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PLENARY PAPER
From the Department of Pediatrics, Section of Pediatric
Hematology/Oncology, Wells Center for Pediatric Research, and the
Department of Medicine, Section of Nephrology, Howard Hughes Medical
Institute, Indiana University School of Medicine, Indianapolis, IN; the
Department of Biochemistry, University of Tennessee, Memphis, TN;
Bonfils Blood Center and the Department of Pediatrics, University of
Colorado School of Medicine, Denver, CO; the Department of Pediatrics,
C. S. Mott Children's Hospital and the Department of Pathology,
Howard Hughes Medical Institute, University of Michigan Medical School;
Ann Arbor, MI.
Rho GTPases control a variety of cellular processes, including
actin polymerization, integrin complex formation, cell adhesion, gene
transcription, cell cycle progression, and cell proliferation. A
patient is described who has recurrent infections and defective neutrophil cellular functions similar to those found in Rac2-deficient mice. Molecular methods were used to clone the expressed Rac2 cDNA from
this patient, and a single base pair change (G Circulating blood neutrophils, which make up the
initial cellular component of inflammatory infiltrates, interact with
endothelium in a multistep process involving selectin-initiated
endothelial capture, integrin-mediated adhesion, and chemotaxis-induced
diapedesis (reviewed in Lowe and Ward1). The importance of
neutrophil movement and function in host defense is demonstrated by
genetic defects affecting neutrophil-endothelial interactions
(leukocyte adhesion deficiency),2 microbial killing
(chronic granulomatous disease),3,4 and movement
(neutrophil actin dysfunction).5
To accomplish cellular responses to environmental signals, cells use
various signaling pathways linking receptors for extracellular stimuli
with effectors in the cytoplasm and nucleus. The Rho family of GTPases,
members of the Ras superfamily of small signaling molecules, plays key
roles in regulating these responses. Rho GTPases are involved in a wide
array of cellular functions, including cytoskeletal (particularly
actin) reorganization, integrin complex formation and cell adhesion,
gene transcription, cell cycle progression, and cell
proliferation.6-8 Rho GTPases, in a fashion similar to
Ras, cycle between an active, GTP-bound state and an inactive, GDP-bound state. Proteins that activate these transitions (guanine nucleotide exchange factors and GTPase-activating proteins) and the
downstream effector proteins that interact with activated Rho-GTPases
have been increasingly identified.9 However, the specific
mechanism(s) by which Rho GTPases control a wide range of cellular
processes and the relationship between the cellular functions
controlled by the same or different Rho GTPases remain(s) largely unknown.
Several distinct members of the Rho family of GTPases have been
characterized, including Rho, Rac, and Cdc42. Each member of the family
appears to control a distinct function of the actin cytoskeleton.9 In fibroblasts, the activation of Rho leads to the formation of actin-myosin stress fibers and integrin-containing adhesion complexes. Cdc42 activation induces spike-like filopodia rich
in F-actin, whereas the activation of Rac is associated with broad
lamellipodia.7 The latter 2 morphologic structures are associated with poorly defined integrin complexes.9
Interestingly, Rac members of the Rho family have also been implicated
in regulation of the NADPH oxidase enzyme complex that generates
superoxide in phagocytes.10-15 Although human gene
mutations encoding other components of the NADPH oxidase complex
(gp91phox, p47phox, p67phox, p40phox) results in absent or reduced
superoxide production and chronic granulomatous disease, no patients
have yet been recognized with Rac-associated abnormalities in the NADPH
oxidase.3,4
Recent work by our laboratory and by other investigators has begun to
elucidate the role of Rho GTPases in phagocyte cell function. Rho has
been demonstrated to mediate actin polymerization in
neutrophils,16-18 and, using dominant negative and
constitutively activated mutants, Rac function has been implicated in
membrane ruffling, lamellipodia formation, and chemotaxis in a
macrophage cell line.19,20 In these same cells, Cdc42
function appears essential for cell polarization mediated by filopodia
toward a chemotactic gradient. Using a genetic approach to generate
mice deficient in the hematopoietic specific Rho GTPase, Rac2, we
demonstrated that Rac2 plays an essential and unique role in neutrophil
rolling via L-selectin, F-actin assembly, and chemotaxis in
response to several chemoattractants and in superoxide generation in
response to some, but not all, agonists.21,22
Rac2-deficient mice display a unique phenotype of immunodeficiency
consisting of the above cellular defects, leukocytosis, and
neutrophilia, but increased susceptibility to fungal infections.
Based on the phenotypic analysis of Rac2-deficient mice, it is possible
to predict a human phenotype associated with functional abnormalities
in Rac. We now characterize the molecular, biochemical, and biologic
defects in a patient with a phenotype similar to that of Rac2-deficient
mice by cloning and sequencing the expressed Rac2 mRNA, and demonstrate
that the mutation in Rac2 acts in a dominant negative fashion in blood
cells. The cDNA mutation is consistent with genomic sequence recently
published23 on this same patient.
Blood and bone marrow isolation and neutrophil separation
Rolling and chemotaxis assays
Chemotaxis was evaluated using 2 methods. Neutrophils were suspended in
Dulbecco minimum essential medium (DMEM) containing 0.1% bovine serum
albumin (Sigma, St Louis, MO) and gradually warmed for 1 hour to
20°C. N-formyl-methionyl-leucyl-phenylalanine (10 In addition, PMN chemotaxis in response to fMLP (0-10 Phagocytosis assays Phagocytosis assays were carried out essentially as previously described.26,27 Antibody-treated erythrocytes (EIgG) were washed 3 times and suspended in the same buffer at 2 to 3 × 109 cells/mL. After incubation, neutrophils were activated with fMLP (100 nmol/L) for 10 minutes at 37°C; EIgG (2 × 106/mL) were added to the activated neutrophils, and incubation was continued for an additional 30 minutes at 37°C. EIgG that were not internalized were lysed with distilled water, returned to isotonicity by the addition of 0.6 mol/L KCl, and fixed with 1% glutaraldehyde. Phagocytosis was quantitated microscopically and expressed as the number of particles ingested per 100 cells (phagocytic index).Measurement of NADPH oxidase activity and NBT test Superoxide production was measured in a quantitative kinetic assay based on the reduction of cytochrome C (Sigma) after the stimulation of cells with fMLP alone or with cytochalasin B or phorbol myristate acetate (PMA) alone (all Sigma). PMN were treated with 5 µg/mL cytochalasin B for 3 minutes and then by 400 nmol/L fMLP for 5 minutes, or fMLP alone, or 10 ng/mL PMA alone for 5 minutes. Data were expressed as mean nmol 02 /106
PMN/5 minutes ± SEM. The nitroblue tetrazolium (NBT) test for superoxide production was performed on PMN derived from the patient or
from healthy donors after expansion with or without transduction in
vitro as described.28 The percentage of NBT-positive cells and the intensity of staining were determined by evaluating
100 cells.
Adhesion to fibrinogen Neutrophils were activated with 107 mol/L fMLP for
5 minutes at 37°C, then washed with and resuspended in Dulbecco
phosphate-buffered saline containing 5 mmol/L glucose. Plates (24 well)
were coated with fibrinogen (50 µg/well) (Chromogenix, Milan, Italy)
for 2 hours at 37°C, washed with Dulbecco phosphate-buffered saline, and blocked with 1% gelatin (Norland Products, New Brunswick, NJ) for
1 hour at 22°C. Plates were again washed, then neutrophils (1 × 105/well) were added to plates at appropriate times
to yield the durations specified. Plates were incubated at 22°C.
Neutrophils that had not adhered were washed away, and adherent cells
were fixed in 1% glutaraldehyde. The number of adherent neutrophils was determined by counting 5 random 200× microscopic fields.
Cloning and sequencing of Rac2 cDNA Total RNA was isolated from expanded BM cells of healthy donors and the patient using TRIZOL reagent as described by the manufacturer (Life Technologies, Rockville, MD). First-strand cDNA synthesis was carried out using Superscript II RNase H reverse
transcriptase (Life Technologies) and oligo (dT)16 as primer (Perkin Elmer, Foster City, CA). Polymerase chain reaction (PCR)
was performed using primers spanning the entire human Rac2 coding
region (5' CCGGAATTCATGCAGGCCATCAAGTGTGTGGTG 3' and 5' CCGCTCGAGCTAGAGGAGGCTGCAGGCGCGCTT C 3'). The 597-bp PCR products were
cloned into pCR4-TOPO vector using the procedures recommended by the
manufacturer (Invitrogen, Carlsbad, CA). Individual colonies were then
picked and sequenced with T3 and T7 primers. Thirteen clones from the
patient and 7 from the normal sample were successfully sequenced.
Construction of retroviral vectors, transfections, and infection of target cells An improved murine stem cell virus (MSCV)-based bi-cistronic retroviral vector, MIEG3, was constructed and used in this study. MIEG3 gives brighter EGFP fluorescence than the parental MSCV and MIG vectors,29 which allows direct visualization of the EGFP-positive cells under a fluorescent microscope. To construct MIEG3, the RSGFP in the MIG retroviral vector was replaced with the EGFP of pEGFP-C1 (Clontech Laboratories, Palo Alto, CA), yielding MIEG vector (Hanenberg H, D.A.W., unpublished data). Subsequently, the BglII-NcoI fragment in MIEG containing the disabled internal ribosome entry site (IRES) was replaced with BamHI-NcoI fragment of the pIRES2-EGFP (Clontech Laboratories), yielding MIEG3 vector. This last maneuver restores the encephalomyocarditis virus (EMCV) IRES element to its original EMCV viral configuration, resulting in more efficient translation of EGFP.Flag epitope tag was introduced into the N-terminal region of the murine wild-type (WT) Rac2 cDNA using a PCR-based technique. The primers used in this study were as follows: 5' CGGAATTCACCATGGACTACAAAGACGATGACGACAAGCAGGCCA-TCATTGTGTGGTGGTGGGTGATG 3' and 5' GCTCGAGCCTAGAGCAGGCTGCAGG GGCGCTTCTGCTG 3'. Similarly, the Kozak consensus translation initiation sequence was incorporated into the normal human Rac2 cDNA and the mutant (D57N) Rac2 cDNA derived from the patient to maximize the translation of the expressed Rac2 genes. The following primers were used: 5' CCGGAATTCCACCATGCAGGCCATCAAGTG TGTGGTG 3' and 5' CCGCTCGAGCTAGAGGAGGCTGCAGGCGCGCTT C 3'. Amplified fragments were cloned into pPCR-Script SK plasmid (Stratagene, La Jolla, CA) and sequenced multiple times. The sequence-confirmed, Flag-tagged murine Rac2, normal human Rac2, and human mutant D57N Rac2 inserts were digested with EcoRI and XhoI and were cloned into the same sites of MIEG3 (see below) yielding MIEG3FR2, HR2WT, and HR2MU vectors, respectively. Viral supernatant collected from the transfected Phoenix-Ampho packaging cell line (obtained from American Type Culture Collection, Manassas, VA) was used to infect the GP+E86 packaging cell line.30 The GFP-expressing cells with high-fluorescence intensity after 2 consecutive infections were selected by fluorescence-activated cell sorter (FACS). The retroviral titer was determined by FACS analysis using a method similar to that reported recently.31 The titer of the stable MIEG3 and MIEG3FR2 E86 clone viral supernatant was approximately 2.5 × 104 cfu/mL and 1.4 × 105 cfu/mL, respectively. Transient retroviral supernatant derived from Phoenix-Ampho (titers of approximately 5 × 105/mL) was also used to directly infect human mononuclear cells using the procedures described previously.24 Infected cells were cultured with 100 ng/mL MGDF, 100 ng/mL hG-CSF, and 100 ng/mL SCF for 5 days. Transduced cells were analyzed for GFP expression by FACScan (Becton Dickinson, Mountain View, CA) 48 hours after the second infection. Flow cytometric analysis, Western analysis, and cell sorting After the initial 5 days of expansion in 3 cytokines (see above), the transduced cells were further expanded 10 to 14 days in G-CSF, SCF (as above), and recombinant human IL-3 (100 ng/mL; Peprotech) to enhance myeloid differentiation. Before functional assays, cells were analyzed for transduction by flow analysis of GFP, and the differentiation of the cells was evaluated by expression of human CD13 on the cell surfaces using a FACScan (Becton Dickinson) after staining the cells with phycoerythrin antihuman CD13 (Pharmingen, San Diego, CA). The GFP-positive cells were isolated by FACS (FACStar Plus; Becton Dickinson) under sterile conditions. Reanalysis of the GFP-sorted cells showed purity of greater than 90%. NIH/3T3 cells were infected twice with the MIEG3, HR2MU, HR2WT, and MIEG3FR2 retroviral supernatants. GFP-positive cells were sorted using FACS. After culturing in DMEM for several days, a portion of the cells was analyzed by FLOW for GFP expression, and the remaining cells were used for Western blot analysis using a 1:5000 dilution of anti-Rac2 antibody (kindly supplied by Gary Bokoch, Scripps Institute, La Jolla, CA). The Western blot was performed as previously described.32Cell shape changes MIEG3, HR2WT, and HR2MU-infected NIH3T3 cells were seeded on gelatinized glass coverslips. Several days after seeding, the cells were serum starved for 20 hours. The cells were then treated with 5 ng/mL platelet-derived growth factor (Peprotech) or mock treated for 8 minutes and fixed in 3.7% paraformaldehyde for 20 minutes at room temperature. Subsequently, the cells were stained with rhodamine-phalloidin as previously described.33Expression, purification, and guanine nucleotide binding of recombinant proteins The cDNA of human D57N mutant Rac2 or WT human Rac2 was cloned into the bacterial expression vector pET-28a (Novagen, Milwaukee, WI) at the EcoRI and XhoI cloning sites. Both WT and D57N Rac2 were expressed in Escherichia coli as a fusion protein with His6 tag. Purification of these proteins was carried out according to the manufacturer's instructions. The purified recombinant proteins were then separated on an SDS-PAGE gel to confirm the purity (greater than 90%) of the expressed proteins. 3H-GDP (10 µmol/L) and 35S- GTP (1 µmol/L) (Amersham/Pharmacia Biotech, Piscataway, NJ) were loaded onto
the recombinant GTPases (1-2 µg) in the presence of 20 mmol/L
Tris-HCl (pH 7.6), 100 mmol/L NaCl, 2 mmol/L EDTA, and 1 mmol/L
dithiothreitol for 20 minutes at room temperature. Reactions were
terminated by the addition of MgCl2 to a final concentration of 10 mmol/L, and the mixture was placed on ice. Binding
was quantitated by vacuum filtration, as described.34
Case history The patient was a 1-year-old boy who had multiple recurrent, life-threatening infections characterized by leukocytosis and notable for the absence of pus in the inflamed tissues. A detailed clinical description of the child's history before transplantation will be presented elsewhere (A. Kurchubasche et al, manuscript submitted).A phagocyte defect was suspected and found by the original treating physicians on the basis of neutrophilia, absence of pus in the wounds, and improved healing in response to granulocyte transfusions of an umbilical infection. Evaluation of neutrophil function at the University of Michigan confirmed the original impression of the referring physicians. Neutrophil functional data revealed that the patient's neutrophils responded normally to PMA in terms of the respiratory burst. In addition, the presence and density of CD11b, CD11c, and CD18 were normal. New data indicated that the expression of P-selectin and CD62L (data not shown) were also normal. Other neutrophil functions are described below. The patient was referred to the University of Michigan Pediatric Bone Marrow Transplantation Program for allogeneic bone marrow transplantation from his HLA-identical older brother. A detailed workup of neutrophil function was completed before transplantation (see below). The patient was conditioned with antithymocyte globulin 90 mg/kg, intravenous busulfan 16 mg/kg, and cyclophosphamide 200 mg/kg. Transplantation was complicated by severe veno-occlusive disease of the liver, necessitating serial paracentesis and prolonged mechanical ventilation. Neutrophils became engrafted on day 11. The only infection-related complication was an infected arterial line that grew Stenotrophomonas maltophilia and Enterobacter cloacae, which responded well to intravenous antibiotic therapy. The patient made a full recovery and is currently thriving at home 4 months after undergoing bone marrow transplantation. He has had no infections since hospital discharge, and he requires no medications. His white blood cell count and differential are normal, and he is well engrafted for red blood cells and platelets, though mild thrombocytopenia persists. Immunologic studies confirm normal T- and B-cell subsets and normal immunoglobulin levels. His bone marrow demonstrates 100% donor cells by a variable number of tandem repeat analyses. Neutrophil function assays The patient had markedly impaired chemotaxis in response to fMLP and IL-8 in modified Boyden chamber analysis (Figure 1A). Patient neutrophils migrating in response to 100 nmol/L fMLP and 5 mmol/L IL-8 were significantly reduced (0.5 ± 0.1 and 0.3 ± 0.1 cells/400× field, respectively) compared to PMN obtained from a healthy donor (77.3 ± 22.1 and 45.5 ± 10.9, respectively; mean ± SEM; n = 5). The lack of movement was confirmed using direct observation of PMN movement in a Zigmond chamber in the presence of a gradient of fMLP (105 mol/L) (see below). Phagocytosis was also impaired,
but not as severely as chemotaxis. Phagocytosis of erythrocytes
opsonized with IgG (EIgG) (Figure 1B) by patient neutrophils stimulated with 100 nmol/L fMLP was 42 ± 6.0 EIgG/100 PMN, significantly higher
(P < .05) than that for unstimulated neutrophils but
lower (P < .001) than the 138.5 ± 17.5 EIgG ingested
per 100 PMN by normal fMLP-stimulated controls. Thus, fMLP-stimulated
phagocytosis was only slightly increased in patient cells, whereas it
was increased 6-fold in control cells. Because a striking
characteristic of neutrophils derived from Rac2-deficient mice
previously described was defective rolling by L-selectin,
the ability of patient neutrophils to tether and subsequently roll on
the L-selectin ligand, GlyCAM-1, was analyzed (Figure 1C).
At 1.26 dynes/cm2, the patient neutrophils demonstrated an
approximately 5-fold reduction in the number of cells that tethered and
rolled on GlyCAM-1. Adhesion of fMLP-stimulated patient PMN to
fibrinogen-coated plates over 10 to 60 minutes tended to be reduced
compared to normal controls (Figure 1D), but this reduction reached
significance only at 30 minutes. In summary, the cellular phenotype of
the patient was remarkably similar to that of the actin-based
neutrophil cellular defects described in the Rac2-deficient
mouse.21
Superoxide production Rac1, Rac2, or both appear to be essential for superoxide production in cell-free systems, and neutrophil NADPH oxidase function is impaired,10-12 but not completely deficient, in neutrophils derived from Rac2-deficient mice.21 To assess NADPH oxidase function in neutrophils from patients and normal controls, cells were stimulated with fMLP and PMA, and superoxide production was measured (Figure 2). Without prior treatment with cytochalasin b, fMLP-elicited O2 production by patient neutrophils was
significantly reduced compared to that of normal neutrophils. After
treatment with cytochalasin b, fMLP still failed to activate
O2 production in the patient's neutrophils,
which produced only 1.7 ± 1.2 nmol superoxide/106 cells
per 5 minutes. In contrast, as expected, a more than 3-fold increase in
O2 production was seen in neutrophils from
healthy donors, which produced 43.7 ± 5.8 nmol
superoxide/106 cells per 5 minutes (mean ± SEM,
n = 5). However, analogous to Rac2-deficient murine neutrophils, PMA
(10 ng/mL) stimulation of the patient's neutrophils yielded
normal production of superoxide. Also as expected, after
successful bone marrow transplantation, O2 production was equivalent on
concurrently analyzed patient and normal neutrophils (Figure
2).
Sequencing of the Rac2 cDNA and identification of a D57N point mutation in the patient Low-density bone marrow (LDBM) cells obtained from the patient before transplantation or from healthy donors were expanded in multiple cytokines and used to prepare RNA for RT-PCR. PCR primers were designed that flanked the entire Rac2 coding sequence. As seen in Figure 3A, the predicted Rac2 PCR product of 597 bp was obtained from the patient's cells and normal BM cells. PCR products from both were cloned and sequenced. As seen in Figure 3B, multiple sequenced cDNAs from the patient demonstrated a G A base
pair change at nucleotide 169. Among 13 individual clones sequenced
from samples from the patient, 8 were found to have the GA mutation,
whereas the remaining 5 were wild type. All 7 clones sequenced from the
normal sample were found to contain the WT sequence. In addition, we
found another single base-pair change (CT) at nucleotide position
477 compared with the GenBank human Rac2 sequence; this mutation does
not change the coding amino acid sequence, and this mutation was
present in all clones sequenced (patient and healthy donor). Northern
blot analysis of total cellular RNA from the patient demonstrated
nearly normal levels of Rac2 message (data not shown). The predicted
amino acid change encoded by the mutant cDNA sequence was a
substitution of asparagine for aspartic acid at position 57 (D57N).
This mutation occurs in a GTP-binding domain that is highly conserved
in all Rho GTPases and in the Ras superfamily.35 Normal
levels of mRNA and normal sequences in half the cloned cDNA from the
patient suggested that the mutant acted in a dominant-negative fashion at the cellular level.
Construction of retrovirus expressing WT and mutant D57N Rac2 cDNA and functional analysis To study the cellular phenotype of the patient and the mutant D57N cDNA in more detail, recombinant retrovirus vectors expressing either the WT (both murine and human) and mutant Rac2 cDNA were constructed (Figure 4A). The retrovirus backbone, MIEG3, is derived from the MSCV retrovirus29 with modifications to improve the expression of GFP. The use of an internal ribosome re-entry site (IRES) in a bi-cistronic vector allows estimates of expression of both GFP and inserted upstream sequences (in this case, Rac2) in a limited number of cells by the intensity of GFP using Flow analysis. To determine that HR2WT and HR2MU both encode Rac2 protein, NIH/3T3 cells, which lack endogenous Rac2 expression, were infected and then sorted for GFP-positive cells (Figure 4B,C). As seen in Figure 4B, most NIH/3T3 cells were GFP positive (the percentages of GFP-positive cells for MIEG3, HR2MU, and HR2WT were 94%, 84%, and 96%, respectively). GFP-sorted NIH/3T3 cells shown in Figure 4B were used for Western blot analysis. As seen in Figure 4C, Western blot analysis confirmed the expression of vector-encoded mutant Rac2 (lane 2) and WT Rac2 (lane 4) and of Flag-tagged murine Rac2 (lane 5) in infected and GFP-sorted cells. WT Flag-tagged murine Rac2 protein is also present, as expected, in the MIEG3FR2 producer cell line. A minor band, likely representing cross-hybridization with Rac1 (see Roberts et al21) is present in uninfected NIH/3T3 cells (lane 1) and 3T3 cells infected with MIEG2 vector (lane 3). NIH/3T3 cells expressing D57N grew more slowly than cells expressing either WT Rac2 or GFP alone (see below).
We found no differences between the functions of murine and human WT
Rac2 (data not shown), which was consistent with the high degree of
homology of the coding sequences between species. Therefore, in initial
studies, either the murine WT or the human Rac2 mutant retroviruses
were used to transduce cells. Patient LDBM cells were transduced with
empty vector (MIEG3) or with the WT murine or human cDNA (MIEG3FR2 or
HR2WT) and sorted for GFP-positive cells after in vitro differentiation
into granulocytic cells. As control, normal LDBM cells were transduced
with the empty vector. Neutrophil NADPH oxidase activity was assessed
using the NBT test, which detects superoxide production in individual
cells. Compared to the empty vector, expression of WT Rac2 in patient
cells derived from transduced LDBM cells failed to increase the
percentage of NBT-positive cells in response to fMLP (4% NBT
positive), though, as previously noted, the patient cells showed
similar levels of NBT-positive cells in response to PMA (52% NBT
positive) (Table 1). In contrast, normal
cells transduced with the control retrovirus expressing only GFP
demonstrated NBT-positive cells after stimulation with either PMA
(91%) or fMLP (55%).
To demonstrate more precisely at the cellular level the
dominant-negative nature of the D57N mutation, normal human LDBM (or umbilical cord blood) cells were infected with either the WT and the
D57N-expressing retrovirus, and the cells were differentiated in vitro
into mature neutrophils. Between 2% and 4% of primary bone marrow
cells were transduced by the empty retrovirus vector (MIEG3), the WT
Rac2 cDNA-containing vector (MIEG3FR2), or the D57N mutant (HR2MU)
(data not shown). After transduction and differentiation for 14 days in
culture, more than 95% of cells displayed a myeloid phenotype, as
assessed by staining with CD13. Transduced GFP-positive cells were
sorted to purity by FACS and analyzed for cell movement and superoxide
generation by NBT test. After sorting, more than 90% of the cells were
found to be CD13 and GFP double positive. As seen in Table
2, either PMA or fMLP activates
superoxide production in normal human cells transduced with either the
vector expressing GFP alone (MIEG3) or WT Rac2 (MIEG3FR2). In contrast,
normal human cells expressing the D57N mutant (HR2MU) have few
NBT-positive cells in response to fMLP but are able to generate
superoxide in response to PMA in a fashion similar to normal.
This phenotype (PMA-, but not fMLP-, induced superoxide production)
mimics that of the patient's cells and provides evidence of the
dominant-negative nature of the D57N mutant at the cellular level.
Effects of D57N on movement of normal transduced myeloid cells To generate adequate numbers of transduced and expanded cells for movement studies, we used umbilical cord blood cells, which have a higher transduction efficiency (20%-40%) than bone marrow cells. Chemotaxis analyzed by modified Boyden chamber demonstrated that GFP+/CD13+ cells expressing either MIEG3 or HR2WT (human WT Rac2) were able to move in response to fMLP (Table 3). The expression of WT Rac2 by retrovirus appeared to enhance movement in response to fMLP (Table 3). In contrast, expression of the D57N mutant cDNA significantly impaired movement of these cells in response to fMLP compared with the expression of WT Rac2 (11% ± 4% mutant vs 100% ± 19% WT, P < .001, and impaired movement when compared with normal cells at 11% ± 4% vs 65% ± 14%, P < .01).
To assess the speed of movement of the patient cells, we analyzed
responses to fMLP in a videomicroscopy chamber (Figure
5A). Compared to normal neutrophils in
which 50% of the cells moved at more than 4 µm/min, patient
neutrophils uniformly moved slowly. Wild-type neutrophils were observed
to move at rates of up to 13 µm/min, whereas the maximum speed of
patient neutrophils recorded was 3.1 µm/min. The expression of WT
Rac2 failed to increase the speed of patient neutrophil movement
(patient Rac2-EGFP). We next analyzed the effect of WT Rac2 expression
vs D57N Rac2 mutant expression on the speed of movement of normal
neutrophils derived from transduced bone marrow cells (Figure 5B).
GFP-positive cells expressing either the retrovirus containing EGFP
alone or WT Rac2 demonstrated fast movement (more than 4 µm/min) in an fMLP gradient, as analyzed by videomicroscopy. In
contrast, expression of the D57N mutant increased the number of
neutrophils that demonstrated slow movement (less than 4 µm/min), and
none of these mutant Rac2-GFP-positive cells were seen moving at a
fast speed. Thus, the differential response of both the NADPH oxidase
to PMA and the fMLP agonist and the movement of mutant neutrophils in
response to fMLP was recapitulated in normal cells by expressing the
patient's mutant cDNA. The data demonstrate that expression of the
patient's mutant allele is associated with abnormal movement and
superoxide production, even in the presence of normal Rac2 protein, and
they confirm the dominant-negative nature of this mutation.
D57N mutant fails to bind GTP and acts as a dominant negative for Rac1 and Rac2 To determine the mechanism of the D57N mutation at the biochemical level, GTP- and GDP-binding assays were performed. As seen in Figure 6A, compared to recombinant WT Rac2 the D57N mutant protein failed to bind GTP. Binding to GDP appeared normal. These data are consistent with the location of the mutation in the Rac sequence and the biochemical abnormalities previously described for the analogous Ras D57 and Rho D59 position mutations.35 Because this sequence and function are highly conserved throughout all members of the Rho GTPase family, and because Rac1 continues to be expressed in myeloid and lymphoid cells, we hypothesized that the D57N phenotype may involve affects on both Rac1 and Rac2 in these lineages. To test the hypothesis that D57N Rac2 can have dominant-negative effects on Rac1, we expressed the D57N mutant in NIH/3T3, which expresses Rac1 but not Rac2. Expression was confirmed by Western blot (Figure 4C). As noted previously, NIH/3T3 cells expressing D57N Rac2 grew more slowly than NIH/3T3 cells expressing either WT Rac2 or GFP only, showing a 2-fold reduction at 72 hours after transduction and a 7-fold reduction 5 days after transduction (T.W., D.A.W., unpublished data). Expression of D57N, but not WT Rac2, significantly altered the morphology of NIH/3T3 cells, generating a cuboidal, epithelial-like shape characterized by loss of cytoplasmic extension (Figure 6B-D), and greatly reduced membrane ruffling in response to platelet-derived growth factor stimulation, as assessed by rhodamine-phalloidin staining (Figure 6E). Morphologic and growth characteristics of these cells closely mimics those of fibroblasts transfected with dominant-negative Rac1 (17N) constructs.36,37 Because Rac2 is not expressed in these cells, these data support the conclusion that the D57N mutant can act in a dominant-negative fashion to interfere with Rac1 and Rac2 function.
Cellular immunity mediated by phagocytic cells is an important component of host defense. Several genetic causes of neutrophil dysfunction have been characterized at both the molecular and the cellular levels. The child described by Ambruso et al23 and further characterized by us displayed cellular abnormalities that overlapped previously described syndromes of chronic granulomatous disease,3,4 leukocyte adhesion deficiency,2 and actin dysfunction,5 and he shared a phenotype that closely mimicked that of a mouse mutant deficient in the Rho GTPase, Rac2. Previously, another child with recurrent infections who shared many of the phenotypic abnormalities demonstrated in Rac2-deficient mice was also identified.38 The important characteristics of Rac2 dysfunction in mice and humans appear to include leukocytosis, impaired neutrophil chemotaxis in vitro and in vivo, impaired capture and rolling via the L-selectin ligand, GlyCAM-1, but not via P-selectin mild to moderate deficiency in adhesion, spreading and phagocytosis of neutrophils, and reduced generation of superoxide in response to some, but not all, agonists. The mammalian Rho GTPase family is made up of at least 7 distinct
protein subfamilies The mutation in the patient described here is located in the
GTP-binding region of Rac. Aspartate at position 57 (in Ras, 59 in Rho)
is invariant and is part of the G-3 region loop that is highly
conserved in all mammalian Rho GTPases. The aspartate is thought to
bind catalytic Mg2+ by a water molecule and, ultimately, by
the The data derived from murine Rac2
We thank Dr J. Panepinto (Rhode Island Hospital) for referring this patient. We thank Dr Dirk Roos and members of our laboratory for helpful discussions and Eva Meunier and Sharon Smoot for assistance in preparation of the manuscript.
Submitted April 4, 2000; accepted May 11, 2000.
Supported by National Institutes of Health grants R01 AI-20065 (L.B.), R01 HL-45635 (M.C.D.), GM60523 (Y.Z.), and 1P01CA71932 (J.B.L.), and by the University of Michigan Clinical Research Center. The Wells Center for Pediatric Research is a Center of Excellence in Molecular Hematology (NIDDK P50DK49218).
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: David A. Williams, Department of Pediatrics, Wells Center for Pediatric Research, Howard Hughes Medical Institute, 1044 West Walnut St, Rm 402, Indianapolis, IN 46202-5225; e-mail: dwilliam{at}iupui.edu.
1. Lowe JB, Ward PA. Therapeutic inhibition of carbohydrate-protein interactions in vivo. J Clin Invest. 1997;99:822-826[Medline] [Order article via Infotrieve]. 2. Arnaout MA, Spits H, Terhorst C, Pitt J, Todd RF III. Deficiency of a leukocyte surface glycoprotein (LFA-1) in two patients with Mo1 deficiency: effects of cell activation on Mo1/LFA-1 surface expression in normal and deficient leukocytes. J Clin Invest. 1984;74:1291-1300.
3.
Roos D, de Boer M, Kuribayashi F, et al.
Mutations in the X-linked and autosomal recessive forms of chronic granulomatous disease.
Blood.
1996;87:1663-1681 |
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Abstract
Introduction
P = .001 patient vs normal. (C) Neutrophil
rolling on the L-selectin ligand, GlyCAM-1. Tethered and
rolling cells were scored and averaged in 10 to 12 different fields of
the GlyCAM-1-coated plate in the second minute of the rolling assay.
Data shown represent the average number of tethered cells, ± SEM,
observed using 2 independently isolated cell preparations and 2 independently prepared GlyCAM-1-substituted chamber surfaces.
*P = .01. (D) Time course of neutrophil adhesion to
fibrinogen. The number of adherent neutrophils was determined by
counting 5 random 200× microscope fields. **P = .05 only
at the 30-minute time point; P > .05 (NS) all
other points.
P = .005. NS, PMA patient vs normal;
fMLP + cytochalasin B after BMT.
Rho, Rac, Cdc42, RhoD, RhoG, RhoE, and TC10.