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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 94 No. 4 (August 15), 1999:
pp. 1409-1417
Demonstration of Frequent Occurrence of Clonal T Cells in the
Peripheral Blood But Not in the Skin of Patients With Small Plaque
Parapsoriasis
By
J. Marcus Muche,
Ansgar Lukowsky,
Jürgen Heim,
Markus Friedrich,
Heike Audring, and
Wolfram Sterry
From the Department of Dermatology, University Hospital
Charité, Humboldt University of Berlin, Germany.
 |
ABSTRACT |
Clinical, immunohistological, and molecular biological data suggest
the chronic dermatosis small plaque parapsoriasis (SPP) to be a
precursor of mycosis fungoides (MF). However, most data are
contradictory and confusing due to inexact definition of SPP. Recently,
clonal T cells were detected in skin and blood samples of early MF.
Because demonstration of identical T-cell clones in skin and blood of
SPP patients would indicate a close relationship of SPP to MF, we
investigated the clonality of skin and blood specimens from 14 well-defined SPP patients. By a polymerase chain reaction (PCR)
amplifying T-cell receptor  rearrangements and subsequent
high-resolution electrophoresis, clonal T cells were detected in 9 of
14 initial and 32 of 49 follow-up blood samples, but in 0 of 14 initial
skin specimens. Even a clone-specific PCR showing the persistence of
the initial blood T-cell clone in 20 of 20 follow-up samples, failed to
detect the T-cell clone in the skin. In 2 patients, the clonal T cells
were shown to be CD4+. For the first time, the majority
of SPP patients was shown to carry a T-cell clone in the peripheral
blood. Although a relation between circulating clonal T cells and SPP
cannot directly be proven by the applied techniques, our results
indicate blood T-cell clonality to be a characteristic feature of SPP
and CTCL because analysis of multiple controls and clinical workup of
our SPP patients excluded other factors simulating or causing a clonal
T-cell proliferation. A sufficient cutaneous antitumor response but
also an extracutaneous origin of the T-cell clones might explain the
failure to detect skin infiltrating clonal T cells.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
THE TERM PARAPSORIASIS was proposed by
Brocq1 in 1902 to describe a group of skin diseases, all of
unknown etiology, chronicity, the failure to respond to therapy, and
lack of subjective symptoms as pruritus.1 He differentiated
three subtypes: parapsoriasis en gouttes, parapsoriasis lichenoide, and
parapsoriasis en plaques (PEP). Today, the first subtype is referred as
pityriasis lichenoides, the second as parakeratosis variegata, and the
last is used analogously to small plaque parapsoriasis
(SPP).2 Discrepancies exist regarding the classification of
large plaque parapsoriasis (LPP). Most authors refer both, LPP and SPP
as PEP,3,4 whereas Lambert and Everett2 group
LPP with parakeratosis variegata and differentiate both from SPP.
Because the term parapsoriasis is used to name the entire group as well
as particular subtypes, data generated in parapsoriatic patients are
confusing.5,6 In particular, the relation of PEP and
cutaneous T-cell lymphoma (CTCL) is discussed controversially.
According to the clinical course, LPP as well as SPP were found to
progress into mycosis fungoides (MF) in 0% to 46% of the cases.2,7-9 Clonal T-cell receptor (TCR)
rearrangements were detected by Kikuchi et al10 using
Southern blot analysis in 4 of 20 LPP cases and by Haeffner et
al11 using a polymerase chain reaction (PCR) assay in two
of three SPP patients. In contrast, other authors12,13
using the same methods failed to detect clonal rearrangements in LPP
and SPP, respectively. Immunohistology showed the skin-infiltrating T
cells of 14 MF and 7 LPP patients to be CD4+ and
Leu8 in all samples as well as
CD7 (Leu9) in 11 MF and 4 LPP
cases, respectively. Because this pattern is uncommon in inflammatory
skin diseases, PEP was considered to be an early form of
MF.14 This hypothesis is supported by the detection of
functionally abnormal blood lymphocytes in both, early MF and
LPP.15 Furthermore, G-banding and interphase cytogenetic assays showed circulating chromosomally abnormal T lymphocytes usually
found in MF and Sezary syndrome patients also in PEP
samples.16
Regarding these data, most authors2,17,18 consider LPP (and
SPP) as the most common precursor of CTCL. Ackerman et
al6,19 even regard SPP, LPP, and parakeratosis variegata as
clinical presentations of MF. In contrast, Burg et al5,20
emphasize that differentiation between LPP and SPP is not always clear
and that most of the data mentioned above refer to LPP. Therefore, they
discuss SPP as a benign process in which an initial DNA defect leads to
the generation of a skin homing T-cell clone not undergoing further
mutations necessary to develop into overt CTCL and suggest the term
abortive CTCL for SPP. The clinical course of the disease should be the
criterion to differentiate between SPP and MF simulating SPP. However,
differentiation between MF simulating SPP and SPP is advisable at the
initial analysis to anticipate the clinical course as well as to treat
the patients in an adequate manner. This view might be supported by the
observation that early stage MF, appropriately treated with topical
mechlorethamine or total skin electron beam therapy, shows survival
rates similar to that of a race-, age-, and sex-matched
population.21
Recently, we could show clonal T cells not only in skin but also in
blood samples of early MF.22 Because SPP might be a precursor of MF, we asked whether T-cell clonality is also detectable in skin and blood specimens of a well-defined group of 14 SPP patients.
Demonstration of identical T-cell clones in both compartments would
indicate the systemic character of SPP and its close relationship to MF
as well as facilitate prediction of the clinical course and selection
of therapy modalities.
| |
MATERIALS AND METHODS |
Patient samples.
The study included 14 SPP patients that were untreated for at least 6 months before the initial analysis. For each patient, a blood and a
skin specimen was collected for the initial analysis, 1 to 8 additional
blood samples were taken during the follow-up of 18 to 47 months.
Detailed data on medical history and follow-up are given in
Tables 1 and 2.
Special attention was paid to conditions reported to be risk factors
for disease progression in CTCL (lower rate of complete remissions with
initial treatment, older age as well as a higher stage of skin lesions
at the initial presentation, elevated levels of  2 microglobulin
and/or lactic dehydrogenase).21,23,24 The diagnosis of SPP
was assessed by two independent investigators according to the
following, previously published criteria2,4,18,25: chronic
persistence (more than 2 years) of asymptomatic, well defined, round to
oval shaped, red to brown colored, nonindurated macules, or thin
plaques measuring less than 5 cm in diameter and displaying a fine
scaling that gives the surface a wrinkled appearance; symmetrical
involvement preferring trunk and proximal extremities, but sparing face
and volar surfaces; compact stratum corneum with moderate acanthosis,
spongiosis, and foci of laminated parakeratosis; edematous papillary
dermis with a sparse, perivascular to band-like infiltrate composed of
lymphohistiocytic cells appearing neither enlarged nor atypical or
neoplastic; few of the bland-appearing lymphocytes infiltrating the
epidermis; size (< 5cm), regular shape, and symmetrical distribution
of the lesions discerning SPP from LPP; mild extent of the
lymphohistiocytic infiltrate and absence of necrotic keratinocytes and
of broad bands of parakeratosis differentiating SPP from pityriasis
lichenoides; absence of so-called Pautrier's microabscesses in the
epidermis containing atypical large lymphocytes discriminating SPP from
MF. Histological criteria are illustrated in
Fig 1. Patients not exactly matching the
diagnostic criteria or not equally rated by both investigators were
excluded from the study.
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| Fig 1.
Biopsy specimens from SPP patient SF illustrating the
diagnostic criteria compact stratum corneum with moderate acanthosis
and foci of spongiosis, slightly edematous papillary dermis with a
sparse band-like infiltrate composed of bland lymphohistiocytic cells,
few of them epidermotropic lymphocytes, and absence of so-called
Pautrier's microabscesses. (A) Hematoxyline-eosine staining (×33);
(B) CD3 staining (×50) showing the T-cell nature of the infiltrating
lymphocytes.
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Ten control blood specimens were derived from patients with active
contact dermatitis (n = 5), and alopecia areata sensitized with topical
diphencyprone to induce a local contact dermatitis (n = 5). Cells of
the JM cell line26 (DSMZ, Braunschweig, Germany) served as
clonal control.
Sample preparation.
Peripheral blood mononuclear cells (PBMC) were prepared from 10 mL of
heparinized blood by density-gradient centrifugation through
Ficoll-HyPaque (Pharmacia, Freiburg, Germany). CD4+ and
CD8+ T cells were purified by
CD4+/CD8+ T cell Isolation Kit (Miltenyi
Biotec, Bergisch Gladbach, Germany) and Midi MACS. Genomic DNA was
prepared from about 1 × 106 cells by a standard
procedure using Proteinase K digestion.27 For preparation
of genomic DNA from the paraffin-embedded skin specimens, the paraffin
of 10 sections per sample (10 µm each) was dissolved with xylene.
After centrifugation, the pellet was washed with ethanol and also
digested by proteinase K.
TCR PCR and determination of clonality.
TCR rearrangements were PCR amplified using primers annealing at the
V and J segments, respectively.22 Primers VG1 (segments V1 to 8; CTACATCCACTGGTACCT), VG9 (V9; ATTGGTATCGAGAGAGAC), VG2
(V10, 11, B, [A]; CACTGGTACKKGCAGAAAC), and JG12-a (J1, 2;
CAACAAGTGTTGTTCCAC) were applied to all specimens (PCR-1), whereas
PCR-2 using primers VG1, VG9, VG2, and JGP12-a (JP1, P2;
CTATGAGCYTAGTCCCTT) was performed in those SPP specimens appearing polyclonal in PCR-1 and in all control samples. PCR reaction mixture included 0.5 to 1 µg (5 µL) of genomic DNA, 1.75 U of Taq
polymerase, 1.5 mmol/L MgCl2, and 7.5 µL 10 × PCR
buffer (Perkin Elmer, Branchburg, NJ), 0.1 mmol/L of each
deoxynucleotide triphosphate (dNTP; Pharmacia, Freiburg, Germany), and
0.6 µmol/L of each primer in a final volume of 75 µL. Amplification
was performed on a thermal cycler (Varius-V; Vers, Hannover, Germany)
by a 4-minute denaturation step at 95°C, followed by 40 cycles
including 1 minute denaturation at 94°C, 1 minute annealing at
58°C, and 1 minute extension at 72°C. Finally, an extension
step of 5 minutes at 72°C was added. Six microliters of the PCR
products were screened for successful amplification on a 2% agarosegel
stained by ethidium bromide.
T-cell clonality was established by detection of a dominant TCR
rearrangement in a heteroduplex loaded temperature gradient gel
electrophoresis (HD-TGGE). Eight microliters of the PCR products were
prepared to form heteroduplices (5 minutes denaturation at 95°C,
gradual cooling to 50°C)28 and separated on the Diagen TGGE-System (Qiagen, Hilden, Germany). Electrophoretic run and subsequent silver staining were performed according to standard protocols.29 Due to the denaturation-renaturation step,
polyclonal (ie, not identical) amplification products form
heteroduplices that appear as a broad smear on the gel. In contrast,
clonal (ie, identical) PCR products form homoduplices migrating as
sharp bands into the high temperature range of the gradient gel
(Fig 2). PCR products derived from the JM
cell line (rearranged V8 and V11) served as positive clonal
control.
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| Fig 2.
Temperature-gradient gel of TCR PCR products. Lane
1-10, PCR products derived from blood and skin samples of SPP patients
HO, ME, RI, SF, and SZ; all blood samples showed clonal PCR products.
Lane 11, Hinc II digest of phi X174. S, skin sample; B, blood sample;
M, marker (Hinc II digest of phi X 174); b, biallelic (or biclonal); m,
monoclonal; p, polyclonal; a, range of polyclonal smears;
b, range of clonal bands.
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Cloning and sequencing of the TCR rearrangements.
For direct sequencing, the distinct HD-TGGE band was cut out and
dissolved in 40 µL 1 × PCR buffer (Perkin Elmer) overnight. Five microliters of the solution were reamplified under the same conditions described above. Primer JG12-i (TGTTGTTCCACTGCCAAA) or
JGP12-i (CCTTYWGCAAAYRTCTTGA) was applied instead of JG12-a or JGP12-a,
respectively. The PCR product was purified by the QIAquick PCR
purification kit (Qiagen) and sequenced on an automated DNA sequencer
(Model 373A, Perkin Elmer Applied Biosystems, Weiterstadt, Germany) by
the Taq cycle sequencing method using primers VG, JG12-i, or JGP12-i.
Sequences were aligned to the published germline sequences of the
TCR V and J segments.30-36
Cloning of the PCR products was performed by applying the TA Cloning
Kit (Invitrogen, Fleek, The Netherlands). Plasmids were sequenced using
the universal forward sequencing primer for M13 as described above.
Clone-specific PCR.
Using the Oligo 5.0 software (National Biosciences, Plymouth, MN),
clone-specific primers were designed so that the 3' end annealed
at the N region of the clonal TCR rearrangement. After PCR
amplification using clone-specific and corresponding VG primer at
standard conditions, PCR products were screened on an agarose gel.
Primer and PCR conditions were considered specific for the clonal
TCR sequence if at least 7 randomly chosen clonal or polyclonal DNA
samples from CTCL patients and healthy volunteers (tester DNAs) yielded
no amplification product (Fig 3A). To
achieve this, the annealing temperature was increased by 1°C steps
until amplification products of the tester DNA disappeared. In the
seminested clone-specific PCR, the clone-specific primer was applied in
combination with an inner VG primer corresponding to the rearranged
V segment: VGseq (V1-8; AGRCCCCACAGCRTCTTC),22 VG10-i
(V10; ATCCGCAGCTCGACGCAGCA), or VG11-i (V11;
CTCAAGATTGCTCAGGTGGG).37
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| Fig 3.
(A) Agarose gel of products of the clone-specific PCR
using primer BE-1. Lane 1, skin sample DNA of BE; 2, blood sample DNA
of BE; 3-5, skin sample DNA from CTCL patients (tester DNA); 6-8, blood
sample DNA from CTCL patients (tester DNA). A specific PCR product
(indicated by the arrow) is observed exclusively in lane 2. The
additional band in lanes 2 and 5 represents a larger PCR product that,
according to the size, is referred to as an unspecific amplificate. (B)
Agarose gel of the sensitivity assay using primer JU-5 and 10-fold
dilution of JM cell line DNA in PBMC from a healthy donor. A specific
PCR product (indicated by the arrow) is found up to the dilution of 10 JM cells in 106 PBMC (10-5) in lane 6. (C)
Results of the analysis of the fractionated clones (1-14, established
from patient HO; 15-20, from BE): fraction A was sequenced directly,
fraction B was amplified by clone-specific PCR using primer HO-2 and
separated on an agarose gel. Nonconcordance between PCR and sequencing
is observed in lane 1 and 19. M, marker (Hinc II digest of phi X 174);
PCR, PCR result; SEQ, result of sequencing; +, PCR
product of expected size/sequence identical to the second allel of
patient HO; , no PCR product of expected size/sequence
not identical to the second allele of patient HO.
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RESULTS |
TCR PCR analysis.
Previous studies proved specificity and sensitivity of the applied
TCR PCR/HD-TGGE in cases of CTCL and several benign conditions. Among 60 control specimens derived from peripheral blood of patients with atopic dermatitis (n = 20), psoriasis vulgaris (n = 20), and
healthy volunteers (n = 20), all but 1 sample derived from a psoriatic
patient showed polyclonal PCR products.22,38 The present
study included 10 additional control specimens derived from peripheral
blood of patients with active contact dermatitis (n = 5), and with
alopecia areata treated by diphencyprone to induce contact dermatitis
(n = 5). Clonal T cells circulating in the peripheral blood were
detected in none of these control samples.
Among the SPP patients (Table 2), TCR PCR/HD-TGGE detected clonally
expanded T cells in 9 of the 14 blood samples taken for initial
analysis. In patient HO, a biallelic rearrangement was found in this
compartment. The clonality of the 9 blood samples was confirmed by
directly sequencing the PCR products. The clonal sequences had never
been found in our laboratory so far, as excluded by data base searches.
Analysis of the follow-up blood specimens showed T-cell clonality in 32 of 34 samples taken from the 9 SPP patients previously shown to carry
circulating clonal T cells, whereas all of the 15 samples collected
from the remaining 6 SPP cases were shown to be polyclonal.
In contrast to the peripheral blood, no DNA sample from a corresponding
SPP skin lesion showed a clonal PCR product (Table 2). Nevertheless,
only simultaneous demonstration of the T-cell clone in both, blood and
skin specimens of a patient would indicate a direct relation between
SPP and the T-cell clonality found in the peripheral blood.
Clone-specific PCR analysis.
To detect the T-cell clone more sensitively than by TCRPCR/HD-TGGE,
a clone-specific PCR was established. Based on the N region of the
clonal TCR sequences, clone-specific primers were designed and
applied in combination with the corresponding VG primer (Table 2). For
the biallelic TCR rearrangement detected in patient HO, two
clone-specific primers were constructed.
Specificity of the clone-specific PCR was achieved by increasing the
annealing temperatures until tester DNA amplification products
disappeared (Fig 3A). By this procedure, a clone-specific PCR could be
established in 6 of the 9 SPP patients with clonal blood samples (Table
2). In patients CZ and BC, the N region was too short for the design of
specific primers. No clone specificity was achieved with primers HO-1
and HL-1.
Furthermore, specificity of the approach was shown by cloning the PCR
products of patient HO and BE. Twenty clones (14 from patient HO, 6 from BE) were picked and each clone was aliquoted into 2 fractions (A
and B). Fraction A was sequenced directly, whereas fraction B was
reamplified by the clone-specific PCR using primer HO-2. Concordance of
sequencing and clone-specific PCR was observed in 18 of the 20 clones
(11 clone positive, 7 clone negative). Two samples were positive by the
clone-specific PCR of fraction B, but the clonal TCR sequence was
not obtained by sequencing fraction A (Fig 3C).
The sensitivity of our clone-specific PCR system was determined by
dilution of clonal T cells (JM cell line) in polyclonal PBMC of a
healthy volunteer. After DNA preparation and PCR by primer VG1 and
JU-5, a distinct electrophoretic band was observed down to a dilution
of 10 JM cells in 1,000,000 PBMC (0.001%; Fig 3B).
By the clone-specific PCR assays, all of the 20 follow-up blood samples
taken from the patients BE, HO, ME, RI, SF, and SZ were shown to carry
the T-cell clone detected and sequenced in the corresponding initial
blood specimen (Table 2). To confirm the putative relation between SPP
and the peripheral blood T-cell clonality, the skin samples of these
patients were also reevaluated using the clone-specific PCR. However,
in none of the 6 cases, a product was observed after PCR of the skin
sample DNA. To further increase the sensitivity, the clone-specific
amplification was performed seminested using clone-specific and
corresponding VG primer in the first round as well as clone-specific
and corresponding VG-i (VGseq) primer in the second round. Again, this
procedure failed to detect the peripheral blood T-cell clone in the
skin specimens.
In patients SZ and HO, CD4+ and CD8+ T cells
were isolated from peripheral blood mononuclear cells (purity 98.26%
and 96.79%, respectively). Using the corresponding clone-specific
assays, an amplification product was observed only in the
CD4+ fraction of both patients.
Clinical investigation.
We analyzed our SPP patients (T-cell clone in the peripheral blood
detected v not detected) for the predominant occurrence of
conditions previously reported to be risk factors for CTCL progression.21,23,24 Although the mean age (68 v 60 years) and the mean duration of the skin lesions (11.8 v 10.8 years) were found to be somewhat higher in the clone-positive group of SPP patients, no statistical significance of these differences was
found (Mann Whitney test, P = .15 and P =
.30, respectively). Evaluation of the other markers (course of the
disease [remission v relapse rate: 6/3 v 3/2],
previous treatment, concomitant disease and medication, laboratory
tests) as well showed no differences between the two groups (Table 1).
In summary, clonal T cells were shown by TCRPCR/HD-TGGE in 9 of 14 blood samples, but surprisingly not in the 14 skin specimens initially
taken from the SPP patients. The data obtained from peripheral blood
were confirmed in 47 of 49 follow-up samples investigated by this
technique. Clone-specific PCR assays could be established in 6 of 9 SPP
cases carrying circulating clonal T cells. These assays showed the
T-cell clone in all follow-up blood samples of the 6 patients. But,
even the use of this high-sensitive technique, including seminested
clone-specific PCR failed to detect the peripheral blood T-cell clone
in the skin. In 2 patients, the circulating clonal T cells were shown
to be CD4+ T cells. The occurrence of T-cell clones in the
peripheral blood of the SPP patients was not correlated with changes in
clinical markers previously found to be risk factors for disease
progression in CTCL.
| |
DISCUSSION |
As the classification of parapsoriasis has been controversial, we
employed a combination of strict clinical and histological criteria,2,4,18,25 applied independently by 2 investigators, to select the patients evaluated in the present study.
To determine the clonality of the samples, a well-established
TCRPCR/HD-TGGE assay was used. The lower detection limit of our test
system was estimated at 103 clonal in 106
polyclonal T cells (0.1%).22 Such a high sensitivity might enable detection of minor clones of reactive lymphocytes in skin lesions of nonspecific dermatitis and cutaneous lymphoid hyperplasia proposed as the "clonal dermatitis" concept.39 Clonal
PCR products were also found repetitively in skin samples of the benign
dermatoses pityriasis lichenoides (chronica and acuta) and
angioimmunoblastic lymphadenopathia40 as well as in the
peripheral blood of posttransplant41 and immunodefiency
patients.42 Some reports described clonal, most notably
CD8+ and  + T cells in the blood of
healthy elderly donors.43 However, all of these conditions
were excluded in our patients (Table 1) and at least in 2 cases, the
circulating clonal T cells belong to the CD4+ fraction that
was not found to be clonally expanded in healthy volunteers by
Fitzgerald et al.43 Moreover, using the applied test system
to investigate 60 blood samples obtained from an age-matched cohort of
healthy volunteers, psoriatic patients, and atopic patients, we
previously found clonal T cells in just 1 blood sample of a patient
with psoriasis vulgaris.22 In the present study,
circulating clonal T cells were not detected in 10 control blood
specimens from patients with active contact dermatitis and with
alopecia areata treated by diphencyprone to induce contact dermatitis
although both circumstances are expected to carry circulating T-cell
clones. Moreover, specificity of the data was also
confirmed by direct sequencing of the 9 blood samples showing T-cell clonality.
Validity of the clone-specific PCR was ensured by nondetection of
tester DNA in all 6 assays, by fractional analysis of a cloned PCR
product of patient HO, and by determination of the sensitivity to
0.001%. The 2 false-positive results in the fractional analysis might
be caused by a contamination of the amplified fraction, but also by an
unspecific annealing of primer HO-2. This might, at least partly,
question the significance of the detection of the initial T-cell clone
in all 20 follow-up blood specimens investigated, but concordant
results of the TCRPCR/HD-TGGE in 18 of the 20 cases support the
validity of the data achieved by the clone-specific PCR. Because
false-negative PCR results were not noticed in the fractional analysis,
the failure of the clone-specific PCR to detect the T-cell clone in the
skin specimens of the 9 SPP patients showing T-cell clonality in the
blood seems not to be caused by technical problems.
Because even the high-sensitive clone-specific techniques failed to
detect the T-cell clone in the skin specimens corresponding to the
clonal blood samples, this clone is either below 0.001 % of all
infiltrating T cells or absent in cutaneous SPP lesions. Due to this
nondetection in the skin, a relation of the circulating clonal T cells
to the pathogenesis of SPP cannot directly be proven by the applied
techniques. Nevertheless, our results indicate the occurrence of T-cell
clonality in the peripheral blood to be a characteristic feature of SPP
and CTCL because analysis of a large cohort of control specimens and
the clinical workup of our SPP patients excluded other factors
simulating or causing a clonal T-cell proliferation in the peripheral
blood. This is in line with previous reports showing functionally and
chromosomally abnormal T lymphocytes in the peripheral blood of both,
PEP and CTCL patients.15,16 For explanation, the following
hypotheses can be discussed.
One scenario would regard the clonal T cells in the peripheral blood of
SPP as already transformed precursors of the malignant skin
infiltrating MF. Assuming transformation in the skin, it remains
unclear why the untransformed but not the transformed T cells migrate
into the skin. Supposing transformation within the peripheral blood,
the skin homing should be induced by a secondary event at least in a
fraction of the clonal cells. However, because cutaneous lesions
already occur in SPP before clonal T cells are detectable in the skin,
this scenario is not supported by our current data and knowledge.
In a second hypothesis, the cutaneous lesions of SPP are regarded as a
successful antitumor response reducing the skin-infiltrating clonal T
cells to an undetectable amount. This would explain the presence of the
clonal T cells in the peripheral blood as well as their nondetection in
the skin lesions. Antitumor responses are likely to occur, because high
frequencies of activated CD8+ T cells, suspected to be
cytotoxic T cells, were observed in the majority of early MF
patients.44 However, at least at some time points, ie, in
some cutaneous specimens, the presence of clonal T cells in the skin
should be postulated. To prove this, investigation of numerous lesions
per patient is required. Accordingly, Haeffner et al,11
also using a PCR assay observed clonal TCR rearrangements even in skin
samples of 2/3 SPP patients.
In conclusion, our data for the first time show the majority of SPP
patients to carry a clonal T-cell population in the peripheral blood
compartment that persists during the course of the disease, and
independently from the administered therapy. Significant differences between clone-positive and clone-negative SPP patients were not found
for markers previously reported to be risk factors for disease progression in CTCL.21,23,24 Although the T-cell clones
were undetectable in the corresponding skin specimens, the blood
clonality is most likely associated with the existence of SPP because
other conditions previously associated with T-cell clonality were
excluded and analysis of multiple control specimens failed to detect
circulating clonal T cells. The nondetection of skin-infiltrating
clonal T cells might be due to a sufficient cutaneous antitumor
response but an extracutaneous origin of the T-cell clones is also
conceivable. Extended follow-up of skin and blood specimens from the
SPP patients carrying circulating clonal T cells will allow further
insight and will indicate a putative correlation of blood clonality and clinical course.
| |
ACKNOWLEDGMENT |
We thank U. Heiduk and S. Richter for their excellent technical
assistance and P. Zambon for critical review.
| |
FOOTNOTES |
Submitted August 10, 1998; accepted April 19, 1999.
Supported by Grant Ste 366/7-1 from the Deutsche Forschungsgemeinschaft
and Grant 70-2091-Lu1 from the Deutsche Krebshilfe-Mildred Scheel Stiftung.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to A. Lukowsky, PhD, Department of
Dermatology, University Hospital Charité, Humboldt University
Berlin, Schumannstrae 20/21, D-10117 Berlin, Germany; email:
ansgar.lukowsky{at}charite.de.
| |
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