INFECTION AND IMMUNITY,0019-9567/99/$04.0010
Sept. 1999, p. 4661–4667 Vol. 67, No. 9
Copyright © 1999, American Society for Microbiology. All Rights
Dominance of Immunoglobulin G2c in the
AntiphosphorylcholineResponse of Rats Infected with Trichinella
PHILIP J. PETERS,1† LUCILLE F. GAGLIARDO,1 ELIZABETH A. SABIN,2‡
ARIN B. BETCHEN,1§KAYA GHOSH,1 JEB B. OBLAK,1 AND JUDITH A.
James A. Baker Institute for Animal Health1 and Department of
Microbiology and Immunology,2
College of Veterinary Medicine, Cornell University, Ithaca, New
Received 12 March 1999/Returned for modification 11 May
1999/Accepted 3 June 1999
The antibody response to the L1 stage of Trichinella spiralis
has been described as biphasic. Worms residentin the intestine
during the first week of infection stimulate an antibody response
against a subset of larvalproteins. L1 larvae in the muscle at the
end stage of infection stimulate a second antibody response
againsttyvelose-bearing glycoproteins. Antityvelose antibodies
protect rats against challenge infection with larvae. Theaim of
this study was to characterize the rat B-cell response against
larval antigens during the intestinal phaseof T. spiralis infection
and to test the antiparasitic effects of such antibodies. Strain
PVG rats were infectedorally with 500 larvae. Antibodies specific
for phosphorylcholine-bearing proteins of L1 larvae first
appearedin serum 9 days postinfection. Absorption experiments
showed that the majority of antilarval antibodiesproduced in rats
16 days after infection with T. spiralis were specific for
phosphorylcholine-bearing proteins.A fraction of these antibodies
bound to free phosphorylcholine. Immunoglobulin G2c (IgG2c)
producing cellsin the mesenteric lymph node dominated this early
antibody response. IgG2c is associated with T-independentimmune
responses in the rat; however, a comparison of athymic rats with
euthymic controls suggested that onlya small fraction of the
phosphorylcholine-related antibody response against T. spiralis was
T independent.Phosphorylcholine is a common epitope in antigens of
bacteria and nematode parasites and has been shownto be a target of
protective immunity in certain bacteria. A monoclonal IgG2c
antibody was prepared frominfected rats and shown to be specific
for phosphorylcholine. Monoclonal phosphorylcholine-specific
IgG2cfailed to protect rats against intestinal infection with T.
spiralis. Therefore, our findings do not support a rolefor
phosphorylcholine-bearing antigens in immune defense against T.
spiralis; however, the potency of theimmune response induced
suggests an immunomodulatory role for the lymphocytes involved.
Trichinella spiralis parasitizes many mammalian species andhas
served as a model for the study of parasitic nematodes andmucosal
immunity. The rat is a natural host for T. spiralis.Infective
larvae (L1 stage) are freed in the stomach when a ratconsumes an
infected carcass. Larvae enter the small intestine,where they
undergo a series of four molts in a period of 30 h.Worms reside in
an epithelial habitat (41). Adult worms re-produce, and newborn
larvae migrate from the gut to striatedmuscle, where they invade
cells and mature to the infectious L1stage (reviewed by Despommier
Different antigens are expressed during each life stage of
T.spiralis. Philipp et al. (28) showed that the proteins
displayedon the epicuticle change with each molt. Jungery and
Ogilvie(16) reported that the host antibody response mirrors
thechanges in antigen expression. In addition to the
antigenicvariation observed in different life stages, Denkers et
al. re-ported that the antibody response directed against L1 larvae
isbiphasic (8). Of the two responses, the one that has beenstudied
most intensively is induced late in infection by muscle-stage L1
larvae. Antibodies are induced by glycans attached to
a number of larval proteins found in the secretory cells,
orstichocytes, as well as on the body surface (4, 12). These
gly-cans are tri- and tetra-antennary structures that are
cappedwith 3,6-dideoxy-D-arabino-hexose (tyvelose) (11, 31, 40).
Ty-velose forms the immunodominant epitope. This response
isimportant to the survival of host and parasite, as shown
bypassive immunization of rats with antityvelose MAbs. Antibod-ies
eliminate up to 99% of an oral challenge dose of infectiveT.
spiralis larvae within a few hours. IgG1 and IgG2c appear tobe the
most protective antibody subclasses (1, 3, 5).
The other phase of the antilarval response is less well
char-acterized, but indirect evidence suggests that it is specific
forphosphorylcholine-bearing glycoproteins (35). Phosphorylcho-line
is widely distributed in tissues, including the cuticle,
epi-dermis, hypodermis, hemolymph, and intestinal gland (15).Ubeira
et al. (36) investigated the splenic B-cell response in-duced by
phosphorylcholine antigens in mice infected withTrichinella;
however, the more relevant, intestinal B-cell re-sponse has not
been studied. Phosphorylcholine immunogenshave been found in
bacteria and nematodes and serve as tar-gets of protective immunity
against certain bacteria (19). Inthis report, we describe the rat
B-cell response against phos-phorylcholine-bearing glycoproteins
during the intestinalphase of T. spiralis infection as well as
results of experiments totest the antiparasitic effects of such
(Some of these data were presented at the 75th AnnualMeeting of
the Conference of Research Workers in AnimalDiseases, Chicago, Ill.
[abstract no. 142], and at the 9th Inter-national Congress on
Immunology, San Francisco, Calif. [pro-gram no. 931].)
* Corresponding author. Mailing address: James A. Baker
Institutefor Animal Health, College of Veterinary Medicine, Cornell
Univer-sity, Ithaca, NY 14853. Phone: (607) 256-5600. Fax: (607)
256-5608.E-mail: [email protected]
† Present address: College of Medicine, Cornell University,
NewYork, NY 10021.
‡ Present address: American Veterinary Medical
Association,Schaumburg, IL 60173-4360.
§ Present address: Department of Biomedical Sciences, San
Fran-cisco, CA 94143.
on March 14, 2020 by guest
MATERIALS AND METHODS
Abbreviations. ASC, antibody-secreting cells; BSA, bovine serum
albumin;CLN, cervical lymph node; DPBS, Dulbecco’s
phosphate-buffered saline; ECL,enhanced chemiluminescence; ELISA,
enzyme-linked immunosorbent assay;ELISPOT, enzyme-linked
immunospot; HRP, horseradish peroxidase; IFN-g,gamma interferon;
Ig, immunoglobulin; IL, interleukin; MAb, monoclonal anti-body;
MLN, mesenteric lymph node; TI-2, thymus-independent type 2.
Rats. Strain AO and strain PVG rats were produced and maintained
underspecific-pathogen-free conditions. Outbred, male NIH-RNU
(athymic) andNIH-RNU/1 (euthymic) rats were obtained from the
Frederick Cancer Re-search and Development Center and were barrier
maintained. All rats werehoused in the James A. Baker Institute
vivarium according to American Asso-ciation for Accreditation of
Laboratory Animal Care guidelines.
Parasite. T. spiralis (pig strain) was maintained in irradiated
adult rats. Infec-tious larvae were harvested from muscle by
digestion in a solution of 1% pep-sin–1% HCl at 37°C. Experimental
rats were infected orally with larvae sus-pended in a volume of 0.2
to 0.5 ml of 0.6% nutrient broth–2% gelatin (2).
Antigen. Crude larval antigen and excretory/secretory antigens
of muscle lar-vae were prepared as previously described (4).
Phosphorylcholine-bearing anti-gens were isolated from crude
antigen by affinity chromatography with MAb 6G3(see below).
Antibodies. Normal rat serum was collected by cardiac puncture
from adultrats anesthetized with ether. Sera were collected at the
times specified (see
Results) from rats infected with 500 or 1,000 T. spiralis
larvae. Monoclonal ratantibody 9E6 (IgG2c) is specific for
tyvelose-capped glycans of T. spiralis (3, 11).
Mouse MAbs specific for rat IgA, IgM, and HRP-conjugated
polyclonalanti-rat IgG2c or IgE were obtained from Serotec (Harlan,
Indianapolis,Ind.). Mouse MAbs specific for rat IgG2a, IgG2b, and
IgG1 were a gift fromT. Springer (Harvard University, Boston,
Monoclonal rat antibody 6G3 was produced by somatic cell
hybridization (17).MLN cells were collected from a PVG rat which
had been infected orally with2,500 larvae 10 days previously. The
MLN cells were processed in the mannerdescribed below (“Preparation
of Cells”). Rat lymphocytes and mouse myelomacells (SP2/0) were
fused at a ratio of 2:1. Fused cells were plated in 96-well
plateson rat peritoneal exudate cells. Hybridomas were screened by
ELISA in order todetect IgG2c specific for crude T. spiralis larval
antigen. Positive hybridomaswere cloned by limiting dilution on rat
peritoneal exudate cells. Ascites fluid wasproduced in nude mice by
Harlan Sprague-Dawley (Indianapolis, Ind.).
ELISA. Crude larval antigen (5 mg/ml, 50 ml/well) was incubated
in polyvinylmicrotiter plates overnight at 4°C. Wells were blocked
with DPBS containing0.2% gelatin for 15 min. The remainder of the
procedure was performed asdescribed previously (3), except that for
hybridoma screening, antibody bindingwas detected with
peroxidase-conjugated rabbit antibody specific for rat
In one experiment, pooled serum (collected from rats 16 days
postinfection) orMAb 6G3 was absorbed with MAb 6G3
affinity-purified antigens or with BSA by
FIG. 1. Kinetics of the T. spiralis-specific ASC response in
three lymphoid tissues: MLN (A), CLN (B), and spleen (C). PVG rats
were infected with an oral doseof 867 T. spiralis muscle larvae.
The numbers of specific ASC were determined by ELISPOT on days 8,
16, 30, and 50 postinfection. Values expressed are the means
1standard deviations (for three rats or, on day 50, two rats). p,
IgE was not assayed in this experiment.
4662 PETERS ET AL. INFECT. IMMUN.
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incubating antibody solutions on antigen-coated (5 mg of
protein/ml) petri dishesfor 2 h at 4°C. Absorbed antibody
preparations were tested by ELISA for bindingto crude larval
antigen. Antibody binding was detected by using
peroxidase-con-jugated goat antibody specific for rat IgG (heavy-
and light-chain reactive) (3).
Preparation of cells. MLNs, CLNs, spleens, and blood were
collected frominfected rats at intervals following infection. Lymph
nodes and spleens weredissected to remove the fat, cut into several
pieces, and pressed through a sterilesieve. Cell suspensions were
filtered through a cell strainer (Becton Dickinson,Lincoln Park,
N.J.), diluted to 25 ml in DPBS–0.05% BSA (Sigma, St. Louis,Mo.),
and centrifuged at 153 3 g for 10 min at 4°C. This step was
repeated twice.The cells were resuspended, counted (Coulter Counter
ZM; Coulter Electronics,Hialeah, Fla.), centrifuged, and
resuspended at 107 cells/ml in RPMI mediumcontaining 5% fetal calf
serum, 50 U of penicillin per ml, 50 mg of streptomycinper ml, and
1 mM sodium pyruvate (all from Gibco, Grand Island, N.Y.), as
wellas 2 mM L-glutamine and 50 mM 2-mercaptoethanol (Sigma).
ELISPOT. Tissue culture plates (96 well) with nitrocellulose
filters (Millipore,Bedford, Mass.) were incubated for 24 h at 4°C
with 50 ml of crude antigen (100mg/ml) in DPBS. The filters were
washed with DPBS and then blocked for 15 minin DPBS–1% dry milk.
The plates were washed twice with DPBS, and 100 ml ofcomplete RPMI
medium was added to each well, together with 100 ml of
theappropriate cell suspension. Cells were tested at three
dilutions. Each dilutionwas tested in duplicate. The plates were
left at room temperature for 5 min andthen incubated for 3 h at
37°C in 5% CO2. The filters were washed three timeswith DPBS and
incubated with unconjugated mouse anti-rat isotype antibodiesor
with HRP-conjugated anti-IgG2c or anti-IgE for 1 h at 4°C with
agitation.Filters were washed three times in DPBS–0.05% BSA. For
detection of IgG1,IgG2a, IgG2b, IgM, and IgA, filters were next
incubated with agitation for 1 h at4°C with goat anti-mouse IgG
conjugated to HRP (Southern Biotech, Birming-ham, Al.) diluted in
DPBS–0.05% BSA plus 10% normal rat serum. For detec-tion of IgE,
the signal was amplified further by an additional incubation with
anHRP-conjugated antiperoxidase antibody. The filters were washed
three times inDPBS–0.05% Tween 20 (Sigma) and incubated with
4-chloro-naphthol (Sigma)for 30 min at room temperature with
Dark purple spots, representing ASC, were counted with a Nikon
dissectingmicroscope (63) fitted with a Sony DXC-107 video camera
attached to a videodisplay. Wells were counted if they contained
between 1 and 200 spots per well.Data were expressed as the mean
numbers of ASC per tissue 6 1 standarddeviation. Hybridoma cells
that produce anti-T. spiralis MAbs of known isotypes(3) were used
as positive controls and to confirm the specificity of the
Polyacrylamide gel electrophoresis and Western blotting. Ten
percent poly-acrylamide gels with 3% stacking gels in standard
format or miniformat (as
noted) were used. Samples were boiled for 3 to 5 min prior to
loading in samplepreparation buffer (0.0625 M Tris–2% sodium
dodecyl sulfate–10% glycerol [pH6.8]) containing 5%
2-mercaptoethanol. Prestained molecular weight
standards(Diversified Biotech, Newton Center, Mass.) were used. Gel
electrophoresis andWestern transfer to nitrocellulose were
performed according to the methodsdescribed by Burnette (7).
Nitrocellulose was blocked with DPBS–0.2% Tween–7.5% dry milk–2%
BSA (Sigma). Antibodies were diluted in DPBS–0.2%Tween–2% dry
milk–0.25% BSA. The nitrocellulose was washed in DPBS–0.2%Tween.
ECL reagent (Amersham Life Science, Little Chalfont,
Buckingham-shire, England) was prepared according to the
manufacturer’s instructions. Inone experiment, primary antibodies
were prepared in the presence of phospho-rylcholine chloride
Passive immunization of rats. Fourteen-day-old AO rats were
injected intra-peritoneally with antibody (2.5 mg/20 g of body
weight) 90 min prior to oralinfection with 185 larvae. Twelve rats
each were treated with serum immuno-globulin from uninfected rats,
antityvelose MAb 9E6 (IgG2c), or MAb 6G3(IgG2c). Worms were
harvested from intestines on days 1, 8, and 10 afterinfection
according to methods that have been described (2).
Statistical analyses. Student’s t test was used to compare means
for treatmentgroups. Analysis of variance and Tukey’s least
significant difference test wereused to detect differences among
three or more means. Differences were con-sidered significant when
P was ,0.05.
IgG2c ASC dominated the early immune response. We usedcrude
larval antigen in the ELISPOT. Repeated attempts todetect ASC
specific for larval excretory/secretory antigens(which are largely
tyvelose-bearing glycoproteins) during theintestinal stage of
infection were unsuccessful. Thus, antigensrecognized in crude
larval homogenates by ASC were distinctfrom those found in larval
excretory/secretory antigens andwere not tyvelose-bearing
glycoproteins. The presence oftyvelose-bearing proteins in crude
antigen was confirmed byELISPOT with specific hybridomas (not
We assayed parasite-specific ASC in three lymphoid tissueson
days 8, 16, 30, and 50 postinfection (Fig. 1). The MLN
FIG. 2. Comparison of the kinetics of the T. spiralis-specific
ASC responses in the MLNs of euthymic (haired) and athymic (nude)
rats. Rats were infected with anoral dose of 500 T. spiralis muscle
larvae. The numbers of specific ASC were determined by ELISPOT on
days 8 and 16 postinfection. Values expressed are the means
1standard deviations (n 5 three rats).
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dominated the antibody response, generating 50 times theASC
found in spleen or CLN. Over 105 T. spiralis-specific ASCwere
detected in MLN 8 and 16 days after infection. The peakactivity in
the spleen and CLN did not occur until day 16. ASCnumbers in all
tissues decreased more than 10-fold betweendays 16 and 30.
IgG2c was the dominant isotype in the MLN, comprising55% of the
total ASC on day 8 and 89% of the total ASC onday 16 (Fig. 1A).
IgG2b made a large contribution on day 8(41% of the total ASC) but
was more variable and declinedquickly. IgA ASC contributed 4.3% of
the total ASC on day 16.IgM, IgG1, and IgG2a ASC together comprised
less than 4%of the total ASC at any given time. Parasite-specific
IgE ASCwere not assayed in this experiment.
The ASC response in the CLN was similar in isotype distri-bution
but muted in comparison to that of the MLN (Fig. 1C).IgG2c
accounted for 60% of the total ASC on day 16. No ASCwere detected
on day 30. On day 50, specific ASC again weredetected, coincident
with the onset of the antityvelose responsedescribed previously
The isotype profile of ASC in the spleen was distinct fromthose
of the MLN and CLN (Fig. 1B). IgA ASC accounted for64% of the total
on day 16. IgA, IgM, and IgG2c ASC declinedfrom day 16 to day 30
and then rebounded on day 50.
The kinetics of the early immune response were confirmedin three
additional experiments. In one experiment, parasite-specific IgE
ASC were rare but detectable in the MLN (lessthan 0.1% of the total
ASC) on day 16. In a third experiment,we detected low numbers of
IgE ASC in the CLN 47 dayspostinfection (data not shown).
The ASC response was largely T dependent. Euthymic NIHrats
produced an antibody response with an isotype distribu-tion similar
to that of the PVG rats, although it was not ofequal magnitude
(Fig. 2). Nude rats mounted a weak (less than10% of the haired-rat
response) but constant ASC response inthe MLN, with relatively
constant numbers of ASC at 8, 15,and 22 days postinfection. The
isotypes produced by nude ratswere the same as those produced by
euthymic rats: IgG2b,IgG2c, and IgA. The overall level of activity
in the spleens ofrats in this experiment was too small to allow any
conclusionsto be drawn (data not shown). When euthymic rats were
in-fected orally with larvae, they eliminated nearly all of
theworms by 15 days postinfection. As expected based on previ-ously
reported studies (27, 37), athymic rats failed to
eliminatesignificant numbers of adult worms by 22 days
postinfection(data not shown), confirming the T-cell dependence of
adultworm rejection and showing that the weak B-cell response
didnot promote worm rejection.
Isotype profiles of specific antibodies in sera from T.
spira-lis-infected PVG rats correlated with ASC numbers.
Westernblots of crude larval antigens were probed with pooled
serumantibodies collected from uninfected rats, rats infected
with1,000 larvae (56 days postinfection), or rats infected with
500larvae (16 days postinfection) (Fig. 3). Antibodies in the
seracollected 16 days postinfection bound to at least three
proteins(reduced) of 40 to 60 kDa. The difference in binding
specific-ities between antibodies collected on days 16 and 56 of
infec-tion illustrates the switch from phosphorylcholine antigens
totyvelose antigens that takes place in the biphasic
antibodyresponse (8). The predominance of IgG1, IgG2a, and
FIG. 3. Isotype profile of specific antibodies in sera of PVG
rats on days 16 and 56 postinfection (PI). Western-blotted T.
spiralis crude antigen was probed withpooled sera from normal rats
(NRS), rats infected with 1,000 larvae 56 days prior to bleeding,
and rats infected with 500 larvae 16 days prior to bleeding.
Antibodybinding was detected with HRP-conjugated antibodies
developed in an ECL system, as described in Materials and Methods.
Molecular mass standards are indicatedin kilodaltons.
Autoradiograph exposure time for each isotype is shown at the
bottom of the figure. The small arrow marks migration of the 49-kDa
phosphorylcholine-bearing protein. The large arrowhead marks
migration of the smallest tyvelose-bearing glycoprotein.
4664 PETERS ET AL. INFECT. IMMUN.
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among antityvelose antibodies detected on day 56 postinfec-tion
corresponds with ELISA data reported previously (3).
The isotype profile of pooled antibodies in sera collected
16days postinfection correlated with ELISPOT data obtainedfrom the
same rats. IgG2c was the predominant isotype. IgG2bhad a weak
signal, correlating with a variable and volatileIgG2b ASC response.
The IgE signal was extremely weak,requiring a 2-h exposure to
develop, compared to the 3- to 12-sexposures for the other
isotypes. The specificities of the iso-types were similar, but the
intensity of binding to differentproteins varied. IgE was
distinctive in that it bound to a singleprotein species.
Serum antibodies from infected rats and MAb 6G3 werespecific for
phosphorylcholine. Based on indirect evidence,Takahashi has
concluded that the antibody response inducedin the first 2 weeks of
infection with T. spiralis is specific forphosphorylcholine-bearing
proteins (35). In order to confirmthe specificity of the antibodies
produced in infected rats, weperformed blocking experiments.
Incubation of MAb 6G3 with10 mM phosphorylcholine blocked binding
of the antibody toWestern blots of crude larval antigen (Fig. 4),
proving thespecificity of the antibody. In contrast,
phosphorylcholine com-pletely blocked serum antibody binding to
only one of severalproteins, suggesting that other epitopes on
these proteins werebeing recognized (Fig. 4). We confirmed the
specificity of se-rum antibodies by absorbing rat sera with
phosphorylcholine-bearing proteins that were affinity purified from
crude antigenby using MAb 6G3. Absorption removed all serum
antibodiescapable of binding crude antigens in ELISA (Fig. 5),
confirm-ing the phosphorylcholine-bearing antigens to be dominant
inthis phase of the antilarval response.
Antiphosphorylcholine MAb 6G3 was not protective. In or-der to
test the effects of antiphosphorylcholine antibody onintestinal T.
spiralis, we used a passive immunization and chal-lenge protocol in
suckling rats. We have shown this system tobe highly sensitive in
revealing protective activity of rat anti-bodies against intestinal
infection with the parasite (3). Passiveimmunization with MAb 6G3
failed to accelerate expulsion ofintestinal larvae or adult worms
from rat pups challenged withT. spiralis (Fig. 6). We used
antityvelose antibody 9E6 (IgG2c)as an isotype-matched, positive
control in this experiment. As
described previously, passive immunization with 9E6 causedrats
to eliminate more than 90% of the challenge larvae within24 h (3).
Antibody 9E6 had no effect on the few adult wormsthat developed in
passively immunized rats.
MLNs are sites of intense B- and T-lymphocyte activity be-tween
8 and 16 days after infection with T. spiralis in rats andmice (13,
22). The increased cellular activity in the MLN iscorrelated with
the expulsion of the adult worms. T lympho-cytes from this tissue
(13) and from lymph vessels draining the
FIG. 4. Soluble phosphorylcholine (pc) blocked binding of MAbs
and poly-clonal antibodies to crude larval antigens. Crude antigens
were resolved on 10%acrylamide minigels, blotted, immunostained,
and developed with ECL. Phos-phorylcholine was titrated in the
presence of the primary antibodies: 6G3 hy-bridoma culture
supernatant (diluted 1:40) (20-s exposure) or pooled serumfrom rats
infected 16 days prior with 500 larvae (diluted 1:5,000) (1-min
expo-sure). Similarly diluted serum from uninfected rats showed no
reaction. Molec-ular mass standards are indicated in
FIG. 5. Serum antibodies in rats infected 16 days with T.
spiralis were specificfor phosphorylcholine-bearing antigens.
Pooled serum collected from rats 16days postinfection or antibody
6G3 was absorbed with phosphorylcholine-bear-ing antigens or BSA as
described elsewhere and then titrated in an ELISA oncrude larval
antigens. Absorbances are graphed for pooled serum collected
fromrats 16 days postinfection with 500 T. spiralis absorbed with
phosphorylcholineantigens (■), absorbed with BSA (F), or unabsorbed
(E); for pooled seracollected from uninfected rats (h); and for MAb
6G3 absorbed with phospho-rylcholine antigens (Œ) or BSA (}).
FIG. 6. Comparison of intestinal worm burdens in rat pups that
were pas-sively immunized with 6G3 MAb (F), 9E6, tyvelose-specific
MAb (E), or serumantibodies from an uninfected rat (h). PVG rats
were infected with an oral doseof 185 T. spiralis muscle larvae.
The numbers of worms per intestine werecounted on 1, 8, and 10 days
postinfection. Values are the means 1 standarddeviations (four or
five rats). p, 9E6-treated rats bore significantly fewer wormsthan
the two other treatment groups (Tukey’s least significant
difference test, P ,0.05).
VOL. 67, 1999 ANTIPHOSPHORYLCHOLINE RESPONSE TO T. SPIRALIS
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intestine (18) are protective in adoptive transfer
experiments.We found that the isotype distribution among ASC in
lymphnodes draining the intestine correlated well with specific
anti-body isotypes in serum, suggesting that in addition to the
T-cellresponse, the MLN is playing a major role in the serum
anti-body response to infection.
Intestinal nematodes have long been known to induce strongIgE
responses in rodents. Although we expected to measurelarge numbers
of IgE ASC in our experiments, such cells wererare in the tissues
examined from infected PVG rats. We ob-tained similar results from
other rat strains, including BrownNorway (data not shown), a strain
that is reported to producestrong allergic responses (42). It is
possible that IgE ASC mayreside in tissues that we did not
evaluate, such as intestinallamina propria. Circulating IgE levels
similarly were barelydetectable. Interpretation of our results is
aided by the findingsof Ramaswamy and coworkers, which showed that
IgE is trans-ported from serum to the gut lumen in T.
spiralis-infected rats(29). This transport depletes the serum of
circulating IgE (butnot IgG) between 4 and 14 days postinfection,
times that cor-relate with our assessments. Thus, we cannot rule
out thepossibility that IgE was produced by B cells in tissues we
didnot assay or that it was cleared from the circulation so that
oursampling methods failed to detect it.
The dominant isotype in the antiphosphorylcholine
antigenresponse against T. spiralis was IgG2c. In the rat,
polysaccha-ride and hapten-polysaccharide antigens (TI-2 antigens)
stim-ulate IgG2c predominantly (9). Our studies of athymic
ratsshowed that less than 10% of the early antibody responseagainst
T. spiralis was T-lymphocyte independent. This resultis compatible
with findings for nude mice, where IgG3 (themouse homologue of rat
IgG2c [6, 9]) responses to TI-2 anti-gens are reduced by more than
90% compared to those ineuthymic controls (32). Biological
activities unique to ratIgG2c are not known.
Other investigations have shown that anti-Ig antibodies
con-jugated to dextran mimic the B-cell-stimulatory activity of
TI-2antigens in mice (reviewed by Mond et al. ). The
influencesof cytokines on this stimulation have been investigated.
Anti-Ig–dextran conjugates trigger B-cell proliferation in vitro,
andaddition of IL-5 induces these cells to produce IgG3.
Further-more, IgG3 production is enhanced 10-fold by the addition
ofIFN-g together with IL-5 (33). These findings can be applied
tothe analysis of the immune response against T. spiralis in
rats.Ramaswamy et al. (30) showed that IFN-g was released intothe
intestinal lymph in a cyclical manner, increasing at 35, 75,and 180
h following infection. IL-5 was detected as early as20 h following
infection and subsequently was present togetherwith IFN-g. Thus,
IL-5 and IFN-g are present in intestinallymphoid tissue at the time
that worms are releasing theirphosphorylcholine-bearing antigens in
the intestine. These cy-tokines may drive the production of IgG2c
against phospho-rylcholine-bearing antigens of T. spiralis.
Phosphorylcholine epitopes in other nematodes have beendescribed
to have immunomodulatory effects. A phosphoryl-choline-bearing
glycoprotein of the filarial nematode Acantho-cheilonema viteae has
been shown to inhibit B-cell activation(14). In addition,
phosphorylcholine has been implicated in theinduction of IL-10
secretion by B-1 cells from mice immunizedwith microfilarial
lysates of Brugia malayi (26). Although wehave not investigated
B-1-cell proliferation or cytokine produc-tion in T.
spiralis-infected rats, the magnitude of the responseto
phosphorylcholine-bearing antigens suggests that it is likelyto
influence the other immune responses induced by the par-asite.
Bacterial organisms incorporate phosphorylcholine into tei-
choic acids (24), lipopolysaccharide (39), protein, and pili
(38).Using mice transgenic for an antiphosphorylcholine
antibody,Lim and Choy demonstrated that phosphorylcholine is a
targetof protective immunity against Streptococcus pneumoniae
(20).In these studies, transgenic adult mice were not resistant to
T.spiralis infection. Because it was found that suckling rats
aremore sensitive to the effects of protective antilarval
antibodiesthan adult rats (3, 5), we tested
phosphorylcholine-specificantibody for protection in suckling rats.
AntiphosphorylcholineIgG2c did not alter the establishment or
development of T.spiralis in the intestine. Previously, it was
shown that protec-tive, antityvelose IgG2c antibodies bind to
tyvelose-bearingsurface and secreted products of L1 larvae (4, 11,
12) and thatsecreted products are the primary targets in protection
(21). Incontrast, phosphorylcholine antigens are sequestered in
thebody of the larva and are not secreted by this stage
(reference25 and unpublished observations). Sequestration would
ex-plain both the failure of these antigens to induce an
immuneresponse during the muscle stage of infection and also
theirfailure to serve as targets in protective intestinal immunity.
Itseems plausible, although not proven, that
phosphorylcholineantigens would be released during molting.
Although phospho-rylcholine antigens have been detected in adult
worms (25), wefound that antiphosphorylcholine IgG2c antibody did
not ac-celerate expulsion of adult worms from the gut. Thus,
phos-phorylcholine antigens are highly immunogenic but do notappear
to serve as targets of antibody-mediated, protectiveimmunity
against intestinal T. spiralis.
In summary, we have described an IgG2c-dominated anti-body
response in the intestines of rats infected with the para-sitic
nematode, T. spiralis. This response is directed againstpreviously
described phosphorylcholine-bearing glycoproteinsproduced by the
first larval stage and is distinct from thewell-characterized
antityvelose response which is induced bymuscle-stage larvae. The
antiphosphorylcholine response islargely T dependent and, although
it is induced in the intestine,does not appear to directly
influence the outcome of intestinalinfection. Unique biological
activities of rat IgG2c are notknown. The strong representation of
IgG2c in two distinctantibody responses directed against T.
spiralis larvae supportsa conclusion that this isotype, as well as
the cells that produceit, has a significant and unique role in host
responses to infec-tion with this nematode.
This work was supported by USPHS grant AI 14490, a grant fromthe
Cooperative State Research, Education and Extension Service,U.S.
Department of Agriculture, under project no. 473401, and aFogarty
Senior International Fellowship (J.A.A.). P.J.P. and A.B.B.were
supported by a grant from the Howard Hughes Medical
Institute(Cornell Hughes Scholars Program), and E.A.S. was
supported by afellowship from the College of Veterinary Medicine,
We thank F. Ubeira for helpful discussion and A. Hesser for
assis-tance in preparing the manuscript.
1. Appleton, J. A., and D. D. McGregor. 1987. Characterization
of the immunemediator of rapid expulsion of Trichinella spiralis in
suckling rats. Immunol-ogy 62:477–484.
2. Appleton, J. A., and D. D. McGregor. 1985. Life-phase
specific induction andexpression of rapid expulsion in rats
suckling Trichinella spiralis-infecteddams. Immunology
3. Appleton, J. A., L. R. Schain, and D. D. McGregor. 1988.
Rapid expulsion ofTrichinella spiralis in suckling rats: mediation
by monoclonal antibodies.Immunology 65:487–492.
4. Appleton, J. A., and L. Usack. 1993. Identification of
potential antigenictargets for rapid expulsion of Trichinella
spiralis. Mol. Biochem. Parasitol.58:53–62.
4666 PETERS ET AL. INFECT. IMMUN.
on March 14, 2020 by guest
5. Bell, R. G., J. A. Appleton, D. A. Negrao-Correa, and L. S.
Adams. 1992.Rapid expulsion of Trichinella spiralis in adult rats
mediated by monoclonalantibodies of distinct IgG isotypes.
6. Brüggemann, M., P. Delmastro-Galfrè, H. Waldmann, and F.
Calabi. 1988.Sequence of a rat immunoglobulin g2c heavy chain
constant region cDNA:extensive homology to mouse g3. Eur. J.
7. Burnette, W. N. 1981. Western blotting: electrophoretic
transfer of proteinsfrom sodium dodecyl sulfate-polyacrylamide gels
to unmodified nitrocellu-lose and radiographic detection with
antibody and radioiodinated protein A.Anal. Biochem.
8. Denkers, E. Y., D. L. Wassom, C. J. Krco, and C. E. Hayes.
1990. The mouseantibody response to Trichinella spiralis defines a
single, immunodominantepitope shared by multiple antigens. J.
9. der Balian, G., J. Slack, B. L. Clevinger, H. Bazin, and J.
M. Davie. 1980.Subclass restriction of murine antibodies. III.
Antigens that stimulate IgG3in mice stimulate IgG2c in rats. J.
Exp. Med. 152:209–218.
10. Despommier, D. D. 1983. Biology, p. 75–80. In W. C. Campbell
(ed.),Trichinella and trichinosis. Plenum Press, New York, N.Y.
11. Ellis, L. A., C. S. McVay, M. A. Probert, J. Zhang, D. R.
Bundle, and J. A.Appleton. 1997. Terminal b-linked tyvelose creates
unique epitopes inTrichinella spiralis glycan antigens.
12. Ellis, L. A., A. J. Reason, H. R. Morris, A. Dell, R.
Iglesias, F. M. Ubeira, andJ. A. Appleton. 1994. Glycans as targets
for monoclonal antibodies thatprotect rats against Trichinella
spiralis. Glycobiology 4:585–592.
13. Grencis, R. K., and D. Wakelin. 1982. Short lived, dividing
cells mediateadoptive transfer of immunity to Trichinella spiralis
in mice. I. Availability ofcells in primary and secondary
infections in relation to cellular changes in themesenteric lymph
node. Immunology 46:443–450.
14. Harnett, W., and M. M. Harnett. 1993. Inhibition of murine B
cell prolifer-ation and down-regulation of protein kinase C levels
by a phosphorylcholine-containing filarial excretory-secretory
product. J. Immunol. 151:4829–4837.
15. Hernández, S., F. Romarı́s, I. Acosta, P. N. Gutiérrez,
and F. M. Ubeira.1995. Ultrastructural colocalization of
phosphorylcholine and a phosphoryl-choline-associated epitope in
first-stage larvae of Trichinella spiralis. Parasi-tol. Res.
16. Jungery, M., and B. M. Ogilvie. 1982. Antibody response to
stage-specificTrichinella spiralis surface antigens in strong and
weak responder mousestrains. J. Immunol. 129:839–843.
17. Kohler, G., and C. Milstein. 1975. Continuous cultures of
fused cells secret-ing antibody of predefined specificity. Nature
18. Korenaga, M., C. H. Wang, R. G. Bell, D. Zhu, and A. Ahmad.
1989.Intestinal immunity to Trichinella spiralis is a property of
OX82OX222T-helper cells that are generated in the intestine.
19. Lim, P. L., and W. F. Choy. 1990. A thymus-independent (type
1) phospho-rylcholine antigen isolated from Trichinella spiralis
protects mice againstpneumococcal infection. Immunology
20. Lim, P. L., W. F. Choy, S. T. H. Chan, D. T. Leung, and S.
S. Ng. 1994.Transgene-encoded antiphosphorylcholine (T151)
antibodies protectCBA/N (xid) mice against infection with
Streptococcus pneumoniae but notTrichinella spiralis. Infect.
21. McVay, C. S., A. Tsung, and J. Appleton. 1998. Participation
of parasitesurface glycoproteins in antibody-mediated protection of
epithelial cellsagainst Trichinella spiralis. Infect. Immun.
22. Molinari, J. A., J. A. Hess, and R. K. Wesley. 1982.
Sequential mesentericlymph node histological response following
Trichinella spiralis infection. Int.Arch. Allergy Appl. Immunol.
23. Mond, J. J., A. Lees, and C. M. Snapper. 1995. T
cell-independent antigenstype 2. Annu. Rev. Immunol.
24. Mosser, J. L., and A. Tomasz. 1970. Choline-containing
teichoic acid as astructural component of pneumococcal cell wall
and its role in sensitivity tolysis by an enzyme. J. Biol. Chem.
25. Ortega-Pierres, M. G., L. Yepez-Mulia, W. Homan, H. R.
Gamble, P. L. Lim,
Y. Takahashi, D. L. Wassom, and J. A. Appleton. 1996. Workshop
on adetailed characterization of Trichinella spiralis antigens: a
platform for futurestudies on antigens and antibodies to this
parasite. Parasite Immunol. 18:273–284.
26. Palanivel, V., C. Posey, A. M. Horauf, W. Solbach, W. F.
Piessens, and D. A.Harn. 1996. B-cell outgrowth and ligand-specific
production of IL-10 corre-late with Th2 dominance in certain
parasitic diseases. Exp. Parasitol. 84:168–177.
27. Perrudet, B. A., A. Y. Boussac, E. J. Ruitenberg, W.
Kruizinga, and A.Elgersma. 1983. Protection to challenge with
Trichinella spiralis after primaryoral infection in congenitally
athymic (nude) mice. J. Parasitol. 69:253–255.
28. Philipp, M., R. M. Parkhouse, and B. M. Ogilvie. 1980.
Changing proteins onthe surface of a parasitic nematode. Nature
29. Ramaswamy, K., J. Hakimi, and R. G. Bell. 1994. Evidence for
an interleukin4-inducible immunoglobulin E uptake and transport
mechanism in the in-testine. J. Exp. Med. 180:1793–1803.
30. Ramaswamy, K., D. Negrao-Correa, and R. Bell. 1996. Local
intestinal im-mune responses to infections with Trichinella
spiralis. J. Immunol. 156:4238–4337.
31. Reason, A. J., L. A. Ellis, J. A. Appleton, N. Wisnewski, R.
B. Grieve, M.McNeil, D. L. Wassom, H. R. Morris, and A. Dell. 1994.
Novel tyvelose-containing tri- and tetra-antennary N-glycans in the
immunodominant anti-gens of the intracellular parasite Trichinella
spiralis. Glycobiology 4:593–603.
32. Slack, J. H., and J. M. Davie. 1982. Subclass restriction of
murine antibodies.V. The IgG plaque-forming cell response to
thymus-independent and thy-mus-dependent antigens in athymic and
euthymic mice. Cell. Immunol.68:139–145.
33. Snapper, C. M., T. M. McIntyre, R. Mandler, L. M. Pecanha,
F. D. Finkel-man, A. Lees, and J. J. Mond. 1992. Induction of IgG3
secretion by inter-feron-gamma: a model for T cell-independent
class switching in response toT cell-independent type 2 antigens.
J. Exp. Med. 175:1367–1371.
34. Springer, T. A., A. Bhattacharya, J. T. Cardoza, and F.
Sanchez-Madrid.1982. Monoclonal antibodies specific for rat IgG1,
IgG2a, and IgG2b sub-classes, and kappa chain monotypic and
allotypic determinants: reagents foruse with rat monoclonal
antibodies. Hybridoma 1:257–273.
35. Takahashi, Y. 1997. Antigens of Trichinella spiralis.
Parasitol. Today 13:104–106.
36. Ubeira, F. M., J. Leiro, M. T. Santamarina, T. G. Villa, and
M. L. Sanmar-tı́n-Durán. 1987. Immune response to Trichinella
epitopes: the antiphospho-rylcholine plaque-forming cell response
during the biological cycle. Parasi-tology 94:543–553.
37. Vos, J. G., E. J. Ruitenberg, N. Van Basten, J. Buys, A.
Elgersma, and W.Kruizinga. 1983. The athymic nude rat. IV.
Immunocytochemical study todetect T-cells, and immunological and
histopathological reactions againstTrichinella spiralis. Parasite
38. Weiser, J. N., J. B. Goldberg, N. Pan, L. Wilson, and M.
Virji. 1998. Thephosphorylcholine epitope undergoes phase variation
on a 43-kilodaltonprotein in Pseudomonas aeruginosa and on pili of
Neisseria meningitidis andNeisseria gonorrhoeae. Infect. Immun.
39. Weiser, J. N., N. Pan, K. L. McGowan, D. Musher, A. Martin,
and J.Richards. 1998. Phosphorylcholine on the lipopolysaccharide
of Haemophi-lus influenzae contributes to persistence in the
respiratory tract and sensitiv-ity to serum killing mediated by
C-reactive protein. J. Exp. Med. 187:631–640.
40. Wisnewski, N., M. McNeil, R. B. Grieve, and D. L. Wassom.
1993. Charac-terization of novel fucosyl- and tyvelosyl-containing
glycoconjugates fromTrichinella spiralis muscle stage larvae. Mol.
Biochem. Parasitol. 61:25–35.
41. Wright, K. D. 1979. Trichinella spiralis: an intracellular
parasite in the intes-tinal phase. J. Parasitol. 65:441–445.
42. Yoo, T. J., and C. Y. Kuo. 1980. Cellular and reaginic
immune responses toragweed antigen E in inbred rats. Int. Arch.
Allergy Appl. Immunol. 61:259–270.
Editor: S. H. E. Kaufmann
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