(Received for publication, September 5, 1995; and in revised form, November 8, 1995)
From the
Acanthocheilonema viteae is a parasitic nematode of rodents. We identified the chitinase of A. viteae infective stage larvae (L3) as the main target of the humoral immune response of jirds, which were protected against challenge infection after vaccination with irradiation attenuated L3. The cDNA of the L3 chitinase has been sequenced, and the deduced amino acid sequence shows significant homologies to chitinases of Brugia malayi microfilariae, insects, yeast, bacteria, and Streptomyces sp. The protein has been characterized by monoclonal antibodies and substrate activity gels. The chitinase of L3 may contribute to degrading the nematode cuticle during molting and thus represents a target of protective immune responses in a phase where the parasite is highly vulnerable. In addition, it has been shown that a similar enzyme exists in uterine microfilariae, which probably has a role in casting the egg shell.
Filarial parasites, such as Onchocerca volvulus, Brugia malayi, and Wuchereria bancrofti, the causative agents of river blindness and lymphatic filariases, affect more than 100 million people throughout the tropics(1) . Infection of the human host occurs by a bite of an infected arthropod. There is evidence that some individuals develop protective immunity naturally, as they do not acquire filarial infections despite high levels of local transmission(2, 3) . The humoral immune response of these patients differentially recognizes antigens of infective third stage larvae (L3) (4) , and it is conceivable that vaccination with such antigens could prevent the infection. As the study of protective immunity in human filariae is hampered by the host specificity of the parasites, we adopted the approach to identify immunorelevant antigens of the rodent filaria Acanthocheilonema viteae, a parasite of the jird (Meriones unguiculatus).
For animal models, the vaccination with irradiation-attenuated infective larvae (L3) has been demonstrated to be the most effective way to induce protective immunity(5) . Immunization of jirds with irradiation-attenuated L3 of A. viteae induces more than 90% protection against a challenge infection(6) . Sera of such vaccinated animals recognize few L3 proteins. We report here on the characterization and the molecular cloning of the immunodominant antigen recognized by the humoral immune response of challenge resistant jirds, a chitinase of A. viteae L3. Chitinases were recently described from microfilariae of B. malayi(7) . It was supposed that the enzymes might have a role in casting the microfilarial sheath, a modified egg shell. This structure was believed to contain chitin on biochemical grounds (8, 9) and due to its analogy with the chitin-containing egg shell of other nematodes. Interestingly, the egg shell is the only structure of nematodes known to contain chitin. However, recent reports deny the presence of chitin in the microfilarial sheath (see (10, 11, 12) ) and call the chitin-degrading role of filarial chitinase into question. Our characterization of L3 chitinase (an active, chitin-degrading enzyme) in a filarial stage that is not yet known to contain chitin suggests that the enzyme has additional substrate specificities. Chitinase-like proteins of vertebrates, a class of animals not producing chitin, were described to degrade extracellular matrix under inflammatory or degenerative conditions and to play a role in the process of fertilization(13, 14) . The close homology of the L3 chitinase with these molecules suggests that the described enzyme degrades N-acetylglucosamine-containing structures of the parasite during molting and might help the parasite to migrate through the host's tissues.
Figure 1: Part A, stage specificity and immunodominance of the A. viteae L3 chitinase. 15 µg of A. viteae L3 (A) and L4 (B) extracts were electrophoresed on 12.5% SDS-PAGE and transblotted. Blots were probed with sera of vaccinated jirds (panel 1), sera of naive jirds (panel 2), mAb 2H2 (panel 3), mAb 24-4 (panel 4), control mAb (IgG1, panel 5). Part B, reaction of mAb 2H2 with A. viteae male (lane 1) and female worms (lane 2), blood MF (lane 3), and infective stage larvae (lane 4). Parasites were extracted as above and electrophoresed on a 4-15% SDS-polyacrylamide gel, and the transblotted proteins were immunostained. Part C, reaction of mAb 2H2 with extracts of vector-derived L3 (lane 1), L3 3 days postinfection (lane 2), L4 16 days postinfection (lane 3), culture supernatants of 1,000 L3 after 24 h cultivation under vertebrate conditions (lane 4), molting supernatants of 1,000 L3 (lane 5). Samples were processed as in part B. Part D, chitinase activity is demonstrated in substrate-SDS/polyacrylamide gels containing glycol chitin for total female worm extract (lane 1) and total L3 extract (lane 2).
To characterize the immunoreactive L3 antigens two mAbs were produced against irradiated L3 and against molting fluid, respectively. mAb 24-4, resulting from fused spleen cells of a mouse immunized with irradiation-attenuated L3, specifically recognized the 205 kDa band of L3 and of L3 culture supernatant and reacted slightly with the 68 kDa band (Fig. 1A, panel 4). mAb 2H2, produced by immunization with molting fluid of L3, recognized in immunoblots the 205 and 68 kDa bands of L3 (Fig. 1A, panel 3). mAb 2H2 also reacted in immunoblots with a 205- and a 68-kDa antigen of L3 culture supernatant, and with a 68 kDa band of culture supernatant collected after the in vitro molting of L3 (Fig. 1C). Both mAbs did not recognize any antigen in L4, indicating that the main target antigens of the immune response of vaccinated jirds are stage-specific.
N-terminal sequence analysis of the 205-kDa protein of L3 generated a single amino acid residue at each cycle. Six amino acids could be determined from the 205-kDa antigen (Fig. 2). A data base search of the EMBL and GenBank data bases revealed an identity with the amino acids of the N terminus of a endochitinase of MF of B. malayi. For the characterization of the 68-kDa protein, it has been purified from a supernatant of A. viteae L3 after extraction with PBS containing 1% CTAB by lectin affinity chromatography using SBA-agarose (see ``Materials and Methods''). N-terminal sequencing of the purified glycoprotein revealed 20 amino acids showing an identity between the 68- and the 205-kDa protein (Fig. 2). To test for chitinase activity, extracts of L3 were electrophoresed under nonreducing conditions in SDS gels containing glycol chitin. Chitinase activity was localized preferentially in the 68 kDa band of L3, and weakly in the 205-kDa protein, which was recognized by both mAbs (Fig. 1D). Extensive boiling of L3 antigens using a sample buffer containing 100 mM dithiothreitol resulted in the disappearance of the 205-kDa protein (Fig. 3). Therefore, we assume that monomeric 68-kDa chitinase can form trimers, which are linked by disulfide bridges that can be reduced by high concentrations of sulfhydryl compounds.
Figure 2: Nucleotide sequence and deduced amino acid sequence of the A. viteae L3 chitinase. Amino acid numbering begins at the initiating methionine, with the mature N terminus beginning at amino acid 18. Lower case letters represent the signal peptide on the amino acid line and untranslated regions on the nucleic acids line. Primers used for PCR amplifications and sequencing are marked as arrows above the nucleic acids line. The consensus polyadenylation signal is underlined. The N-terminal amino acid sequence derived from the purified 205- and 68-kDa proteins is boxed and shadowed, and the amino acids resulting from the 205-kDa protein are in italics. Potential myristilation sites are indicated by an inversed Gly residue. The attachment site for glycosaminoglycan (amino acids 234-237) is shaded. One potential N-glycosilation site is doubly underlined. Conserved Cys residues within the carboxyl-terminal domain are indicated in boldface. The four repeated amino acid sequences in the C-terminal domain are boxed and shadowed.
Figure 3: Reaction of mAb 2H2 with extracts of vector derived A. viteae L3. Antigens were electrophoresed on a 4-15% SDS-polyacrylamide gel under reducing conditions using a sample buffer containing 5% (v/v) 2-mercaptoethanol (lane 1) and 100 mM dithiothreitol (lane 2), respectively.
As mAb 24-4 was directed against an epitope stable under a variety of conditions, it served to localize the filarial chitinase in L3. Immunogold staining of ultrathin sections of L3 with mAb 24-4 revealed the presence of A. viteae chitinase in the cellular cytoplasm and in the lumen of the glandular oesophagus of vector-derived L3 (Fig. 4A). No chitinase could be detected in muscle, cuticle, or on the outermost surface of L3. Neither live L3 nor formaldehyde-fixed L3 studied under a variety of conditions by IFAT carried detectable amounts of chitinase on the surface.
Figure 4:
Immunogold staining of ultrathin sections
of A. viteae L3 (A) and A. viteae MF in
utero (B) with mAb 24-4. c, cuticle; e,
epidermis; es, egg shell; go, glandular oesophagus; mc, muscle cell; mf, microfilaria; ul,
uterine lumen. (L3 32,000; MF
48,000; bars represent 0.5 µm).
The obtained sequence was 1,670 base pairs long and contained an open reading frame coding for 520 amino acids with a theoretical molecular mass of 58 kDa (Fig. 2). The 5`-end consisted of a potential signal sequence of 17 residues(27) , indicating that L3 chitinase is secreted. This sequence was followed by the N terminus of the mature protein, which corresponded exactly to the N-terminal amino acid sequence of the protein as obtained by protein sequencing (Fig. 2). The sequence showed one potential N-glycosylation site, six myristylation sites, and one attachment site for glycosaminoglycan. The noncoding 3`-end showed a consensus signal for polyadenylation(28) . The 5`-end of the mRNA did not contain a spliced leader sequence, as confirmed by DNA sequencing and by PCR with phage DNA of the L3 library using an oligonucleotide primer corresponding to the nematode spliced leader sequence (29) and the sequence-specific primer P3.
The cDNA exhibited high homology (69% nucleotide identity over the whole sequence) to the MF chitinase of B. malayi(7) and weaker similarities to chitinases of W. bancrofti(23) , insects, bacteria, Streptomyces sp., fungi, and plants (30, 31, 32) . The domain structure was similar to the one of B. malayi MF chitinase. The N-terminal signal sequence is followed by the well conserved catalytic domain, spanning amino acids 18-370 (31, 32, 33) . The third domain is less conserved and contains 35% Ser and Thr residues. It could be a target of extensive O-glycosilation(7, 31) . This region comprises four imperfect repeats of 14 amino acids in length between positions 370 and 440 of the mature protein. The carboxyl-terminal end of the protein is closely related in structure to the B. malayi and insect chitinases as the 6 Cys residues are perfectly matched, suggesting that they could be involved in disulfide bridges. However, these regions are different from the chitin-binding domains described for yeast and class III plant chitinases.
To study the presence of chitinase mRNA, cDNA from male worms, female worms, blood MF, L3, and L4 was amplified with degenerate primers P1 and P2. In all stages, except male worms, specific amplification products were obtained, indicating that chitinase mRNA is present in most filarial stages (not shown).
Immunogold staining of ultrathin sections of female worms with mAb 24-4 revealed that chitinase was localized in the epidermis and in the laminated layer of the cuticle of most MF present in the uteri (Fig. 4B). Only a small proportion of the MF carried the antigen on the outermost surface. The egg shell itself and other compartments of the MF did not contain detectable amounts of chitinase. To study the presence of chitinase on the surface of uterine MF, we performed IFAT with mAbs 24-4 and 2H2 using cryostat sections and preparations of uterine content of female A. viteae, where both mAbs immunostained exactly the same structures. IFAT analysis revealed that the target epitopes were accessible to the mAbs on nearly mature uterine MF, where chitinase was evenly distributed on the surface. Most of these Ag-bearing MF were still within the egg shell (Fig. 5, A and B). In contrast, no more chitinase was present on the surface of fully mature, hatched uterine MF, which are more slender than the nearly mature forms and have pointed heads. However, MF, which had degenerated within the female worms around the time of molting and were stumpy, immotile, and sometimes broken, carried detectable amounts of chitinase on their surface. An IFAT study of newborn MF revealed that such degenerated MF bearing chitinase on their surface represented around 10% of the MF production released by in vitro cultured female A. viteae (Fig. 5, C and D).
Figure 5: Localization of A. viteae chitinase in intrauterine MF by IFAT with mAb 24-4 (left panels) and corresponding light microscope photographs (right panels). Uterine contents of gravid female A. viteae with nearly mature MF inside the egg shell, younger embryonic stages and a mature, hatched MF (A and B). Newborn MF (C and D). Note that fluorescent microfilariae are swollen and stumpy. es, embryonic stages; nmf, nearly mature microfilariae; hmf, hatched microfilariae. Bars represent 50 µm.
The most effective way to induce protective immunity against
filarial infections in experimental animals is the immunization with
irradiation-attenuated L3 (see (5) and (6) ). Our
study reveals that L3 chitinase is the immunodominant antigen
recognized by the humoral immune response of jirds vaccinated against A. viteae infection. This protein has several properties of a
vaccine candidate antigen; in particular it is exported during the
initial phase of the infection and during molting of L3, which are
phases of key importance for the attrition of filarial
larvae(34, 35) . Furthermore, L3 culture supernatants
and molting fluid, which both contain chitinase, induce protection
against a challenge infection with A. viteae in
jirds(6) . ()The findings of Freedman et
al.(3) , describing a 43-kDa chitinase-like filarial
protein to be recognized by the sera of persons resistant against
infection with the lymphatic filaria W. bancrofti(23) , support the notion of chitinase being a protein
with protective properties. Immunization studies in our animal model
will evaluate whether immune responses against L3 chitinase can prevent
an infection with A. viteae.
Chitinases are enzymes that
hydrolyze chitin (poly--(1-4)-linked GlcNAc). Well known
examples of chitinases are the enzymes of insects (Manduca),
which have a role in degrading the chitinous exoskeleton during the
molt and the chitin-degrading enzymes of fungi and
streptomycetes(30, 31, 32) . However,
chitinases can cleave substrates other than chitin and can have
activities as trans-glycosidases(36) . The role of chitinase in
L3 is unclear, and the fact that the enzyme is released by the L3
during different phases suggests that it might have a broad substrate
specificity. First, L3 chitinase is exported into the culture medium
immediately after the larvae are cultured under vertebrate conditions.
This time point corresponds to the early infection where L3 migrate
through host tissues(37) . An enzyme exported during this phase
could help to degrade host tissues, provided that it has an appropriate
substrate specificity. Interestingly, chitinase-like molecules were
recently described from the cartilago and the oviduct of vertebrates (13, 14, 38, 39, 40) ,
which are supposed to contribute to the degradation of extracellular
matrix under inflammatory or degenerative conditions or to
fertilization of vertebrates. Second, L3 chitinase is produced during
molting, a phase where the nematodes' cuticle is reorganized and
finally cast. It is not clear whether chitin is a target substrate
during this phase, as the only stage of nematodes that has been
demonstrated to contain chitin is the egg(41, 42) .
However, recent studies described chitotriosides in the cuticle of
adult Haemonchus contortus, which is indicative for chitin
being a structural component of the nematode cuticle(43) .
Therefore, further studies have to show whether the target substrate of
L3 chitinase is chitin or whether other substrates are converted as
-(1-4)-linked N-acetylglucosamine oligomers, which
were shown to occur in the filarial cuticle by lectin binding studies
or enzymatic studies (19, 43) . Irrespective of the
substrate, the localization of the enzyme and the timing of release
suggest that L3 chitinase is involved in the process of molting. The
source of released L3 chitinase are the pharyngeal glands, where the
enzyme was localized by immunoelectron microscopy. Pharyngeal gland
products, which are exported through the buccal cavity, were described
to have a role in molting in other nematodes (44) and are
released around the time of molting by L3 of O.
volvulus(45) . Molting products of nematodes are so far
poorly described and the only pharyngeal gland component described on
the molecular level is a metalloprotease of H. contortus(46) . Recent data
indicate that the
production of L3 chitinase is regulated by steroid hormones, which is
an analogy to the tighly regulated expression of insect chitinases
during molting (30) .
Our study suggests that the chitinase of uterine MF has a role in cleaving chitinous structures of the eggshell of A. viteae. Chitin was detected in the eggs of intestinal helminths Ascaris suum and Heligmosomoides polygyrus(41, 47) and it was described to be a component of the egg shell of the filariae Onchocerca gibsoni and O. volvulus(42) . Chitinase is considered as a relevant factor for the casting of the egg shell of A. suum(47) . The strict coincidence of the occurrence of chitinase on the surface of uterine MF of A. viteae and the time point of molting suggests that A. viteae chitinase contributes to hatching. It is conceivable that chitinase detected within the cuticle of immature uterine MF represents a storage form, which is transported to the MF surface prior to the hatching. The lack of chitinase in blood MF of A. viteae suggests a tightly regulated expression of the protein at the required timepoint. The contrasting presence of chitinase in blood MF of B. malayi could be explained by the fact that MF of this filarial species remain surrounded by the modified egg shell, which is not cast until the blood MF have entered the arthropod host(48) .
The cDNA
of the A. viteae L3 chitinase shows 69% nucleotide identity
with the cDNA of B. malayi MF chitinase(7) . The
chitinase of the tobacco hornworm Manduca sexta(30) is also closely related to the filarial chitinases as
shown by the similarity of the enzyme domain structure and by the amino
acid composition (28% of the amino acids are identical, and about 70%
are similar), whereas the described chitinases of streptomycetes,
bacteria, and fungi are less related to our molecule. The sequence of
the chitinase-like 43-kDa molecule of W.
bancrofti(23) , which is derived from genomic DNA, has two
small regions of homology in common with A. viteae L3
chitinase and B. malayi MF-chitinase but is otherwise
relatively distinct. The homology between these filarial chitinases is
highest (86% nucleotide identity) in the N-terminal signal sequence and
the adjacent 370-amino acid region, the catalytic domain. The catalytic
domain of A.viteae L3 chitinase is identical to the one of
uterine MF of A. viteae and very closely related to the one of
uterine MF of O. volvulus. Downstream follows a
Ser/Thr-rich domain, which encompasses three nearly perfect repeats of
14 amino acids in the B. malayi gene, whereas this region
shows four imperfect repeats of each 14 amino acids in the A.
viteae gene. The Ser/Thr-rich domain is 21 amino acids longer in
the A. viteae cDNA. The carboxyl-terminal Cys-rich domain
shows similarities between the filarial chitinases and the enzyme of M. sexta. The 6 Cys residues are perfectly matched, indicating
a conserved function of this domain, which is potentially important for
intra- and intermolecular bridging.
The fact that the open reading
frame of the L3 chitinase cDNA codes for a protein with a theoretical
molecular mass of 58 kDa suggests that the protein backbone is
posttranslationally modified. The dicrepancy between theoretical and
actual molecular mass is probably due to extensive O-glycosylation of the Ser/Thr-rich domain and the very acidic
composition of this region, which could be responsible for an anomaly
in the migration in SDS gels(7, 31) . Deglycosilation
of native A. viteae L3 chitinase using O-glycanase
resulted in a shift of molecular weight. Expression of the cDNA in Escherichia coli yielded a 58-kDa protein, which hydrolized
chitin, indicating that the activity of the enzyme is not dependent on
glycosylation. Our experiments show that the 205 kDa band
of L3 is a multimeric form of chitinase, derived from 68-kDa monomers
by disulfide bridging. Forming of multimers was associated with an
alteration of the antigenicity and of the chitinase activity, since mAb
24-4 did not bind to the 68 kDa band and the 205 kDa protein was shown
to be less active in substrate gels.
The presence of chitinase in
two distinct stages of the parasite's life cycle and the stage
specific localization of the enzyme suggest that the expression of
chitinases of L3 and MF is specifically regulated. Furthermore, the
differences of molecular weight between L3 chitinase and MF chitinase
show a specific structure of the molecules. Sequencing of the C
terminus of the cDNAs of both stages revealed sequence variation, being suggestive for the presence of several chitinase genes.
However, it is also possible that stage-specific differential splicing
or differential posttranslational modifications contribute to the
observed differences between L3 chitinase and MF chitinase of A. viteae. It will be interesting to study which
specific properties are related to the timing of expression, the
localization of the enzyme within the parasites, and the substrate
specificity of chitinases of various stages and species of filarial
parasites.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBank(TM)/EMBL Data Bank with accession number(s) U14638[GenBank].