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CLINICAL OBSERVATIONS, INTERVENTIONS, AND THERAPEUTIC TRIALS
From the First Department of Internal Medicine,
Fukushima Medical University, Fukushima; Shionogi Biomedical
Laboratories, Osaka; Department of Clinical Laboratory Science, Osaka
University Medical School, Osaka; and Otsuka Assay Laboratories, Tokyo,
Japan.
It is unclear how a paroxysmal nocturnal hemoglobinuria (PNH) clone
expands in bone marrow, although immune mechanisms involving cytotoxic
T lymphocytes, autosomal proliferation, and apoptosis resistance have
been hypothesized. To clarify aspects of immune mechanisms and
proliferation of PNH cells, we investigated HLA-DRB1, -DQA1, and -DQB1
alleles by polymerase chain reaction (PCR)-based genotyping and
expression of the Wilms' tumor gene, WT1, by
real-time reverse transcriptase-PCR (RT-PCR) in 21 PNH and 21 aplastic
anemia (AA) patients. HLA genotyping indicated that the frequency of DRB1*1501, DQA1*0102, and DQB1*0602 alleles in PNH patients and of
DQB1*0602 allele in AA patients was significantly higher than in 916 Japanese controls, and that the HLA-DRB1*1501-DQA1*0102-DQB1*0602 haplotype, found in 13 of 21 PNH patients, 5 of 7 AA-PNH syndrome patients, and 7 of 21 AA patients showed significant differences compared with healthy individuals. RT-PCR analysis showed that the mean
values of WT1 RNA were 3413, 712, and 334 copies/µg RNA in PNH, AA,
and healthy individuals, respectively. The values for PNH patients were
significantly higher than for AA patients and healthy volunteers and
were correlated with the proportion of CD16b Recently, paroxysmal nocturnal hemoglobinuria (PNH)
has been considered to be an acquired stem cell disorder affecting all hematopoietic lineages, which lack glycosylphosphatidylinositol (GPI)-anchored membrane proteins because of abnormalities in the phosphatidylinositol glycan-class A (PIG-A) gene.1 It is
well known that deficiencies in CD55 and CD59 expression on
erythrocytes cause complement-mediated hemolysis in
PNH.2,3 It has been hypothesized that expansion of a PNH
clone with a PIG-A gene abnormality is associated with an immunologic
mechanism involving cytotoxic T cells,4 autosomal
proliferation of a PNH clone,5 and resistance to apoptosis
in a PNH clone.6 Some reports7,8 support the
concept that loss of GPI-anchored proteins results in less efficient
killing of GPI-deficient hematopoietic target cells by putative
autoimmune T-cell clones, causing expansion of a PNH clone. In
addition, some researchers also reported abnormalities of T cells
derived from PNH patients.9,10 Nevertheless, a wide
variety of immunologic assays have not demonstrated a difference between normal and PIG-A-mutated target cells.11 Although
resistance of PNH cells to apoptosis has been
reported,12,13 this is a property of both GPI-deficient
cells with PIG-A gene abnormalities and GPI-positive cells, suggesting
that apoptosis resistance does not underlie the expansion only of PNH
clones. In fact, it is well known that there is no known molecular
mechanism by which GPI-anchored proteins influence apoptosis, and there
are conflicting data concerning the apoptosis resistance of PNH
cells.6,11,14 Moreover, in chimeric knockout mice, the
cells without the PIG-A gene constitute a minor, static fraction of the
bone marrow or circulating hematopoietic compartments, and
PIG-A-negative and normal cells behave similarly in paired tissue
culture assays,5,15 negating autosomal proliferation of a
PNH clone. At present, it is unclear how a PNH clone in bone
marrow expands.
The transformation of aplastic anemia (AA) into PNH is well known as
the AA-PNH syndrome, described by Lewis and Dacie.16 Recently, several reports17-22 have indicated that PNH
eventually develops in 29% to 45% of patients with AA who have been
treated with immunosuppressive agents and/or androgens and that some
patients with PNH respond to immunosuppressive therapy. These facts
suggest an immunologic linkage between AA and PNH, with the common
feature of bone marrow failure.4 Several studies on HLA
class I and II antigens and alleles in AA patients have been
reported,23-27 but the relationship between AA and HLA
class I and II antigens or alleles is controversial because of
conflicting data, mainly due to differences in statistical
analysis.28 Ustariz et al29 reported that
there was no significant increase in any of the 26 HLA antigens
corresponding to loci A and B in 10 Cuban patients with PNH.
Nevertheless, from many clinical and basic studies of AA patients, it
is certain that cytotoxic T lymphocytes and/or helper/inducer T
lymphocytes are implicated in some of the immune mechanisms involved in
the occurrence of AA.28 Recently, Zeng et al30
found sequence identity for complementarity determining region 3 among
a majority of CD4 clones from a patient with AA, from which the
dominant clone showed a Th1 secretion pattern, lysed autologous
CD34+ cells, and inhibited their hematopoietic colony
formation. Thus, we considered the possibility that HLA class II
alleles and haplotypes may be related to the immunologic pathogenesis
of AA and PNH because it is known that antigen-specific T
helper/inducer cells, when present at very low levels, are capable of
rapid clonal expansion during antigenic challenge, and the specificity
of this response provides for the activation and expansion of a very
select cohort of T cells.31
The Wilms' tumor gene, WT1, is responsible for the
childhood renal neoplasm, Wilms' tumor, and is located at chromosome
11p13.32 Although the kind of hematopoietic cells
expressing WT1 RNA in normal bone marrow is controversial, it is
generally believed that the WT1 gene is expressed in
CD34+ hematopoietic progenitor cells, but not in
CD34 In the present study, to clarify some characteristics in the
pathophysiology of PNH and AA, we investigated HLA-DRB1, -DQA1, and
-DQB1 alleles and haplotypes, and WT1 RNA expression in bone marrow, in
21 PNH patients and 21 AA patients, including 2 with pure red cell aplasia.
Patients and controls
As controls, peripheral blood samples were obtained after informed
consent from 20 healthy volunteers for single-color flow cytometric
analysis of CD59 expression on erythrocytes and CD16b expression on
granulocytes, and bone marrow samples were obtained from an additional
20 healthy volunteers for quantitation of WT1 RNA expression by
real-time reverse transcriptase-polymerase chain reaction (RT-PCR). In
addition, peripheral blood and bone marrow samples were taken from 21 AA patients, including 2 with pure red cell aplasia. Based on the
criteria of the Research Committee of Idiopathic Hematopoietic
Disturbances of the Ministry of Health and Welfare of
Japan,50 the diagnosis and grading of the severity of AA
were determined by peripheral blood and/or bone marrow cytomorphologic findings following exclusion of the other disorders presenting with
pancytopenia. Severe, moderate, or mild AA was established when at
least 2 of the following criteria were met: absolute numbers of
reticulocytes, neutrophils, or platelets less than
20 × 109/L, 0.5 × 109/L, or
20 × 109/L; less than 60 × 109/L,
1 × 109/L, or 50 × 109/L; or more than
60 × 109/L, 1 × 109/L, or
50 × 109/L, respectively.
Clinical and hematologic parameters
CD59 expression on erythrocytes and CD16b expression on granulocytes by flow cytometry Immunofluorescent staining and flow cytometric analysis of CD59 expression on erythrocytes from all the patients and 20 healthy volunteers were performed as described previously.3,47 Analysis of CD16b expression on granulocytes was also performed with cells from 21 PNH patients and healthy volunteers as described previously.51 Mouse monoclonal antibodies to CD59 (3E1, IgG1)52 and to CD16b (ID3, IgM, ) labeled with
fluorescein isothiocyanate (Immunotech, Marseille, France) were used,
and irrelevant monoclonal antibodies of the same subclass were used as
negative controls.3,47,51
Analyses of HLA-DRB1, -DQA1, and -DQB1 alleles and haplotypes High-molecular-weight DNA was extracted from leukocytes of patients by standard methods.53 The second exons of the HLA-DRB1 and -DQB1 genes and the first 3 exons of the HLA-DQA1 gene were amplified using PCR with group-specific primers and sequence-specific primers, respectively. Each allele was typed using the restriction fragment length polymorphism method and/or the PCR sequence-specific primers method, as described previously,54-56 with the modification of adding several restriction enzymes or several primers to detect the DRB1 and DQB1 alleles or the DQA1 alleles, respectively, the nomenclature of which was officially updated by the World Health Organization Nomenclature Committee. An HLA class II haplotype in these patients was determined on the basis of the frequency of HLA class II haplotypes reported in unrelated Japanese individuals.57,58 The frequency of an HLA class II haplotype in these patients was compared with that in a control population consisting of 916 unrelated Japanese individuals.57 This control population was considered valid because both populations originated from the same area in Japan.Quantitation of WT1 RNA by RT-PCR WT1 RNA levels in bone marrow samples from 21 PNH and AA patients were quantified by RT-PCR. Total RNA was extracted from bone marrow mononuclear cells using a QIAamp RNA Blood Mini Kit (Qiagen GmbH, Hiden, Germany) according to the manufacturer's instruction. Two micrograms of total RNA was converted into cDNA in 45 µL reaction mixture containing 200 units of reverse-transcriptase SuperScript II (Gibco BRL, Rockville, MD), 1 mM of each deoxynucleotide triphosphate, 500 ng random hexamers (Gibco BRL), and 40 U RNase inhibitor (Gibco BRL). RT-PCR59,60 was performed in a MicroAmp optical 96-well plate with 2.3 µL of the above cDNA solution, 7.5 pmol forward and reverse primers, 5 pmol TaqMan probe, and 12.5 µL 2X TaqMan Universal PCR Master Mix (PE Biosystems, Foster City, CA). The sequences of forward and reverse primers and TaqMan probe for the quantitation of the expression levels of the WT1 and GAPDH genes were as follows: the WT1 gene, 5'-GATAACCACACAACGCCCATC-3' (forward primer), 5'-CACACGTCGCACATCCTGAAT-3' (reverse primer), 5'-FAM-ACACCGTGCGTGTGTATTCTGTATTGG-TAMRA-3' (TaqMan probe); the glyceraldehyde-3-phosphate dehydrogenase (GAPDH) gene, 5'-CACCAGGGCTGCTTTTAACTC-3' (forward primer), 5'-GAAGATGGTGATGGGATTTC-3' (reverse primer), 5'-FAM-CATGGGTGGAATCATATTGGAA-TAMRA-3' (TaqMan probe). The above reaction mixture was heated at 50°C for 2 minutes and then at 95°C for 10 minutes to activate the polymerase. PCR was then performed using an ABI Prism 7700 Sequence Detector System (PE Applied Biosystems) with 45 cycles, one of which consisted of denaturation at 95°C for 15 seconds, and annealing and extension at 60°C for 1 minute. Known copy numbers of WT1 cDNA (102, 104, and 106 copies/well) and GAPDH cDNA (104, 106, and 108 copies/well) were simultaneously amplified by PCR to make standard curves for measuring WT1 and GAPDH expression levels in the samples. These standard cDNAs were prepared by cloning the PCR-amplified cDNA fragments into PCR II vector (Invitrogen, Carlsbad, CA), amplifying the vector, and cutting out the cDNA fragments from the vector. The DNA contents of standard cDNA solutions were quantitated by their absorbance at 260 nm, and copy numbers were calculated by dividing the amounts of DNA by molecular weights. Copy numbers per microgram RNA of WT1 and GAPDH RNA in the patients' samples were calculated using a standard curve. To normalize the differences in RNA loading for RT-PCR and in RNA degradation for individual samples, the values (WT1/GAPDH) of the WT1 RNA levels (copy number/µg RNA) divided by the GAPDH RNA levels (copy number/µg RNA) were calculated according to a procedure described previously,61 and (WT1/GAPDH) × (6.1 × 109) (the mean copy numbers/µg RNA of GAPDH in 47 normal human peripheral blood mononuclear cells, which were determined to have the corrected values of WT1/GAPDH using copy numbers) were defined as copy numbers/µg RNA of WT1 RNA in the samples. To check the reproducibility of the precision of the WT1 RNA assay, we simultaneously PCR-amplified 103 copies of GAPDH and WT1 cDNA with the samples and confirmed with every WT1 RNA assay that their copy numbers were within the mean ± 2 SDs.Statistical analysis The various hematologic and laboratory parameters of the 2 disorders were compared statistically by the Student t test. The proportions of CD59 erythrocytes among PNH
patients, AA patients, and healthy volunteers were examined
statistically by a one-way analysis of variance (Bonferroni/Dunn)
model. The proportions of CD16b granulocytes in PNH
patients and healthy volunteers were compared statistically by the
Student t test. Differences in HLA class II allele or
haplotype frequencies between patients with PNH or AA and
controls57 or between the disorders were calculated by  2 analysis or Fisher exact test. Various hematologic and
laboratory findings, including the proportions of CD59
erythrocytes and CD16b granulocytes, and the values for
WT1 RNA levels, in the HLA-DRB1*1501-DQA1*0102-DQB1*0602 haplotype
group and another haplotype group in the PNH patients were compared
statistically by the Student t test. The WT1 RNA levels
quantitated by RT-PCR were investigated statistically among PNH
patients, AA patients, and healthy volunteers by a one-way analysis of
variance (Bonferroni/Dunn) model. In addition, the relationship between
the values for WT1 RNA and the proportion of CD59
erythrocytes or CD16b granulocytes in 21 PNH patients was
analyzed using correlation coefficients.
Clinical, hematologic, and laboratory findings in patients with PNH and AA Male-to-female ratio and mean age of the AA patients were 11:10 and 56.7 years (range, 18-80 years), respectively. Among 19 AA patients, there were 3 severe, 6 moderate, and 10 mild cases of disease at the time of examination. The clinical, hematologic, and laboratory findings described in "Materials and methods" at the time of examination were compared between patients with PNH and AA. The reticulocyte counts (mean, 106 × 109/L; range, 20-261 × 109/L) and LDH (1506 IU/L; range, 348-6070 IU/L) in PNH patients (Table 1) were significantly higher than those in AA patients (50 × 109/L; range, 3-116 × 109/L; P < .001; and 413 IU/L; range, 171-697 IU/L; P < .002, respectively), but there were no differences between them in the other parameters. The percentages of 21 PNH and AA patients with absolute neutrophil counts below 0.5 × 109/L or reticulocyte counts greater than 50 × 109/L were 9.5% and 9.5% or 85.7% and 47.6%, respectively.The treatment and transfusion regimens of the PNH patients are summarized in Table 1. Twelve of 21 AA patients were treated with immunosuppressive therapy using antithymocyte globulin and/or cyclosporine prior to or at the time of examination. Three of 12 patients received granulocyte colony-stimulating factor (G-CSF) in addition to immunosuppressive therapy, and one received oxymetholone at the time of examination. Six of 21 AA patients received prednisolone and/or androgens, including metenolone and fluoxymesterone, and one of them also received G-CSF at the time of examination. Two and one of the 21 patients received only G-CSF or blood transfusions at the time of examination, respectively. CD59 expression on erythrocytes and CD16b expression on granulocytes in patients with PNH and AA The phenotypes and proportions of each population of erythrocytes in 21 PNH patients are summarized in Table 1. Flow cytometric profiles of CD59 expression on erythrocytes from 21 AA patients and 20 healthy volunteers showed that the erythrocytes consisted of a single positive population. The mean proportions of negative populations determined by flow cytometry were 19.0% (range, 1.4% to 86.9%), 0.43% (0.07% to 0.87%), and 0.39% (0.10% to 0.80%) in 20 PNH, 21 AA, and 20 healthy individuals, respectively. The proportion of CD59 populations in PNH patients was higher than that in
AA (P < .0001) and healthy individuals
(P < .0001). The proportion of CD16b
granulocytes in 21 PNH patients is summarized in Table 1. It was
significantly higher in the PNH patients (mean ± SD,
57.5% ± 39.8%) than in the healthy volunteers (0.46% ± 0.38%,
P < .001). Flow cytometric profiles of CD16b expression
on granulocytes from 20 healthy volunteers showed a single
positive population.
HLA-DRB1, -DQA1, and -DQB1 alleles and haplotypes in patients with PNH and AA The frequencies of the HLA-DRB1, -DQA1, and -DQB1 alleles in 21 PNH and AA patients are presented in Table 2. The frequencies of each allele were statistically compared between the disorders. The frequency of all the alleles did not show significant differences between PNH and AA patients. In addition, the frequency of each HLA-DRB1, -DQA1, and -DQB1 allele in the 2 disorders was statistically compared with that in 916 Japanese individuals.57 The frequencies of HLA-DRB1*1501, -DQA1*0102, and -DQB1*0602 alleles in PNH patients and of HLA-DQB1*0602 allele in AA patients were significantly different than in controls (P < .005, P < .005, P < .005, and P < .025, respectively).
The HLA class II haplotypes in 21 PNH patients are presented in Table
3. Thirteen of 21 PNH patients showed the
DRB1*1501-DQA1*0102-DQB1*0602 haplotype. Five of 7 AA-PNH syndrome
patients and 8 of the 14 primary PNH patients showed this haplotype. In
contrast, the same haplotype was found in 7 of 21 AA patients. The
frequency of this haplotype in 21 PNH patients was significantly higher
than in Japanese healthy volunteers
(P < .005).57 The frequency of this haplotype in patients with primary PNH and AA-PNH syndrome was also
higher than in healthy controls (P < .005 and
P < .005, respectively). In addition, various
hematologic and laboratory findings, including the proportions of
CD59
Quantitation of WT1 RNA in patients with PNH and AA The results of RT-PCR analysis of WT1 RNA levels in bone marrow mononuclear cells from 21 PNH and AA patients and 20 healthy volunteers are presented in Figure 1, and the values for the PNH patients are shown in Table 3. The values were statistically compared among PNH patients, AA patients, and healthy individuals. The values in PNH patients (mean ± SD, 3413 ± 5149 copies/µg RNA) were significantly higher than in AA patients (712 ± 647 copies/µg RNA; P < .005) and in healthy individuals (333 ± 170 copies/µg RNA; P < .002). In contrast, the values in patients with AA did not show significant differences compared with those in healthy volunteers. Five AA patients with more than 1000 copies/µg of WT1 RNA had no specific clinical and hematologic features, including response to immunosuppressive therapy, compared with the other AA patients. In addition, the relationship between the WT1 RNA levels and the proportion of CD59 erythrocytes or CD16b
granulocytes in 21 PNH patients was statistically analyzed. The WT1 RNA
levels were significantly correlated with the proportion of
CD16b granulocytes (r = 0.5207,
P < .02), but not with that of CD59
erythrocytes.
To our knowledge, this is the first investigation of HLA class II
alleles or haplotypes in patients with PNH. We found that HLA-DRB1*1501, -DQA1*0102, and -DQB1*0602 alleles and
HLA-DRB1*1501-DQA1*0102-DQB1*0602 haplotype or HLA-DQB1*0602 allele and
HLA-DRB1*1501-DQA1*0102-DQB1*0602 haplotype, determining the
presentation of HLA-DR2 in the Japanese, were characteristic in
Japanese patients with PNH or AA, respectively, compared with those in
healthy individuals. Nakao et al62 reported that the
cyclosporine-dependent response of AA is closely related to an HLA
class II haplotype of DRB1*1501, DQA1*0102, and DQB1*0602. In addition,
the frequency of this haplotype found in 5 of 7 AA-PNH syndrome
patients in our study appears to be similar to the frequency of
occurrence of GPI-deficient cells in AA patients.63 These facts strongly suggest that a linkage exists in the pathophysiology or
pathogenesis between PNH and AA. They also suggest that immune mechanisms in an HLA-restricted manner play an important role in the
pathogenesis of primary PNH and AA-PNH syndrome, especially a
relationship between the immune mechanisms involving cytotoxic T
lymphocytes and negative selection of a PNH clone with a PIG-A gene
mutation rather than proliferation of a PNH
clone.4,11 Therefore, the specific haplotype in
PNH patients was not related to the proportion of CD59 We found that the expression of WT1 RNA in 21 PNH patients was
significantly higher than in AA patients and healthy individuals. Recently, there has been increasing evidence that WT1 is
strongly expressed in normal CD34+ bone marrow stem cells
but is only weakly expressed or not expressed in normal mature blood
cells.33-35 Moreover, a more recent report66 indicated that WT1 expression in human bone marrow and
peripheral blood cells is biphasic. WT1 RNA is present in
CD34+CD38 Subsequently, we found that the levels of WT1 RNA in bone marrow cells
from our PNH patients were correlated with the proportion of
CD16b In conclusion, the HLA-DRB1*1501-DQA1*0102-DQB1*0602 haplotype was
found with high frequency in PNH patients, which suggests that immune
mechanisms in an HLA-restricted manner play an important role in the
pathogenesis of PNH and AA-PNH syndrome. In addition, the higher
amounts of WT1 RNA in PNH patients compared with AA patients and
healthy individuals were correlated with CD16
We are grateful to Dr Hideyoshi Noji, Dr Kazuei Ogawa, and Dr Toshiyuki Ishibashi (Fukushima Medical University, Japan), Dr Yoshiyuki Kamiyama, Dr Yurie Saitoh, Dr Hiroyuki Kambayashi, Dr Tetsugoroh Tanaka, and Dr Shin Matsuda (Ohta Nishino-uchi Hospital, Japan), Dr Yutaka Shiga and Dr Hideo Kimura (General Hobara-chuoh Hospital, Japan), Dr Rokuo Abe (Fukushima-ken Taiyo-no-kuni Hospital, Japan), Dr Masayuki Mita (Hoshi General Hospital, Japan), Dr Kenichi Nakamura (Shirakawa-Kohsei Hospital, Japan), Dr Toshiaki Sai (Iwaki-Kyoritsu Hospital, Japan), Dr Junichi Kameoka (Tohoku University, Japan), Dr Kazuyasu Endoh (Sendai-City Hospital, Japan), and Dr Kyoko Kaneda (National Miyagi Hospital, Japan) for providing the samples from patients with PNH and AA. Also, we wish to thank Dr Yuuji Sugita (Showa University, Japan), who provided the monoclonal antibody to CD59/membrane attack complex-inhibitory factor.
Submitted June 20, 2001; accepted February 5, 2002.
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.
Reprints: Yukio Maruyama, First Department of Internal Medicine, Fukushima Medical University, 1 Hikariga-oka, Fukushima, Fukushima 960-1295, Japan; e-mail: t-shichi{at}fmu.ac.jp.
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Abstract
Introduction