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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 91 No. 6 (March 15), 1998:
pp. 1852-1857
Clinical-Grade Functional Dendritic Cells From Patients With Multiple
Myeloma Are Not Infected With Kaposi's Sarcoma-Associated Herpesvirus
By
Karin Tarte,
Sonja J. Olsen,
Zhao Yang Lu,
Eric Legouffe,
Jean-François Rossi,
Yuan Chang, and
Bernard Klein
From the Institute for Molecular Genetics, CNRS, Montpellier, France;
the Division of Epidemiology, School of Public Health, Columbia
University, New York, NY; the Unit for Cellular Therapy, CHU
Montpellier, Hopital Saint Eloi, Montpellier, France; the Service des
Maladies du Sang B, CHU Montpellier, Hopital Lapeyronie, Montpellier,
France; and the Department of Pathology, College of Physicians and
Surgeons, Columbia University, New York, NY.
 |
ABSTRACT |
Bone marrow dendritic cells (DC) from patients with multiple myeloma
(MM) were recently reported to be infected with Kaposi's sarcoma-associated herpesvirus (KSHV). Because immunotherapy strategies using DC are very promising in this disease, we looked for KSHV DNA in
clinical-grade DC generated in vitro from MM patients. Adherent
apheresis cells from MM patients were maintained for 7 days in
clinical-grade X-VIVO 15 culture medium supplemented with
granulocyte-macrophage colony-stimulating factor, interleukin-4, or
interleukin-13. Tumor necrosis factor  was added for the last 2 days. We obtained a cell population with a DC phenotype able to
endocytose fluorescein isothiocyanate (FITC)-dextran and efficiently activate resting allogenic T lymphocytes. To detect KSHV DNA, we used
polymerase chain reaction (PCR) followed by Southern blotting of PCR
product with a sensitivity detecting a few copies of viral DNA. All the
PCR were repeated in a blinded fashion three times, on 1 µg and 0.2 µg of genomic DNA, in two different laboratories. Clinical-grade DC
from 10 (91%) of 11 patients were not infected with KSHV. The
apheresis cells and the purified CD34+ cells from the
same patients were also negative. A very weak PCR band was detected
with DC from one patient, but the initial apheresis cells were
negative. The detection of KSHV infection in 1 (9%) of 11 MM patients
probably represents background seroprevalence. It seems likely that
functional and clinical-grade DC from MM patients can safely be used in
clinical trials.
| |
INTRODUCTION |
BONE MARROW dendritic cells (DC) from
patients with multiple myeloma (MM) were found by Rettig et
al1 to be infected with Kaposi's sarcoma-associated herpes
virus (KSHV) in contrast to those from non-MM patients. Additionally,
these cells were reported to express viral interleukin-6 (vIL-6) ,which
is known to share functional homology with human IL-6.2-4
vIL-6 supports the proliferation of B9 murine hybridoma cells,
activates human gp130 independently of binding to
IL-6R,5 and promotes the survival and proliferation of
myeloma cells (unpublished results). The ability of KSHV
to code for vIL-6 and other potential oncogenic
proteins6-10 suggests that KSHV, which is involved in the
pathogenesis of Kaposi's sarcoma, a subset of Castleman disease and
body-cavity based lymphoma,11-13 could also be associated
with MM, an IL-6-related disease.14
Several studies have demonstrated that DC efficiently induce
antigen-specific antitumoral immunity in vitro and in
vivo.15-18 As recently described, clinical trials with DC
pulsed with monoclonal Ig gave promising results in MM.19
DC from MM patients infected with KSHV is a major impediment to the
future clinical use of these cells.
We recently described a simple method to obtain a virtually pure
population of functional dendritic cells from apheresis cells from
patients with MM using a combination of granulocyte-macrophage colony-stimulating factor (GM-CSF), IL-4, and tumor necrosis factor- (TNF).20 Several studies have shown that IL-4 and IL-13
share several biologic activities. In particular, they induce monocytes to differentiate into dendritic cells.21,22 Moroever, the
IL-13 receptor, which comprises the common IL-4 receptor chain,23 is able to transduce a signal for both
cytokines.24-27 In this report, we extend our method to
clinical-grade culture conditions and compare these two related
cytokines, IL-4 and IL-13, for their ability to promote DC development.
We demonstrate that clinical-grade functional DC from most patients
with MM do not harbor KSHV DNA, making their use in vivo possible. In
addition, we were unable to detect any KSHV DNA in apheresis cells or
in purified CD34+ progenitors from these patients.
| |
PATIENTS, MATERIALS, AND METHODS |
Patients.
We evaluated 11 patients with MM (median age, 55 years). All patients
exhibited active disease and, according to Durie and Salmon
classification, there was 1 stade IA, 1 stade IIIB, and 9 stade IIIA.
Five patients had an IgG , 2 an IgG , 3 an IgA, and 1 an IgA
monoclonal component. After written informed consent was obtained,
patients were treated with a three VAD (Vincristine, Adriamycine,
Dexamethasone) chemotherapy regimen followed by high-dose cyclophosphamide (4 g/m2) and recombinant human granulocyte
colony-stimulating factor (G-CSF; filgrastim; NEUPOGEN; Amgen-Roche,
Neuilly sur Seine, France; daily injection of 5 µg/kg from day 2 after chemotherapy until hematologic recovery). The peripheral blood
CD34+ cell count was monitored every day and peripheral
blood cells were collected at the peak of CD34+ cell count
during a 5-hour apheresis. The mean leucocyte count was 21.3 ± 15.2 × 103/µL (range, 2.5 to 69 × 103/µL) and the median time of collection after
cyclophosphamide injection was 10 days (range, 9 to 12 days).
Peripheral blood from two healthy volunteers was collected under
heparin after written informed consent was obtained.
Cell lines.
XG-1 is an IL-6-dependent human MM cell line.28 BCBL-1 is
a body cavity lymphoma cell line latently infected by KSHV but not by
Epstein-Barr virus (EBV).29
Preparation and characterization of DC.
DC were generated from apheresis cells (AC) as previously
described.20 Briefly, 3 × 106 AC/mL were
grown in RPMI 1640 supplemented with 2 mmol/L of L-glutamine and 10% fetal calf serum (FCS) or in X-VIVO 15 serum-free medium from
Biowittaker (Walkersville, MD) (ie, clinical-grade DC). After 2 hours,
nonadherent cells were discarded and adherent cells were cultured in
the same media with 100 ng/mL of GM-CSF (LEUCOMAX; Sandoz, Basel,
Switzerland) and 25 ng/mL of IL-4 (Genzyme, Cambridge, MA) or IL-13
(Sanofi, Labège, France). After 5 days of culture, TNF (R&D
Systems, Minneapolis, MN) was added at 20 ng/mL. The same phenotypic
and functional analyses were performed on DC obtained in RPMI-FCS and
X-VIVO 15 media. On day 5 of culture, before TNF addition, we
studied the capacity of these cells for endocytosis through the mannose
receptor using lysine-fixable fluorescein isothiocyanate
(FITC)-dextran, molecular weight equal to 40,000 (Molecular Probes Inc,
Eugene, OR), as previously described.30 AC-derived DC were
incubated for 7, 15, and 30 minutes at 37°C with 1 mg/mL of
FITC-dextran in RPMI-10% FCS-25 mmol/L HEPES. After 4 washes with cold
phosphate-buffered saline (PBS)-1% FCS-0.02% NaN3 , cells
were analyzed on a FACScan apparatus (Becton Dickinson, San Jose,
CA). Background uptake was measured by incubating cells at
4°C. On day 7, cells maintained in the different media and cytokine
conditions were collected and labeled with the following monoclonal
antibodies (MoAbs): CD4 and CD83 conjugated with phycoerythrin (PE) and
CD14, CD54, CD58, CD80, and HLA DR conjugated with FITC (Immunotech,
Marseille, France). FITC-anti-CD1a, FITC-anti-CD40, and FITC-anti-CD86
MoAbs were purchased from Pharmingen (San Diego, CA). Intracellular
staining for CD68 was performed using EBM11 MoAb (Dako, Glostrup,
Denmark) and Fix and Perm kit (Caltag Laboratories, Burlingame, CA).
Negative controls were performed with corresponding irrelevant matched
murine MoAbs (Immunotech). Finally, these DC were used in an allogenic
mixed lymphocyte reaction (MLR) to stimulate T cells purified from
healthy volunteers' peripheral blood by 2 cycles of negative selection
using anti-CD14, -CD19, -CD56, -CD16, and -HLA DR MoAbs (Immunotech)
and goat-antimouse Ig magnetic microbeads (Dynal, Oslo, Norway) as
previously reported.20 Graded numbers of irradiated DC (30 Gy) were added to 1.5 × 105 allogenic T cells in 200 µL of RPMI-5% serum AB culture medium. After 5 days of coculture,
cells were pulsed with 1 µCi of tritiated thymidine (Amersham,
Buckingham, UK) for 12 hours and radioactive incorporation was counted.
Results are expressed as mean counts per minute (cpm) ± SD
determined in sextuplet culture wells.
KSHV DNA detection.
For patients no. 1 to 11, two different cell populations were tested
for KSHV infection: AC and clinical-grade DC derived from these AC. In
addition, for patients no. 1 through 8, we purified CD34+
cells. AC were collected after centrifugation on Ficoll-Hypaque (Biowittaker). CD34+ AC were purified by the clinical-grade
methodology from Baxter (Isolex 300; Baxter International Inc,
Deerfield, IL) according to the manufacturer's instructions. This
allowed the selection of a highly pure population of CD34+
cells (median, 92%; range, 85% to 98%) as assessed by labeling with
a pool of three PE-conjugated CD34 MoAbs that recognized three
different epitopes of the CD34 molecule (Immunotech). DC were collected
after 7 days of culture in X-VIVO 15 medium with GM-CSF, IL-4, and
TNF . AC, CD34+ cells and clinical-grade DC were frozen
and stored at  80°C until DNA extraction. DNA was extracted
with a Promega kit (Madison, WI) according to the manufacturer's
instructions. The positive control was genomic DNA from the BCBL-1 cell
line. To assess the integrity of genomic DNA, we performed a polymerase
chain reaction (PCR) with primers for  -globin (forward primer,
5 -CAACTTCATCCACGTTCACC-3; reverse primer,
5-GAAGAGCCA AGGACAGGTAC-3). The 268-bp sized -globin
fragment could be detected in all the DNA samples (data not shown). For
sensitivity assay, 1 µg to 0.1 pg of BCBL-1 DNA was diluted in the
DNA of the KSHV-negative XG-1 MM cell line. All PCR were performed 3 times with 1 µg of DNA (corresponding to approximatively 150,000 cells) using the primers described by Chang et al11 that
amplify a 233-bp DNA fragment. A total of 40 cycles of PCR
amplification was used and PCR products (10 µL) were electrophoresed
on a 3% agarose gel impregnated with ethidium bromide. They were then
blotted onto a positive nylon membrane and hybridized, as
described,11 with the 25-bp internal probe, labeled with
32P using the T4 polynucleotide kinase (GIBCO BRL, Paisley,
UK). Autoradiographs were developed after 2 and 6 hours of exposure; the membranes were also exposed for 24 hours to PhosphorImager to
exclude the presence of weak bands. The same procedure was performed on
0.2 µg of DNA in the laboratory of Dr Chang.
vIL-6 immunohistochemistry.
Expression of KSHV vIL-6 was evaluated in two DC cultures (patients no.
1 and 10) using a polyclonal rabbit antiserum raised against vIL-6
peptides that does not cross-react with human IL-6. DC were harvested
and embedded in 1% agarose plugs that were formalin-fixed and then
processed in paraffin. Four-micrometer sections were cut on coated
slides and immunostaining was performed with the avidin-biotin complex
(ABC) method using a previously published protocol.2 BCP-1
(KSHV-infected) and P3HR-1 (EBV-infected) cell lines prepared in the
same way were used as positive and negative controls, respectively.
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RESULTS |
Generation of clinical-grade DC from apheresis from MM patients.
Adherent AC from patients with MM were cultured with RPMI 1640 and 10%
FCS or with clinical-grade X-VIVO 15 culture medium with GM-CSF and
IL-4 or GM-CSF and IL-13 for 5 days. TNF was added on day 5 for 2 additional days. In agreement with our previous report,20
the various culture conditions resulted in the generation of cells with
phenotypic characteristics of DC (Table 1).
In particular, these cells were always CD14 and HLA
DR+. Cells cultured with X-VIVO 15 medium expressed less
CD1a and CD4 than those cultured with RPMI and 10% FCS. A complete
phenotypic picture of clinical-grade DC cultured with GM-CSF and IL-13
is shown in Fig 1. It was similar to the
phenotype of DC cultured in X-VIVO 15 medium with GM-CSF and IL-4 we
previously reported.20 Despite these phenotypic
differences, cells generated in X-VIVO 15 medium have the same ability
to endocytose FITC-dextran as those obtained in RPMI-FCS
(Fig 2). Again, no difference was
found for DC generated with GM-CSF and IL-4 or GM-CSF and
IL-13 (Fig 2). Furthermore, these cells efficiently present antigens to
resting allogenic T cells (Fig 3).
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| Fig 1.
Dendritic phenotype of adherent AC cultured in X-VIVO 15 medium with GM-CSF and IL-13. AC were maintained in X-VIVO 15 clinical-grade culture medium with GM-CSF and IL-13 for 5 days. TNF
was added for 2 additional days. Flow cytometric analysis was performed at day 7. Isotype-matched murine MoAbs were used as negative controls (dotted lines).
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| Fig 2.
Comparison of FITC-dextran endocytosis by DC obtained in
X-VIVO 15 and in RPMI-10% FCS. DC generated after 5 days of culture in
RPMI-10 % FCS (A) or in X-VIVO 15 medium (B) supplemented with GM-CSF
and IL-13 or GM-CSF and IL-4 were incubated for various lengths of time
in medium containing 1 mg/mL of FITC-dextran and fluorescence was
analyzed by flow cytometry after extensive washing. Solid lines are the
background uptake at 4°C. Dotted lines are 7 minutes at 37°C.
Broken lines are 15 minutes at 37°C. Bold lines are 30 minutes at
37°C.
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| Fig 3.
DC obtained in X-VIVO 15 medium stimulate allogenic
T-lymphocyte proliferation. DC were generated in clinical-grade culture medium using 7 days of culture of AC in the presence of GM-CSF and IL-4
or GM-CSF and IL-13. TNF was added for the last 2 days. After 3 washes and irradiation (30 Gy), they were used in graded numbers as
stimulator cells for 1.5 × 105 allogenic T cells. Cell
proliferation was evaluated by a 12-hour pulse with 3H-TdR
after 5 days of coculture. Data are expressed as the mean ± SD of
sextuplet culture wells. 3H-TdR incorporation rates of
irradiated DC or purified T cells were less than 600 cpm.
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In conclusion, clinical-grade culture medium allowed the generation of
functional DC from patients with MM. In addition, IL-13 was as
efficient as IL-4 in these culture conditions.
Lack of KSHV DNA in clinical-grade DC from patients with MM.
We investigated whether DC generated from MM patients in X-VIVO 15 supplemented with GM-CSF and IL-4 contained KSHV DNA. Using 40 cycles
of PCR with KSHV 330 primers, Southern blotting, and hybridization with
an internal radiolabeled probe, we were able to detect the presence of
KSHV DNA in 1 pg of genomic DNA of the KSHV-infected BCBL-1 cell line
diluted in 1 µg of genomic DNA of the KSHV-negative XG-1 myeloma cell
line (Fig 4). Because every BCBL-1 cell
contained an average of 30 copies of the KSHV genome,31 1 pg of BCBL-1 DNA corresponded to about 5 KSHV copies. This was in
agreement with the previously reported high sensitivity of this
PCR.32 We failed to detect KSHV DNA in 1 µg of DNA (ie, 150,000 cells) from MM patients' clinical-grade DC in 10 (91%) of 11 cases (Fig 5A and B). For patient no. 8, a
weak band could be detected, indicating that few KSHV copies were
present. All PCR were repeated with identical results 3 times in
Montpellier on 1 µg of DNA and in New York using 0.2 µg of DNA. We
obtained exactly the same PCR data when starting from DNA of DC
obtained in RPMI-10% FCS (data not shown). In addition, none of the 11 patients' apheresis cells was positive for KSHV DNA (Fig 5C). Because
CD34+ cells could also be used to generate DC in
vitro,33-35 CD34+ cells from 8 of 11 patients
were purified (85% to 98% of CD34+ cells; median, 92%)
and tested. These cells were also negative for KSHV DNA (Fig 5D).
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| Fig 4.
Sensitivity of KSHV PCR. Lanes 1 through 8 contain
10-fold dilutions of BCBL-1 DNA from 1 µg (lane 1) to 0.1 pg (lane
8). BCBL-1 DNA was diluted in the KSHV-negative XG-1 MM cell line DNA
so that all the PCR were run on 1 µg of total DNA. (A) Ethidium bromide-stained agarose gel of the 233-bp amplification products. (B)
Specific hybridization of the PCR products to a
32P-end-labeled internal probe after transfer to a nylon
membrane.
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| Fig 5.
PCR amplification of DNA from clinical-grade DC (A and
B), from corresponding AC (C), and from CD34+ cells (D)
from MM patients. (A) Ethidium bromide-stained agarose gel of the
233-bp amplification products. Lane M was a molecular size marker. (B,
C, and D) Specific hybridization of the PCR products to a
32P-end-labeled internal probe after transfer to a nylon
membrane. The positive control (lane C) was the PCR product from the
BCBL-1 cell line.
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DISCUSSION |
We have previously reported the generation of DC by culturing apheresis
cells from patients with MM with GM-CSF and IL-4 for 7 days. These
cells had the phenotype of DC (CD1+, CD4+,
CD14, HLA DR+, CD80+,
CD86+), were able to endocytose FITC-dextran, and were able
to present soluble antigens to naive T cells. We used RPMI 1640 culture
medium and FCS and presented preliminary data showing that
clinical-grade serum-free culture medium could be used.20
In the present study, we reinforce these data and show that pure and
functional DC could be generated with clinical-grade serum-free culture
medium. Adherent apheresis cells were collected and cultured with
X-VIVO 15 culture medium for 7 days with GM-CSF and IL-4. The
replacement of IL-4 with IL-13 yielded similar results. These cells had
a DC phenotype with slight differences suggestive of a less mature
phenotype when compared with those generated with RPMI 1640 and 10% of
FCS. However, their endocytic capacity was similar. We previously
reported that addition of TNF for the last 2 days of culture was
necessary to get an efficient antigen-presenting ability,20
and this was confirmed in this study (data not shown). Thus, we show
here that fully functional DC can be generated from MM patients.
To address the recent concern that bone marrow DC are infected with
KSHV and express vIL-6 gene,1 we investigated whether DC
generated from apheresis cells in our culture conditions might be
infected with KSHV. Using a very sensitive PCR and Southern blotting,
it is possible to amplify KSHV DNA in 1 pg of DNA from KSHV-infected
BCBL-1 cells. We failed to detect any KSHV amplification in 1 µg of
DNA from clinical-grade DC from 10 patients with MM. PCR was performed
blindly in two independent laboratories using KS330233
primers. Furthermore, we failed to find expression of vIL-6 by
immunostaining with anti-vIL-6 antibody in two patients with negative
PCR results (data not shown). The initial apheresis cells (n = 11) as
well as pure CD34+ cells (n = 8) from which the DC
precursors originated were also negative. For one patient, we found a
weak PCR amplification that was seen only after Southern blotting,
suggesting less than 5 KSHV copies in 1 µg of DNA. No PCR
amplification was seen using 100 ng of DC DNA (data not shown). These
data indicate that only a few cells of 150,000 cells were infected with
KSHV. The detection of KSHV in 1 (9%) of 11 patients is within the
range (0% to 25%) of background KSHV seroprevalence in several
studies.36-39 It is of note that total AC and
CD34+ purified cells from this patient were negative for
KSHV PCR. It is possible that the rare cells infected with KSHV were
retained by the adherence step used to generate DC from total AC,
resulting in sufficient enrichment for the presence of KSHV DNA and
detection with our very sensitive method. It has been recently
suggested that monocytes were productively infected by KSHV in
Kaposi's sarcoma lesions and KSHV was detected by PCR in the adherent
monocytic cell population obtained after 8 days of culture of
peripheral blood mononuclear cells of acquired immunodeficiency
syndrome-associated Kaposi's sarcoma patients.40
We do not use the culture conditions described by Rettig et
al1 because nobody has yet shown that such a method is
successful to obtain DC. Moreover, Rettig et al1 presented
no evidence that their infected cells were really DC in terms of
endocytosis, expression of costimulatory molecules, antigen
presentation, and activation of T lymphocytes. Thus, the present study
may ascertain that fully characterized DC from most MM patients are not
infected with KSHV. In addition, when we cultured stromal cells
according to the protocol described by Rettig et al,1 we
obtained a mixed population of fibroblastic and monocytic cells that
were not infected with KSHV (unpublished results).
However, one cannot exclude the possibility that a rare bone marrow
cell that can be expanded under very specific conditions could be
infected by KSHV in MM patients. We may only note that two groups
failed to find KSHV seropositivity in patients with MM41,42
in agreement with the results of Dr Chang (unpublished
results). Our data, along with the lack of KSHV detection
in tumoral samples 43, do not favor a causal role of KSHV
in MM. In conclusion, this study shows that clinical-grade DC
from most patients with MM are not infected with KSHV.
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FOOTNOTES |
Submitted October 14, 1997;
accepted December 11, 1997.
Supported by grants from ARC (Paris, France), LFNC (Paris, France), and
AFS (Paris, France).
Address reprint requests to Bernard Klein, PhD, Institute for Molecular
Genetics, CNRS, 1919 Route de Mende, 34033 Montpellier, France.
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.
| |
ACKNOWLEDGMENT |
The authors appreciate the generous gift of rhIL-13 from A. Minty (Sanofi, Labège, France). We thank G. Cathala for expert technical advice, thank G. Fiol and M-C. Delteil for their help in
generating DC samples, and acknowledge L. Milligan for her assistance
in the preparation of the manuscript.
| |
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Abstract
Introduction
Abstract