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Blood, Vol. 94 No. 11 (December 1), 1999:
pp. 3872-3882
By
From the Division of Research Immunology and Bone Marrow
Transplantation, Childrens Hospital Los Angeles, Los Angeles, CA.
Wiskott-Aldrich syndrome (WAS) is an X-linked recessive disorder
characterized by thrombocytopenia, eczema, and a progressive deterioration of immune function. WAS is caused by mutations in an
intracellular protein, WASP, that is involved in signal transduction and regulation of actin cytoskeleton rearrangement. Because immune dysfunction in WAS may be due to an accelerated destruction of lymphocytes, we examined the susceptibility to apoptosis of resting primary lymphocytes isolated from WAS patients in the absence of
exogenous apoptogenic stimulation. We found that unstimulated WAS
lymphocytes underwent spontaneous apoptosis at a greater frequency than
unstimulated normal lymphocytes. Coincident with increased apoptotic
susceptibility, WAS lymphocytes had markedly attenuated Bcl-2
expression, whereas Bax expression did not differ. A negative correlation between the frequency of spontaneous apoptosis and the
level of Bcl-2 expression was demonstrated. These data indicate that
accelerated lymphocyte destruction by spontaneous induction of
apoptosis may be one pathogenic mechanism by which the progressive immunodeficiency in WAS patients develops.
WISKOTT-ALDRICH SYNDROME (WAS) is a rare
X-linked recessive disorder.1,2 Affected males are
clinically characterized by eczema, thrombocytopenia with platelets of
reduced size, immunodeficiency, and an increased susceptibility to
hematopoietic malignancy and autoimmunity (Remold-O'Donnell et
al3 and references therein). WAS patients suffer from
chronic and recurrent opportunistic and viral infections as a
consequence of immunologic abnormalities. A progressive deterioration
of T-lymphocyte function in affected children accompanies the
development of T lymphopenia by 6 years of age.4
Cell-mediated immunity is further compromised by poor macrophage and
neutrophil motility and chemotactic responses.4-7 WAS
patients commonly have low isohemagglutinin titers and a depressed response to polysaccharide antigens3; but, their impaired
humoral immunity has been at least partially attributed to T-lymphocyte dysfunction.8 WAS T lymphocytes have diminished
proliferative responses to mitogens, some specific antigens, and
allogeneic stimulation.3 Moreover, WAS T lymphocytes fail
to proliferate or upregulate interleukin-2 gene expression in response
to treatment with immobilized anti-CD3 antibody.9,10
Deformation and deregulation of the actin cytoskeleton is likely to be
the basis for many of the clinical defects associated with WAS,
particularly immune dysfunction.3,10,11 WAS
lymphocytes are frequently morphologically abnormal, with irregular and
bulbous cellular projections,10,12 a paucity of
microvilli,13 and a poorly delineated actin
cortex.10 CD3-mediated stimulation of WAS T-lymphoblastoid
cell lines results in abnormal actin polymerization, marked by the
absence of specific cytoskeleton rearrangements that occur in normal
T-lymphoblastoid cell lines.10 Epstein-Barr
virus-transformed B-cell lines from WAS patients also have marked
abnormalities in actin distribution and polymerization.11
The WAS gene was mapped to the short arm of the X chromosome at
Xp11.2214 and identified in 1994.15 The gene
encodes a 66-kD intracellular protein, WASP, that is expressed
exclusively in blood cells15,16 throughout
hematopoiesis.17 More than 138 unique mutations to WASP
have been described,18 which include frameshifts and
substitutions that generally nullify expression by causing mRNA
instability and protein truncation.11,19-21 There is
considerable variation in WAS severity; even members of the same
kindred may have markedly different phenotypes,22 possibly indicating polygenic or environmental contributions. WAS severity may
be partially determined by the level at which WASP is stably expressed.11,19-21
The function of WASP is unknown; however, it has been surmised that
WASP is involved in the regulation of actin polymerization and actin
cytoskeleton organization from observations that WAS lymphocytes,
platelets, granulocytes, and monocytes are defective in regulating the
cortical actin cytoskeleton (Remold-O'Donnell et al3 and
references therein). In support of this presumption, N-WASP, a
ubiquitous mammalian homolog of WASP, has been demonstrated to directly
induce actin depolymerizaton and actin cytoskeleton reorganization in
nerve growth factor-stimulated cells.23 Although WASP
apparently lacks catalytic activity, the protein colocalizes with actin
and interacts with WIP24 and PSTPIP,25 2 proteins that regulate actin polymerization. The Rho family small
guanosine triphosphatases (GTPases), Cdc42 and Rac, which have been
shown to specifically control actin cytoskeleton
reorganization,26 interact with WASP.27-30 WASP
also associates with numerous other proteins, including various
tyrosine kinases (Btk, Itk, Tec, Fyn, and c-Src), adaptor proteins (Nck
and Grb2), and phospholipase C A WASP deficiency or the expression of dysfunctional WASP is probably
detrimental to the development and survival of hematopoietic cell
lineages.3 A uniformly nonrandom pattern of X-chromosome inactivation in all blood cells from WAS obligate heterozygous carriers and platelet loss in WAS patients suggests that
mutations to WASP probably confer a growth and/or survival disadvantage to affected cells.3 Consequently, we reasoned that the
development of lymphopenia and other progressive immune defects
associated with WAS may be caused by an inherently decreased potential
for survival and the accelerated destruction of peripheral lymphocytes.
In this study, we compared the levels of spontaneous apoptosis in
unstimulated resting lymphocytes isolated from WAS patients and from
normal, healthy controls. We found that WAS lymphocytes underwent
spontaneous apoptosis at a greater frequency than did normal
lymphocytes. Bcl-2 family members regulate the onset of apoptosis,
acting as either agonists or inhibitors of programmed cell
death.38 Because the deregulation of apoptosis in WAS
lymphocytes might be caused by aberrant expression of one or more of
the Bcl-2 family members, we determined that the level of Bcl-2
expression in WAS lymphocytes was attenuated. We suggest that the
accelerated destruction of lymphocytes by spontaneous apoptosis could
account for the progressive deterioration of immune function in WAS patients.
Patient samples.
Samples from 5 male patients previously diagnosed with WAS were used in
this study. At the time of analysis, patients no. 1 and 2 were 2.5 and
4 years of age, respectively. Patients no. 3 and 4 are siblings and
were 4.5 and 14 years of age, respectively. Patient no. 5 was 14 months
of age. Diagnosis of WAS was initially based on family history and
presentation of immunodeficiency with recurrent infection,
thrombocytopenia, platelets with reduced volume, and eczema. To define
the WAS phenotype of the patients, clinical scores were assigned to
each according to published recommendations.18,21 Patients
no. 1, 2, 3, 4, and 5 were assigned clinical scores of 4, 4, 3, 3, and
3, respectively. The genotypes of 4 of 5 patients have been determined.
Patient no. 1 has a point mutation, C290T, resulting in an amino acid
substitution of R86C in exon 2. The genotype of patient no. 2 was not
determined. Patients no. 3 and 4, the siblings, have a deletion
mutation in exon 8 (John Bastian, personal communication, July
1999). Patient no. 5 has a splice site mutation (4-bp
deletion in intron 8), resulting in the deletion of exon 8; patient no.
5 does not express WASP. Patients no. 1, 2, and 5 were treated at
Childrens Hospital (CHLA) in Los Angeles, CA; each has since received
bone marrow transplantations. Patients no. 3 and 4 were treated at
Children's Hospital in San Diego, CA. These studies were performed in
accord with protocols approved by the Committee on Clinical
Investigations of the Institutional Review Board at CHLA. None of the
patients had been splenectomized. Normal controls (8) were obtained
from either healthy volunteer adult donors (5) or healthy pediatric
patients (3) at CHLA. No significant difference between the normal
pediatric and normal adult means was noted (P = .978). Whenever possible, patient and normal blood was collected and
processed under paired conditions. For the comparison of normal
pediatric and normal adult blood, specimens were collected and
processed under paired conditions.
Isolation and in vitro treatment of peripheral blood lymphocytes
(PBL).
Peripheral blood was collected from WAS patients or normal, healthy
individuals in heparinized Vacutainer tubes (Becton Dickinson, Franklin
Lakes, NJ). Within 8 hours, mononuclear cells were isolated from whole
blood by Ficoll-Hypaque (Pharmacia Biotech, Piscataway, NJ) density
gradient centrifugation. Mononuclear cells were washed in Hanks'
balanced salt solution (HBSS; BioWhittaker, Walkersville, MD) and
resuspended in R-10 medium (RPMI-1640 supplemented with 10%
heat-inactivated human A+ serum [CHLA Blood Bank, Los
Angeles, CA], 2 mmol/L L-glutamine [Gemini BioProducts, Calabasas,
CA], 1 × 10 Detection of apoptosis.
Either of 2 methods, terminal deoxynucleotidyl transferase-mediated
dUTP nicked end labeling (TUNEL) or annexin V-labeling, was used to
mark apoptotic cells. For TUNEL,39 cells were harvested and
washed in 3 mL phosphate-buffered saline (PBS; Sigma). At least
105 cells were stained on ice and in the dark with
fluorochrome-conjugated antihuman antibodies recognizing specific cell
surface markers to be used in immunophenotyping. Incubations with the
antibodies were performed in PBS containing 0.1% human intravenous Ig
(Sandoz Pharmaceuticals, East Hanover, NJ) for 15 minutes. All
allophycocyanin- and phycoerythrin-conjugated antibodies were used
according to the supplier's recommendations (Becton Dickinson
Immunocytometry Systems [BDIS], San Jose, CA). Stained cells were
washed in 3 mL PBS and fixed in 0.5 mL 2% paraformaldehyde (Sigma) on
ice for 15 minutes. Cells were washed in 3 mL PBS and permeabilized in
0.5 mL PBS containing 0.5% Tween-20 (Sigma) and 0.2% bovine serum
albumin (BSA; fraction V; Sigma). Samples were mixed gently and placed
on ice for 15 minutes. Cells were washed in 3 mL PBS and resuspended in
50 µL TUNEL reaction mix containing 5 U terminal deoxynucleotidyl
transferase (TdT; Promega, Madison, WI), 1× TdT reaction buffer
(Promega; containing 100 mmol/L cacodylate buffer [pH 6.8], 1 mmol/L
cobalt chloride, and 0.1 mmol/L dithiothreitol), and 10 µmol/L
biotin-16-dUTP (Boehringer Mannheim Biochemicals, Indianapolis, IN).
Cells were incubated in the reaction mix at 37°C for 30 minutes.
Cells were washed in 3 mL PBS and resuspended in 100 µL labeling
solution containing 4× SSC, 0.1% Triton X-100 (Sigma), 10%
nonfat dry milk (Carnation, Glendale, CA), and 0.1% sodium azide (Sigma). Ten microliters of avidin-fluorescein (BDIS) was
mixed with the cells. Samples were kept at room temperature for 30 minutes in the dark. Cells were washed in 3 mL PBS containing 0.1%
Triton X-100. Cells were resuspended in 0.5 mL PBS containing 5 µg/mL
propidium iodide (PI; Sigma) and 0.1% DNase heat-inactivated RNase A
(Sigma). Negative control samples were treated in an identical manner
except that TdT was omitted from the TUNEL reaction mix. Labeled cells
were analyzed promptly by flow cytometry.
Determination of relative Bcl-2 expression levels.
The relative level of Bcl-2 expression in PBL was determined by
intracellular staining using a hamster antihuman monoclonal antibody,
an assay similar to that used by von Freeden-Jeffry et
al.41 Briefly, at least 105 cells were
harvested and stained with immunophenotyping antibodies, as described
above. Cells were washed in 3 mL PBS and fixed in 0.5 mL 2%
paraformaldehyde for 15 minutes on ice. Cells were washed in 3 mL PBS
and permeabilized in 200 µL PBS containing 0.3% saponin (Sigma) and
0.2% BSA. All subsequent steps were performed in this buffer. Samples
were placed on ice for 15 minutes. Two micrograms of anti-Bcl-2
antibody (clone 6C8; Pharmingen, San Diego, CA) was mixed with the
cells. The negative control for measuring background fluorescence was
prepared in a similar manner, except that 2 µg hamster IgG isotype
control polyclonal antibody (Pharmingen) was added to the cells.
Samples were placed on ice for 20 minutes. Cells were washed in 3 mL
PBS and resuspended in 100 µL of the buffer. One microgram of
fluorescein-conjugated mouse antihamster IgG antibody (clone G70-204;
Pharmingen) was mixed with the cells. Samples were placed on ice in the
dark for 20 minutes. Cells were washed in 3 mL PBS and resuspended in
0.5 mL 2% paraformaldehyde. Samples were stored at 4°C until
analysis was performed by flow cytometry.
Flow cytometry and data analysis.
Analysis of TUNEL samples was performed using a FACSVantage flow
cytometer (BDIS) equipped with an argon laser tuned to 488 nm and a
helium-neon (HeNe) laser tuned to 633 nm. A 610 nm short-pass splitter
was used to divert PI fluorescence to FL-3. After electronic compensation, FL-3 fluorescence was measured using both linear and
logarithmic amplification; FL-1 (fluorescein), FL-2 (phycoerythrin), and FL-4 (allophycocyanin) fluorescences were measured using
logarithmic amplification. The flow rate was not permitted to exceed
200 events per second. The data from at least 5 × 104
events were collected for analysis of freshly isolated cells. For all
other experiments, the data from at least 104 events were
collected. The data were analyzed using CELLQuest software (BDIS). For
analysis of TUNEL data, fragmented cells and debris were electronically
excluded. Another region was set to electronically eliminate multiplet
events from the analysis. A negative control sample (no TdT added) was
used to establish TUNEL-negative and -positive regions, such that at
least 99% of the events were in the lower TUNEL-negative region.
Background events were then subtracted from the TUNEL-positive events.
Statistical analysis of results.
Number Crunching Statistical Systems software (Dr Gerry L. Hintze,
Kayszille, UT) was used for the statistical analysis of data. Where
applicable, an unbalanced repeated measures analysis of variance
(ANOVA) was used to determine the significance of differences between
normal and WAS sample values. This method eliminates bias possibly
associated with analysis of unbalanced repeated measures. The TUNEL
data were also analyzed as a set of randomly chosen duplicate repeated
measures to test the validity of the first method of analysis. This
analysis showed that the first method was indeed valid, confirming the
statistical significance of the data. A probability level (P)
of WAS PBL undergo spontaneous apoptosis in vitro at a greater rate than
normal PBL.
The levels of spontaneous apoptosis in unstimulated, resting PBL were
determined in this study. To compare the frequency of apoptosis in WAS
PBL and normal PBL, we determined the fraction of cells undergoing
apoptosis by 2 quantitative methods. TUNEL39 and annexin
V-labeling40 measure oligonucleosomal DNA degradation and
externalization of phosphatidylserine, respectively, 2 hallmarks of
apoptosis.42,43 Flow cytometry was used to quantify marked cells undergoing apoptosis immediately after PBL isolation or after a
period of in vitro incubation. Importantly, PBL were not treated with
cytokines or any other mitogenic or antigenic stimuli and neither was
an extrinsic stimulus used to induce apoptosis.
Both WAS B and T lymphocytes are more susceptible to apoptosis than
normal B and T lymphocytes.
WAS patients have defects in both humoral and cell-mediated immunity;
thus, we sought to determine if both T and B lymphocytes have increased
susceptibility to spontaneous apoptosis. We measured the frequencies of
spontaneous apoptosis in resting T and B lymphocytes isolated from WAS
patients (no. 1 through 4) and normal controls after incubation in
vitro. Figure 3A shows that unstimulated
WAS CD3+ T lymphocytes undergo spontaneous apoptosis at a
greater rate than do unstimulated normal CD3+ T
lymphocytes. In vitro incubation augmented the level of spontaneous apoptosis in unstimulated T lymphocytes. Over time, the frequency of
spontaneous apoptosis in WAS T lymphocytes increased more rapidly than
occurred normally (P = .024). Unstimulated WAS
CD19+ B lymphocytes also underwent spontaneous apoptosis at
an increasing frequency during in vitro incubation (Fig 3B). The mean
level of spontaneous apoptosis in WAS B lymphocytes was higher than in
normal B lymphocytes at each time point. This trend was consistent, although statistical significance could not be demonstrated due to the
presence of very low numbers of circulating B lymphocytes in our WAS
patients. In vitro incubation also augmented spontaneous apoptosis in
unstimulated WAS B lymphocytes by 2-fold over normal.
Bcl-2 expression is attenuated in WAS PBL.
Because Bcl-2 family members have been shown to regulate apoptosis in
hematopoietic cells, the accelerated spontaneous apoptosis in WAS PBL
could result from deregulated expression of regulatory factors, such as
Bcl-2,45 that repress apoptosis. We examined the relative
levels of Bcl-2 expression in WAS and normal PBL by measuring the MFI
of cells stained indirectly with anti-Bcl-2 antibody in situ.
Measurements were made either immediately after isolation of PBL
(Table 1) or after in vitro incubation
(Table 2). WAS PBL from 5 different
patients had a significantly reduced level of Bcl-2 compared with
normal PBL from 8 different individuals (Fig 4). The relative levels of Bcl-2
expressed in isolated PBL did not change significantly over time during
in vitro incubation. Wide ranges in Bcl-2 expression are evident, both
in PBL isolated from WAS patients and from normal controls. Analysis
showed that PBL from patients no. 2 and 5 had Bcl-2 levels that
differed significantly from normal (P = .00477 and .0254, respectively) at the initial time point. There was no significant
difference between the levels of Bax46 expression in WAS
and normal PBL (data not shown). Therefore, we have demonstrated that
Bcl-2, an inhibitor of apoptosis, but not Bax, an activator of
apoptosis, is aberrantly expressed in WAS PBL relative to normal PBL.
Correlation between the level of Bcl-2 expression and susceptibility
to induction of spontaneous apoptosis in PBL.
The relationship between Bcl-2 expression and susceptibility to
induction of spontaneous apoptosis was studied in freshly isolated PBL
from 5 different WAS patients and from 4 normal, healthy individuals.
Relative Bcl-2 expression was measured immediately after isolation
without an extended in vitro incubation. At the same time and using the
same sample, we measured the level of apoptosis in the PBL population
after TUNEL. As illustrated in Fig 5, there
was an inverse relationship between Bcl-2 expression and the
susceptibility to induction of spontaneous apoptosis
(r2 = .744). Unstimulated normal PBL that expressed
relatively high levels of Bcl-2 were less susceptible to induction of
spontaneous apoptosis; the frequency of apoptosis in normal populations
was reduced when compared with the frequency of apoptosis in WAS
populations. In contrast, unstimulated WAS PBL that expressed
relatively low levels of Bcl-2 tended to be more susceptible to
induction of spontaneous apoptosis than were normal PBL.
We have shown that, in the absence of extrinsic apoptotic stimuli,
resting WAS PBL undergo spontaneous apoptosis at a significantly greater frequency than resting normal PBL. We also have found that the
expression of Bcl-2 is attenuated in WAS PBL. Because Bcl-2 inhibits
the onset of apoptosis, insufficient levels of Bcl-2 expression could
diminish the survival potential and shorten the life span of
lymphocytes. We suggest then that peripheral T and B lymphocytes in WAS
patients undergo accelerated spontaneous apoptosis as a consequence of
a partial Bcl-2 deficiency. The accelerated destruction of peripheral T
lymphocytes by spontaneous apoptosis could account for the progressive
deterioration of T-lymphocyte function and lymphopenia in WAS patients.
Moreover, a poor humoral response to certain specific antigens by WAS
patients may not be strictly a consequence of T-lymphocyte
abnormalities, as has been previously suggested.3,8 The
accelerated destruction of memory B lymphocytes with diminished
survival potential may preclude secondary immune
responses.47 Therefore, chronic and recurrent infection in
WAS patients may result from a failure to develop immunologic memory
caused by accelerated spontaneous apoptosis of peripheral T and B lymphocytes.
The authors express our gratitude to Dr Leo Mascarenhas and Dr John
Bastian (Children's Hospital, San Diego, CA) for their assistance in
procuring blood samples from WAS patients, and Dr Hans Ochs for
providing the genotypic information. We appreciate the efforts of Earl
Leonard, our biostatistician. We thank Dr Donald Durden, Dr Jane
Fountain, and Brile Chung for the critical reading of this manuscript.
We are grateful to Flavia Thiemann and Matthieu DeClerck for their
technical assistance. We gratefully acknowledge the CHLA Research
Institute, and the Achievement Rewards for College Scientists
Foundation (Los Angeles Chapter) for the support of S.L.R., the
recipient of the John H. Richardson & Margaret Kersten Ponty Endowed
Fellowship through the Norris Comprehensive Cancer Center, University
of Southern California, Los Angeles, CA.
Submitted January 18, 1999; accepted July 29, 1999.
Supported in part by Grants No. AI40581, HL54729, and HL54850
(Specialized Center of Research in Stem Cell Biology) from the National
Institutes of Health. S.L.R. is a fellow of the Achievement Rewards for
College Scientists Foundation and Childrens Hospital Los Angeles
Research Institute.
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 Kenneth I. Weinberg, MD, Division of
Research Immunology and Bone Marrow Transplantation, Mail Stop #62,
Childrens Hospital Los Angeles, 4650 Sunset Blvd, Los Angeles, CA
90027; e-mail: kweinberg{at}chla.usc.edu.
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TOP
ABSTRACT
INTRODUCTION
1, all of which are involved in
lymphoid signal transduction.
6 mol/L 2-mercaptoethanol [Sigma,
St Louis, MO], and 1× penicillin-streptomycin [Gemini
BioProducts]). Cells were plated at a density of 5 × 106 cells/mL in 100-mm tissue culture dishes (Corning,
Corning, NY) and placed in a 37°C humidified incubator conditioned
with 5% CO2 for 1 hour. Nonadherent PBL were collected and
resuspended in R-10. PBL were plated at a density of 0.5 × 106 cells/mL in 24-well flat-bottomed dishes (Costar,
Cambridge, MA) and maintained in a humidified atmosphere with 5%
CO2 at 37°C for up to 10 days. Some experiments were
performed after maintaining PBL in R-10 containing 10%
heat-inactivated fetal calf serum (Summit Biotechnology, Fort Collins,
CO) instead of human serum, but no differences in the apoptotic
frequencies of PBL were apparent. PBL were not stimulated in vitro.
.05 is termed significant. An analysis of correlation between
Bcl-2 expression and level of apoptosis was performed using logarithmic
transformation. A correlation coefficient (r2) was
determined using the nonparametric Spearman method. Increases in the
frequency of apoptosis over time were determined by linear regression.
