Loss of Complement Activation and Leukocyte Adherence as Nippostrongylus brasiliensis Develops within the Murine Host
http://www.100md.com
感染与免疫杂志 2005年第11期
School of Molecular and Biomedical Science, University of Adelaide, Adelaide, Australia
Department of Microbiology and Infectious Diseases, Flinders Medical Centre, Adelaide, Australia
Rheumatology Section, Imperial College, Hammersmith Campus, London, United Kingdom
ABSTRACT
Complement activation and C3 deposition on the surface of parasitic helminths may be important for recruitment of leukocytes and for damage to the target organism via cell-mediated mechanisms. Inhibition of complement activation would therefore be advantageous to parasites, minimizing damage and enhancing migration through tissues. The aim of this study was to determine ex vivo if complement activation by, and leukocyte adherence to, the nematode Nippostrongylus brasiliensis change as the parasite matures and migrates through the murine host. Pathways of activation of complement and the mechanism of adherence of leukocytes were also defined using sera from mice genetically deficient in either C1q, factor B, C1q and factor B, C3, or C4. Substantive deposition of C3 and adherence of eosinophil-rich leukocytes were seen with infective-stage (L3) but not with lung-stage (L4) larvae. Adult intestinal worms had low to intermediate levels of both C3 and leukocyte binding. For L3 and adult worms, complement deposition was principally dependent on the alternative pathway. For lung-stage larvae, the small amount of C3 detected was dependent to similar degrees on both the lectin and alternative pathways. The classical pathway was not involved for any of the life stages of the parasite. These results suggest that in primary infections, the infective stage of N. brasiliensis is vulnerable to complement-dependent attack by leukocytes. However, within the first 24 h of infection, N. brasiliensis acquires the ability to largely avoid complement-dependent immune responses.
INTRODUCTION
The rapid recruitment of leukocytes to infected tissues is a crucial early step in resistance to parasitic helminths. In order to damage the target organism, many effector cells make close contact with the parasite and this can occur via complement- and antibody-dependent mechanisms (16, 29, 47). Although other mediators may also play minor roles (4, 37, 45), in the absence of parasite-specific antibody and especially in the early stages of primary infection, complement is likely to be the most important factor facilitating leukocyte recruitment and attachment to the parasite. C3b and its cleavage products facilitate attachment of leukocytes to the helminth, while C3a and C5a may act as early chemotactic factors. Eosinophils are major effector cells in host resistance to infection with a range of parasitic helminth species (14, 38, 44, 47) and adherent eosinophils release products that can damage, immobilize, and/or kill the parasite (6, 10).
Using physical or chemical strategies to deplete complement activity in vitro, parasites have been shown to activate complement by either the alternative or classical pathway (19, 28, 31, 36, 42). Although sera from humans and animals with spontaneous mutations in genes encoding individual complement proteins have also been used in studies with parasites (32, 40, 50), it has not previously been possible to comprehensively analyze each complement pathway in a single host species.
A number of pathogenic species of bacteria, viruses, and fungi have evolved strategies to evade activation of complement to ensure survival within the host (17, 23, 27, 34). Some helminths have also developed strategies to avoid complement recognition in the early stages of infection. Soon after entry into the host, Schistosoma mansoni becomes resistant to serum-dependent killing by shedding a strongly antigenic coat, acquiring host proteins, including blood group antigens and the complement regulatory protein decay accelerating factor (39, 41), and secreting proteins that bind to and inhibit complement proteins (13, 30, 32). In the early stages of infection, parasitic nematodes may also undergo changes that inhibit C3 deposition (35) and reduce the capacity of leukocytes to attach (1, 5).
Nippostrongylus brasiliensis is a nematode infecting mice and rats and has a life cycle similar to that of the human pathogen Strongyloides stercoralis. N. brasiliensis eggs hatch in feces, and after maturation to the infective stage, larvae (L3) enter the host through the skin. In experimental studies, most larvae have exited the skin within 4 h of subcutaneous injection, arriving in the lungs 18 to 48 h later and maturing into L4 (9, 24). Larvae then migrate via the trachea and esophagus to the small intestine, where they mature into adult worms. Infective N. brasiliensis L3 are susceptible to complement activation and eosinophil-dependent damage (19, 45). In the presence of serum, mouse eosinophils adhere to N. brasiliensis L3 in vitro and degranulate (29), leaving them temporarily immobilized, and when injected into mice, many fail to migrate to the lungs (8).
Interleukin-5 (IL-5) transgenic mice, which exhibit constitutive eosinophilia, are resistant to infection with N. brasiliensis and other helminths (14, 21, 44). When N. brasiliensis L3 are injected into the skin of these mice, eosinophils are recruited in large numbers and within 2 hours of infection, many are strongly adherent to the larvae (9). Larvae are trapped in the skin for an extended period and, relative to wild-type mice, few reach the lungs (9). However, those L3 that escape the skin and migrate to the lungs to develop into L4, do not elicit a strong inflammatory response, and for at least 24 to 48 h postinfection, few eosinophils are either recruited or adhere to lung-stage larvae (9).
In the first stage of this study we determined that N. brasiliensis develops a strategy to resist complement-dependent immunity as it matures from L3 to L4. C3 deposition and eosinophil-rich leukocyte adherence were compared on infective-stage L3 collected from in vitro cultures with parasites recovered from the skin, the lungs, and the small intestine. In the second stage of this study, we used sera from mice genetically deficient in either C1q, factor B, C1q and factor B (double mutant), C3, or C4 to determine that the pathways of complement activation differ as the parasite matures and migrates through the murine host.
MATERIALS AND METHODS
Animals. Mice were bred in-house under clean barrier conditions at the University of Adelaide and Imperial College, London, and handled according to institutional animal ethics committee guidelines.
Culture medium and sera. Leukocytes were cultured with parasites in RPMI 1640 culture medium (Institute of Medical and Veterinary Science, Adelaide, Australia) supplemented with 2 mM L-glutamine, 12 μg/ml gentamicin, 16 μg/ml penicillin, and 10% heat-inactivated (56°C for 30 min) fetal calf serum (Multiser, Trace Biosciences, Sydney, Australia). Serum was collected from mice by cardiac puncture under pentobarbitone sodium anesthesia. Where indicated, serum complement activity was selectively inhibited by heating for 30 min at 56°C (heat-inactivated mouse serum [MS]). Sera from wild-type CBA/Ca and C57BL/6 mice (normal mouse serum [NMS]) and from C57BL/6 mice deficient in either C1q (classical pathway deficient) (3), factor B (alternative pathway deficient) (33), C1q and factor B (classical and alternative pathway deficient), C3 (deficient in all complement pathways) (49), or C4 (classical and lectin pathway deficient) (18) were used as indicated.
Preparation of larvae and worms. Infective-stage N. brasiliensis L3 were obtained from fecal culture after passage through female Hooded Wistar rats aged 6 to 8 weeks as previously described (9). L3 were used between 9 and 28 days after establishment of fecal cultures. L3 were washed three times in phosphate-buffered saline (PBS), pH 7.4, and concentrated by centrifugation at 50 x g for 5 min. Larvae were then resuspended in PBS (immunofluorescence studies) or in culture medium (leukocyte coculture). Skin air pouches were generated as previously described (9); 750 L3 in 100 μl PBS were injected into each pouch and 30 or 150 min later, larvae were recovered by lavage and washed three times in PBS.
Lung-stage (L4) larvae were obtained after subcutaneous infection of wild-type CBA/Ca or C57BL/6 mice with approximately 750 L3 in 100 μl PBS. Either 24 or 48 h postinfection, mice were sacrificed by CO2 asphyxiation and cervical dislocation. Lungs were resected, washed in saline, minced with scissors, and incubated for 2 h at 37°C to facilitate migration of L4 out of the tissue. L4 were isolated from lung tissue by filtration through a plastic mesh strainer (approximately 1.0-mm-square mesh). Blood was removed by five washes in saline, each time allowing L4 to settle under unit gravity. L4 were then separated from the remaining finer pieces of lung tissue under a dissecting microscope using a 3.5-ml plastic transfer pipette and washed three times in PBS or culture medium, as required. Adult worms were recovered from the small intestine 7 days after subcutaneous infection of mice with 750 L3 in 100 μl PBS. Mice were sacrificed as described above, the intestine was resected, opened longitudinally with fine scissors, incubated in saline for 1 h at 37°C and worms were collected with a transfer pipette and washed five times by gravity sedimentation in PBS or culture medium.
Measurement of larvae and worms. To allow quantitative comparison of immunofluorescence of L3, L4 and intestinal worms that are substantially different in size, the surface area of each stage was estimated. Parasites were dotted on microscope slides and photographs were taken of over 20 of each stage using a 4x objective lens on an Olympus BH-2 light microscope, an Olympus C-35AD-4 camera and Konica Color Centuria 400 ISO/ASA 35-mm print film (Konica Corporation, Japan). Photographic prints were enlarged 100% using a photocopier (final magnification, 111x) and length and width of parasites were measured with a curvimetre map measure (Kartenmesser, Germany). For each parasite stage, a mean was calculated from duplicate measurements of each parasite and the overall mean and standard error of the mean were calculated from these values. Parasite surface area was estimated by calculating the area of the curved surface of a cylinder: surface area of parasite = x diameter of parasite x length of parasite.
The length, diameter, and estimated surface area of parasites life stages are shown in Table 1. The ratio of surface areas for L3, L4, and worms was approximately 1:2:14; hence in experiments where L3, L4, and worms are compared, 120 L3 or 60 L4 or 8 worms were added to each well of a microtiter tray. Since L4 taken from the lungs 24 h postinfection were only slightly smaller (approximately 10%) than those from 48 h postinfection, in each case, the same number of L4 were used per well.
Immunofluorescence microscopy. Aliquots of suspensions of parasites were dotted onto glass slides, placed under a glass coverslip and examined under UV illumination with an Olympus BH-2 microscope at 100x magnification. Photomicrographs were taken using Fujichrome Sensia 400 ISO/ASA 35-mm film (Fuji Photo Film Co. Ltd, Tokyo, Japan) with an exposure time of 16 s for anti-C3 fluorescence, or 40 s for carboxyfluorescein diacetate succinimidyl ester (CFSE)-labeled cell fluorescence and printed at a final magnification of 47x. All immunofluorescence microscopy was conducted after scanning with a phosphorimager.
Detection of C3. Using an adaptation of techniques described recently (19), parasites were incubated with PBS alone or with 50 μl of 20% NMS, MS, or serum from complement-deficient mice for 1 h at 37°C with 5% CO2 in air in a clear 96-well microtiter plate (Greiner Bio-One, Omega Scientific, CA) or in 10-ml plastic test tubes. Unbound serum components were then removed by 10 washes with 200 μl of PBS/0.05% vol/vol Tween 20 (PBST) in 96-well plates using a multichannel pipettor, or by two washes in 5 ml PBST where test tubes were used. Parasites were allowed to settle under unit gravity (96-well plates) or were centrifuged at 18 x g for 5 min (test tubes) between each wash. Parasites were then incubated with 50 μl of a 1/50 dilution of fluorescein isothiocyanate-conjugated goat anti-mouse C3 antibody (ICN/Cappel, catalog no. 55510) for 1 h at room temperature in the dark. Unbound antibody was removed by washing as described above. Finally, parasites were resuspended in 200 μl of 25% vol/vol glycerol in PBS in 96-well plates, or where parasites were incubated in test tubes, aliquots of 200 μl/well of parasite suspensions were added to the wells of a flat-bottomed, solid black 96-well plate (Costar, catalog no.# 3915, Corning Inc.).
Measurement of fluorescence intensity. Fluorescence in individual wells of solid black or clear 96-well microtiter plates was quantified in arbitrary units with a Molecular Imager FX phosphorimager (Bio-Rad USA, catalog no. 170-7800) as described previously (19). In some experiments, fluorescence intensity per parasite was calculated to reduce variation due to differences in the exact number of parasites per well.
CFSE labeling of leukocytes. Peritoneal cells from male or female heterozygous interleukin-5 (IL-5) transgenic C57BL/6 or CBA/Ca mice (15), aged 7 to 10 weeks, were used as an eosinophil-rich (55 to 75%) source of leukocytes. Mice were killed by carbon dioxide asphyxiation and peritoneal cells were harvested by lavage with mouse-osmolality PBS (MPBS) (43). Cells were washed twice in MPBS, twice in RPMI 1640 containing 0.1% vol/vol fetal calf serum and then resuspended in RPMI 1640/0.1% fetal calf serum at 107 cells/ml. Cells were incubated in 2.5 μM CFSE for 10 min at 37°C in 5% CO2 and excess CFSE was removed by washing in culture medium.
Analysis of leukocyte adherence to parasites. Adherence of CFSE-labeled leukocytes to parasites was quantitated using an adaptation of methods described previously (19). Aliquots (30 μl) of approximately 120 L3, 60 L4, or 8 worms in culture medium were mixed with 100 μl of CFSE-labeled leukocytes (106 total) and 70 μl of either culture medium, NMS, MS, or complement deficient serum (7% final concentration) was then added. After incubation at 37°C in 5% CO2 for 2 h, unbound cells were removed by nine washes in MPBS and fluorescence was measured using the phosphorimager, as described above. UV microscopy was used to visualize fluorescent cells attached to parasites.
Statistical analysis. Data for individual experiments, with each sample assayed in duplicate or triplicate are presented as mean ± standard error of the mean and were analyzed using Student's unpaired t test and GraphPad Prism software (version 3.03, GraphPad Software Inc.).
RESULTS
Detection of C3 on N. brasiliensis. The extent of C3 deposition on L3 collected from fecal culture plates was compared to that for L4 recovered from the lungs of CBA/Ca mice either 24 or 48 h postinfection and to adult worms recovered 7 days postinfection from the small intestine. Using immunofluorescence microscopy, it was apparent that C3 deposition in the presence of NMS from CBA/Ca mice was much greater on L3 than on L4 collected either 24 or 48 h after infection (Fig. 1a [i to iii]). C3 was deposited at very high levels over the entire surface of L3. In contrast, the small proportion of L4 that did exhibit some labeling had a very different and irregular pattern of deposition (Fig. 1a [ii and iii]). Intermediate but evenly distributed C3 deposition was observed on intestinal worms (Fig. 1a [iv]).
Quantitative analysis of C3 deposition (i.e., immunofluorescence/total parasite surface area) confirmed the large differences in C3 binding between L3 and L4 opsonized with NMS (Fig. 1b). Larvae typically showed relatively high autofluorescence. Nevertheless, the level of fluorescence detected on L4 incubated with NMS was no greater than that seen with L4 opsonized with serum heat-treated to inactivate C3 (i.e., MS). There were no qualitative or quantitative differences between 24- and 48-h lung larvae. Adult worms showed significant levels of C3 deposition after opsonization with NMS (Fig. 1b), but not at the same intensity as seen on L3. Host-derived C3 was not detected on either L4 or intestinal worms, as fluorescence of PBS-opsonized parasites exposed to anti-C3 antibody was no greater than background (i.e., where anti-C3 antibody was omitted).
To determine if inhibition of C3 deposition occurred soon after infection and at a time when many larvae are migrating to other tissues, we assessed fixation of endogenous and exogenous C3 on parasites recovered from skin air pouches 30 to 150 min after injection. It was difficult to accurately count and dispense larvae recovered from the skin because they tended to coalesce and so quantitative analysis was not performed. However, when assessed visually, skin larvae recovered 30 min postinjection demonstrated clear evidence of in vivo C3 deposition (Fig. 1c [iv]). When subsequently opsonized with NMS, further increases in C3 deposition were evident (Fig. 1c [vi]) similar to that seen on L3 from cultures (Fig. 1c [iii]). Skin larvae recovered 150 min postinfection were comparable to those retrieved after only 30 min (data not shown).
Adherence of leukocytes to N. brasiliensis. The adherence of eosinophil-rich CFSE-labeled peritoneal leukocytes to L3, L4 and intestinal worms opsonized with serum was determined. Large numbers of leukocytes adhered to the surface of NMS-opsonized L3 (Fig. 2a [i]), however, few if any cells bound to similarly treated lung larvae (Fig. 2a [ii and iii]). In samples stained with Vital New Red and Alcian Blue, 92% of leukocytes in the first layer of cells adhering to L3 were eosinophils, with some macrophages (6%) and the occasional mast cell also present. Quantitative analysis of leukocyte adherence using a phosphorimager clearly demonstrated the disparity between leukocyte adherence on L3 and L4 (Fig. 2b).
Large numbers of CFSE-labeled leukocytes bind to L3 and, as reported previously (19), visual assessment revealed that this was reduced to negligible levels when MS or culture medium were substituted for NMS (Fig. 2b and data not shown), indicating that leukocyte adherence was complement mediated. In contrast, few leukocytes adhered to either 24- or 48-hour L4, regardless of the opsonization conditions (Fig. 2b). Some leukocytes adhered to intestinal worms, but unlike L3, this was largely restricted to the posterior end (Fig. 2a [iv]). When assessed quantitatively, leukocyte binding to adult worms was at most only modest (Fig. 2b) but decreased significantly if MS was used instead of NMS, indicating that it was also complement-mediated.
C3 deposition and leukocyte adherence on N. brasiliensis L3 are largely dependent on the alternative complement pathway. Until recently, it has only been possible to use physical or chemical methods of depleting complement activity and these are sometimes only partially effective or may not be specific for complement proteins. It is now possible to use sera from mice that have been genetically manipulated to be deficient in one or more proteins of the complement pathway. We performed a comprehensive analysis of C3 deposition and leukocyte adherence on infective-stage larvae in the presence of NMS and MS from wild-type C57BL/6 mice and most particularly with sera from syngeneic mice genetically deficient in either C1q, factor B, C1q and factor B (double mutant), C3, or C4 (Fig. 3).
There was no significant difference in C3 deposition on L3 exposed to NMS or C1q-deficient serum (Fig. 3a), indicating that C1q and the classical pathway play no role in mediating C3 deposition on N. brasiliensis L3. In contrast, C3 deposition was almost completely abrogated in the absence of factor B (with and without C1q), indicating that the alternative pathway is paramount in this experimental system. A small amount of factor B-independent C3 binding was evident, since C3 deposition was significantly higher in the absence of factor B than it was when larvae were exposed to C3-deficient sera (C3–/– and MS). This is consistent with the observation that C3 deposition was slightly reduced when C4 was ablated, indicating a possible minor role for the C4-dependent lectin pathway.
The adherence of peritoneal leukocytes was similarly assessed in the presence of culture medium, NMS, MS or sera from complement-deficient mice. C3 was required for adherence, which was not affected by an absence of C1q (Fig. 3b). However, adherence was significantly reduced in the absence of factor B, again highlighting the importance of the alternative pathway. There were no significant differences in leukocyte adherence with factor B-deficient serum when compared to sera deficient in C3 (C3–/– and MS) and in the presence of culture medium only. The absence of C4 had no effect on cell adherence, indicating that while the lectin pathway may contribute slightly to complement deposition on L3, for complement-mediated leukocyte adherence, the alternative pathway is sufficient.
Pathway of complement activation changes as N. brasiliensis matures. We next compared C3 deposition on L3, L4 (48 h postinfection), and adult (7 days postinfection) worms in the presence of NMS and MS from wild-type C57BL/6 mice and sera from syngeneic mice genetically deficient in complement proteins (i.e., C1q, factor B, or C4; Fig. 4). MS was substituted for C3–/– serum in this experiment because both performed similarly in previous experiments.
As already demonstrated (Fig. 3a), C3 fixation on L3 was dependent on the alternative pathway; however in this experiment (Fig. 4a), the minor contribution from the lectin pathway was not evident. When comparing sera from CBA/Ca and C57BL/6 mice, we noticed a higher level of C3 deposition on L4 treated with NMS from the latter (mean fluorescence/well ± standard error of the mean, CBA/Ca versus C57BL/6, 3.29 ± 0.09 x 103 versus 6.72 ± 0.38 x 103 respectively, P < 0.001, n = 3). Interestingly, this difference was not evident when screening L3 (mean fluorescence/well ± standard error of the mean, CBA/Ca versus C57BL/6, 1.25 ± 0.08 x 104 and 1.29 ± 0.02 x 104 respectively, not significant, n = 3). Although there was a measurable difference in the level of complement activation on L4 treated with sera from these two strains, this was not associated with differences in the numbers of either lung larvae (day 2) or worms (day 7) recovered (Mean no. of parasites ± standard error of the mean for C57BL/6 and CBA mice [n = 3 to 4] respectively. L4: 323 ± 47 versus 289 ± 44, P > 0.05. Intestinal worms: 254 ± 99 versus 164 ± 33, P > 0.05.).
Although still modest compared to C3 fixation on L3 analyzed in the same experiment (Fig. 4a), the complement deposition on L4 treated with NMS from wild-type C57BL/6 mice was greatly diminished when sera from mice deficient in either factor B or C4 were used (Fig. 4b), indicating that both the alternative and lectin pathways contribute to C3 activation on lung-stage larvae. In three separate experiments, C3 deposition on L4 was significantly less with serum from C4–/– than from factor B–/– mice (Fig. 4b and data not shown) and so for lung-stage larvae, the lectin pathway may be at least marginally more important. The pathway of complement activation for adult worms was similar to that for L3, since deposition of C3 was in both instances significantly reduced in sera from factor B–/– mice (Fig. 4c). For adult worms, some C3 deposition would appear to be independent of the alternative pathway. For all of the life stages assessed, the classical pathway (C1q) was not involved.
DISCUSSION
Evasion of potentially damaging host immune responses is essential for helminths to be successful parasites. In one sense, the parasite is likely to be at its most vulnerable upon entry into the host, principally because it must quickly change from a form suited either to a free-living stage or to residence in another host. Conversely, a nave host is particularly vulnerable in the first few days of infection, since it is totally or largely dependent on innate mechanisms of resistance. In this and our previous studies (9, 14, 19), we have shown the infective-stage L3 of N. brasiliensis to be extremely vulnerable to complement-dependent attack by eosinophils. Complement activation on N. brasiliensis L3 occurs primarily via the alternative pathway. However within the first 24 h of infection, during maturation from L3 to lung-stage L4, N. brasiliensis acquires the ability to evade complement-dependent immune responses. This may explain why the inflammatory response at the site of initial infection, the skin, is rapid and substantive, whereas that in the lungs is slow to develop. The eosinophil chemotactic factors C3a and C5a may be responsible for rapid recruitment of eosinophils into the skin, but are less likely to be available when the lungs become infected. Interestingly, there are major changes in the pathways of complement activation as N. brasiliensis matures, with the lectin pathway becoming more important for the limited levels of complement activation on lung-stage larvae.
Innate resistance of wild-type mice to primary infection with N. brasiliensis is typically very low for most strains, with a large proportion of the infecting dose (>50%) migrating through the lungs and colonizing the small intestine (14). In addition, since the parasite has a very short transit time through the host (approximately 9 to 14 days), the life cycle is completed before an effective adaptive immune response can be generated. Our studies suggest that provided sufficient and appropriate effector cells can be recruited, innate resistance against N. brasiliensis is very effective (9, 14). Eosinophils appear to be a potent force in trapping larvae in the skin in primary infections (9) and this study indicates that C3 is almost exclusively responsible for facilitating stable attachment of these and other leukocytes to the target. Our data also suggest that complement may be important for recruitment of leukocytes, since in the absence of C3 deposition on L4 (Fig. 1), few leukocytes are to be found in the lungs until after the majority of larvae have migrated to the gut (7, 9, 48). Although chemokines such as the eosinophil-specific factor eotaxin are produced in the lungs, it is clear that these do not initially compensate for the failure of complement-dependent recruitment of eosinophils or other leukocytes into this organ.
We have yet to determine how long the larvae are in the host before acquiring the ability to avoid complement activation and leukocyte adherence, though clearly it occurs sometime between 2.5 and 24 h postinfection. Some tissue-invasive parasitic helminths inhibit adherence of leukocytes after a single maturational stage (1, 5, 11, 22, 26). Leukocytes adhere to L3 but not to L4 of Dirofilaria immitis, possibly due to acquisition of host-derived inhibitory proteins (1). The exact mechanism of how lung-stage N. brasiliensis larvae avoid fixing complement or binding leukocytes is not fully understood. N. brasiliensis L3 and L4 exsheath during maturation and the third larval molt occurs within the lungs approximately 24 to 32 h postinfection (24). One or more of these exsheathments may remove parasite-derived molecules that activate the alternative pathway or expose additional binding sites for complement regulatory proteins such as factor H, which has been shown to be important for evasion of complement by Onchocerca volvulus (35). Finally, excretory/secretory proteins produced by L4 could also interfere with host defenses. Treatment of serum with excretory/secretory proteins from the nematode Toxocara canis has previously been shown to inhibit complement deposition on the surface of T. canis larvae (2). The potential mechanism(s) used by N. brasiliensis L4 to inhibit C3 deposition is the subject of further investigation in our laboratory.
Using sera from mice genetically deficient in one or more of the complement proteins, we have comprehensively and for the first time defined which complement pathways are most important for both complement activation and leukocyte adherence on a parasitic helminth. The alternative pathway is of prime importance for both C3 deposition and leukocyte adherence to L3 in vitro, but also makes a significant contribution to the smaller amounts of C3 found on both L4 and adult worms. The broader relevance of C3 deposition on adult worms is questionable, since complement may not be available or active in the gut, except perhaps where parasites attach to, erode, and/or feed on the gut wall. Activation of the classical pathway had no role in mediating C3 deposition on N. brasiliensis L3, L4, or intestinal worms, indicating that molecules capable of directly binding C1q and activating this pathway are not expressed by N. brasiliensis.
Although largely unexplored for helminths, activation via the lectin pathway has been detected on Trichinella spiralis (20) and mannan-binding lectin adheres to schistosomes (25). Complement activation by the lectin pathway was responsible for only a small proportion of the C3 deposited on N. brasiliensis L3 and adult worms. However, of the small amount of C3 detected on L4, the greatest contribution came from the lectin pathway. This stage-specific change in the relative importance of this pathway is novel and warrants further investigation.
Since L4 did not appear to fix C3 in vivo, it is curious that a small amount could be detected after incubation ex vivo with C57BL/6 serum. This occurred regardless of whether the L4 were derived from C57BL/6 or CBA hosts (data not shown). It is also odd that L4 did not acquire C3 when exposed to CBA/Ca serum in vitro. These may be in vitro phenomena of little biological significance and any explanation we might offer is purely speculative. However, complement inhibitors (host or parasite derived) may be more active with CBA serum and/or in vivo in both strains. C57BL/6 serum may have more available C3, but this seems unlikely, since these differences were not evident when L3 were tested. Importantly, we and others have not detected major differences in parasite burdens when infections in CBA and C57 strains are directly compared (12, 46).
In conclusion, we have demonstrated that infective-stage larvae of N. brasiliensis activate the alternative pathway of complement, mediating a high level of leukocyte adherence to these larvae. Complement fixation at this stage of the parasite's life cycle may also be important for recruitment of eosinophils and other leukocytes into the initial site of infection, the skin. Entrapment of larvae may then follow, provided that sufficient effector cells are present. However, within 24 h of infection, the parasite develops the capacity to inhibit deposition of C3, and this could explain the relative absence of an early inflammatory response to L4 in the lungs. In acquiring resistance at this level, larvae may more readily migrate through tissues to undergo further maturation. Interestingly, there is a shift in pathways through which the small amount of complement is activated by lung-stage L4, with the lectin pathway now being a relatively major contributor. As the parasite enters the gut and undergoes further maturation to the adult worm stage, the alternative pathway again becomes more important for complement activation. However, both innate and early adaptive immune responses may be of less consequence to a parasite that resides in the lumen of the gut.
Although we have established in vitro that complement is at least transiently important, a critical role in innate resistance to parasitic helminths in vivo has yet to be conclusively determined. More definitive proof will require investigations of the kinetics of infections in the various complement-deficient mice described here.
ACKNOWLEDGMENTS
This work was supported in part by the National Health and Medical Research Council (Australia) and School and Faculty Strategic Research Funds from the University of Adelaide.
We thank Bruce Lyons for assistance in establishing CFSE labeling techniques.
REFERENCES
1. Abraham, D., R. B. Grieve, and M. Mika-Grieve. 1988. Dirofilaria immitis: surface properties of third- and fourth-stage larvae. Exp. Parasitol. 65:157-167.
2. Badley, J. E., R. B. Grieve, J. H. Rockey, and L. T. Glickman. 1987. Immune-mediated adherence of eosinophils to Toxocara canis infective larvae: the role of excretory-secretory antigens. Parasite Immunol. 9:133-143.
3. Botto, M., C. Dell'Agnola, A. E. Bygrave, E. M. Thompson, H. T. Cook, F. Petry, M. Loos, P. P. Pandolfi, and M. J. Walport. 1998. Homozygous C1q deficiency causes glomerulonephritis associated with multiple apoptotic bodies. Nat. Genet. 19:56-59.
4. Brattig, N. W., A. Z. Abakar, F. Geisinger, and T. F. Kruppa. 1995. Cell-adhesion molecules expressed by activated eosinophils in Onchocerca volvulus infection. Parasitol. Res. 81:398-402.
5. Brattig, N. W., F. W. Tischendorf, G. Strote, and C. E. Medina-de la Garza. 1991. Eosinophil-larval-interaction in onchocerciasis: heterogeneity of in vitro adherence of eosinophils to infective third and fourth stage larvae and microfilariae of Onchocerca volvulus. Parasite Immunol. 13:13-22.
6. Butterworth, A. E. 1984. Cell-mediated damage to helminths. Adv. Parasitol. 23:143-235.
7. Daly, C. M. 1999. Ph.D. thesis. University of Adelaide, Adelaide, Australia.
8. Daly, C. M., B. Li, J. Forbes, P. R. Giacomin, and L. A. Dent. 2004. Eosinophils as mediators of early resistance in mice to the nematode Nippostrongylus brasiliensis. Clin. Investig. Med. 27:99D.
9. Daly, C. M., G. Mayrhofer, and L. A. Dent. 1999. Trapping and immobilization of Nippostrongylus brasiliensis larvae at the site of inoculation in primary infections of interleukin-5 transgenic mice. Infect. Immun. 67:5315-5323.
10. David, J. R., A. E. Butterworth, and M. A. Vadas. 1980. Mechanism of the interaction mediating killing of Schistosoma mansoni by human eosinophils. Am. J. Trop. Med. Hyg. 29:842-848.
11. Davis, S. W., and B. Hammerberg. 1990. Taenia taeniaeformis: evasion of complement-mediated lysis by early larval stages following activation of the alternative pathway. Int. J. Parasitol. 20:587-593.
12. Dehlawi, M. S., P. K. Goyal, A. W. Stadnyk, P. J. McElroy, J. Gauldie, and A. D. Befus. 2003. Responses of inbred mouse strains to infection with intestinal nematodes. J. Helminthol. 77:119-124.
13. Deng, J., D. Gold, P. T. LoVerde, and Z. Fishelson. 2003. Inhibition of the complement membrane attack complex by Schistosoma mansoni paramyosin. Infect. Immun. 71:6402-6410.
14. Dent, L. A., C. M. Daly, G. Mayrhofer, T. Zimmerman, A. Hallett, L. P. Bignold, J. Creaney, and J. C. Parsons. 1999. Interleukin-5 transgenic mice show enhanced resistance to primary infections with Nippostrongylus brasiliensis but not primary infections with Toxocara canis. Infect. Immun. 67:989-993.
15. Dent, L. A., M. Strath, A. L. Mellor, and C. J. Sanderson. 1990. Eosinophilia in transgenic mice expressing interleukin 5. J. Exp. Med. 172:1425-1431.
16. Desakorn, V., P. Suntharasamai, S. Pukrittayakamee, S. Migasena, and D. Bunnag. 1987. Adherence of human eosinophils to infective filariform larvae of Necator americanus in vitro. Southeast Asian J. Trop. Med. Public Health 18:66-72.
17. Duthy, T. G., R. J. Ormsby, E. Giannakis, A. D. Ogunniyi, U. H. Stroeher, J. C. Paton, and D. L. Gordon. 2002. The human complement regulator factor H binds pneumococcal surface protein PspC via short consensus repeats 13 to 15. Infect. Immun. 70:5604-5611.
18. Fischer, M. B., M. Ma, S. Goerg, X. Zhou, J. Xia, O. Finco, S. Han, G. Kelsoe, R. G. Howard, T. L. Rothstein, E. Kremmer, F. S. Rosen, and M. C. Carroll. 1996. Regulation of the B cell response to T-dependent antigens by classical pathway complement. J. Immunol. 157:549-556.
19. Giacomin, P. R., H. Wang, D. L. Gordon, and L. A. Dent. 2004. Quantitation of complement and leukocyte binding to a parasitic helminth species. J. Immunol. Methods 289:201-210.
20. Gruden-Movsesijan, A., M. Petrovic, and L. Sofronic-Milosavljevic. 2003. Interaction of mannan-binding lectin with Trichinella spiralis glycoproteins, a possible innate immune mechanism. Parasite Immunol. 25:545-552.
21. Herbert, D. R., J. J. Lee, N. A. Lee, T. J. Nolan, G. A. Schad, and D. Abraham. 2000. Role of IL-5 in innate and adaptive immunity to larval Strongyloides stercoralis in mice. J. Immunol. 165:4544-4551.
22. Hong, Y., C. W. Kim, and B. Ghebrehiwet. 1992. Trichinella spiralis: activation of complement by infective larvae, adults, and newborn larvae. Exp. Parasitol. 74:290-299.
23. Horstmann, R. D., H. J. Sievertsen, J. Knobloch, and V. A. Fischetti. 1988. Antiphagocytic activity of streptococcal M protein: selective binding of complement control protein factor H. Proc. Natl. Acad. Sci. USA 85:1657-1661.
24. Kassai, T. 1982. Handbook of Nippostrongylus Brasiliensis (nematode). Commonwealth Agricultural Bureau, Slough, United Kingdom.
25. Klabunde, J., J. Berger, J. C. Jensenius, M. Q. Klinkert, U. E. Zelck, P. G. Kremsner, and J. F. Kun. 2000. Schistosoma mansoni: adhesion of mannan-binding lectin to surface glycoproteins of cercariae and adult worms. Exp. Parasitol. 95:231-239.
26. Lawson, B. W., Q. D. Bickle, and M. G. Taylor. 1993. Mechanisms involved in the loss of antibody-mediated adherence of macrophages to lung-stage schistosomula of Schistosoma mansoni in vitro. Parasitology 106:463-469.
27. Lindahl, G., U. Sjobring, and E. Johnsson. 2000. Human complement regulators: a major target for pathogenic microorganisms. Curr. Opin. Immunol. 12:44-51.
28. Linder, E., and G. Huldt. 1983. Antibody-independent binding and activation of complement by Schistosoma mansoni adult worms. Parasite Immunol. 5:183-194.
29. Mackenzie, C. D., M. Jungery, P. M. Taylor, and B. M. Ogilvie. 1981. The in-vitro interaction of eosinophils, neutrophils, macrophages and mast cells with nematode surfaces in the presence of complement or antibodies. J. Pathol. 133:161-175.
30. Marikovsky, M., R. Arnon, and Z. Fishelson. 1988. Proteases secreted by transforming schistosomula of Schistosoma mansoni promote resistance to killing by complement. J. Immunol. 141:273-278.
31. Marikovsky, M., F. Levi-Schaffer, R. Arnon, and Z. Fishelson. 1986. Schistosoma mansoni: killing of transformed schistosomula by the alternative pathway of human complement. Exp. Parasitol. 61:86-94.
32. Marikovsky, M., M. Parizade, R. Arnon, and Z. Fishelson. 1990. Complement regulation on the surface of cultured schistosomula and adult worms of Schistosoma mansoni. Eur. J. Immunol. 20:221-227.
33. Matsumoto, M., W. Fukuda, A. Circolo, J. Goellner, J. Strauss-Schoenberger, X. Wang, S. Fujita, T. Hidvegi, D. D. Chaplin, and H. R. Colten. 1997. Abrogation of the alternative complement pathway by targeted deletion of murine factor B. Proc. Natl. Acad. Sci. USA 94:8720-8725.
34. Meri, T., A. M. Blom, A. Hartmann, D. Lenk, S. Meri, and P. F. Zipfel. 2004. The hyphal and yeast forms of Candida albicans bind the complement regulator C4b-binding protein. Infect. Immun. 72:6633-6641.
35. Meri, T., T. S. Jokiranta, J. Hellwage, A. Bialonski, P. F. Zipfel, and S. Meri. 2002. Onchocerca volvulus microfilariae avoid complement attack by direct binding of factor H. J. Infect. Dis. 185:1786-1793.
36. Prowse, S. J., P. L. Ey, and C. R. Jenkin. 1979. Alternative pathway activation of complement by a murine parasitic nematode (Nematospiroides dubius). Aust. J. Exp. Biol. Med. Sci. 57:459-466.
37. Quaissi, M. A., J. Cornette, D. Afchain, A. Capron, H. Gras-Masse, and A. Tartar. 1986. Trypanosoma cruzi infection inhibited by peptides modeled from a fibronectin cell attachment domain. Science 234:603-607.
38. Rainbird, M. A., D. Macmillan, and E. N. Meeusen. 1998. Eosinophil-mediated killing of Haemonchus contortus larvae: effect of eosinophil activation and role of antibody, complement and interleukin-5. Parasite Immunol. 20:93-103.
39. Ramalho-Pinto, F. J. 1987. Decay accelerating factor (DAF) as the host antigen with protective activity to complement killing of schistosomula. Mem. Inst. Oswaldo Cruz 82(Suppl. 4):213-216.
40. Santoro, F., P. J. Lachmann, A. Capron, and M. Capron. 1979. Activation of complement by Schistosoma mansoni schistosomula: killing of parasites by the alternative pathway and requirement of IgG for classical pathway activation. J. Immunol. 123:1551-1557.
41. Santoro, F., M. A. Ouaissi, J. Pestel, and A. Capron. 1980. Interaction between Schistosoma mansoni and the complement system: binding of C1q to schistosomula. J. Immunol. 124:2886-2891.
42. Shaio, M. F., S. C. Hou, J. G. Chen, C. C. Wu, K. D. Yang, and F. Y. Chang. 1990. Immunoglobulin G-dependent classical complement pathway activation in neutrophil-mediated cytotoxicity to infective larvae of Angiostrongylus cantonensis. Ann. Trop. Med. Parasitol. 84:185-191.
43. Sheridan, J. W., and J. J. Finlay-Jones. 1977. Studies on a fractionated murine fibrosarcoma: a reproducible method for the cautious and a caution for the unwary. J. Cell Physiol. 90:535-552.
44. Shin, E. H., Y. Osada, J. Y. Chai, N. Matsumoto, K. Takatsu, and S. Kojima. 1997. Protective roles of eosinophils in Nippostrongylus brasiliensis infection. Int. Arch. Allergy Immunol. 114:45-50.
45. Shin, E. H., Y. Osada, H. Sagara, K. Takatsu, and S. Kojima. 2001. Involvement of complement and fibronectin in eosinophil-mediated damage to Nippostrongylus brasiliensis larvae. Parasite Immunol. 23:27-37.
46. Stadnyk, A. W., P. J. McElroy, J. Gauldie, and A. D. Befus. 1990. Characterization of Nippostrongylus brasiliensis infection in different strains of mice. J. Parasitol. 76:377-382.
47. Venturiello, S. M., G. H. Giambartolomei, and S. N. Costantino. 1995. Immune cytotoxic activity of human eosinophils against Trichinella spiralis newborn larvae. Parasite Immunol. 17:555-559.
48. Voehringer, D., K. Shinkai, and R. M. Locksley. 2004. Type 2 immunity reflects orchestrated recruitment of cells committed to IL-4 production. Immunity 20:267-277.
49. Wessels, M. R., P. Butko, M. Ma, H. B. Warren, A. L. Lage, and M. C. Carroll. 1995. Studies of group B streptococcal infection in mice deficient in complement component C3 or C4 demonstrate an essential role for complement in both innate and acquired immunity. Proc. Natl. Acad. Sci. USA 92:11490-11494.
50. Yates, J. A., G. I. Higashi, A. Lowichik, T. C. Orihel, R. C. Lowrie, Jr., and M. L. Eberhard. 1985. Activation of the alternative complement pathway in normal human serum by Loa loa and Brugia malayi infective stage larvae. Acta Trop. 42:157-163.(Paul R. Giacomin, Hui Wan)
Department of Microbiology and Infectious Diseases, Flinders Medical Centre, Adelaide, Australia
Rheumatology Section, Imperial College, Hammersmith Campus, London, United Kingdom
ABSTRACT
Complement activation and C3 deposition on the surface of parasitic helminths may be important for recruitment of leukocytes and for damage to the target organism via cell-mediated mechanisms. Inhibition of complement activation would therefore be advantageous to parasites, minimizing damage and enhancing migration through tissues. The aim of this study was to determine ex vivo if complement activation by, and leukocyte adherence to, the nematode Nippostrongylus brasiliensis change as the parasite matures and migrates through the murine host. Pathways of activation of complement and the mechanism of adherence of leukocytes were also defined using sera from mice genetically deficient in either C1q, factor B, C1q and factor B, C3, or C4. Substantive deposition of C3 and adherence of eosinophil-rich leukocytes were seen with infective-stage (L3) but not with lung-stage (L4) larvae. Adult intestinal worms had low to intermediate levels of both C3 and leukocyte binding. For L3 and adult worms, complement deposition was principally dependent on the alternative pathway. For lung-stage larvae, the small amount of C3 detected was dependent to similar degrees on both the lectin and alternative pathways. The classical pathway was not involved for any of the life stages of the parasite. These results suggest that in primary infections, the infective stage of N. brasiliensis is vulnerable to complement-dependent attack by leukocytes. However, within the first 24 h of infection, N. brasiliensis acquires the ability to largely avoid complement-dependent immune responses.
INTRODUCTION
The rapid recruitment of leukocytes to infected tissues is a crucial early step in resistance to parasitic helminths. In order to damage the target organism, many effector cells make close contact with the parasite and this can occur via complement- and antibody-dependent mechanisms (16, 29, 47). Although other mediators may also play minor roles (4, 37, 45), in the absence of parasite-specific antibody and especially in the early stages of primary infection, complement is likely to be the most important factor facilitating leukocyte recruitment and attachment to the parasite. C3b and its cleavage products facilitate attachment of leukocytes to the helminth, while C3a and C5a may act as early chemotactic factors. Eosinophils are major effector cells in host resistance to infection with a range of parasitic helminth species (14, 38, 44, 47) and adherent eosinophils release products that can damage, immobilize, and/or kill the parasite (6, 10).
Using physical or chemical strategies to deplete complement activity in vitro, parasites have been shown to activate complement by either the alternative or classical pathway (19, 28, 31, 36, 42). Although sera from humans and animals with spontaneous mutations in genes encoding individual complement proteins have also been used in studies with parasites (32, 40, 50), it has not previously been possible to comprehensively analyze each complement pathway in a single host species.
A number of pathogenic species of bacteria, viruses, and fungi have evolved strategies to evade activation of complement to ensure survival within the host (17, 23, 27, 34). Some helminths have also developed strategies to avoid complement recognition in the early stages of infection. Soon after entry into the host, Schistosoma mansoni becomes resistant to serum-dependent killing by shedding a strongly antigenic coat, acquiring host proteins, including blood group antigens and the complement regulatory protein decay accelerating factor (39, 41), and secreting proteins that bind to and inhibit complement proteins (13, 30, 32). In the early stages of infection, parasitic nematodes may also undergo changes that inhibit C3 deposition (35) and reduce the capacity of leukocytes to attach (1, 5).
Nippostrongylus brasiliensis is a nematode infecting mice and rats and has a life cycle similar to that of the human pathogen Strongyloides stercoralis. N. brasiliensis eggs hatch in feces, and after maturation to the infective stage, larvae (L3) enter the host through the skin. In experimental studies, most larvae have exited the skin within 4 h of subcutaneous injection, arriving in the lungs 18 to 48 h later and maturing into L4 (9, 24). Larvae then migrate via the trachea and esophagus to the small intestine, where they mature into adult worms. Infective N. brasiliensis L3 are susceptible to complement activation and eosinophil-dependent damage (19, 45). In the presence of serum, mouse eosinophils adhere to N. brasiliensis L3 in vitro and degranulate (29), leaving them temporarily immobilized, and when injected into mice, many fail to migrate to the lungs (8).
Interleukin-5 (IL-5) transgenic mice, which exhibit constitutive eosinophilia, are resistant to infection with N. brasiliensis and other helminths (14, 21, 44). When N. brasiliensis L3 are injected into the skin of these mice, eosinophils are recruited in large numbers and within 2 hours of infection, many are strongly adherent to the larvae (9). Larvae are trapped in the skin for an extended period and, relative to wild-type mice, few reach the lungs (9). However, those L3 that escape the skin and migrate to the lungs to develop into L4, do not elicit a strong inflammatory response, and for at least 24 to 48 h postinfection, few eosinophils are either recruited or adhere to lung-stage larvae (9).
In the first stage of this study we determined that N. brasiliensis develops a strategy to resist complement-dependent immunity as it matures from L3 to L4. C3 deposition and eosinophil-rich leukocyte adherence were compared on infective-stage L3 collected from in vitro cultures with parasites recovered from the skin, the lungs, and the small intestine. In the second stage of this study, we used sera from mice genetically deficient in either C1q, factor B, C1q and factor B (double mutant), C3, or C4 to determine that the pathways of complement activation differ as the parasite matures and migrates through the murine host.
MATERIALS AND METHODS
Animals. Mice were bred in-house under clean barrier conditions at the University of Adelaide and Imperial College, London, and handled according to institutional animal ethics committee guidelines.
Culture medium and sera. Leukocytes were cultured with parasites in RPMI 1640 culture medium (Institute of Medical and Veterinary Science, Adelaide, Australia) supplemented with 2 mM L-glutamine, 12 μg/ml gentamicin, 16 μg/ml penicillin, and 10% heat-inactivated (56°C for 30 min) fetal calf serum (Multiser, Trace Biosciences, Sydney, Australia). Serum was collected from mice by cardiac puncture under pentobarbitone sodium anesthesia. Where indicated, serum complement activity was selectively inhibited by heating for 30 min at 56°C (heat-inactivated mouse serum [MS]). Sera from wild-type CBA/Ca and C57BL/6 mice (normal mouse serum [NMS]) and from C57BL/6 mice deficient in either C1q (classical pathway deficient) (3), factor B (alternative pathway deficient) (33), C1q and factor B (classical and alternative pathway deficient), C3 (deficient in all complement pathways) (49), or C4 (classical and lectin pathway deficient) (18) were used as indicated.
Preparation of larvae and worms. Infective-stage N. brasiliensis L3 were obtained from fecal culture after passage through female Hooded Wistar rats aged 6 to 8 weeks as previously described (9). L3 were used between 9 and 28 days after establishment of fecal cultures. L3 were washed three times in phosphate-buffered saline (PBS), pH 7.4, and concentrated by centrifugation at 50 x g for 5 min. Larvae were then resuspended in PBS (immunofluorescence studies) or in culture medium (leukocyte coculture). Skin air pouches were generated as previously described (9); 750 L3 in 100 μl PBS were injected into each pouch and 30 or 150 min later, larvae were recovered by lavage and washed three times in PBS.
Lung-stage (L4) larvae were obtained after subcutaneous infection of wild-type CBA/Ca or C57BL/6 mice with approximately 750 L3 in 100 μl PBS. Either 24 or 48 h postinfection, mice were sacrificed by CO2 asphyxiation and cervical dislocation. Lungs were resected, washed in saline, minced with scissors, and incubated for 2 h at 37°C to facilitate migration of L4 out of the tissue. L4 were isolated from lung tissue by filtration through a plastic mesh strainer (approximately 1.0-mm-square mesh). Blood was removed by five washes in saline, each time allowing L4 to settle under unit gravity. L4 were then separated from the remaining finer pieces of lung tissue under a dissecting microscope using a 3.5-ml plastic transfer pipette and washed three times in PBS or culture medium, as required. Adult worms were recovered from the small intestine 7 days after subcutaneous infection of mice with 750 L3 in 100 μl PBS. Mice were sacrificed as described above, the intestine was resected, opened longitudinally with fine scissors, incubated in saline for 1 h at 37°C and worms were collected with a transfer pipette and washed five times by gravity sedimentation in PBS or culture medium.
Measurement of larvae and worms. To allow quantitative comparison of immunofluorescence of L3, L4 and intestinal worms that are substantially different in size, the surface area of each stage was estimated. Parasites were dotted on microscope slides and photographs were taken of over 20 of each stage using a 4x objective lens on an Olympus BH-2 light microscope, an Olympus C-35AD-4 camera and Konica Color Centuria 400 ISO/ASA 35-mm print film (Konica Corporation, Japan). Photographic prints were enlarged 100% using a photocopier (final magnification, 111x) and length and width of parasites were measured with a curvimetre map measure (Kartenmesser, Germany). For each parasite stage, a mean was calculated from duplicate measurements of each parasite and the overall mean and standard error of the mean were calculated from these values. Parasite surface area was estimated by calculating the area of the curved surface of a cylinder: surface area of parasite = x diameter of parasite x length of parasite.
The length, diameter, and estimated surface area of parasites life stages are shown in Table 1. The ratio of surface areas for L3, L4, and worms was approximately 1:2:14; hence in experiments where L3, L4, and worms are compared, 120 L3 or 60 L4 or 8 worms were added to each well of a microtiter tray. Since L4 taken from the lungs 24 h postinfection were only slightly smaller (approximately 10%) than those from 48 h postinfection, in each case, the same number of L4 were used per well.
Immunofluorescence microscopy. Aliquots of suspensions of parasites were dotted onto glass slides, placed under a glass coverslip and examined under UV illumination with an Olympus BH-2 microscope at 100x magnification. Photomicrographs were taken using Fujichrome Sensia 400 ISO/ASA 35-mm film (Fuji Photo Film Co. Ltd, Tokyo, Japan) with an exposure time of 16 s for anti-C3 fluorescence, or 40 s for carboxyfluorescein diacetate succinimidyl ester (CFSE)-labeled cell fluorescence and printed at a final magnification of 47x. All immunofluorescence microscopy was conducted after scanning with a phosphorimager.
Detection of C3. Using an adaptation of techniques described recently (19), parasites were incubated with PBS alone or with 50 μl of 20% NMS, MS, or serum from complement-deficient mice for 1 h at 37°C with 5% CO2 in air in a clear 96-well microtiter plate (Greiner Bio-One, Omega Scientific, CA) or in 10-ml plastic test tubes. Unbound serum components were then removed by 10 washes with 200 μl of PBS/0.05% vol/vol Tween 20 (PBST) in 96-well plates using a multichannel pipettor, or by two washes in 5 ml PBST where test tubes were used. Parasites were allowed to settle under unit gravity (96-well plates) or were centrifuged at 18 x g for 5 min (test tubes) between each wash. Parasites were then incubated with 50 μl of a 1/50 dilution of fluorescein isothiocyanate-conjugated goat anti-mouse C3 antibody (ICN/Cappel, catalog no. 55510) for 1 h at room temperature in the dark. Unbound antibody was removed by washing as described above. Finally, parasites were resuspended in 200 μl of 25% vol/vol glycerol in PBS in 96-well plates, or where parasites were incubated in test tubes, aliquots of 200 μl/well of parasite suspensions were added to the wells of a flat-bottomed, solid black 96-well plate (Costar, catalog no.# 3915, Corning Inc.).
Measurement of fluorescence intensity. Fluorescence in individual wells of solid black or clear 96-well microtiter plates was quantified in arbitrary units with a Molecular Imager FX phosphorimager (Bio-Rad USA, catalog no. 170-7800) as described previously (19). In some experiments, fluorescence intensity per parasite was calculated to reduce variation due to differences in the exact number of parasites per well.
CFSE labeling of leukocytes. Peritoneal cells from male or female heterozygous interleukin-5 (IL-5) transgenic C57BL/6 or CBA/Ca mice (15), aged 7 to 10 weeks, were used as an eosinophil-rich (55 to 75%) source of leukocytes. Mice were killed by carbon dioxide asphyxiation and peritoneal cells were harvested by lavage with mouse-osmolality PBS (MPBS) (43). Cells were washed twice in MPBS, twice in RPMI 1640 containing 0.1% vol/vol fetal calf serum and then resuspended in RPMI 1640/0.1% fetal calf serum at 107 cells/ml. Cells were incubated in 2.5 μM CFSE for 10 min at 37°C in 5% CO2 and excess CFSE was removed by washing in culture medium.
Analysis of leukocyte adherence to parasites. Adherence of CFSE-labeled leukocytes to parasites was quantitated using an adaptation of methods described previously (19). Aliquots (30 μl) of approximately 120 L3, 60 L4, or 8 worms in culture medium were mixed with 100 μl of CFSE-labeled leukocytes (106 total) and 70 μl of either culture medium, NMS, MS, or complement deficient serum (7% final concentration) was then added. After incubation at 37°C in 5% CO2 for 2 h, unbound cells were removed by nine washes in MPBS and fluorescence was measured using the phosphorimager, as described above. UV microscopy was used to visualize fluorescent cells attached to parasites.
Statistical analysis. Data for individual experiments, with each sample assayed in duplicate or triplicate are presented as mean ± standard error of the mean and were analyzed using Student's unpaired t test and GraphPad Prism software (version 3.03, GraphPad Software Inc.).
RESULTS
Detection of C3 on N. brasiliensis. The extent of C3 deposition on L3 collected from fecal culture plates was compared to that for L4 recovered from the lungs of CBA/Ca mice either 24 or 48 h postinfection and to adult worms recovered 7 days postinfection from the small intestine. Using immunofluorescence microscopy, it was apparent that C3 deposition in the presence of NMS from CBA/Ca mice was much greater on L3 than on L4 collected either 24 or 48 h after infection (Fig. 1a [i to iii]). C3 was deposited at very high levels over the entire surface of L3. In contrast, the small proportion of L4 that did exhibit some labeling had a very different and irregular pattern of deposition (Fig. 1a [ii and iii]). Intermediate but evenly distributed C3 deposition was observed on intestinal worms (Fig. 1a [iv]).
Quantitative analysis of C3 deposition (i.e., immunofluorescence/total parasite surface area) confirmed the large differences in C3 binding between L3 and L4 opsonized with NMS (Fig. 1b). Larvae typically showed relatively high autofluorescence. Nevertheless, the level of fluorescence detected on L4 incubated with NMS was no greater than that seen with L4 opsonized with serum heat-treated to inactivate C3 (i.e., MS). There were no qualitative or quantitative differences between 24- and 48-h lung larvae. Adult worms showed significant levels of C3 deposition after opsonization with NMS (Fig. 1b), but not at the same intensity as seen on L3. Host-derived C3 was not detected on either L4 or intestinal worms, as fluorescence of PBS-opsonized parasites exposed to anti-C3 antibody was no greater than background (i.e., where anti-C3 antibody was omitted).
To determine if inhibition of C3 deposition occurred soon after infection and at a time when many larvae are migrating to other tissues, we assessed fixation of endogenous and exogenous C3 on parasites recovered from skin air pouches 30 to 150 min after injection. It was difficult to accurately count and dispense larvae recovered from the skin because they tended to coalesce and so quantitative analysis was not performed. However, when assessed visually, skin larvae recovered 30 min postinjection demonstrated clear evidence of in vivo C3 deposition (Fig. 1c [iv]). When subsequently opsonized with NMS, further increases in C3 deposition were evident (Fig. 1c [vi]) similar to that seen on L3 from cultures (Fig. 1c [iii]). Skin larvae recovered 150 min postinfection were comparable to those retrieved after only 30 min (data not shown).
Adherence of leukocytes to N. brasiliensis. The adherence of eosinophil-rich CFSE-labeled peritoneal leukocytes to L3, L4 and intestinal worms opsonized with serum was determined. Large numbers of leukocytes adhered to the surface of NMS-opsonized L3 (Fig. 2a [i]), however, few if any cells bound to similarly treated lung larvae (Fig. 2a [ii and iii]). In samples stained with Vital New Red and Alcian Blue, 92% of leukocytes in the first layer of cells adhering to L3 were eosinophils, with some macrophages (6%) and the occasional mast cell also present. Quantitative analysis of leukocyte adherence using a phosphorimager clearly demonstrated the disparity between leukocyte adherence on L3 and L4 (Fig. 2b).
Large numbers of CFSE-labeled leukocytes bind to L3 and, as reported previously (19), visual assessment revealed that this was reduced to negligible levels when MS or culture medium were substituted for NMS (Fig. 2b and data not shown), indicating that leukocyte adherence was complement mediated. In contrast, few leukocytes adhered to either 24- or 48-hour L4, regardless of the opsonization conditions (Fig. 2b). Some leukocytes adhered to intestinal worms, but unlike L3, this was largely restricted to the posterior end (Fig. 2a [iv]). When assessed quantitatively, leukocyte binding to adult worms was at most only modest (Fig. 2b) but decreased significantly if MS was used instead of NMS, indicating that it was also complement-mediated.
C3 deposition and leukocyte adherence on N. brasiliensis L3 are largely dependent on the alternative complement pathway. Until recently, it has only been possible to use physical or chemical methods of depleting complement activity and these are sometimes only partially effective or may not be specific for complement proteins. It is now possible to use sera from mice that have been genetically manipulated to be deficient in one or more proteins of the complement pathway. We performed a comprehensive analysis of C3 deposition and leukocyte adherence on infective-stage larvae in the presence of NMS and MS from wild-type C57BL/6 mice and most particularly with sera from syngeneic mice genetically deficient in either C1q, factor B, C1q and factor B (double mutant), C3, or C4 (Fig. 3).
There was no significant difference in C3 deposition on L3 exposed to NMS or C1q-deficient serum (Fig. 3a), indicating that C1q and the classical pathway play no role in mediating C3 deposition on N. brasiliensis L3. In contrast, C3 deposition was almost completely abrogated in the absence of factor B (with and without C1q), indicating that the alternative pathway is paramount in this experimental system. A small amount of factor B-independent C3 binding was evident, since C3 deposition was significantly higher in the absence of factor B than it was when larvae were exposed to C3-deficient sera (C3–/– and MS). This is consistent with the observation that C3 deposition was slightly reduced when C4 was ablated, indicating a possible minor role for the C4-dependent lectin pathway.
The adherence of peritoneal leukocytes was similarly assessed in the presence of culture medium, NMS, MS or sera from complement-deficient mice. C3 was required for adherence, which was not affected by an absence of C1q (Fig. 3b). However, adherence was significantly reduced in the absence of factor B, again highlighting the importance of the alternative pathway. There were no significant differences in leukocyte adherence with factor B-deficient serum when compared to sera deficient in C3 (C3–/– and MS) and in the presence of culture medium only. The absence of C4 had no effect on cell adherence, indicating that while the lectin pathway may contribute slightly to complement deposition on L3, for complement-mediated leukocyte adherence, the alternative pathway is sufficient.
Pathway of complement activation changes as N. brasiliensis matures. We next compared C3 deposition on L3, L4 (48 h postinfection), and adult (7 days postinfection) worms in the presence of NMS and MS from wild-type C57BL/6 mice and sera from syngeneic mice genetically deficient in complement proteins (i.e., C1q, factor B, or C4; Fig. 4). MS was substituted for C3–/– serum in this experiment because both performed similarly in previous experiments.
As already demonstrated (Fig. 3a), C3 fixation on L3 was dependent on the alternative pathway; however in this experiment (Fig. 4a), the minor contribution from the lectin pathway was not evident. When comparing sera from CBA/Ca and C57BL/6 mice, we noticed a higher level of C3 deposition on L4 treated with NMS from the latter (mean fluorescence/well ± standard error of the mean, CBA/Ca versus C57BL/6, 3.29 ± 0.09 x 103 versus 6.72 ± 0.38 x 103 respectively, P < 0.001, n = 3). Interestingly, this difference was not evident when screening L3 (mean fluorescence/well ± standard error of the mean, CBA/Ca versus C57BL/6, 1.25 ± 0.08 x 104 and 1.29 ± 0.02 x 104 respectively, not significant, n = 3). Although there was a measurable difference in the level of complement activation on L4 treated with sera from these two strains, this was not associated with differences in the numbers of either lung larvae (day 2) or worms (day 7) recovered (Mean no. of parasites ± standard error of the mean for C57BL/6 and CBA mice [n = 3 to 4] respectively. L4: 323 ± 47 versus 289 ± 44, P > 0.05. Intestinal worms: 254 ± 99 versus 164 ± 33, P > 0.05.).
Although still modest compared to C3 fixation on L3 analyzed in the same experiment (Fig. 4a), the complement deposition on L4 treated with NMS from wild-type C57BL/6 mice was greatly diminished when sera from mice deficient in either factor B or C4 were used (Fig. 4b), indicating that both the alternative and lectin pathways contribute to C3 activation on lung-stage larvae. In three separate experiments, C3 deposition on L4 was significantly less with serum from C4–/– than from factor B–/– mice (Fig. 4b and data not shown) and so for lung-stage larvae, the lectin pathway may be at least marginally more important. The pathway of complement activation for adult worms was similar to that for L3, since deposition of C3 was in both instances significantly reduced in sera from factor B–/– mice (Fig. 4c). For adult worms, some C3 deposition would appear to be independent of the alternative pathway. For all of the life stages assessed, the classical pathway (C1q) was not involved.
DISCUSSION
Evasion of potentially damaging host immune responses is essential for helminths to be successful parasites. In one sense, the parasite is likely to be at its most vulnerable upon entry into the host, principally because it must quickly change from a form suited either to a free-living stage or to residence in another host. Conversely, a nave host is particularly vulnerable in the first few days of infection, since it is totally or largely dependent on innate mechanisms of resistance. In this and our previous studies (9, 14, 19), we have shown the infective-stage L3 of N. brasiliensis to be extremely vulnerable to complement-dependent attack by eosinophils. Complement activation on N. brasiliensis L3 occurs primarily via the alternative pathway. However within the first 24 h of infection, during maturation from L3 to lung-stage L4, N. brasiliensis acquires the ability to evade complement-dependent immune responses. This may explain why the inflammatory response at the site of initial infection, the skin, is rapid and substantive, whereas that in the lungs is slow to develop. The eosinophil chemotactic factors C3a and C5a may be responsible for rapid recruitment of eosinophils into the skin, but are less likely to be available when the lungs become infected. Interestingly, there are major changes in the pathways of complement activation as N. brasiliensis matures, with the lectin pathway becoming more important for the limited levels of complement activation on lung-stage larvae.
Innate resistance of wild-type mice to primary infection with N. brasiliensis is typically very low for most strains, with a large proportion of the infecting dose (>50%) migrating through the lungs and colonizing the small intestine (14). In addition, since the parasite has a very short transit time through the host (approximately 9 to 14 days), the life cycle is completed before an effective adaptive immune response can be generated. Our studies suggest that provided sufficient and appropriate effector cells can be recruited, innate resistance against N. brasiliensis is very effective (9, 14). Eosinophils appear to be a potent force in trapping larvae in the skin in primary infections (9) and this study indicates that C3 is almost exclusively responsible for facilitating stable attachment of these and other leukocytes to the target. Our data also suggest that complement may be important for recruitment of leukocytes, since in the absence of C3 deposition on L4 (Fig. 1), few leukocytes are to be found in the lungs until after the majority of larvae have migrated to the gut (7, 9, 48). Although chemokines such as the eosinophil-specific factor eotaxin are produced in the lungs, it is clear that these do not initially compensate for the failure of complement-dependent recruitment of eosinophils or other leukocytes into this organ.
We have yet to determine how long the larvae are in the host before acquiring the ability to avoid complement activation and leukocyte adherence, though clearly it occurs sometime between 2.5 and 24 h postinfection. Some tissue-invasive parasitic helminths inhibit adherence of leukocytes after a single maturational stage (1, 5, 11, 22, 26). Leukocytes adhere to L3 but not to L4 of Dirofilaria immitis, possibly due to acquisition of host-derived inhibitory proteins (1). The exact mechanism of how lung-stage N. brasiliensis larvae avoid fixing complement or binding leukocytes is not fully understood. N. brasiliensis L3 and L4 exsheath during maturation and the third larval molt occurs within the lungs approximately 24 to 32 h postinfection (24). One or more of these exsheathments may remove parasite-derived molecules that activate the alternative pathway or expose additional binding sites for complement regulatory proteins such as factor H, which has been shown to be important for evasion of complement by Onchocerca volvulus (35). Finally, excretory/secretory proteins produced by L4 could also interfere with host defenses. Treatment of serum with excretory/secretory proteins from the nematode Toxocara canis has previously been shown to inhibit complement deposition on the surface of T. canis larvae (2). The potential mechanism(s) used by N. brasiliensis L4 to inhibit C3 deposition is the subject of further investigation in our laboratory.
Using sera from mice genetically deficient in one or more of the complement proteins, we have comprehensively and for the first time defined which complement pathways are most important for both complement activation and leukocyte adherence on a parasitic helminth. The alternative pathway is of prime importance for both C3 deposition and leukocyte adherence to L3 in vitro, but also makes a significant contribution to the smaller amounts of C3 found on both L4 and adult worms. The broader relevance of C3 deposition on adult worms is questionable, since complement may not be available or active in the gut, except perhaps where parasites attach to, erode, and/or feed on the gut wall. Activation of the classical pathway had no role in mediating C3 deposition on N. brasiliensis L3, L4, or intestinal worms, indicating that molecules capable of directly binding C1q and activating this pathway are not expressed by N. brasiliensis.
Although largely unexplored for helminths, activation via the lectin pathway has been detected on Trichinella spiralis (20) and mannan-binding lectin adheres to schistosomes (25). Complement activation by the lectin pathway was responsible for only a small proportion of the C3 deposited on N. brasiliensis L3 and adult worms. However, of the small amount of C3 detected on L4, the greatest contribution came from the lectin pathway. This stage-specific change in the relative importance of this pathway is novel and warrants further investigation.
Since L4 did not appear to fix C3 in vivo, it is curious that a small amount could be detected after incubation ex vivo with C57BL/6 serum. This occurred regardless of whether the L4 were derived from C57BL/6 or CBA hosts (data not shown). It is also odd that L4 did not acquire C3 when exposed to CBA/Ca serum in vitro. These may be in vitro phenomena of little biological significance and any explanation we might offer is purely speculative. However, complement inhibitors (host or parasite derived) may be more active with CBA serum and/or in vivo in both strains. C57BL/6 serum may have more available C3, but this seems unlikely, since these differences were not evident when L3 were tested. Importantly, we and others have not detected major differences in parasite burdens when infections in CBA and C57 strains are directly compared (12, 46).
In conclusion, we have demonstrated that infective-stage larvae of N. brasiliensis activate the alternative pathway of complement, mediating a high level of leukocyte adherence to these larvae. Complement fixation at this stage of the parasite's life cycle may also be important for recruitment of eosinophils and other leukocytes into the initial site of infection, the skin. Entrapment of larvae may then follow, provided that sufficient effector cells are present. However, within 24 h of infection, the parasite develops the capacity to inhibit deposition of C3, and this could explain the relative absence of an early inflammatory response to L4 in the lungs. In acquiring resistance at this level, larvae may more readily migrate through tissues to undergo further maturation. Interestingly, there is a shift in pathways through which the small amount of complement is activated by lung-stage L4, with the lectin pathway now being a relatively major contributor. As the parasite enters the gut and undergoes further maturation to the adult worm stage, the alternative pathway again becomes more important for complement activation. However, both innate and early adaptive immune responses may be of less consequence to a parasite that resides in the lumen of the gut.
Although we have established in vitro that complement is at least transiently important, a critical role in innate resistance to parasitic helminths in vivo has yet to be conclusively determined. More definitive proof will require investigations of the kinetics of infections in the various complement-deficient mice described here.
ACKNOWLEDGMENTS
This work was supported in part by the National Health and Medical Research Council (Australia) and School and Faculty Strategic Research Funds from the University of Adelaide.
We thank Bruce Lyons for assistance in establishing CFSE labeling techniques.
REFERENCES
1. Abraham, D., R. B. Grieve, and M. Mika-Grieve. 1988. Dirofilaria immitis: surface properties of third- and fourth-stage larvae. Exp. Parasitol. 65:157-167.
2. Badley, J. E., R. B. Grieve, J. H. Rockey, and L. T. Glickman. 1987. Immune-mediated adherence of eosinophils to Toxocara canis infective larvae: the role of excretory-secretory antigens. Parasite Immunol. 9:133-143.
3. Botto, M., C. Dell'Agnola, A. E. Bygrave, E. M. Thompson, H. T. Cook, F. Petry, M. Loos, P. P. Pandolfi, and M. J. Walport. 1998. Homozygous C1q deficiency causes glomerulonephritis associated with multiple apoptotic bodies. Nat. Genet. 19:56-59.
4. Brattig, N. W., A. Z. Abakar, F. Geisinger, and T. F. Kruppa. 1995. Cell-adhesion molecules expressed by activated eosinophils in Onchocerca volvulus infection. Parasitol. Res. 81:398-402.
5. Brattig, N. W., F. W. Tischendorf, G. Strote, and C. E. Medina-de la Garza. 1991. Eosinophil-larval-interaction in onchocerciasis: heterogeneity of in vitro adherence of eosinophils to infective third and fourth stage larvae and microfilariae of Onchocerca volvulus. Parasite Immunol. 13:13-22.
6. Butterworth, A. E. 1984. Cell-mediated damage to helminths. Adv. Parasitol. 23:143-235.
7. Daly, C. M. 1999. Ph.D. thesis. University of Adelaide, Adelaide, Australia.
8. Daly, C. M., B. Li, J. Forbes, P. R. Giacomin, and L. A. Dent. 2004. Eosinophils as mediators of early resistance in mice to the nematode Nippostrongylus brasiliensis. Clin. Investig. Med. 27:99D.
9. Daly, C. M., G. Mayrhofer, and L. A. Dent. 1999. Trapping and immobilization of Nippostrongylus brasiliensis larvae at the site of inoculation in primary infections of interleukin-5 transgenic mice. Infect. Immun. 67:5315-5323.
10. David, J. R., A. E. Butterworth, and M. A. Vadas. 1980. Mechanism of the interaction mediating killing of Schistosoma mansoni by human eosinophils. Am. J. Trop. Med. Hyg. 29:842-848.
11. Davis, S. W., and B. Hammerberg. 1990. Taenia taeniaeformis: evasion of complement-mediated lysis by early larval stages following activation of the alternative pathway. Int. J. Parasitol. 20:587-593.
12. Dehlawi, M. S., P. K. Goyal, A. W. Stadnyk, P. J. McElroy, J. Gauldie, and A. D. Befus. 2003. Responses of inbred mouse strains to infection with intestinal nematodes. J. Helminthol. 77:119-124.
13. Deng, J., D. Gold, P. T. LoVerde, and Z. Fishelson. 2003. Inhibition of the complement membrane attack complex by Schistosoma mansoni paramyosin. Infect. Immun. 71:6402-6410.
14. Dent, L. A., C. M. Daly, G. Mayrhofer, T. Zimmerman, A. Hallett, L. P. Bignold, J. Creaney, and J. C. Parsons. 1999. Interleukin-5 transgenic mice show enhanced resistance to primary infections with Nippostrongylus brasiliensis but not primary infections with Toxocara canis. Infect. Immun. 67:989-993.
15. Dent, L. A., M. Strath, A. L. Mellor, and C. J. Sanderson. 1990. Eosinophilia in transgenic mice expressing interleukin 5. J. Exp. Med. 172:1425-1431.
16. Desakorn, V., P. Suntharasamai, S. Pukrittayakamee, S. Migasena, and D. Bunnag. 1987. Adherence of human eosinophils to infective filariform larvae of Necator americanus in vitro. Southeast Asian J. Trop. Med. Public Health 18:66-72.
17. Duthy, T. G., R. J. Ormsby, E. Giannakis, A. D. Ogunniyi, U. H. Stroeher, J. C. Paton, and D. L. Gordon. 2002. The human complement regulator factor H binds pneumococcal surface protein PspC via short consensus repeats 13 to 15. Infect. Immun. 70:5604-5611.
18. Fischer, M. B., M. Ma, S. Goerg, X. Zhou, J. Xia, O. Finco, S. Han, G. Kelsoe, R. G. Howard, T. L. Rothstein, E. Kremmer, F. S. Rosen, and M. C. Carroll. 1996. Regulation of the B cell response to T-dependent antigens by classical pathway complement. J. Immunol. 157:549-556.
19. Giacomin, P. R., H. Wang, D. L. Gordon, and L. A. Dent. 2004. Quantitation of complement and leukocyte binding to a parasitic helminth species. J. Immunol. Methods 289:201-210.
20. Gruden-Movsesijan, A., M. Petrovic, and L. Sofronic-Milosavljevic. 2003. Interaction of mannan-binding lectin with Trichinella spiralis glycoproteins, a possible innate immune mechanism. Parasite Immunol. 25:545-552.
21. Herbert, D. R., J. J. Lee, N. A. Lee, T. J. Nolan, G. A. Schad, and D. Abraham. 2000. Role of IL-5 in innate and adaptive immunity to larval Strongyloides stercoralis in mice. J. Immunol. 165:4544-4551.
22. Hong, Y., C. W. Kim, and B. Ghebrehiwet. 1992. Trichinella spiralis: activation of complement by infective larvae, adults, and newborn larvae. Exp. Parasitol. 74:290-299.
23. Horstmann, R. D., H. J. Sievertsen, J. Knobloch, and V. A. Fischetti. 1988. Antiphagocytic activity of streptococcal M protein: selective binding of complement control protein factor H. Proc. Natl. Acad. Sci. USA 85:1657-1661.
24. Kassai, T. 1982. Handbook of Nippostrongylus Brasiliensis (nematode). Commonwealth Agricultural Bureau, Slough, United Kingdom.
25. Klabunde, J., J. Berger, J. C. Jensenius, M. Q. Klinkert, U. E. Zelck, P. G. Kremsner, and J. F. Kun. 2000. Schistosoma mansoni: adhesion of mannan-binding lectin to surface glycoproteins of cercariae and adult worms. Exp. Parasitol. 95:231-239.
26. Lawson, B. W., Q. D. Bickle, and M. G. Taylor. 1993. Mechanisms involved in the loss of antibody-mediated adherence of macrophages to lung-stage schistosomula of Schistosoma mansoni in vitro. Parasitology 106:463-469.
27. Lindahl, G., U. Sjobring, and E. Johnsson. 2000. Human complement regulators: a major target for pathogenic microorganisms. Curr. Opin. Immunol. 12:44-51.
28. Linder, E., and G. Huldt. 1983. Antibody-independent binding and activation of complement by Schistosoma mansoni adult worms. Parasite Immunol. 5:183-194.
29. Mackenzie, C. D., M. Jungery, P. M. Taylor, and B. M. Ogilvie. 1981. The in-vitro interaction of eosinophils, neutrophils, macrophages and mast cells with nematode surfaces in the presence of complement or antibodies. J. Pathol. 133:161-175.
30. Marikovsky, M., R. Arnon, and Z. Fishelson. 1988. Proteases secreted by transforming schistosomula of Schistosoma mansoni promote resistance to killing by complement. J. Immunol. 141:273-278.
31. Marikovsky, M., F. Levi-Schaffer, R. Arnon, and Z. Fishelson. 1986. Schistosoma mansoni: killing of transformed schistosomula by the alternative pathway of human complement. Exp. Parasitol. 61:86-94.
32. Marikovsky, M., M. Parizade, R. Arnon, and Z. Fishelson. 1990. Complement regulation on the surface of cultured schistosomula and adult worms of Schistosoma mansoni. Eur. J. Immunol. 20:221-227.
33. Matsumoto, M., W. Fukuda, A. Circolo, J. Goellner, J. Strauss-Schoenberger, X. Wang, S. Fujita, T. Hidvegi, D. D. Chaplin, and H. R. Colten. 1997. Abrogation of the alternative complement pathway by targeted deletion of murine factor B. Proc. Natl. Acad. Sci. USA 94:8720-8725.
34. Meri, T., A. M. Blom, A. Hartmann, D. Lenk, S. Meri, and P. F. Zipfel. 2004. The hyphal and yeast forms of Candida albicans bind the complement regulator C4b-binding protein. Infect. Immun. 72:6633-6641.
35. Meri, T., T. S. Jokiranta, J. Hellwage, A. Bialonski, P. F. Zipfel, and S. Meri. 2002. Onchocerca volvulus microfilariae avoid complement attack by direct binding of factor H. J. Infect. Dis. 185:1786-1793.
36. Prowse, S. J., P. L. Ey, and C. R. Jenkin. 1979. Alternative pathway activation of complement by a murine parasitic nematode (Nematospiroides dubius). Aust. J. Exp. Biol. Med. Sci. 57:459-466.
37. Quaissi, M. A., J. Cornette, D. Afchain, A. Capron, H. Gras-Masse, and A. Tartar. 1986. Trypanosoma cruzi infection inhibited by peptides modeled from a fibronectin cell attachment domain. Science 234:603-607.
38. Rainbird, M. A., D. Macmillan, and E. N. Meeusen. 1998. Eosinophil-mediated killing of Haemonchus contortus larvae: effect of eosinophil activation and role of antibody, complement and interleukin-5. Parasite Immunol. 20:93-103.
39. Ramalho-Pinto, F. J. 1987. Decay accelerating factor (DAF) as the host antigen with protective activity to complement killing of schistosomula. Mem. Inst. Oswaldo Cruz 82(Suppl. 4):213-216.
40. Santoro, F., P. J. Lachmann, A. Capron, and M. Capron. 1979. Activation of complement by Schistosoma mansoni schistosomula: killing of parasites by the alternative pathway and requirement of IgG for classical pathway activation. J. Immunol. 123:1551-1557.
41. Santoro, F., M. A. Ouaissi, J. Pestel, and A. Capron. 1980. Interaction between Schistosoma mansoni and the complement system: binding of C1q to schistosomula. J. Immunol. 124:2886-2891.
42. Shaio, M. F., S. C. Hou, J. G. Chen, C. C. Wu, K. D. Yang, and F. Y. Chang. 1990. Immunoglobulin G-dependent classical complement pathway activation in neutrophil-mediated cytotoxicity to infective larvae of Angiostrongylus cantonensis. Ann. Trop. Med. Parasitol. 84:185-191.
43. Sheridan, J. W., and J. J. Finlay-Jones. 1977. Studies on a fractionated murine fibrosarcoma: a reproducible method for the cautious and a caution for the unwary. J. Cell Physiol. 90:535-552.
44. Shin, E. H., Y. Osada, J. Y. Chai, N. Matsumoto, K. Takatsu, and S. Kojima. 1997. Protective roles of eosinophils in Nippostrongylus brasiliensis infection. Int. Arch. Allergy Immunol. 114:45-50.
45. Shin, E. H., Y. Osada, H. Sagara, K. Takatsu, and S. Kojima. 2001. Involvement of complement and fibronectin in eosinophil-mediated damage to Nippostrongylus brasiliensis larvae. Parasite Immunol. 23:27-37.
46. Stadnyk, A. W., P. J. McElroy, J. Gauldie, and A. D. Befus. 1990. Characterization of Nippostrongylus brasiliensis infection in different strains of mice. J. Parasitol. 76:377-382.
47. Venturiello, S. M., G. H. Giambartolomei, and S. N. Costantino. 1995. Immune cytotoxic activity of human eosinophils against Trichinella spiralis newborn larvae. Parasite Immunol. 17:555-559.
48. Voehringer, D., K. Shinkai, and R. M. Locksley. 2004. Type 2 immunity reflects orchestrated recruitment of cells committed to IL-4 production. Immunity 20:267-277.
49. Wessels, M. R., P. Butko, M. Ma, H. B. Warren, A. L. Lage, and M. C. Carroll. 1995. Studies of group B streptococcal infection in mice deficient in complement component C3 or C4 demonstrate an essential role for complement in both innate and acquired immunity. Proc. Natl. Acad. Sci. USA 92:11490-11494.
50. Yates, J. A., G. I. Higashi, A. Lowichik, T. C. Orihel, R. C. Lowrie, Jr., and M. L. Eberhard. 1985. Activation of the alternative complement pathway in normal human serum by Loa loa and Brugia malayi infective stage larvae. Acta Trop. 42:157-163.(Paul R. Giacomin, Hui Wan)