Basophil contribution to the IL-4 pool in filarial infections was assessed using PBMC from 20 patients with active filarial infections and from 9 uninfected subjects. Patient basophils released histamine in response to Brugia malayi Ag (BmAg). They also released IL-4 within 2 h after exposure to BmAg, as assessed by intracellular cytokine flow cytometry. This IL-4 induction was Ag specific, as IL-4 was not detected in BmAg-exposed basophils obtained from uninfected subjects. Although there were, on average, 64 times more CD4+ T cells than basophils in the peripheral circulation of filaria-infected patients, the absolute numbers of basophils and CD4+ T cells producing IL-4 per 100,000 PBMC were equivalent (geometric mean: 16 IL-4-producing basophils/100,000 PBMC vs 22 IL-4-producing CD4+ T cells/100,000 PBMC). Basophils also released IL-4 in response to both low and high concentrations of BmAg, whereas CD4+ T cells released IL-4 only after incubation with a high concentration of BmAg, raising the possibility that basophils, due to their lower threshold for activation, may actually release IL-4 more frequently than CD4+ T cells in vivo. Furthermore, IL-4 production in vitro by Ag-stimulated purified basophils or CD4+ T cells provided evidence that basophils release greater quantities of IL-4 per cell than CD4+ T cells in response to BmAg. These results suggest that, when Ag-specific IgE is present in a filaria-infected individual, basophils function to amplify the ongoing Th2 response by releasing IL-4 in greater amounts and possibly more frequently than CD4+ T cells in response to filarial Ag.
A growing body of evidence suggests basophils are important components of the human immune response to helminth infection. Helminth infections are well known to be associated with high IgE levels (1, 2, 3). Basophils bind IgE through high affinity receptors (FcεRI) on their surface and degranulate and release histamine and other inflammatory mediators after cross-linking of these receptors (4). Basophils from patients infected with Toxocara, Ascaris, Onchocerca, Wuchereria, Strongyloides, and Schistosoma have been shown to release histamine in response to parasite Ag (5, 6, 7, 8, 9), with the amount of histamine released proportional to the serum concentration of Ag-specific IgE (5).
In addition to high levels of IgE, human helminth infections are associated with increased production of IL-4, a prototypical Th2 cytokine (10, 11). Of all PBMC, only basophils have the ability to produce IL-4 early in response to a stimulus (12), releasing preformed IL-4 within 5–10 min of IgE surface cross-linking (13). After activation, basophils also synthesize IL-4 and IL-13 de novo, with time-course experiments showing a second peak of IL-4 release after 4 h and a first peak of IL-13 release at 24 h (13). Recently, basophils from filaria-infected patients have been shown to release IL-4 in response to L3 larval and adult filarial Ag (14). Of interest, basophils from normal (uninfected, parasite naive) individuals have been shown to release IL-4 after stimulation with Schistosoma egg Ag (15, 16), raising the possibility that basophils could be the initial trigger for the Th2 response in some helminth infections.
Infection with the pathogenic human filariae (Wuchereria bancrofti, Brugia malayi, Onchocerca volvulus, and Loa loa), tissue-dwelling helminths that cause, respectively, lymphatic filariasis, onchocerciasis (river blindness), and loiasis (African eyeworm), causes responses in humans associated with eosinophilia, elevated serum levels of Ag-specific and polyclonal IgE, and increases in T cell production of IL-4, IL-5, and IL-13 (10, 17, 18, 19). Although the initial trigger for this response is not yet known, much of this response is most likely driven by IL-4, as it causes CD4+ T cells to differentiate into Th2 cells and can, along with CD40-CD40 ligand interaction, cause B cells to switch to IgE production (20).
Although previous studies have shown that CD4+ T cells are a significant contributor to the IL-4 pool in filarial infections (11), the contribution that basophils make has not yet been investigated. Although basophils make up only a small percentage of peripheral leukocytes (usually ∼0.5%), they have the ability to recognize many different Ag specifically by virtue of IgE Ab of varied specificities present on their surface. Consequently, it is possible that the number of basophils specifically recognizing and activated by helminth Ag may equal or even exceed the number of Ag-specific T cells responding to helminth Ag. In asthmatic patients, for example, basophils producing IL-4 have been shown to outnumber IL-4-producing CD4+ T cells in response to dust mite Ag (21).
Our goal was to define further the role played by basophils in the immune response to filarial infections in humans. Specifically, we sought to: 1) confirm that basophils from filaria-infected patients are activated by filarial Ag; 2) determine whether Brugia malayi Ag (BmAg)2 can nonspecifically activate basophils from uninfected individuals; and 3) assess the contribution of basophils to the IL-4 pool in filaria-infected patients relative to CD4+ T cells in terms of both frequency of IL-4-producing cells and actual amounts of IL-4 produced.
Materials and Methods
Filaria-infected patients and uninfected individuals were recruited from the Clinical Parasitology Unit, Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health (Bethesda, MD) and from the Department of Transfusion Medicine, National Institutes of Health, respectively, under protocols approved by the National Institute of Allergy and Infectious Diseases Institutional Review Board. Filarial infection was diagnosed using: 1) positive identification of appropriate parasite or parasite DNA in blood, skin snips, or tissue biopsy by microscopy or PCR; 2) positive circulating Ag test for W. bancrofti; 3) positive antifilarial serology plus Calabar swellings and response to therapy for L. loa; 4) positive antifilarial serology plus either Mazzoti reaction or opthalmologic exam consistent with onchocercal eye disease for O. volvulus (22); or meeting all the established criteria for tropical pulmonary eosinophilia (23). In total, 29 patients were included in the study: 20 filaria-infected patients (14 with L. loa, 4 with O. volvulus, and 2 with W. bancrofti) and 9 uninfected controls. Of the 20 filaria-infected patients, 11 were expatriates and 9 were immigrants from countries endemic for filarial infections. BmAg-specific IgE levels in filaria-infected patients ranged from 0.1 to 254 ng/ml with a geometric mean (GM) of 7.3 ng/ml, and total IgE levels ranged from 7 to 11,362 IU/ml with a GM of 376. None of the normal controls had detectable levels of BmAg-specific IgE.
Preparation of BmAg
Soluble BmAg was made from B. malayi adult worms (provided by J. McCall, University of Georgia, Athens, GA). Male and female adult worms (5000 each) were directly harvested from the peritoneal cavities of infected jirds and frozen at −70°C. Worms were then thawed, washed in PBS (BioWhittaker, Walkersville, MD), refrozen in liquid nitrogen, and then lyophilized overnight. Lyophilized samples were resuspended in PBS, stirred with a magnetic bar overnight at 4°C, and then spun at 750 × g for 10 min at 4°C with no brake. The supernatant was collected and the pellet was resuspended in PBS and again stirred at 4°C overnight. Following centrifugation at 1200 × g for 10 min at 4°C, supernatant was again collected. Both sets of supernatants were then pooled and spun at 12,000 × g for 30 min at 4°C. This supernatant was collected and stored in aliquots at −70°C. Endotoxin level of final soluble BmAg was <0.1 EU/ml using the QCL-1000 Chromogenic LAL Test Kit (BioWhittaker).
Aliquots (200 μl) of heparinized whole blood diluted 1/7 with histamine release buffer (Beckman Coulter, Marseilles, France) were incubated for 30 min at 37°C with buffer alone and with 10 different concentrations (1 × 10−7 to 100 μg/ml) of BmAg. In five patients, cells were also incubated with and without recombinant bactericidal/permeability-increasing protein (rBPI; XOMA, San Diego, CA) at 5 μg/ml or polymyxin B (Sigma-Aldrich, St. Louis, MO) at 20 μg/ml. One aliquot of blood from each patient was frozen and thawed twice to give a measure of total histamine stores. Tubes were then centrifuged for 5 min at 900 × g at 4°C, and supernatants were collected. Histamine concentrations were then measured using a histamine enzyme immunoassay kit (Immunotech, Marseilles, France). Values obtained under different conditions were divided by total histamine to obtain percentages of total histamine released.
Cell preparation and fixation for intracellular flow cytometry
Heparinized venous blood was collected, and PBMC were separated using a Ficoll diatrizoate gradient at 4°C (lymphocyte separation media; ICN Biomedicals, Aurora, OH). Cells were washed and then cultured at 2 × 106/ml in Iscove’s Dulbecco modified medium (Biofluids, Rockville, MD) supplemented with 10% heat-inactivated FCS (HyClone Laboratories, Logan, UT), 1% l-glutamine (Biofluids), 1% insulin-transferrin-selenium medium (Biofluids), and 80 μg/ml gentamicin (BioWhittaker). Cells were cultured with BmAg at 10 μg/ml, BmAg at 0.1 μg/ml, ionomycin (Calbiochem, La Jolla, CA) at 1 μg/ml, Staphylococcus enterotoxin B (SEB; Toxin Technologies, Sarasota, FL) at 10 μg/ml, or in medium alone. Brefeldin A (Sigma-Aldrich) was added at 20 μg/ml at time zero for 2-h incubations and at 2 h for 6- and 24-h incubations. After 2, 6, and 24 h, DNase (Sigma-Aldrich) was added at 666 Dornase U/ml and allowed to incubate for 5 min at 37°C. Cells were then transferred to 15-ml conical tubes, washed twice in PBS (BioWhittaker), and then fixed in PBS containing 4% paraformaldehyde (Sigma-Aldrich) for 5 min. Cells were then washed in PBS containing 0.1% BSA (Sigma-Aldrich), resuspended in PBS/10% DMSO (Fischer Scientific, Fair Lawn, NJ), and cryopreserved at −70°C.
Fixed cells were thawed, washed with PBS/0.1% BSA, and then permeabilized and blocked for 1 h by incubating in PBS containing 0.1% saponin (Calbiochem)/1% BSA at 4°C. Cells were then stained with R-PE-conjugated rat IgG1 anti-human IL-13 (BD PharMingen, San Diego, CA), allophycocyanin-conjugated rat IgG1, anti-human IL-4 (BD PharMingen), and either CyChrome-conjugated mouse IgG1 anti-human CD4 for CD4+ T cell analysis or, for basophil analysis, FITC-conjugated goat anti-human IgE (BioSource International, Camarillo, CA), CyChrome-conjugated mouse IgG1 anti-human CD2 (BD PharMingen), tricolor-conjugated mouse IgG2a anti-human CD14 (Caltag, Burlingame, CA), tricolor-conjugated mouse IgG1 anti-human CD16 (Caltag), and CyChrome-conjugated mouse IgG1κ anti-human CD19 (BD PharMingen). R-PE-conjugated rat IgG1κ (BD PharMingen) and allophycocyanin-conjugated rat IgG1 (BD PharMingen) were used as isotype controls for IL-13 and IL-4 staining, respectively. After staining, cells were washed twice with PBS/0.1% saponin, resuspended in PBS, and analyzed using a FACSCalibur flow cytometer (BD Biosciences, Mountain View, CA) and CellQuest software (BD Biosciences). Before running patient samples, all Ab were individually titrated with positive and negative control cells for optimal sensitivity and specificity.
Basophil and CD4+ T cell purification and culture
Concurrent purification of basophils and CD4+ T cells was done on three patients. Each patient underwent apheresis using a CS 3000+ system (Baxter Healthcare, Deerfield, IL). PBMC were then further purified using a Ficoll diatrizoate gradient, as described above. Between 5 and 7 × 109 PBMC were isolated from each patient.
Basophils were isolated from PBMC by negative selection using a MACS basophil isolation kit (Miltenyi Biotec, Auburn, CA). Concentrations of the blocking reagent, hapten-Ab cocktail, and MACS antihapten microbeads were used at double the manufacturer’s suggested concentration, and no more than 100 × 106 cells were passed per LS-type MACS magnetic separation column (Miltenyi Biotec). Basophil isolation was done on 2 × 109 PBMC from each patient, resulting in 5.2 × 106, 7.6 × 106, and 8.6 × 106 cells with >85% purity of basophils, as measured by both flow cytometry and manual counts of 1000 cells stained with May-Grunwald-Giemsa (Sigma-Aldrich) on cytocentrifuged slides.
CD4+ T cells were also isolated from PBMC by negative selection, as described previously (24). CD4+ T cell isolation was done on 400 million PBMC from each patient, resulting in 48 × 106, 120 × 106, and 64 × 106 cells with >95% purity for all samples by flow cytometry.
After purification, PBMC, CD4+ T cells, and basophils were cultured in medium, as described above, at 1.4 × 106 cells/ml with BmAg at 10 μg/ml, BmAg at 0.1 μg/ml, ionomycin, SEB, or medium at 37°C. Irradiated autologous PBMC were added to the CD4+ T cell wells at a final concentration of 350,000/ml to serve as APC. In two patients, purified CD4+ T cell populations were also cultured under the same conditions at 2 × 106 cells/ml. After 24 h, cells were centrifuged and supernatants were collected for cytokine measurement.
Calculation of estimated per cell IL-4 production
The amount of IL-4 produced per basophil or CD4+ T cell was approximated using the following calculation: (amount of IL-4 produced in culture)/((number of cells in culture) × (percentage of patient cells releasing IL-4 under given conditions as calculated by flow cytometry)).
Quantification of IL-4, IL-13, total IgE, and BmAg-specific IgE, IgG, and IgG4
Measurements of IL-4 and IL-13 were performed by ELISA. Flat-bottom plates (Immulon 4; Dynatech, Chantilly, VA) were coated overnight at 4°C with either mouse anti-human IL-4 (8D4-8; BD PharMingen) or rat IgG1 anti-human IL-13 (JES10 5A2; BD PharMingen) in PBS, followed by washing with PBS and 0.05% Tween (Sigma-Aldrich). Plates were then blocked with PBS, 5% BSA, and 0.05% Tween for 2 h at 37°C and washed again. Culture supernatants were added and incubated overnight at 4°C. Plates were then washed and incubated with either biotinylated rat anti-human IL-4 (MP4-25D2; BD PharMingen) or biotinylated polyclonal goat anti-human IL-13 (R&D Systems, Minneapolis, MN) in PBS, 1% BSA, and 0.05% Tween 20 (ELISA diluent) for 2 h at 37°C. Following washing, 1 μg/ml of alkaline phosphatase-conjugated streptavidin (Jackson ImmunoResearch Laboratories, West Grove, PA) in ELISA diluent was added for 1 h at 37°C. Plates were again washed, and p-nitrophenyl phosphate disodium (Sigma-Aldrich) was added at 1 mg/ml in sodium carbonate buffer (KD Medical, Columbia, MD). Colorimetric development was detected at 405 nM using a microplate reader (Molecular Devices, Sunnyvale, CA).
BmAg-specific IgE measurements were similarly performed using an ELISA. Before measuring BmAg-specific IgE, IgG was adsorbed by incubating serum samples with GammaBind G Sepharose (Pharmacia Biotech, Uppsula, Sweden) overnight at 4°C. BmAg-specific IgE levels were then measured by ELISA using plates initially coated with BmAg at 10 μg/ml and then using biotinylated goat anti-human IgE (Fortron Bioscience, Morrisville, NC) for detection. Quantification was based on standardized curves using recombinant cytokines and purified human BmAg-specific IgE.
BmAg-specific IgG and IgG4 were measured by ELISA, as described previously (25). Total serum IgE levels were measured by the Laboratory of Clinical Pathology (Clinical Center, National Institutes of Health, Bethesda, MD) using an Immulite 2000 Immunoassay System (Diagnostic Products, Los Angeles, CA).
Comparisons between groups of unpaired data were performed using the nonparametric Mann-Whitney unpaired rank test. The Wilcoxon signed rank test was used to compare paired data, Fisher’s exact test was used for comparisons of frequency, and correlations were derived using the Spearman rank test. All statistics were performed using StatView5 (SAS Institute, Cary, NC).
Basophils from filaria-infected patients specifically release histamine in response to BmAg
To assess the ability of basophils from filaria-infected and uninfected individuals to respond to parasite Ag, whole blood samples from 16 filaria-infected patients and 7 uninfected controls were incubated for 30 min alone or with serial 10-fold dilutions of BmAg (from 100 to 1 × 10−7 μg/ml) and assayed for released histamine. Whereas 15 of 16 filaria-infected patients had a maximum histamine release of greater than 10% of total histamine stores in response to BmAg (range 4–67%, GM 37%), only 1 of the 7 uninfected controls (range of maximum histamine release 1–16%, GM 4%) released histamine in response to BmAg (p < 0.01; Fig. 1,A). For most filaria-infected patients, the threshold for induction of histamine release occurred at a BmAg concentration of 0.1 μg/ml and peaked at 10 μg/ml (Fig. 1 B). BmAg-specific histamine release correlated strongly with serum levels of BmAg-specific IgE (ρ = 0.615, p = 0.02) in the filaria-infected patients, but not with levels of Ag-specific IgG or IgG4 or with polyclonal IgE. As expected, none of the uninfected subjects had measurable serum BmAg-specific IgE.
To assess whether the LPS of the Wolbachia bacteria present in filariae augment histamine release in filaria-infected patients, histamine release assays were done in the presence or absence of rBPI or polymyxin B in five patients. No significant differences in histamine release were found when either rBPI or polymyxin B was added to BmAg (p > 0.2 when comparing maximum histamine release of BmAg alone vs either BmAg + rBPI or BmAg + polymyxin B).
Basophils from filaria-infected patients specifically release IL-4 in response to BmAg
To determine the contribution basophils make to the production of IL-4 in circulating leukocytes, PBMC from 20 filaria-infected patients and 9 uninfected controls were incubated with medium, BmAg at 0.1 or 10 μg/ml, or ionomycin for 2 h, fixed, and analyzed for cytokine production by intracellular flow cytometry. Basophil populations were identified and gated based on surface IgE positivity and negativity for CD2, CD14, CD16, and CD19 (Fig. 2 A).
Basophils from filaria-infected patients released IL-4 at greater frequencies than basophils from uninfected controls at baseline (GM 0.9 vs 0.15%, p < 0.01) and in response to BmAg at 0.1 μg/ml (GM 3.9 vs 0.17%, p < 0.01) and 10 μg/ml (GM 3.2 vs 0.24%, p < 0.01). These findings were not due to intrinsic differences in the ability of basophils from these two populations to release IL-4, as there was no difference in the frequency of IL-4+ basophils between filaria-infected patients and uninfected controls after stimulation with ionomycin (GM 28%, range 0.1–78.9% vs GM 43%, range 18–66.2%, p > 0.9) (Fig. 2 B).
The frequencies of basophils from filaria-infected patients releasing IL-4 increased significantly after stimulation with BmAg at 0.1 μg/ml compared with medium alone (GM 3.9 vs 0.9%, p < 0.01) and after stimulation with BmAg at 10 μg/ml compared with medium (3.2 vs 0.9%, p < 0.01). There was no significant difference between the frequency of basophils releasing IL-4 after stimulation with 10 μg/ml as compared with 0.1 μg/ml (GM 3.2 vs 3.9%, p = 0.8).
When enough patient cells were available, additional cultures for basophil flow cytometry were fixed at 6 and 24 h as well as at 2 h and then analyzed. There was no significant difference in frequency of IL-4+ basophils at 2 vs 6 h (15 patients, GM 3.2 vs 2.4%, p = 0.29) or at 2 vs 24 h (3 patients, 7.4 vs 4.5%, p = 0.3) after stimulation with BmAg at 10 μg/ml. Culture with medium, BmAg at 0.1 μg/ml, or ionomycin also had no significant differences in terms of frequencies of IL-4+ basophils at the different time points. These data suggest that the release of IL-4 is rapid and has a relatively low (in terms of Ag concentration) threshold for release.
The frequencies of basophils releasing IL-4 after stimulation with BmAg at 0.1 μg/ml correlated strongly with Ag-specific histamine release (ρ = 0.618, p = 0.02) (Fig. 3,A) as well as with serum levels of BmAg-specific IgE (ρ = 0.500, p = 0.03) (Fig. 3 B) and BmAg-specific IgG4 (ρ = 0.583, p = 0.01), but not with Ag-specific IgG or polyclonal IgE. Of note, none of the uninfected controls had detectable quantities of Ag-specific IgE.
CD4+ T cells from filaria-infected patients also specifically release IL-4 in response to BmAg, but require a higher concentration to induce such release and do so at significantly lower frequencies than do basophils
PBMC from 19 filaria-infected patients and 9 uninfected controls were incubated with medium, BmAg at 0.1 or 10 μg/ml, or SEB for 6 h, fixed, and analyzed for cytokine production by intracellular flow cytometry. CD4+ T cells from filaria-infected patients released IL-4 at greater frequencies than did CD4+ T cells from uninfected controls after stimulation with BmAg at 0.1 μg/ml (GM 0.031 vs 0.003%, p = 0.004) and BmAg at 10 μg/ml (GM 0.080 vs 0.003%, p < 0.01), but not when incubated with medium alone (GM 0.032 vs 0.015%, p = 0.35). CD4+ T cell populations from filaria-infected patients and uninfected controls had the same potential to release IL-4 as equivalent percentages of CD4+ T cells released IL-4 after stimulation with SEB (1.43 vs 1.03%, p = 0.33).
Unlike basophils from filaria-infected patients, in which IL-4 release was triggered with as little as 0.1 μg/ml of BmAg, CD4+ T cells from filaria-infected patients required a BmAg concentration of 10 μg/ml to induce IL-4+CD4+ frequencies above those of PBMC cultures left unstimulated (medium alone). The percentage of CD4+ T cells from filaria-infected patients that produced IL-4 was significantly greater after incubation with BmAg at 10 μg/ml compared with incubation with BmAg at 0.1 μg/ml (GM 0.080 vs 0.031%, p < 0.01) or medium (GM 0.080 vs 0.032%, p < 0.01) (Fig. 4).
Although CD4+ T cells definitely released IL-4 in response to high concentrations of BmAg, the percentages of CD4+ T cells that did so were much lower than the percentages of basophils that did so in the same patients (GM 0.080 vs 3.2%, p < 0.01).
Absolute numbers of basophils and CD4+ T cells from filaria-infected patients that release IL-4 in response to BmAg are equivalent
By flow cytometry, 32% (GM) of PBMC in filaria-infected patients were CD4+ T cells and 0.05% (GM) were basophils. Thus, to compare directly the numbers of basophils and CD4+ T cells in the peripheral circulation that released IL-4 in response to BmAg, the flow cytometry data were analyzed in terms of cells producing IL-4 per 100,000 PBMC. Per this analysis, the actual numbers of basophils and CD4+ T cells in the peripheral circulation of filaria-infected patients that release IL-4 in response to BmAg were equivalent. After 6 h of incubation with BmAg at 10 μg/ml, the optimal concentration for stimulation of CD4+ T cells, a GM of 22 CD4+ T cells produces IL-4 per 100,000 PBMC. This compares to a GM of 16 basophils per 100,000 PBMC producing IL-4 after incubation for 2 h with 0.1 μg/ml of BmAg (Fig. 5) and 13 basophils per 100,000 PBMC producing IL-4 after incubation with BmAg at 10 μg/ml (p > 0.1 for IL-4+CD4+ T cells/100,000 PBMC at 10 μg/ml vs IL-4+ basophils/100,000 PBMC at 10 and 0.1 μg/ml). At a BmAg concentration of 0.1 μg/ml, fewer CD4+ T cells released IL-4 than did basophils per 100,000 PBMC (7 vs 16), but this difference was not statistically significant (p > 0.4).
Basophils from filaria-infected patients release more IL-4 per cell than do CD4+ T cells in response to BmAg
PBMC, basophils, and CD4+ T cells (along with irradiated PBMC to serve as APC) were isolated from three patients with filarial infections and cultured at 1.4 × 106 cells/ml for 24 h. Purified CD4+ T cells were cocultured with irradiated autologous PBMC at 350,000 cells/ml to serve as APC. Although there was no measurable IL-4 produced by purified CD4+ T cell cultures after incubation with BmAg, purified basophil cultures released more IL-4 than did PBMC cultures in the two patients whose PBMC released measurable IL-4 (Table I). Because an increase in the number of basophils in culture resulted in more IL-4 release than from PBMC culture, and because an increase in CD4+ T cells in culture did not result in a similar increase, one can conclude that basophils are the primary contributors to the total IL-4 pool in filaria-infected patients.
|Patient .||IL-4 Production in pg/mlb .||.||.||.||.||.||.||.||.|
|.||By PBMC .||.||.||By CD4+ T cells .||.||.||By Basophils .||.||.|
|.||A .||B .||C .||A .||B .||C .||A .||B .||C .|
|Patient .||IL-4 Production in pg/mlb .||.||.||.||.||.||.||.||.|
|.||By PBMC .||.||.||By CD4+ T cells .||.||.||By Basophils .||.||.|
|.||A .||B .||C .||A .||B .||C .||A .||B .||C .|
Amounts of IL-4 released into the supernatant by PBMC, purified CD4+ T cells, and purified basophil cultures were measured by ELISA after 24-h incubation with BmAg, ionomycin, SEB, or media. The lower limit of detection for ELISA was 48 pg/ml.
BmAg-IgE levels for patients A, B, and C were 254, 9, and 11.5 ng/ml, respectively, and total IgE levels for these patients were 11,362, 508, and 598 IU/ml, respectively.
nd, Not done.
These conditions resulted in no measurable IL-4 even when cells were cultured at 2 × 106/ml.
While no IL-4 was produced from the third patient when purified CD4+ T cells were at a concentration of 1.4 × 106/ml with SEB, IL-4 was produced at a concentration of 66 pg/ml when these cells were cultured at 2 × 106/ml with SEB.
This difference in IL-4 contribution in response to BmAg is not due to different maximum potentials of releasable IL-4 from these patient populations. By combining the individual frequency data of the percentage of each cell type releasing IL-4 in response to a particular culture condition with the results from the purification studies, we were able to calculate the approximate IL-4 produced per cell under various conditions. Both cell types have the ability to release similar quantities of IL-4, as the purified basophil populations released a GM of 9.8 fg of IL-4 per cell (range 5.5–14.2) after stimulation with ionomycin and the purified CD4+ T cell populations released a mean of 10.6 fg of IL-4 per cell (range 5.9–13) after stimulation with SEB. In response to BmAg, the purified basophil populations from two patients each released an average of 2.6 fg of IL-4 per cell. The amount of IL-4 produced per CD4+ T cell in response to BmAg could not be calculated, as the amount of IL-4 produced by these cell cultures was below the level of detection.
IL-13 production by basophils and CD4+ T cells
Expression of IL-13 was also assessed in basophils and CD4+ T cells in flow cytometry experiments. After stimulation with BmAg at 0.1 μg/ml, only 4 of 20 patients had detectable IL-13 from basophils after 2 h of incubation; only 5 of 16 had detectable IL-13 after 6 h; and 0 of 3 had detectable IL-13 after 24 h. Frequencies of IL-13+ basophils in those with detectable IL-13 were very low, with none having >0.9% of basophils producing IL-13. These basophils had the capability to produce IL-13, as basophils from all but one individual had detectable IL-13 after stimulation with ionomycin for 2 h (GM 3.2%, range 0–21.6%). The potential of basophils from filaria-infected patients to produce IL-13 was similar to that of uninfected patients, as the frequencies of basophils releasing IL-13 after stimulation with ionomcyin were not statistically different between infected and uninfected patients (GM 3.2 vs 2.6%).
Similarly, CD4+ T cells had very low frequencies of IL-13 expression, with IL-13 detected in only 5 of 19 patients after 6 h of incubation with BmAg at 10 μg/ml (GM 0.12%, range 0–1.94%). All patient CD4+ T cell populations had the potential to produce IL-13, as each had some IL-13 expression after stimulation with SEB for 6 h (GM 2.0%, range 0.47–7.39%). The frequencies of CD4+ T cells releasing IL-13 after stimulation with SEB were not significantly different between infected and uninfected patients (GM 2.0 vs 1.7%, p = 0.87).
Additionally, no measurable IL-13 was detected by ELISA in the supernatants of PBMC, purified basophils, or purified CD4+ T cells after stimulation with BmAg at 0.1 and 10 μg/ml for 24 h from any of the three patients tested, although these cell cultures all released IL-13 after stimulation with ionomycin or SEB.
Although they were initially described over a century ago by Ehrlich (26), basophils are enigmatic cells. Although much is known of their basic biology and their role in allergic responses, what beneficial physiologic function they perform in humans, if any, remains unknown. A growing body of evidence, however, suggests that basophils may be important in the immune response to helminth infections.
In this study, we have demonstrated that circulating basophils from filaria-infected patients specifically release histamine and IL-4 in response to soluble BmAg. The amounts of histamine released and the frequencies of basophils releasing IL-4 correlated with BmAg-specific IgE levels, suggesting that release of histamine and IL-4 from basophils in filaria-infected patients occurs primarily through IgE-mediated activation.
Using flow cytometry, we have shown that although basophils make up only a small percentage of the cells in the peripheral circulation, the actual numbers of basophils that release IL-4 in response to BmAg in filaria-infected patients are equivalent to the number of CD4+ T cells that do so. This most likely occurs because, whereas individual CD4+ T cells are specific for only a single Ag, individual basophils, by virtue of their ability to bind IgE molecules of many specificities, can theoretically specifically recognize and be activated by hundreds of different Ag. In our study, an average of >3% of all basophils from filaria-infected patients released IL-4 in response to BmAg, whereas the average percentage of CD4+ T cells that released IL-4 under similar conditions was less than 0.1%. Consequently, although CD4+ T cells were far more numerous than basophils in these patients, the total numbers of each of these cell types releasing IL-4 in response to BmAg were equivalent.
Notably, even though most filaria-infected patients had a maximum histamine release of >50% in response to BmAg, suggesting that most of the basophils in these patients were activated by BmAg, the GM percentage of IL-4+ basophils was only 3.9% after incubation with BmAg. This suggests that either some basophils produce histamine and not IL-4 after BmAg activation or that most basophils produce IL-4 after BmAg stimulation, but do so at levels below detection of current intracellular flow cytometry techniques.
Two of our findings suggest that basophils may be the principal IL-4-producing cell in the response to filarial infections. First, basophils were able to release IL-4 after exposure to low concentrations of BmAg (0.1 μg/ml), whereas CD4+ T cells only released significant amounts of IL-4 after exposure to high concentrations of BmAg (10 μg/ml). Because of this lower threshold of activation for IL-4 release, basophils most likely release IL-4 more frequently in response to filarial infections than do CD4+ T cells in vivo, in which cells are exposed to a wide range of Ag concentrations. Second, as highly purified cultures of basophils from filaria-infected patients released more IL-4 than PBMC cultures, and because CD4+ T cell cultures did not release any measurable IL-4 in response to BmAg, our study strongly suggests that basophils make greater amounts of IL-4 per cell in response to BmAg than do CD4+ T cells. Together, these findings suggest that in vivo basophils may release IL-4 more frequently and in greater amounts than do CD4+ T cells in filaria-infected patients.
Although it is well documented that basophils release IL-13 as well as IL-4 upon IgE-mediated activation, we found only low frequencies of IL-13 release from activated basophils in filaria-infected patients by flow cytometry and no IL-13 release by ELISA. Although this may be a valid finding, it is also possible that the conditions we used were insensitive for the detection of IL-13. In particular, we may have used insufficient duration of cell culture time, as IL-13 release from basophils may not peak until at least 24 h after basophil activation (13).
Of interest, stimulation of basophils from uninfected controls did not result in substantial histamine or IL-4 release. Although we did not expect basophils from the control patients to degranulate through parasite Ag/IgE interactions, as none of the uninfected patients had measurable amounts of Ag-specific IgE, we did think it was theoretically possible that BmAg could cause nonspecific basophil activation, because nonspecific basophil activation by helminth Ag has been demonstrated previously. A glycoprotein from Schistosoma mansoni eggs can cause basophils to degranulate and release IL-4 by binding and cross-linking IgE nonspecifically (15, 16), and homologues of mammalian translationally controlled tumor protein (TCTP), a calcium-binding protein that directly stimulates histamine release from basophils, have been cloned recently from S. mansoni (27) and the filarial parasites W. bancrofti and B. malayi (28). As the B. malayi TCTP homologue is maximally expressed in adult stage filariae (28), and because the filarial Ag we used consisted of solubilized adult B. malayi worms, the lack of nonspecific basophil activation in our study suggests that B. malayi worms do not produce sufficient quantities of TCTP homologue to trigger basophil degranulation.
The lack of IL-4 release from basophils of uninfected individuals in our study provides strong evidence that basophils do not serve as an initial source of IL-4 in filarial infections and thus do not initiate the Th2 response to filariae. Once a Th2 response and Ag-specific IgE are present, however, our results provide evidence that basophils gain the ability to specifically release IL-4 in response to BmAg. Moreover, because basophils have preformed IL-4, these cells can release IL-4 very quickly in response to an appopriate stimulus. Although we have demonstrated that basophils release IL-4 as quickly as 2 h after exposure to BmAg (the shortest time period we evaluated), basophils have been documented to release IL-4 within 5–10 min of IgE cross-linking (13). Also, because basophils can maintain IgE-mediated memory for at least 1 mo even in the absence of ongoing de novo IgE production (29), basophils may be responsible for quickly re-establishing a Th2 response against future filarial exposures in patients who have been infected previously with filariae.
In conclusion, our study provides evidence that once a specific IgE response is made, basophils become the predominant contributors of IL-4 in the immune response to filarial infection. Recognizing BmAg at low concentrations and then rapidly releasing IL-4 in large amounts, basophils appear to be the primary cells that release IL-4 in response to filarial infections, serving to amplify ongoing Th2 responses to filarial infections and possibly serving to quickly re-establish the Th2 response to future exposure to filariae.
We thank Dr. Calman P. Prussin for advice on analyzing basophils by flow cytometry and Brenda Rae Marshall for help in preparation of this manuscript.
Abbreviations used in this paper: BmAg, Brugia malayi Ag; BPI, bactericidal/permeability-increasing protein; GM, geometric mean; SEB, Staphylococcus enterotoxin B; TCTP, translationally controlled tumor protein.