence. Frozen sections were stained by
either double or triple immunofluorescence techniques as
described previously (7), with fluorochrome-conjugated antibodies to a panel of dendritic cell, T cell, and B cell markers.
Staining was done sequentially by combining antibodies to
specific cell phenotypes. These included markers for T cells
(CD3, CD4, CD8, CD45RA, CD45RO), B cells (CD20,
CD23), FDCs (CD21, CNA.42), and HEVs (HECA 452). For
a nuclear counterstain, we used 4⬘,6-diamidino-2-phenylindole
(DAPI). Slides were analyzed using fluorescence microscopy
(Olympus BX51 and Olympus DP70 Microscope Digital Camera; Olympus, Melville, NY) and read independently by at
least 2 different observers without access to clinical data.
Immunohistochemistry. Expression of CXCL13 (or B
cell–attracting chemokine 1 [BCA-1]) and its receptor CXCR5
was examined by double staining according to a standard
protocol using biotinylated streptavidin conjugated to horseradish peroxidase (CTS002, CTS003; R&D Systems, Minneapolis, MN). To verify specificity of immunolabeling, primary
antibodies were omitted from the staining procedure in a set of
control sections (not shown).
Quantitative imaging. Whole cross-sections of tissues
were systematically examined at high-power magnification.
Fluorescent images were selected randomly, focused, captured
under a DAPI filter, and recaptured using other filters (fluorescein isothiocyanate autofluorescence and Texas Red) without changing focus to ensure the same plane of focus, and the
number of positively staining cells was counted using ImageJ
software (NIH Image, National Institutes of Health, Bethesda,
MD; online at: The number of positive cells per mm2 was calculated for each high-power field
(hpf), and the mean values were calculated. The mean area of
a muscle section was 3.0 mm2.
Detection of microchimerism in muscle sections by
fluorescence in situ hybridization (FISH). We investigated for
tissue chimerism on 7-␮m frozen muscle sections from 6 boys
with juvenile DM, by FISH analysis using the CEP X spectrum
orange/Y spectrum green DNA probe kit (Vysis, Downers
Grove, IL) as previously described (24). Three nonmyositic
muscle samples from male subjects were used as controls.
To further assess phenotypes of maternal XX cells, the
combination of immunoperoxidase staining and FISH was
performed on frozen muscle sections as described previously
(27). The slides were viewed using both fluorescence and
confocal laser microscopy.
Table 1. Demographic and clinical characteristics at disease onset in the children with juvenile
Age at onset, mean (range) years
Cutaneous ulcers
Nailfold changes
Raynaud’s phenomenon
Mechanic’s hands
GI involvement
Avascular necrosis
Elevated ESR
ANA positive
Myositis-specific antibodies, anti–Jo-1
Muscle strength, mean†
Muscle-associated enzymes, mean‡
Physician’s global assessment of disease
activity (0–10-cm VAS), mean
Duration of symptoms to time of
biopsy, mean (range) months
(n ⫽ 7)
(n ⫽ 11)
Lymphoid follicle–like
(n ⫽ 5)
16.5 (11–22)
5 (71.4)
0 (0.0)
1 (14.3)
0 (0.0)
0 (0.0)
0 (0.0)
2 (28.6)
0 (0.0)
1 (14.3)
0 (0.0)
0 (0.0)
0 (0.0)
7 (100)
1 (14.3)
0 (0.0)
7.3 (2–13)
8 (72.7)
1 (9.1)
11 (100)
2 (18.2)
1 (9.1)
2 (18.2)
3 (27.3)
0 (0.0)
2 (18.2)
0 (0.0)
1 (9.1)
3 (27.3)
11 (100)
2 (18.2)
0 (0.0)
5.6 (4–6)
4 (80)
3 (60)
3 (60)
3 (60)
2 (40)
1 (20)
2 (40)
3 (60)
1 (20)
1 (20)
1 (20)
1 (20)
5 (100)
1 (20)
2 (40)
5.7 (2–12)
4.8 (1–12)
1.8 (1–3)
* Except where indicated otherwise, values are the number (%) of patients. GI ⫽ gastrointestinal; ESR ⫽
erythrocyte sedimentation rate; ANA ⫽ antinuclear antibody; VAS ⫽ visual analog scale.
† On the Manual Muscle Test 8, with a highest possible total score of 80.
‡ Mean number of enzymes with abnormal levels (aldolase, creatine kinase, lactate dehydrogenase,
aspartate aminotransferase, alanine aminotransferase).
Detection of microchimerism in peripheral blood. Isolation of CD34⫹ cells. Peripheral blood samples (10 ml) from
male patients with juvenile DM were positively separated by
CD34⫹ magnetic-activated cell sorting (Miltenyi Biotec, Bergisch Gladbach, Germany). The purity of CD34⫹ blood stem
cells was ⬎96% as determined by flow cytometry.
FISH of positively selected CD34⫹ cells. Positively selected CD34⫹ blood stem cells were probed for X and Y
chromosomes as described previously (24). Only nonoverlapping cells were examined and counted for X and/or Y
chromosome–fluorescent signals. Results were expressed as
the number of cells per slide.
Laser capture microdissection and RNA extraction.
Laser capture microdissection from muscle tissue was done
using the Veritas Microdissection Instrument (Arcturus,
Mountain View, CA) as described previously (7). Laser capture microdissection was carried out only after detailed histologic analysis as described above. Sections used in the histologic studies or serial sections from the same tissue blocks were
prepared and then used to capture cellular infiltrates.
Real-time quantitative reverse transcription–polymerase
chain reaction (RT-PCR) experiments. For relative quantitative analysis, total RNA isolated was reverse-transcribed to
complementary DNA (cDNA) using the Reaction Ready kit
(SuperArray Bioscience, Frederick, MD). First-strand cDNA
was synthesized from 25 ng of total RNA. PCR was performed
in duplicate using RT2 real-time SYBR Green/ROX qPCR
Master Mix (PA-012; SuperArray Bioscience). The housekeeping gene ␤-actin was amplified to normalize expression levels
of target genes between different samples and to monitor assay
reproducibility (28). The reaction mixtures without template
cDNA were used as negative controls. Melting curve analysis
was performed to verify that there was a single PCR product
and a lack of primer dimers.
For all of the genes measured (i.e., CXCL13, lymphotoxin ␣ [LT␣], LT␤), real-time PCR assays were run with
serially diluted standards that were plotted to calculate the
exact number of transcripts. Amplification efficiencies of the
target and references were equivalent.
Statistical analysis. Quantitative data from replicate
measurements were expressed as the mean ⫾ SD; data were
analyzed using the R statistical package (
Differences between control muscle samples and the 3 histologic types of juvenile DM muscle were examined by KruskalWallis one-way analysis of variance. Pairwise between-group
comparisons (e.g., control versus each histologic type) were
examined by Mann-Whitney U test. P values less than 0.05
were considered significant.
Figure 1. Histologic patterns of cellular aggregates in muscle from patients with juvenile dermatomyositis (DM). Serial sections of muscle biopsy
samples from patients with juvenile DM and control tonsil tissue were stained with hematoxylin and eosin (H&E) and with antibodies to T cells
(CD3⫹; green) and B cells (CD20⫹; red). In one-third of patients examined, T and B cells were distributed as ring-shaped structures resembling
lymphoid follicles (A and B) with a B cell–rich center surrounded by T cells (B). Eleven of 23 patients had dense infiltrates of lymphoid cells, but
follicle-like structures were absent (C and D). The third group of patients exhibited fewer inflammatory cells, which were diffusely distributed
throughout the tissue (E and F). In B, D, and F, arrowheads indicate B cell aggregates; arrows indicate T cells. (Original magnification ⫻ 40.)
Organization of cellular infiltrates in juvenile
DM muscle. All patients had a confirmed diagnosis of
juvenile DM (Table 1). Histopathologic evaluation of
muscle biopsy samples showed extensive cellular infiltra-
tion of perimysial and perivascular areas. Figure 1 shows
3 patterns of organization of infiltrates based on the
topographic organization of T and B cells as determined
by CD3 and CD20 staining, respectively. These were 1)
dense inflammatory infiltrates resembling lymphoid
Figure 2. Architecture of lymphoid follicle–like structures in inflamed muscle from patients with juvenile DM. Muscle tissues with H&E staining
patterns of lymphoid follicle–like structures were examined further by multicolor immunofluorescence and immunohistochemistry. A, Tissue sections
were stained for the distribution of CD4⫹ T cells (green), CD8⫹ T cells (red), and CD20⫹ B cells (blue). A compact collection of CD4⫹ T cells
(arrow) intermingled with CD20⫹ B cells (notched arrowhead) and CD8⫹ T cells (arrowhead) was observed in muscles with follicle-like
organization. B, There was also colocalization of CD20 (red) and CD23 (green) with the nuclear stain 4⬘,6-diamidino-2-phenylindole (DAPI; blue)
(arrowheads), indicating an advanced differentiation state of B cells. C, Colocalization of CNA.42 (green) and CD21L (red) with DAPI (blue)
indicated the presence of follicular dendritic cells (FDCs). Many of the CNA.42⫹CD21L⫹ FDCs (arrow) had fine elongated processes (inset) that
formed interdigitating networks between neighboring cells. D, All follicle-like structures were found to contain high endothelial venules (HEVs),
identified by staining with HECA 452 (red, in contrast to the blue nuclear stain). Two patterns of HECA 452 staining were observed in juvenile DM
muscle: HECA 452 positive (arrow) and HECA 452 negative (arrowhead). (Original magnification ⫻ 100 in A–D; ⫻ 200 in inset.) See Figure 1 for
other definitions.
follicle–like structures (21.7% of tissues) (Figures 1A
and B), 2) dense lymphocytic aggregates lacking folliclelike organization (47.8% of tissues) (Figures 1C and D),
and 3) diffuse distribution of inflammatory cells (30.5%
of tissues) (Figures 1E and F). The T and B cell
compartmentalization in lymphoid-like structures was
reminiscent of that seen in tonsil (results not shown).
For the majority of muscle sections examined, lymphoid
follicle–like structures were located preferentially in the
perimysium. None of these patterns was observed in
control muscle samples without myositis (not shown). In
2 of 23 examined patients with juvenile DM, interposing
follicle-like structures and lymphocytic aggregates were
observed. These 2 patients had a higher number of
follicle-like microstructures covering the whole area of
analyzed tissue. Serial sections from the entire 2-cm
open biopsy sample were examined.
Muscle-infiltrating lymphoid cells were identified
by their reactivity to antibodies to CD3, CD4, CD20,
CD23, and CD8 (Figures 1 and 2) as well as by their
reactivity to antibodies to CD45RA, CD45RO, CD11c,
and CD123. Overall, cellular infiltrates consisted predominantly of CD3⫹ T cells (mean ⫾ SD 59.3 ⫾
13.41%), CD20⫹ B cells (22.7 ⫾ 14.98%), and
CD123⫹CD11c–CD4⫹ PDCs (17.7 ⫾ 3.5%). We previously reported the morphology, location, and distribution of PDCs in inflamed juvenile DM muscle; PDCs
were situated adjacent to blood vessels in close proximity to T and B cell areas (7,11). In the present study,
follicle-like structures were found in muscle samples
from 5 of 23 patients. These structures were characterized by large numbers of CD3⫹ cells distributed as a
circumferential ring around B cell and PDC aggregates
(Figure 1B). The percentage of B cells and the CD4⫹:
Figure 3. Naive and memory T cells in inflamed muscle from patients with juvenile dermatomyositis (DM). Phenotypes of T cells were examined
further by immunofluorescence staining for CD4 (green) (A), the memory marker CD45RO (blue) (B), and the naive marker CD45RA (red) (C).
Arrows in A and B indicate CD45RO⫹CD4⫹ T cells. Arrowheads in A and C indicate CD45RA⫹CD4⫹ T cells. There was a predominance of
CD45RO⫹CD4⫹ T cells (arrow in D, merged image) surrounded by a small but distinguishable rim of CD45RA⫹CD4⫹ T cells (arrowhead in D,
merged image). Naive CD45RA⫹CD4⫹ T cells were found in all juvenile DM muscle with lymphoid follicle–like structures and more occasionally
in muscle with lymphocytic aggregates, but not in muscle with diffuse infiltrates. In some cases, naive CD45RA⫹CD4⫹ T cells were found inside
blood vessels. (Original magnification ⫻ 100.)
CD3⫹ cell ratio were significantly higher in juvenile DM
muscle with lymphoid follicle–like structures than in
muscle samples having either lymphocytic aggregates or
diffuse infiltrates. Muscles with follicle-like structures
also had higher proportions of CD8⫹ T cells (Figure
Similarly, cellular infiltrates in juvenile DM muscles with diffuse infiltrates (Figures 1E and F) or lymphocytic aggregates (Figures 1C and D) were predominantly CD3⫹ T cells and CD20⫹ B cells, but there was
no discrete organization of the clustering of T cells and
B cells. Diffuse infiltrates showed significantly fewer
cells, at ⱕ50 cells per hpf; most of these were
CD4⫹CD123⫹ PDCs and a few scattered CD20⫹ B
cells and CD4⫹ (or CD8⫹) CD3⫹ T cells. Lymphoid
cells were generally absent in control muscle (not
shown). As expected, tonsil tissues had abundant lymphoid follicles, consisting of CD20⫹ B cells and CD3⫹
T cells found mostly in GCs and in the interfollicular
areas, respectively (results not shown).
Advanced differentiation state of CD20ⴙ cells in
juvenile DM muscle. Since production of autoantibodies
to muscle is pathognomonic for juvenile DM (29,30), we
examined the differentiation state of the infiltrating B
cells. In normal B cells, CD23, the low-affinity IgE
receptor, indicates B cell maturation and differentiation
to plasma cells (31). Hence, we examined costaining for
CD23 and CD20. Results showed high-intensity staining
for CD23 in ⬃30% of all CD20⫹ B cells in muscles with
lymphoid follicle–like structures (Figure 2B). Costaining
for CD23 and CD20 was observed in only 2 muscle
specimens with dense lymphocytic aggregates but lacking follicle-like organization. All of the remaining cases
(including muscles with diffuse infiltrates) and all of the
control muscle samples were CD23–.
FDCs in follicle-like microstructures in juvenile
DM muscle. True lymphoid follicles found in secondary
lymphoid organs are characterized by networks of FDCs
that serve as a reservoir for antigens for intranodal
activation of B and T cells (32). Hence, we examined the
distribution of FDCs in juvenile DM muscle for the
expression of the long form of CD21 (CD21L) and the
FDC antigen CNA.42 (33,34). The results showed highintensity staining for these FDC antigens (Figure 2C).
CD21L⫹CNA.42⫹ cells showed a dendritic morphology
with cytoplasmic projections or elongations. FDCs were
found exclusively in juvenile DM muscles with lymphoid
follicle–like structures. As expected, tonsil tissue stained
strongly for CD21L and CNA.42 (not shown). All control muscle was CD21L–CNA.42–.
HEVs in follicle-like microstructures in juvenile
DM muscle. HEVs are also considered to be a feature of
normal lymphoid follicles (35). We therefore examined
patterns of staining for HECA 452, an adhesion molecule expressed on endothelial cells of HEVs (36). In
tonsil there was strong HECA 452 staining of blood
vessels, as expected (not shown). In juvenile DM muscles, 2 patterns of staining were observed, namely,
HECA 452 positive and HECA 452 negative (Figure
2D). Overall, HECA 452–positive vessels were found in
5 of 18 juvenile DM muscle samples, 3 of which had
many lymphoid follicle–like structures. HECA 452–
positive vessels were observed in 2 juvenile DM muscle
tissue samples with dense lymphocyte infiltrates. Juvenile DM muscle with diffuse infiltrates displayed little, if
any, HECA 452 staining. HECA 452–positive vessels
were absent in control muscle.
Naive T cells in follicle-like microstructures in
juvenile DM muscle. While naive T cells are found in
circulation, they home to, and are residents of, regional
lymph nodes (37). Except for ␥/␦ T cells, classic naive
CD4⫹ and CD8⫹ T cells are not known to reside in
nonlymphoid tissue but have been reported in the
peripheral blood of adults with myositis (38). Hence, we
further examined the phenotype of T cells in juvenile
DM muscles, since the vast majority of CD3⫹ cells were
also CD4⫹ (Figure 3). Using antibodies to CD45RO
and CD45RA (classic memory and naive markers, respectively), we observed that CD4⫹ T cells within
lymphoid follicle–like structures were mostly CD45RO⫹
(Figures 3B and D). In addition, there were significant
numbers of CD45RA⫹CD4⫹ T cells within these extranodal follicles (Figures 3C and D). CD45RA⫹ cells
constituted 28.7 ⫾ 6.1% of all CD4⫹ T cells in these
microstructures. Naive CD45RA⫹CD4⫹ T cells were
observed predominantly in perivascular areas and, in
some cases, within blood vessels, which suggests that
they may have been either infiltrating into, or emigrating
out of, the inflammatory lesion (not shown).
CD45RA⫹CD4⫹ T cells were also frequently observed
in small clusters within CD123⫹ PDC–rich areas (not
shown). CD45RA⫹CD4⫹ T cells were sporadically seen
in juvenile DM muscle with diffuse infiltrates or with
lymphocytic aggregates. These cells were absent in control muscle.
Expression of CXCL13, LT␣, and LT␤ in folliclelike microstructures in juvenile DM muscle. The notion
of lymphoneogenesis in juvenile DM came from our
original observation that CCL19 and CCL21, 2 chemokines involved in the formation of lymphoid follicles, are
up-regulated in cellular infiltrates in muscle (7). In the
present study, we examined other humoral factors involved in lymphoid organogenesis. We investigated for
BCA-1/CXCL13 and CXCR5, a ligand–receptor pair
that promotes lymphoid organogenesis through the induction of LT␤ in FDCs (13,15). Lymphoid infiltrates
from juvenile DM muscle sections were isolated by laser
capture microdissection, and messenger RNA (mRNA)
was prepared and used in real-time quantitative RTPCR. While nonmyositic control muscle showed few or
no chemokine transcripts, we found that juvenile DM
muscle had uniformly high levels of BCA-1/CXCL13
(3.7-fold), LT␣ (2.6-fold), and LT␤ (3.5-fold) transcripts. We found differential levels of expression of
these molecules among the 3 histologic types of juvenile
DM muscle. Figure 4A shows that diffuse infiltrates had
the lowest level of specific mRNA for each of these
molecules. Follicle-like structures showed higher levels
of up-regulation of CXCL13 and LT␤ transcripts than
were found in muscle samples having either lymphocytic
aggregates or diffuse infiltrates. Muscles with lymphocytic aggregates showed slightly higher levels of LT␣
mRNA than those with follicle-like organization.
To complement the quantitative RT-PCR assays,
we examined staining patterns of CXCL13 and CXCR5
in muscle tissue. Figure 4B shows colocalization of these
2 molecules only in follicular structures and lymphocytic
aggregates in juvenile DM muscle. As noted above, we
observed the highest number of CXCL13⫹ cells within
Figure 4. Expression of CXCL13, lymphotoxin ␣ (LT␣), and LT␤. A, Cellular infiltrates in frozen muscle sections for each of the 3 histologic types
were harvested by laser capture microdissection. Messenger RNA was isolated and used in quantitative reverse transcription–polymerase chain
reaction (PCR) assays for CXCL13, LT␣, and LT␤. Levels of amplicons for each of the 3 molecules in muscle from patients with juvenile
dermatomyositis (JDM) are expressed as the mean fold difference compared with levels in nonmyositic control muscle; data were normalized to the
internal housekeeping gene ␤-actin. The amounts of CXCL13, LT␣, and LT␤ were significantly higher in juvenile DM muscle with either dense
infiltrates or lymphoid follicle–like structure than in nonmyositic control muscle (P ⬍ 0.05). Juvenile DM muscle with diffuse infiltrates also had
higher levels of CXCL13 and LT␣ than did nonmyositic muscle, but the difference was not statistically significant (P ⫽ 0.054). Also shown are
agarose gels of fractionated specific PCR products with the indicated amplicon sizes. B, Expression of the receptor-ligand pair, CXCL13 and
CXCR5, was verified by immunostaining of frozen sections of juvenile DM muscle with lymphoid follicle–like structures, lymphocytic aggregates,
or diffuse infiltrates. Dense staining was observed for CXCL13 (brown; arrows) and CXCR5 (purple; arrowheads) in follicular structures and
lymphocytic aggregates. Nonmyositic muscle biopsy samples did not stain for this receptor-ligand pair (not shown). (Original magnification ⫻ 40.)
Figure 5. Microchimerism of lymphoid follicle–like structures in muscle and in peripheral blood from patients with juvenile dermatomyositis (DM).
Muscle tissue from male juvenile DM patients with follicle-like structures (A–D) or lymphocytic aggregates (E and F) and peripheral blood CD34⫹
cells (G) were subjected to fluorescence in situ hybridization (FISH). FISH probes for X and Y chromosomes were labeled with spectrum orange
(red) and fluorescein isothiocyanate (green), respectively. The merged image in D shows a female XX cell (2 red spots in a single nucleus with intact
borders; arrowhead) surrounded by male XY cells (arrow indicates Y chromosome). Shown in E and F is the colocalization of CD20 and the
X-chromosome probe. Similar colocalization of the X-chromosome probe and the plasmacytoid dendritic cell marker CD123 was also found (not
shown). Arrowheads in B, E, and F show female XX cells. Depicted in G are CD34⫹ cells from the blood of a male juvenile DM patient showing
the presence of a female XX cell (2 red spots in a single nucleus with intact borders; arrowhead) along with male XY cells (1 green and 1 red signal;
arrow). Nuclei werre counterstained with 4⬘,6-diamidino-2-phenylindole (blue). Immunophenotypes of XX cells were better discriminated in
lymphocytic aggregates due to lower cell density and less overlap of cells relative to follicle-like structure. (Original magnification ⫻ 250 in A–D;
⫻ 100 in E–G.)
follicle-like structures (Figure 4A). Neither CXCL13 nor
CXCR5 was detected in nonmyositic control muscle.
Microchimerism. Investigators in our group have
reported that juvenile DM is characterized by microchimerism, in which maternal cells can be identified in the
inflammatory lesions in the muscle as well as in circulating leukocytes (39). Microchimerism has been postulated to be one of the driving forces of autoimmunity in
juvenile DM (39,40).
Among the 23 children studied, we analyzed 6
boys. Using FISH, maternal cells, identified by 2 X
chromosomes, were detected in muscle biopsy samples
from these boys. We observed maternal XX cells in the
center of lymphoid follicle–like structures (Figures
5A–D) and throughout the diffuse infiltrates.
The phenotype of the maternal XX cells was
determined using FISH for X and Y chromosomes
simultaneously with immunohistochemical staining for
CD20, CD45, and CD123 (Figures 5E and F). The
results showed that 33% of maternal XX cells expressed
CD20 and 25% expressed CD123. None of the cells
expressed CD45. The rest were uncharacterized female
Chimerism was further examined in blood from 3
boys with juvenile DM by FISH analysis of sorted
CD34⫹ blood cells (Figure 5G). The total number of
XX cells in the blood was 1–10 cells per 2,500 CD34⫹
cells with distinct X and Y chromosomes (0.04–0.42%)
(data not shown). These data suggest that XX cells
found in blood from boys with juvenile DM come from
a hematopoietic precursor that is maternally derived.
Whether maternal XX stem cells are themselves found
in muscle or are derived from muscle stem cells is
unknown at this time, since muscle tissues were not
available for muscle stem cell analysis.
Clinical features associated with the prevalence
of follicle-like microstructures in juvenile DM muscle.
The 17 female patients and 6 male patients with juvenile
DM all manifested characteristic skin rash, symmetric
proximal weakness, elevated levels of muscle-derived
enzymes, electromyographic evidence of myopathy, and
classic perifascicular atrophy with capillary occlusion on
muscle biopsy. Table 1 shows demographic and clinical
characteristics and laboratory findings in all patients
with juvenile DM, distributed according to the histologic
pattern of cellular infiltrates in the muscle.
For the purpose of our study, severe disease was
defined as muscle strength ⬍68/80 by Manual Muscle
Test 8 (MMT 8) scoring (26), a score of ⱖ5 on a 10-cm
visual analog scale (VAS) for physician’s and patient’s
global assessment of disease activity, and evidence of
cutaneous and muscle vasculopathy with cutaneous ulcerations. The disease activity was scored at diagnosis
and after 6 months of followup. We found that patients
with lymphocytic aggregates were more likely to have
severe disease, but that they responded to standard
therapy with oral and intravenous corticosteroids and
methotrexate. In contrast, patients with well-defined
lymphoid follicle–like structures had severe disease and
still needed additional disease-remitting agents such as
cyclosporine, intravenous Ig, or rituximab after 6 months
of standard treatment. Patients with juvenile DM who
were diagnosed as having milder disease (defined as an
MMT 8 score of ⱖ70, no cutaneous ulcerations or
periungual telangiectasias, physician’s and patient’s
global assessment VAS scores of ⬍2.5, and responsiveness to treatment) all had diffuse infiltrates. These
observations suggest that outcome and response to
treatment are correlated with the presence of ectopic
lymphoid structures in muscle lesions.
Lymphocytic aggregates are characteristic of inflammatory myopathies in both adults and children
(1,6,41). Our data show that some of these cellular
aggregates organize as extranodal lymphofollicular
structures in the muscle of children with newly diagnosed juvenile DM. Dense aggregates of CD3⫹ T cells
and CD20⫹ B cells were found in almost all juvenile
DM muscle biopsy samples examined. Approximately
one-third of all biopsy samples had the following cellular
and molecular components reminiscent of lymph nodes:
1) abundant lymphoid infiltrates with discrete compartmentalization of CD3⫹ T cells and CD20⫹ B cells,
2) accumulation of naive CD45RA⫹CD4⫹ T cells,
3) HEVs, 4) activated CD23⫹ B cells, 5) CD8⫹ T cells,
6) FDC networks, and 7) the lymphoid-specific chemokines CXCL13, LT␣, and LT␤. Complementing our
previous report of high levels of expression of CCL19
and CCL21 (7), the present data categorically indicate
formation of ectopic lymphoid microstructures. The
muscle inflammatory process is focal and can at times
spare closely associated tissue or have disparate effects
on muscles in close proximity. In the diagnosis of
juvenile DM, biopsy samples from multiple muscles are
generally not clinically obtained. In the present study, all
specimens were open biopsy samples with an average
yield of a 2-cm3 sample, minimizing sampling error.
With this single random biopsy approach, it is noteworthy that we were able to identify discrete histologic forms
of lymphoid organization in juvenile DM muscle that
even correlated with clinical disease severity.
Lymphocytic aggregates with characteristics of
extranodal lymphoid structures in inflammatory lesions
have been reported for 2 other autoimmune diseases,
Sjögren’s syndrome (inflamed salivary glands) and rheumatoid arthritis (the synovium) (15,42). In both of these
diseases of adults, the follicle-like structures in the
lesions are populated with T cells, B cells, and FDCs
similar to what we report here for children with newonset juvenile DM (43,44). The presence of dendritic
cells in extranodal follicular structures in these 2 adult
diseases has not been validated. In contrast, our present
data show that PDCs and FDCs are components of
lymphoid follicle–like structures in juvenile DM.
Although the presence of classic GCs has yet to
be determined, our data indicate that lymphoid-like
microstructures in juvenile DM muscle likely represent
extranodal sites of immune activation. They are populated by activated PDCs in close proximity to T cells and
B cells. Accumulation of naive CD45RA⫹CD4⫹ T cells
strongly suggests bypassing of immune activation and
perhaps even immune regulation that normally occur in
regional lymph nodes. While the overexpression of
CXCL13 suggests active homing of naive and memory
T cells to the inflammatory lesions, it is not yet clear
whether antigen-driven activation and expansion of T
cells occur in situ.
Investigators in our group previously demonstrated maternal chimerism in juvenile DM (24,39) and
other autoimmune disorders. In maternal chimerism,
akin to graft-versus-host disease, it is thought that
maternally derived T and B cells could be reactive with
the host (the child with juvenile DM) (45). An alternative view is that chimerism could be beneficial to the
host. In this case, maternally derived cells could actively
induce tolerance in the host.
In the present study, we found maternal XX cells
in the muscle of boys with juvenile DM, particularly in B
cell–rich areas, and in follicle-like structures in general.
Our data show that these maternal cells express antigens
that tag B cells (CD20) and PDCs (CD123, CD4). We
also identified CD34⫹ XX cells in the blood of boys with
juvenile DM, indicating that maternal hematopoietic
stem cells are present in the blood in juvenile DM. It is
not known whether XX cells found in the juvenile DM
muscle lesion could also be derived from maternal
muscle stem cells.
Perhaps the most compelling evidence for lymphoneogenesis in juvenile DM is provided by our observations of the high levels of expression of CXCL13, LT␣,
and LT␤, as well as the presence of HEVs; these findings
complement our previously reported findings on the
expression of CCL19 and CCL21 (7). CXCL13, CCL19,
and CCL21 are well-known chemokines for homing of
lymphocytes and dendritic cells to normal lymphoid
organs (46). LT␣ and LT␤ are critical factors for lymphoid organogenesis, so much so that targeted deletion
of the encoding genes in mice yields animals with a tiny
amount of or completely absent lymphoid tissue (22).
Further, notwithstanding the role of HEVs in intranodal
trafficking and peripheral export of lymphocytes, HEVs
are essential for lymphoid organogenesis since they form
the conduit through which precursor cells of FDCs and
stromal cells enter the developing lymphoid organ (46).
HEV staining was not seen in all the ectopic lymphoid
structures in juvenile DM muscle. The reason for this is
unknown, but it could be related to angiogenic processes
of HEV formation, which remain to be examined in the
context of lymphoneogenesis. Nevertheless, high levels
of expression of CXCL13, CCL19, CCL21, and HEVs in
juvenile DM muscle suggest a local microenvironment
that is permissive for extranodal lymphoid development
in new-onset disease. It could be that development of
these structures is among the initiating events that
ultimately lead to muscle inflammation.
An important finding of the present study is the
association of lymphoid follicle–like structures with severe disease. In clinical practice, evaluation of muscle
biopsy in the diagnosis of juvenile DM has become less
common. Our histologic and molecular data, in conjunction with clinical data on muscle strength and vascular
changes, underscore the importance of tissue histopathology in ascertaining juvenile DM prognosis (3,47).
We report the unique observation that muscle lymphoid
follicle–like organization is found among patients with
more severe and difficult-to-treat disease, while those
with milder disease have a scattering of inflammatory
cells. Large numbers of naive and memory T cells, B
cells, FDCs, PDCs, and HEVs in these follicle-like
structures indicate that they are extranodal foci of
immune activation that bypass the normal regulatory
controls in the regional lymph nodes. It could be that
these structures represent inflammatory foci that are
more difficult to disrupt; hence our observation that
patients having these structures required additional
therapeutic agents. Thus, it will be of interest to examine
whether indications of clinical improvement in difficultto-treat juvenile DM with anti–B cell therapy using
rituximab (48,49) might involve disruption of the folliclelike structures via the in situ depletion of B cells. Having
identified the cellular and molecular components of
these extranodal structures, we suggest that they are
potential targets of improved immune-based therapies
especially for juvenile DM.
We thank Molly S. Wagner for assistance with chimerism analysis; Anthony Blahnik and Jim Tarara for assistance
with laser capture microdissection and confocal microscopy,
respectively; Dr. Andrew Engel (Mayo Clinic Muscle Laboratory) for providing additional specimens; and José R. Padilla
for assistance with the creation of the figures.
Dr. Reed had full access to all of the data in the study and
takes responsibility for the integrity of the data and the accuracy of the
data analysis.
Study design. López de Padilla, Vallejo, Reed.
Acquisition of data. López de Padilla, Lacomis, McNallan, Reed.
Analysis and interpretation of data. López de Padilla, Vallejo, Lacomis, McNallan, Reed.
Manuscript preparation. López de Padilla, Vallejo, Lacomis, Reed.
Statistical analysis. López de Padilla.
1. Banker BQ, Victor M. Dermatomyositis (systemic angiopathy) of
childhood. Medicine (Baltimore) 1966;45:261–89.
2. Pachman LM. Juvenile dermatomyositis: immunogenetics, pathophysiology, and disease expression. Rheum Dis Clin North Am
3. Miles L, Bove KE, Lovell D, Wargula JC, Bukulmez H, Shao M,
et al. Predictability of the clinical course of juvenile dermatomyositis based on initial muscle biopsy: a retrospective study of 72
patients. Arthritis Rheum 2007;57:1183–91.
4. Ramanan AV, Feldman BM. Clinical features and outcomes of
juvenile dermatomyositis and other childhood onset myositis syndromes. Rheum Dis Clin North Am 2002:28:833–57.
5. Dalakas MC. Immunopathogenesis of inflammatory myopathies.
Ann Neurol 1995:37 Suppl 1:S74–86.
6. De Bleecker JL, Engel AG, Butcher EC. Peripheral lymphoid
tissue-like adhesion molecule expression in nodular infiltrates in
inflammatory myopathies. Neuromuscul Disord 1996:6:255–60.
Lopez de Padilla CM, Vallejo AN, McNallan KT, Vehe R, Smith
SA, Dietz AB, et al. Plasmacytoid dendritic cells in inflamed
muscle of patients with juvenile dermatomyositis. Arthritis Rheum
Cavanagh LL, Boyce A, Smith L, Padmanabha J, Filgueira L,
Pietschmann P, et al. Rheumatoid arthritis synovium contains
plasmacytoid dendritic cells. Arthritis Res Ther 2005;7:R230–40.
Jahnsen FL, Farkas L, Lund-Johansen F, Brandtzaeg P. Involvement of plasmacytoid dendritic cells in human diseases. Hum
Immunol 2002;63:1201–5.
Cella M, Jarrossay D, Facchetti F, Alebardi O, Nakajima H,
Lanzavecchia A, et al. Plasmacytoid monocytes migrate to inflamed lymph nodes and produce large amounts of type I interferon. Nat Med 1999;5:919–23.
Lopez de Padilla CM, Reed AM. Involvement of dendritic cells in
autoimmune diseases in children. Pediatr Rheumatol Online J
Luther SA, Bidgol A, Hargreaves DC, Schmidt A, Xu Y, Paniyadi
J, et al. Differing activities of homeostatic chemokines CCL19,
CCL21, and CXCL12 in lymphocyte and dendritic cell recruitment
and lymphoid neogenesis. J Immunol 2002;169:424–33.
Hjelmstrom P. Lymphoid neogenesis: de novo formation of lymphoid tissue in chronic inflammation through expression of homing chemokines. J Leukoc Biol 2001;69:331–9.
Weyand C, Kurtin P, Goronzy J. Ectopic lymphoid organogenesis:
a fast track for autoimmunity. Am J Pathol 2001;159:789–93.
Takemura S, Klimiuk PA, Braun A, Goronzy JJ, Weyand CM.
Lymphoid neogenesis in rheumatoid synovitis. J Immunol 2001;
Page G, Lebecque S, Miossec P. Anatomic localization of immature and mature dendritic cells in an ectopic lymphoid organ:
correlation with selective chemokine expression in rheumatoid
synovium. J Immunol 2002;168:5333–41.
Amft N, Curnow SJ, Scheel-Toellner D, Devadas A, Oates J,
Crocker J, et al. Ectopic expression of the B cell–attracting
chemokine BCA-1 (CXCL13) on endothelial cells and within
lymphoid follicles contributes to the establishment of germinal
center–like structures in Sjögren’s syndrome. Arthritis Rheum
Armengol MP, Juan M, Lucas-Martin A, Fernandez-Figueras MT,
Jaraquemada D, Gallart T, et al. Thyroid autoimmune disease:
demonstration of thyroid antigen-specific B cells and recombination-activating gene expression in chemokine-containing active
intrathyroidal germinal centers. Am J Pathol 2001;159:861–73.
Weninger W, Carlsen HS, Goodarzi M, Moazed F, Crowley MA,
Baekkevold ES, et al. Naive T cell recruitment to nonlymphoid
tissues: a role for endothelium-expressed CC chemokine ligand 21
in autoimmune disease and lymphoid neogenesis. J Immunol
Aust G, Sittig D, Becherer L, Anderegg U, Schutz A, Lamesch P,
et al. The role of CXCR5 and its ligand CXCL13 in the compartmentalization of lymphocytes in thyroids affected by autoimmune
thyroid diseases. Eur J Endocrinol 2004;150:225–34.
Drayton DL, Ying X, Lee J, Lesslauer W, Ruddle NH. Ectopic
LT␣␤ directs lymphoid organ neogenesis with concomitant expression of peripheral node addressin and a HEV-restricted sulfotransferase. J Exp Med 2003;197:1153–63.
Drayton DL, Liao S, Mounzer RH, Ruddle NH. Lymphoid organ
development: from ontogeny to neogenesis. Nat Immunol 2006;7:
Aloisi F, Pujol-Borrel R. Lymphoid neogenesis in chronic inflammatory diseases. Nat Rev Immunol 2006;6:205–17.
Reed AM, McNallan K, Wettstein P, Vehe R, Ober C. Does
HLA-dependent chimerism underlie the pathogenesis of juvenile
dermatomyositis? J Immunol 2004;172:5041–6.
Bohan A, Peter JB. Polymyositis and dermatomyositis (first of two
parts). N Engl J Med 1975;292:344–7.
Oddis CV, Rider LG, Reed AM, Ruperto N, Brunner HI, Koneru
B, et al, for the International Myositis Assessment and Clinical
Studies Group. International consensus guidelines for trials of
therapies in the idiopathic inflammatory myopathies. Arthritis
Rheum 2005;52:2607–15.
McNallan KT, Aponte C, el-Azhary R, Mason T, Nelson AM, Paat
JJ, et al. Immunophenotyping of chimeric cells in localized scleroderma. Rheumatology (Oxford) 2007;46:398–402.
Livak KJ, Schmittgen TD. Analysis of relative gene expression
data using real-time quantitative PCR and the 2⫺⌬⌬CT method.
Methods 2001;25:402–8.
Love LA, Leff RL, Fraser DD, Targoff IN, Dalakas M, Plotz PH,
et al. A new approach to the classification of idiopathic inflammatory myopathy: myositis-specific autoantibodies define useful homogeneous patient groups. Medicine (Baltimore) 1991;70:360–74.
Wedderburn LR, McHugh NJ, Chinoy H, Cooper RG, Salway F,
Ollier WE, et al, on behalf of the Juvenile Dermatomyositis
Research Group (JDRG). HLA class II haplotype and autoantibody associations in children with juvenile dermatomyositis and
juvenile dermatomyositis–scleroderma overlap. Rheumatology
(Oxford) 2007;46:1786–91.
Delespesse G, Sarfati M, Wu CY, Fournier S, Letellier M. The
low-affinity receptor for IgE. Immunol Rev 1992;125:77–97.
Van Nierop K, de Groot C. Human follicular dendritic cells:
function, origin and development. Semin Immunol 2002;14:251–7.
Liu YJ, de Bouteiller O, Parham CL, Grouard G, Djossou O,
de Saint-Vis B, et al. Follicular dendritic cells specifically express
the long CR2/CD21 isoform. J Exp Med 1997;185:165–70.
Raymond I, Al Saati T, Tkaczuk J, Chittal S, Delsol G. CNA.42, a
new monoclonal antibody directed against a fixative-resistant
antigen of follicular dendritic reticulum cells. Am J Pathol 1997;
Girard JP, Springer TA. High endothelial venules (HEVs): specialized endothelium for lymphocyte migration. Immunol Today
Duijvestijn AM, Horst E, Pals ST, Rouse BN, Steere AC, Picker
LJ, et al. High endothelial differentiation in human lymphoid and
inflammatory tissues defined by monoclonal antibody HECA-452.
Am J Pathol 1988;130:147–55.
Lee Y, Chin RK, Christiansen P, Sun Y, Tumanov AV, Wang J,
et al. Recruitment and activation of naive T cells in the islets by
lymphotoxin ␤ receptor-dependent tertiary lymphoid structure.
Immunity 2006;25:499–509.
O’Hanlon TP, Messersmith WA, Dalakas MC, Plotz PH, Miller
FW. ␥␦ T cell receptor gene expression by muscle-infiltrating
lymphocytes in the idiopathic inflammatory myopathies. Clin Exp
Immunol 1995;100:519–28.
Reed AM, Picornell YJ, Harwood A, Kredich DW. Chimerism in
children with juvenile dermatomyositis. Lancet 2000;356:2156–7.
Artlett CM, Ramos R, Jiminez SA, Patterson K, Miller FW, Rider
LG. Childhood Myositis Heterogeneity Collaborative Group.
Chimeric cells of maternal origin in juvenile idiopathic inflammatory myopathies. Lancet 2000;356:2155–6.
Arahata K, Engel AG. Monoclonal antibody analysis of mononuclear cells in myopathies. I. Quantitation of subsets according to
diagnosis and sites of accumulation and demonstration and counts
of muscle fibers invaded by T cells. Ann Neurol 1984;16:193–208.
Stott DI, Hiepe F, Hummel M, Steinhauser G, Berek C. Antigendriven clonal proliferation of B cells within the target tissue of an
autoimmune disease: the salivary glands of patients with Sjögren’s
syndrome. J Clin Invest 1998;102:938–46.
Nordmark G, Alm GV, Ronnblom L. Mechanisms of disease:
primary Sjögren’s syndrome and the type I interferon system. Nat
Clin Pract Rheumatol 2006;2:262–9.
Sarkar S, Fox DA. Dendritic cells in rheumatoid arthritis. Front
Biosci 2005;10:656–65.
Kremer Hovinga IC, Koopmans M, Baelde HJ, de Heer E, Bruijn
JA, Bajema IM. Tissue chimerism in systemic lupus erythematosus
is related to injury. Ann Rheum Dis 2007;66:1568–73.
Muller G, Lipp M. Concerted action of the chemokine and
lymphotoxin system in secondary lymphoid-organ development.
Curr Opin Immunol 2003;15:217–24.
Wedderburn LR, Varsani H, Li CK, Newton KR, Amato AA,
Banwell B, et al, on behalf of the UK Juvenile Dermatomyositis
Research Group. International consensus on a proposed score
system for muscle biopsy evaluation in patients with juvenile
dermatomyositis: a tool for potential use in clinical trials. Arthritis
Rheum 2007;57:1192–201.
48. Noss EH, Hausner-Sypek DL, Weinblatt ME. Rituximab as therapy for refractory polymyositis and dermatomyositis. J Rheumatol
49. Cooper MA, Willingham DL, Brown DE, French AR, Shih FF,
White AJ. Rituximab for the treatment of juvenile dermatomyositis:
a report of four pediatric patients. Arthritis Rheum 2007;56:3107–11.
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muscle, severity, inflamed, dermatomyositis, lymphoid, disease, juvenile, extranodal, microstructure, new, onset
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