Bone marrow progenitor cell reserve and function and stromal cell function are defective in rheumatoid arthritis: evidence for a tumor necrosis factor alpha-mediated effect.

Abstract

Based on previous reports for impaired hematopoiesis in rheumatoid arthritis (RA), and in view of the current interest in exploring the role of autologous stem cell transplantation (ASCT) as an alternative treatment in patients with resistant disease, we have evaluated bone marrow (BM) progenitor cell reserve and function and stromal cell function in 26 patients with active RA. BM progenitor cells were assessed using flow cytometry and clonogenic assays in short-term and long-term BM cultures (LTBMCs). BM stroma function was assessed by evaluating the capacity of preformed irradiated LTBMC stromal layers to support the growth of normal CD34(+) cells. We found that RA patients exhibited low number and increased apoptosis of CD34(+) cells, defective clonogenic potential of BM mononuclear and purified CD34(+) cells, and low progenitor cell recovery in LTBMCs, compared with healthy controls (n = 37). Patient LTBMC stromal layers failed to support normal hematopoiesis and produced abnormally high amounts of tumor necrosis factor alpha (TNF alpha). TNF alpha levels in LTBMC supernatants inversely correlated with the proportion of CD34(+) cells and the number of colony-forming cells, and positively with the percentage of apoptotic CD34(+) cells. Significant restoration of the disturbed hematopoiesis was obtained following anti-TNF alpha treatment in 12 patients studied. We concluded that BM progenitor cell reserve and function and BM stromal cell function are defective in RA probably due, at least in part, to a TNF alpha-mediated effect. The role of these abnormalities on stem cell harvesting and engraftment in RA patients undergoing ASCT remains to be clarified.

Full text

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doi:10.1182/blood.V99.5.1610
2002 99: 1610-1619




Boumpas and George D. Eliopoulos
Helen A. Papadaki, Heraklis D. Kritikos, Claudia Gemetzi, Helen Koutala, Judith C. W. Marsh, Dimitrios T.

mediated effect
−
alpha
are defective in rheumatoid arthritis: evidence for a tumor necrosis factor
Bone marrow progenitor cell reserve and function and stromal cell function

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HEMATOPOIESIS
Bonemarrowprogenitorcellreserveandfunctionandstromalcellfunction
aredefectiveinrheumatoidarthritis:evidenceforatumornecrosis
factoralpha–mediatedeffect
HelenA. Papadaki, Heraklis D. Kritikos, Claudia Gemetzi, Helen Koutala, Judith C. W. Marsh,
Dimitrios T. Boumpas, and George D. Eliopoulos
Based on previous reports for impaired
hematopoiesis in rheumatoid arhrtitis
(RA), and in view of the current interest in
exploring the role of autologous stem cell
transplantation (ASCT) as an alternative
treatment in patients with resistant dis-
ease, we have evaluated bone marrow
(BM) progenitor cell reserve and function
and stromal cell function in 26 patients
with active RA. BM progenitor cells were
assessedusingflowcytometryandclono-
genic assays in short-term and long-term
BM cultures (LTBMCs). BM stroma func-
tion was assessed by evaluating the ca-
pacity of preformed irradiated LTBMC
stromal layers to support the growth of
normal CD34?cells. We found that RA
patients exhibited low number and in-
creased apoptosis of CD34?cells, defec-
tive clonogenic potential of BM mono-
nuclear and purified CD34?cells, and low
progenitor cell recovery in LTBMCs, com-
pared with healthy controls (n ? 37). Pa-
tient LTBMC stromal layers failed to sup-
port normal hematopoiesis and produced
abnormally high amounts of tumor necro-
sis factor alpha (TNF?). TNF? levels in
LTBMC supernatants inversely correlated
with the proportion of CD34?cells and
the number of colony-forming cells, and
positivelywiththepercentageofapoptotic
CD34?cells. Significant restoration of the
disturbed hematopoiesis was obtained
following anti-TNF? treatment in 12 pa-
tients studied. We concluded that BM
progenitor cell reserve and function and
BM stromal cell function are defective in
RA probably due, at least in part, to a
TNF?-mediated effect. The role of these
abnormalitiesonstemcellharvestingand
engraftment in RA patients undergoing
ASCT remains to be clarified. (Blood.
2002;99:1610-1619)
© 2002 by TheAmerican Society of Hematology
Introduction
Rheumatoid arthritis (RA) is a systemic autoimmune disease
characterized mainly by chronic destructive polyarthritis.Although
the main target tissue of the disease is the synovium, evidence has
been accumulated that bone marrow (BM) may be also actively
involved in the pathophysiology of RA by providing immune and
inflammatory cells in the affected synovial membranes.1-3It has
been suggested that during the inflammatory stages of the disease,
BM progenitor cells or even BM-derived immature mesenchymal
cells enter the joint microenvironment through mechanisms similar
to those of mature inflammatory cells, and gradually replace
normal synovial cells.4,5Moreover, on the basis of findings from
experimental models, it has been proposed that RA might even
begin within the BM and might be regarded as a BM disorder.6,7
However, this hypothesis has not been adequately addressed
in humans.
It is now well established that lymphocytes, macrophages, and
fibroblastlike cells found in rheumatoid synovium originate from or
have their counterpart in the BM. These cells are capable of
producing a variety of cytokines that may act not only as mediators
of inflammation but also as regulators of hematopoiesis. It is then
possible that BM, even if not primarily affected, may display
impaired function in RAas a consequence of abnormal interactions
between immune and hematopoietic cells or as a result of an
aberrant, local or systemic, cytokine production potentially affect-
ing the survival, proliferation, and optimal growth of hematopoietic
cells. Indeed, previous studies have shown an altered phenotype
and abnormal growth activity of BM myeloid progenitor cells in
RA, at least at sites adjacent to the affected joints.2,8-10It has also
beenreportedthatelevatedlevelsoflocallyproducedproinflamma-
tory cytokines, such as tumor necrosis factor alpha (TNF?), may
modulate the frequency of erythroid progenitor cells in the BM and
may contribute to the pathogenesis of anemia frequently seen in
patients with RA.11The observation that BM stromal cell lines
established from RApatients displayed abnormal support of B-cell
growth, is additional evidence for a possible defect of BM
microenvironment in RA.12
However, stem cells per se, in terms of their number and
functional characteristics, and BM microenvironment, in terms of
its capacity to support hematopoietic progenitor cell growth, have
not been extensively studied in RA. In view of the current interest
in exploring the use of high-dose immunosuppression followed by
autologous hematopoietic stem cell transplantation (ASCT) in
patients with severe, resistant autoimmune diseases including
RA,13-16it is important to know whether BM stem cell compart-
ment and/or BM microenvironment are already affected by the
inflammatory process in these patients. The aim of the present
study was to evaluate hematopoietic progenitor cell reserve and
From the Departments of Hematology and Rheumatology, University Hospital
of Crete School of Medicine, Heraklion, Greece; and the Department of
Hematology, St George’s Hospital Medical School, London, United Kingdom.
Submitted June 4, 2001; accepted October 26, 2001.
Supported by University Hospital of Heraklion, Greece.
Reprints: Helen A. Papadaki, Department of Hematology, University Hospital
of
epapadak@med.uoc.gr.
Heraklion,POBox1352,Heraklion,Crete,Greece; e-mail:
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.
© 2002 by TheAmerican Society of Hematology
1610BLOOD, 1 MARCH 2002?VOLUME 99, NUMBER 5
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function and BM stromal cell hematopoiesis supporting capacity in
patients with severe, active RA.
Patients, materials, and methods
Patients
We studied 26 white patients with RA, 18 females and 8 males, aged 28 to
71 years (median, 53.5 years). All patients satisfied the 1987 revised
diagnostic criteria of the American College of Rheumatology for RA17and
had evidence of active disease according to established criteria.18Health-
related quality of life and functional status of the patients were determined
using the HealthAssessment Questionnaire (HAQ) index.19Patient charac-
teristics are summarized in Table 1.At the time of study, 6 patients were on
treatment only with nonsteroidal anti-inflammatory drugs (NSAIDs) and
had never been previously treated with corticosteroids or disease-modifying
antirheumatic drugs (DMARDs), 4 patients were on a low-dose oral
corticosteroid regimen receiving 5 mg to 10 mg prednisolone daily, 18 were
taking methotrexate at a dose ranging from 15 mg to 20 mg weekly, and 14
were receiving other DMARDs including 400 mg hydroxychloroquine
daily and/or 2000 mg sulfasalazine-A daily, or 150 mg cyclosporine-A
daily. Patients had discontinued any medication for at least 24 hours prior to
BM aspiration. Of the patients, 12 were also studied 6 to 8 months after
initiation of treatment with cA2 (Remicade, Infliximab; Schering-Plough,
Kenilworth, NJ), a human/murine chimeric monoclonal antibody (Mab) of
IgG1? isotype, with specificity for recombinant and natural humanTNF?.20
Patients were receiving the antibody in a 2-hour intravenous infusion at a
dose of 3 mg/kg on weeks 0, 2, and 6, and then every 2 months, without
discontinuing previous medication.As controls, 37 hematologically healthy
subjects undergoing surgery for lumbar fixation or healthy volunteers, age-
and sex-matched with the patients, were studied. Institutional ethics
committee approval was granted prior to the study. Informed consent was
provided according to the Declaration of Helsinki.
BM samples
BM cells obtained from posterior iliac crest aspirates from patients and
healthy controls, were immediately diluted 1:1 in Iscoves modified
Dulbecco medium (IMDM; Gibco BRL Life Technologies, Paisley, Scot-
land), supplemented with 100 IU/mL penicilline-streptomycin (PS; Gibco-
BRLLifeTechnologies) and 10 IU/mLpreservative-free heparin (Sigma, St
Louis, MO). Diluted BM samples were centrifuged on Lymphoprep (Nycomed
PharmaAS,Oslo,Norway)(density1.077g/cm3)at400gfor30minutesatroom
temperature to obtain the bone marrow mononuclear cells (BMMCs). Cell
number and viability were assessed after staining with trypan blue.
Immunophenotyping and 7-amino-actinomycin D staining
An indirect immunofluorescence technique was used to quantitate BM
CD34?cells. In brief, 1 ? 106BMMCs were stained with phycoerythrin
(PE)–conjugated mouse antihuman CD34 Mab (QBEND-10; Immunotech,
Marseille,France)andfluoresceinisothiocyanate(FITC)–conjugatedmouse
antihuman CD38 Mab (T16; Immunotech) for 30 minutes on ice. PE- and
FITC-conjugated mouse IgG isotype-matched controls were used as
negative controls. Cells were washed twice in phosphate buffered saline
(PBS) containing 1% fetal calf serum (FCS; Gibco BRL, Life Technolo-
gies) and 0.05% azide, and fixed in 500 ?L 2% paraformaldeyde solu-
tion (Sigma).
Aliquots of 1 ? 106BMMCs stained with PE-conjugated anti-CD34
Mab and FITC-conjugated mouse antihuman Fas (CD95) Mab (LOB 3/17;
Serotec, United Kingdom) as above, were further stained prior to fixation
with 7-amino-actinomycin D (7AAD; Calbiochem-Novabiochem, La Jolla,
CA) as previously described.21In brief, 100 ?L 7AAD solution (200 ?g/
mL) was added to the cells, suspended in 1 mL PBS, and incubated for 20
minutes on ice protected from the light. Following centrifugation, the
supernatant was removed and cells were fixed as above. Unstained fixed
cells were used as negative controls.
Cell samples were analyzed on an Epics Elite model flow cytometer
(Coulter, Miami, FL) within 30 minutes of fixation. Data were acquired and
processed on 500 000 events. After creating a scattergram combining
forward and right-angle light scatter for the whole population, a region was
drawn around BMMCs (low forward and low right scatter properties) for
CD34?and CD38?cell estimation. When cells were stained for CD34
antigen, Fas antigen, and 7AAD, a scattergram was created by combining
right-angle light scatter with CD34 fluorescence in the gate of BMMCs and
a second scattergram was generated by combining CD34 and Fas fluores-
cence in the gate of CD34?cells. Finally, a scattergram was generated by
combining forward light scatter with 7AAD fluorescence to quantitate
7AAD-negative (live), -dim (apoptotic), and -bright (dead) cells in the gate
of CD34?, CD34?/Fas?, and CD34?/Fas?cell populations. Regions were
drawn around clear-cut populations, and the proportion of cells within each
region was calculated excluding cell debris (Figure 1).
Purification of CD34?cells
CD34?cells were isolated from BMMCs by indirect magnetic labeling
(magnetic activated cell sorting; MACS isolation kit, Mitenyi Biotec
GmbH, Bergisch Gladbach, Germany) according to the manufacturer’s
protocol. In each experiment, purity of CD34?cells was more than 96% as
estimated by flow cytometry.
Clonogenic progenitor cell assays
BMMCs or CD34?BM cells were cultured in 1 mL IMDM supplemented
with 30% FCS, 1% bovine serum albumin (BSA; Gibco BRL Life
Technologies),10?4Mmercaptoethanol(Sigma),0.075%sodiumbicarbon-
ate (Gibco BRL Life Technologies), 2 mM L-glutamine (Sigma), 0.9%
methylcellulose (StemCell Technologies, Vancouver, BC, Canada), in the
presence of 5 ng granulocyte macrophage–colony-stimulating factor (GM-
CSF; R&D Systems, Minneapolis, MN), 50 ng interleukin-3 (IL-3; R&D
Systems), and 2 IU erythropoietin (EPO; Janssen-Ciliag, Bucks, United
Kingdom), at a concentration of 105BMMCs or 3 ? 103CD34?cells/mL
of culture medium. Cultures were set up in duplicate in 35-mm petri dishes
and incubated at a 37°C, 5% CO2, fully humidified atmosphere. On day 14,
colonies were scored and classified as granulocyte colony-forming units
(CFU-G), macrophage colony-forming units (CFU-M), granulocyte-
macrophage colony-forming units (CFU-GM), erythroid burst-forming
units (BFU-E) and colonies containing both granulocyte-macrophage and
erythroid elements (CFU-GEM) according to established criteria.22Results
were expressed as total numbers of CFU-GM (CFU-G plus CFU-M plus
CFU-GM) and total numbers of BFU-E (BFU-E plus CFU-GEM).
Long-term bone marrow cultures
Long-term bone marrow cultures (LTBMCs) from 107BMMCs were grown
according to the standard technique23,24in 10 mL IMDM supplemented
with 10% FCS, 10% horse serum (Gibco BRL Life Technologies), 100
IU/mL PS, 2 mmol L-glutamine and 10?6mol hydrocortisone sodium
succinate (Sigma), and incubated at a 33°C, 5% CO2, fully humidified
atmosphere. At weekly intervals, cultures were examined for stromal layer
formation using an inverted microscope and fed by removing half of the
medium and replacing it with equal volume of fresh IMDM supplemented
as above. Nonadherent cells were counted and assayed for CFU-GM and
BFU-E committed progenitor cells as described above. Colony results were
expressed as total numbers of colony-forming cells (CFCs) (CFU-GM plus
BFU-E). When mentioned, a mouse antihuman TNF? monoclonal neutral-
izing antibody (R&D Systems) was added weekly to the cultures at a
concentration of 1.8 ?g/mL.According to the manufacturer, the neutraliza-
tion dose50for this antibody is 0.02 ?g/mLto 0.04 ?g/mLin the presence of
0.25 ng/mL TNF?. On confluence (week 3-4), cell-free supernatants were
harvested and stored at ?70°C for TNF? quantification by means of a
commercially available enzyme-linked immunosorbent assay (ELISA) kit
(Biosource International, Camarillo, CA). According to the manufacturer,
the sensitivity of this assay is less than 0.09 pg/mL.
PROGENITORAND STROMALCELLS IN RA1611 BLOOD, 1 MARCH 2002?VOLUME 99, NUMBER 5
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Page 4

Table 1. Clinical and laboratory data of the patients studied
Patient
Age/Sex
Duration
(mo)
Arthritis
HAQ score
CRP
(mg/dL)
ESR
(mm/h)
RF
(mg/dL)
Peripheral blood counts†
Bone
marrow†
Medication‡
Hgb
(g/dL)
WBC
Neutro
Lympho
Mono
Plt
NSAID
MTX
PDN
Other
(DMARDs)
No. tender/swollen joints
(?109/L)
Cellularity
1
71/F
180
56/12
ND
8.6
61
166
11.1
9.7
7.1
1.8
0.6
252
Normal
No
Yes
5 mg/d
None
2
37/M
5
40/22
ND
4.6
50
153
14.2
8.4
5.7
1.8
0.8
255
Normal
Yes
No
No
None
3*
35/M
12
22/10
ND
0.6
2
55
15.0
6.4
3.4
2.2
0.5
244
Increased
No
20 mg/w
No
HQ
4
53/F
96
26/20
ND
0.7
42
293
12.2
5.7
3.6
1.6
0.4
210
Normal
No
No
No
HQ
5*
55/F
24
55/37
0.600
0.9
20
155
12.0
6.9
6.2
0.5
0.1
239
Increased
No
20 mg/w
10 mg/d
No
6*
55/M
276
27/20
1.750
0.6
42
137
12.4
8.6
6.1
1.7
0.6
383
Increased
No
15 mg/w
No
CSA
7*
58/F
60
58/50
1.125
0.6
58
51
14.1
9.3
4.2
4.1
0.7
458
Normal
No
15 mg/w
No
SSZ
8*
30/F
10
39/39
0.375
0.1
45
20
12.6
7.6
5.5
1.4
0.4
400
Normal
No
20 mg/w
No
SSZ, HQ
9
67/F
96
49/47
1.500
0.8
41
41
11.4
8.4
5.0
2.5
0.7
383
Decreased
No
20 mg/w
No
SSZ
10*
55/F
48
57/54
1.250
1.2
17
37
14.2
6.6
4.6
1.3
0.5
108
Increased
No
15 mg/w
No
SSZ
11
44/F
240
53/46
0.500
1.3
45
43
13.0
6.8
3.8
2.4
0.4
256
Normal
No
15 mg/w
No
SSZ, HQ
12
49/F
120
58/55
1.250
0.4
73
49
10.4
4.8
2.1
2.0
0.5
371
Normal
No
20 mg/w
No
SSZ
13
60/M
96
8/8
ND
0.4
24
51
13.3
8.0
5.8
1.5
0.6
363
Normal
No
15 mg/w
No
No
14
31/M
120
32/31
1.250
0.7
43
50
10.1
7.2
2.6
1.5
1.9
153
Decreased
No
15 mg/w
No
CSA
15*
54/M
24
43/31
0.375
0.2
67
21
9.9
7.4
4.7
1.7
0.8
430
Increased
No
15 mg/w
No
SSZ
16*
54/F
96
55/38
1.250
0.8
55
31
11.3
4.6
3.1
1.1
0.3
320
Decreased
No
15 mg/w
No
SSZ, HQ
17
49/F
12
62/21
0.250
6.3
62
292
11.3
8.6
5.9
2.0
0.3
292
Normal
Yes
No
No
None
18*
62/F
22
40/22
0.750
0.8
69
111
12.5
7.8
5.0
2.2
0.5
293
Normal
No
15 mg/w
No
SSZ, HQ
19*
33/F
84
36/26
0.250
0.8
58
230
11.8
9.8
7.1
1.7
0.9
221
Increased
No
15 mg/w
No
None
20*
68/F
72
22/18
1.375
1.2
23
51
12.5
6.8
3.5
2.4
0.7
253
Decreased
No
15 mg/w
5 mg/d
SSZ, HQ
21
71/F
120
21/19
ND
0.3
21
68
13.7
5.2
3.5
1.2
0.3
229
Decreased
No
15 mg/w
No
None
22
49/M
6
40/12
0.250
1.1
128
786
15.0
6.3
3.5
2.4
0.4
185
Normal
Yes
No
No
None
23
47/F
6
35/31
0.250
1.6
33
25
11.3
3.0
1.5
1.3
0.2
186
Normal
Yes
No
No
None
24
28/F
12
33/20
0.375
1.2
85
110
12.3
5.0
2.8
1.6
0.6
220
Normal
Yes
No
No
None
25
35/M
10
22/12
0.375
0.8
43
137
13.0
6.8
3.1
2.4
0.8
250
Normal
Yes
No
No
None
26*
65/F
240
50/45
1.375
0.9
85
155
11.2
5.7
3.6
1.6
0.5
125
Decreased
No
15 mg/w
5 mg/d
None
HAQ indicates Health Assessment Questionnaire; CRP, C-reactive protein; ESR, erythrocyte sedimentation rate; RF, rheumatoid factor; Hgb, hemoglobin; WBC, white blood cells; Neutro, neutrophils; Lympho, lymphocytes; Plt,
platelets; Mono, monocytes; ND, not done; NSAID, nonsteroidal anti-inflammatory drugs; MTX, methotrexate; PDN, prednizone; DMARDs, disease-modifying antirheumatic drugs; HQ, hydroxychloroquine (400 mg/d); SSZ, sulfasalazine-A
(2 g/d); CSA, cyclosporin-A(150 mg/d).
*These patients were also studied 6 to 8 months after initiation of anti-TNF? treatment.
†At the time of study.‡During the last 6 months.
1612 PAPADAKI et alBLOOD, 1 MARCH 2002?VOLUME 99, NUMBER 5
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Page 5

Assessment of BM stromal cell function
A 2-stage culture procedure was used to test the capacity of BM stromal
layers to support normal hematopoiesis. Confluent stromal layers from
patients and healthy controls, grown in standard LTBMCs, were irradiated
(10 Gy) and recharged with 5 ? 104normal allogeneic CD34?BM cells as
previously described.25In each experiment, flasks were recharged in
triplicate and the CD34?cells from the same healthy control were used to
test patient and healthy control cultures. Cultures were fed weekly by
demidepopulation and supernatants were monitored by determining the
total number of nonadherent cells and CFC frequency.
Statistical analysis
Data were analyzed in the GraphPad Prism statistical PC program (GraphPad
Software, San Diego, CA) by means of the nonparametric Mann-Whitney test,
the Student t test for paired samples, and the Pearson coefficient of correlation
test.Standard2-wayanalysisofvariancetestwasusedtodefinedifferencesinthe
number of nonadherent cells and the number of CFCs generated in LTBMCs in
thegroupstested.TheFvaluerepresentstheratioofvariancebetweenthegroups
and the total variance of values in the study. Statistically significant differences
for a certain F value are given in specific tables on the basis of the respective
degreesoffreedom.26
Results
Flow-cytometric data
The percentage of BM CD34?cells and their subpopulations in
patients and healthy controls are presented in Table 2. Patients with
RA displayed significantly lower proportions of CD34?cells
within the BMMCs, compared with the controls (P ? .0016). This
decrease probably reflected the reduction of the committed CD34?/
CD38?cells (P ? .0017), because no statistically significant
difference was found between patients and healthy controls in the
number of the more primitive CD34?/CD38?cells (P ? .1890).
To explore whether the decrease of CD34?cells in patients with
RA was due to increased apoptotic cell death, we evaluated the
proportion of apoptotic cells within the CD34?cell fraction using
7AAD staining (Table 2). We found that patient CD34?cells
contained significantly higher numbers of apoptotic (7AADdim)
cells compared with the controls (P ? .0136). In contrast, no
statistically significant difference was found between patients
(6.96% ? 5.86%, n ? 23) and control subjects (6.69% ? 5.61%,
n ? 24) in the percentage of apoptotic cells detected in the
non-CD34?BMMC fraction (P ? .6630).
Because Fas antigen expression has been associated with
apoptosis of BM hematopoietic progenitor cells,27we evaluated the
expression of this molecule on patients’CD34?cells (Table 2). We
found that the proportion of Fas?cells in the gate of CD34?cells
was significantly higher in the group of patients than in the group of
controls (P ? .0210). A highly significant positive correlation was
noted between the proportion of Fas?cells and the percentage of
apoptotic cells within the CD34?cell population in the entire group
of subjects studied (r ? 0.611, P ? .0001). Moreover, the in-
creased apoptosis found in patients’CD34?cell compartments was
due mainly to the increased proportion of apoptotic cells within the
CD34?/Fas?cell fraction (P ? .0310), because no statistically
significant difference could be demonstrated between patients and
healthy controls in the proportion of apoptotic cells within the
CD34?/Fas?cell fraction (P ? .2940). Notably, the percentage of
apoptotic cells was significantly higher among the CD34?/Fas?
cells than among the CD34?/Fas?cells in both the patient
(P ? .0001) and control (P ? .0001) groups. Taken together, these
data suggest that Fas antigen expression may account for the
increased apoptosis detected in patients’ CD34?cell compart-
ments. Moreover, a highly significant inverse correlation was noted
between the percentage of CD34?cells and the proportion of both
Fas?
cells (r ? ?0.462, P ? .0018) and apoptotic cells
(r ? ?0.326, P ? .0375), indicating that Fas antigen expression
and increased apoptosis are probably involved in the aforemen-
tioned reduction of CD34?cells in patients with RA.
Because the majority of the patients had been previously treated
with cytotoxic and/or immune suppressant agents, a subset analysis
in the number of CD34?, Fas?, and apoptotic cells in the groups of
previously treated (n ? 20) and untreated (n ? 6) patients was
performed, to exclude the possibility of drug-induced damage.
Statistical analysis was similar in treated and untreated patients
(Table 2), suggesting that drug-induced damage of patient progeni-
tor cells is unlikely.
Colony-forming cells
The frequency of clonogenic progenitor cells in the BMMC
fraction of patients with RA(n ? 25) and healthy controls (n ? 25)
is depicted in Figure 2. The mean number of CFU-GM and BFU-E
obtained by 107BMMCs was significantly lower in the patients
(4648 ? 2197 and 1923 ? 1286, respectively) compared with the
controls (7680 ? 3603 and 4196 ? 1859, respectively; P ? .002
and P ? .0001 respectively). Further analysis showed that com-
pared with the healthy controls, the numbers of CFU-GM and
BFU-E were significantly lower in both the previously treated
(n ? 19) (5284 ? 2079 per 107BMMCs; P ? .0007 and 2169 ?
1368 per 107BMMCs; P ? .0005, respectively) and untreated
Figure 1. Flow cytometric analysis of normal BMMCs stained with anti-CD34
antibody, anti-Fas (CD95) antibody, and 7AAD. (A) Scattergram of forward light
scatter (FSC) versus right-angle light scatter (SSC), to allow gating on the BMMCs
(low FSC and low SSC properties) (R1). (B) Scattergram of anti-CD34 fluorescence
versus SSC gated on R1, to allow gating on CD34?cells (R2). (C) Scattergram of
FSC versus 7AAD fluorescence gated on R2, showing 7AADbright(dead), 7AADdim
(apoptotic), and 7AAD?(live) cells. (D) Scattergram of anti-CD34 versus anti-Fas
fluorescence gated on R2, showing Fas?and Fas?CD34?cells. Similar staining
patterns were obtained for RABM samples.
PROGENITORAND STROMALCELLS IN RA 1613BLOOD, 1 MARCH 2002?VOLUME 99, NUMBER 5
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