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ABSTRACT. Stable resistance to the anthelmintic hycan- thone can be produced in the human blood fluke Schistosoma mansoni by exposing immature ...

Proc. Natl. Acad. Sci. USA

Vol. 88, pp. 7754-7758, September 1991 Medical Sciences

Characterization of a programmed alteration in an 18S ribosomal gene that accompanies the experimental induction of drug resistance in Schistosoma mansoni (schistosmes/hycanthone/rRNA-encoding DNA)

PAUL J. BRINDLEY*, STEPHEN HEATH, ANDREW P. WATERS, THOMAS F. MCCUTCHAN, AND ALAN SHER Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892

Communicated by William Trager, May 15, 1991

related drug oxamniquine, and the parasites derived show cross-resistance to both compounds. Resistance is induced by exposure of the 27- to 29-day-old parasite to HC, when schistosomes are developmentally insusceptible to the anthelmintic (6). Nevertheless, this procedure leads to drug resistance in subsequent generations in the developmentally drug-susceptible adult parasite (4). The remarkable feature of HC resistance is that it occurs within a single generation and thus does not appear to be the result of selection by the drug, unlike the appearance of resistance in other parasites (e.g., ref. 7), because all of the progeny of the schistosomes exposed to HC exhibit resistance and because no drug pressure is applied to the drugsensitive stages of these parasites. To define the molecular changes underlying the appearance of HC resistance in this model, genomic DNA from the drug-resistant schistosomes was compared to that of the sensitive parent parasites. Alterations, detectable as restriction fragment length polymorphisms (RFLPs) with a ribosomal gene probe, were reported to accompany the appearance of drug resistance, and one, an -3.6-kilobase (kb) BamHI fragment, was consistently associated with resistant parasites (5). To investigate the molecular events responsible for the generation of this marker, we have cloned and sequencedt the resistance-associated RFLP and used oligonucleotide probes diagnostic for this fragment to assess its presence and determine its genomic location in the drugsensitive parent population. The results of these analyses are reported here.

Stable resistance to the anthelmintic hycanABSTRACT thone can be produced in the human blood fluke Schistosoma mansoni by exposing immature parasites in mice to the drug. Within a single generation, genomic rearrangements, detected as rRNA-encoding DNA restriction fragment length polymorphisms (RFLPs), accompany the appearance of resistance in this model. One of these RFLPs, an ;"3.6-kilobase BamHI fragment, was shown previously to associate consistently with resistance in independent generations of the JHU strain of S. mansoni. To characterize the genetic changes responsible for this RFLP, the fragment was cloned and sequenced. A comparison of the cloned fragment with a normal 18S rRNA gene demonstrated that the drug resistance-associated RFLP fragment arises through the addition of 732 base pairs into an 18S rRNA gene, 134 base pairs downstream of the junction of the intergenic spacer and the mature 18S rRNA gene. The mutation is nonrandom, targets one, or a few only, of the 100 or so copies of the ribosomal genes, and may represent the incomplete duplication of the gene since the inserted element is identical in sequence to the region contiguous to it. The sequence spanning the junction of the insertion and the original 18S rRNA gene was used as a specific primer for the BamHI RFLP in PCR experiments. The analysis conclusively demonstrated that the mutation is induced rather than selected by the drug since the junctional sequence was not detectable in the drug-sensitive parent population of schistosomes. In addition, analysis of four, independently derived, resistant lines indicated that the same region of the gene was mutated each time. Together, these data demonstrate that reproducible changes are induced during the acquisition of resistance in schistosomes and suggest that the resistant phenotype is induced rather than selected from preexisting forms.

MATERIALS AND METHODS Parasites and Induction of Drug Resistance. Drug resistance was produced experimentally in the JHU strain of S. mansoni by injecting mice with 80 mg of HC (base) per kg of body weight at 28 days after infection with 100 cercariae (5). Lines of schistosomes were established with eggs obtained from the tissues of one to three of the HC-treated mice. Using these

Schistosomiasis is caused by various species of blood flukes of the genus Schistosoma, which as adults live and lay eggs in the vasculature of the intestines and bladder. Chemotherapy is the major method of control for schistosomiasis, and its use is increasing (1). Widespread application of schistosomicidal compounds is expected to lead to increased levels of drug resistance as has occurred with other parasites. Indeed, resistance to thioxanthenones has been detected in natural populations of Schistosoma mansoni (2) and can be derived experimentally (3-5). Nonetheless, the molecular basis of the resistance is poorly understood. In one model developed by Bueding and colleagues (4), resistance to thioxanthenones appears in S. mansoni after drug treatment of mice infected with immature schistosomes. This phenomenon is parasite strain dependent, but it is seen with schistosomes of disparate geographical origin. The resistance can be produced with hycanthone (HC) or the

procedures, we derived a number of lines of schistosomes from the parent JHU and NMRI strains. [The JHU and NMRI are laboratory-maintained strains of Puerto Rican origin (4, 8).] Resistance to HC was produced in the JHU strain but not in NMRI worms. Parent lines of these two strains are sensitive to the drug and are termed JHU.A and NMRI.A, respectively. JHU.C, JHU.D, JHU.F, and JHU.G are drug-resistant lines derived from the HC-sensitive parent line JHU.A. In contrast, NMRI.B and NMRI.C are drugAbbreviations: HC, hycanthone; RFLP, restriction fragment length polymorphism; rDNA, rRNA-encoding DNA. *To whom reprint requests should be addressed at: Laboratory of Parasitic Diseases, Building 4, Room 126, National Institutes of Health, Bethesda, MD 20892. tThe sequences reported in this paper have been deposited in the GenBank data base (accession nos. M62652 and M63432).

The publication costs of this article were defrayed in part by page charge payment. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact. 7754

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Brindley et al.

sensitive lines; they are the progeny of HC-treated NMRI.A line worms (5). Pools of genomic DNAs were prepared from 103 adult schistosomes from each of the JHU.D, JHU.F, JHU.G, NMRI.B, and NMRI.C lines; from 3 x 103 worms from the JHU.A and JHU.C lines; and from -1.2 x 104 NMRI.A parasites. In addition, DNA was extracted from individual adult schistosomes. Molecular Probes. The recombinant plasmid pSM 389 contains a BamHI fragment of the 18S rRNA gene plus some of the intergenic spacer from the NMRI strain of S. mansoni (see Fig. 2). Other plasmid probes (pSM HCR0, pSM HCR2, pSM HCR3, and pSM HCR4) contain fragments of the 18S rRNA gene from the JHU strain and the drug-resistanceassociated RFLP, as described below. Synthetic oligonucleotides were purchased from Synthecell (Gaithersburg, MD). Plasmids were labeled by nick-translation using [a-32P]dCTP. Southern hybridizations to genomic DNA extracted from schistosomes as well as autoradiography were performed as described (5). Comparative strengths of Southern hybridization signals were determined by laser densitometry. Radiochemicals were obtained from Amersham. Cloning of Ribosomal Gene Fragments. Genomic DNA from the drug-resistant JHU.G line of S. mansoni (8) was digested with BamHI, and the fragments were separated by agarose gel electrophoresis. DNA fragments ranging in size from -4.5 to -3.3 kb were electroeluted from the gel, purified by affinity chromatography (9), and ligated into BamHI-linearized, dephosphorylated plasmid pUC 13 DNA. Bacterial colonies resulting from transformation of Escherichia coli strain DH5a were screened by hybridization with the radiolabeled insert of pSM 389. Recombinant plasmid DNA was extracted from cultures of the positive colonies of bacteria using alkali/detergent lysis and purified by CsCI ultracentrifugation. A similar procedure using the insert of pSM 389 to probe another partial library comprising BamHI fragments ofgenomic DNA ranging in size from -3.2 to -2.8 kb was employed to clone an =3.0-kb fragment, which represents part of the tandemly repeated 18S rRNA gene (10) from the resistant JHU.G line (5). Restriction and DNA modifying enzymes and their buffers as well as bacterial host cells were purchased from BRL. Nudeotid Sequencing and Sequendng Strategies. Nucleotide sequencing reactions were performed on recombinant pUC 13 templates using the dideoxynucleotide chain-termination method (11). 17 polymerase (Sequenase version 2.0; United States Biochemical), deoxyadenosine 5'-[a-[35S]thio]triphosphate, and pUC plasmid universal primers (Promega) or other oligonucleotide primers were employed in the sequencing reactions. The strategy involved double-stranded sequencing of the inserts of pSM HCR3 and pSM HCR4, which were subcloned from pSM HCR2 (see Fig. 2). Subsequent to sequencing these inserts by using polylinker primers, other primers based on the sequenced regions were used to sequence remaining regions of the plasmids. These same primers were used to determine the sequence of regions of interest in pSM HCR2, pSM HCRO, and pSM 389 (see Fig. 2). PCR. PCR (12) was performed by using recombinant Taq polymerase (Amplitaq; Perkin-Elmer) and a programmable thermal cycler (MJ Research, Watertown, MA; model PTC100). Ten or 100 ng of target DNAs, recombinant plasmid or genomic DNA, was hybridized with the primer oligonucleotides (10 nM) at 64°C for 1 min, extended for 3 min at 72°C, and melted at 94°C for 1 min. Each PCR proceeded through 30 thermal cycles. The products were analyzed by electrophoresis through 0.7% agarose gels in Tris/borate/EDTA buffer, the gels were stained with ethidium bromide, and the products were sized by comparison with DNA standards (BRL).

Proc. Natl. Acad. Sci. USA 88 (1991)


RESULTS Distribution of the BamHI RFLP Sequence in HC-Sensitive vs. HC-Resistant Genomes. Our previous results (5) revealed that a cloned fragment of the ribosomal gene of S. mansoni, pSM 389, identifies a BamHI RFLP of -3.6 kb in Southern blots of DNA from drug-resistant parasites that was absent from the sensitive parent line (JHU.A) of S. mansoni. DNA fragments containing this RFLP were purified by electrophoresis and cloned into pUC 13. A clone (pSM HCR2) containing the RFLP was isolated by screening the resulting partial genomic library with the insert of pSM 389. To locate the sequence homologous to pSM HCR2 in the genome of the parent drug-sensitive parasites before the experimental production of HC resistance, Southern blots of BamHI-digested genomic DNAs of drug-sensitive and -resistant schistosomes were hybridized with 32P-labeled pSM 389 and then stripped and reprobed with 32P-labeled pSM HCR2. The hybridization signals obtained with pSM HCR2 were nearly identical to those obtained with pSM 389 (Fig. 1; compare a and b). In brief, a major band of hybridization at -3.0 kb (which represents the repeated copies of the 18S rRNA gene) and several minor bands are evident. A minor RFLP band at -3.6 kb [similar to that described previously using pSM 389 (5)] was seen in the genome of the drug-resistant parasites, which was absent in the drug-sensitive parasites (Fig. 1, thick arrow), indicating that pSM HCR2 detects the same drug resistance-associated RFLP that hybridizes to pSM 389. In addition, a reduction in intensity of an RFLP of -5 kb (Fig. 1, thin arrow) was seen with the drug-resistant DNA. These results suggested that the insert of pSM HCR2 contains 18S ribosomal gene sequences and that the pSM HCR2 and pSM 389 sequences are highly homologous. In addition, Southern analysis of restricted pSM HCR2 fragments probed with labeled pSM 389 (data not shown) indicated that the inserts of these plasmids are close to identical, despite the difference in their size. Structural Analysis of the Drug Resistance-Associated RFLP. Restriction maps of pSM HCRO (the JHU strain homolog of pSM 389) and pSM HCR2 were constructed to characterize the relationship of their inserts. This analysis a


1 2 3 4 1 2 3 4




pSM 389 pSM HCR2 FIG. 1. Comparative Southern blot analyses of hybridization signals obtained with the ribosomal gene probe pSM 389 (a) vs. pSM HCR2 (b) when these probes were hybridized sequentially to BamHI-digested DNA from drug-sensitive JHU.A (lanes 1 and 2) and drug-resistant JHU.G schistosomes (lanes 3 and 4). The drug resistance-associated RFLP is indicated in lanes 3 and 4 by the thick arrow, at about 3.8 kb. The thin arrow shows another RFLP (-5 kb), which is much less intense in lanes 3 and 4 compared to lanes 1 and 2. The major band of hybridization at -3.0 kb represents the 100 or so copies of the 18S ribosomal gene.


Medical Sciences: Brindley et al.

Proc. NatL Acad Sci. USA 88 (1991)

demonstrated the presence of Xba I and Pst I sites in pSM HCR2 that were absent from pSM HCRO and that pSM HCR2 contains rDNA sequences encompassing the 5' end of the 18S rRNA gene (Fig. 2). Therefore, the nucleotide sequences of the inserts of pSM HCR2 and pSM HCRO were compared to determine the structural differences between them. For this, Xba I-Xba I and Pst I-Pst I fragments, which are each =0.8 kb in length, of the insert of pSM HCR2 were subcloned as pSM HCR3 and pSM HCR4 and sequenced (Fig. 2). In addition, the sequences of substantial tracts of pSM HCRO, pSM HCR2, and pSM 389, including their 5' and 3' termini, were determined. The compiled sequence data demonstrate that pSM HCR2 differs from pSM HCRO by the presence of a 732-base-pair (bp) insert located 134 bp downstream from the junction of the intergenic spacer and the 5' end of the 18S rRNA gene. The junction of the intergenic spacer and the 5' end of the mature 18S rRNA gene was deduced by comparison to the nucleotide sequence of an 18S RNA gene of Plasmodium falciparum (14) (data not shown). The 732-bp insertion has the same sequence as the region contiguous to it. pSM HCR2 is identical to pSM HCRO except for this insertion. The nucleotide sequences in the region of the mutation are presented in Fig. 3. The nucleotide sequence allowed a more accurate calculation of the size ofthe insert ofpSM HCR2 as =3.8 kb, rather than 3.6 kb as reported previously for the drug resistanceassociated RFLP (5), since pSM HCR2 is 732 bp longer than pSM HCRO, and the insert of the latter (and of pSM 389) is =3 kb (Fig. 2) (13). Densitometric comparison of the hybridization signals of pSM 389 to the drug resistance-associated BamHI RFLP with the major band of hybridization at r3.0 kb (see Fig. la, lanes 3 and 4) showed that the former accounts for 1-4% of the combined signals of these two bands. Since there are about 100 rDNA copies in the NMRI strain of S. mansoni (13), the -3.8-kb HC resistanceassociated RFLP represents one, or at most a few, mutated 18S ribosomal gene(s). The Mutant rRNA Gene Is Absent from the Drug-Sensitive Parent Schistosome Population. PCR analyses were performed to determine whether the drug resistance-associated RFLP was induced in the genome of the parasites or whether it was present at low frequency in the drug-sensitive parent population and subsequently selected as a result of the HC treatment. An antisense oligonucleotide (25-mer), PJB6C (5 '-CTACAGTTATCCATGTTCAAAGTAA-3'), which straddles the 3' junction of the 732-bp insert and the 18S

rDNA unit


B 1 L-







11pSM 38' -








B !











46 91 136 181 226

r C

__ _C ___

271 316 361 406 451 496 541

___ ~ C mm

586 631 676 721 766 811 856 901 946 991 1036 1081 1126


IC= _

_ C













1216 1261 1306 1351 1396



1486 1531 1576 1621 1666

FIG. 3. Nucleotide sequence of a portion of the drug resistanceassociated BamHI RFLP cloned as pSM HCR2. The RFLP represents a 18S ribosomal gene mutated through the insertion of an extra 732 bp. The inserted sequence is in boldfaced type. The remaining uppercase letters represent the sequence of the 18S ribosomal gene sequence, which is identical to the sequence of pSM HCRO. The lowercase sequence represents spacer 5' to the start of the 18S rRNA gene. The positions of three oligonucleotides are shown.




I -



I X 1


P 1 1

I\i _1-J1




732 bp insertional

mutation x

P x

pSM HCR3 P x









oligonucleodde PJB6C

"mutation junctional sequence"

FIG. 2. Structural analysis of the drug resistance-associated BamHI RFLP, determined by comparison of restriction maps of the inserts of plasmids pSM HCRO and pSM HCR2. pSM HCR3 and pSM HCR4 were subcloned from pSM HCR2. Recognition sites of restriction enzymes BamHI (B), HindIII (H), Xba I (X), and Pst I (P) are shown. The physical map of S. mansoni rRNA-encoding DNA (rDNA) was prepared from refs. S and 13. The sequences and positions of oligonucleotides PJB1A and PJB6C are shown.

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rRNA gene (Figs. 2 and 3), and therefore uniquely identifies the RFLP sequence, was synthesized. The size of the expected PCR product defined by the junctional oligonucleotide PJB6C and by the sense-strand oligonucleotide (23-mer) PJB1A, which represents part of the regular rRNA gene (Figs. 2 and 3), is 529 nucleotides. A PCR product of this size was amplified from genomic DNA pooled from -1000 drugresistant schistosomes (JHU.G line). In contrast, no amplification was evident from DNA from pools of 3000 worms from the sensitive parent line JHU.A or from >10,000 NMRI.A parasites, a strain in which drug resistance could not be induced (Fig. 4a) (5). To examine the sensitivity of this PCR, we diluted drug-resistant genomic DNA into drugsensitive DNA. A target of 529 nucleotides was amplified with as little as 20 pg of JHU.C DNA mixed into 100 ng of drug-sensitive DNA, showing that this PCR has a sensitivity equal to or greater than 1:5000 (moles of drug-resistant DNA to moles of drug-sensitive DNA) (Fig. 4b). To determine whether the same insertional mutation occurs reproducibly during the induction of resistance, oligonucleotides PJB1A and PJB6C were employed as primers in PCR analyses of DNA from four independently derived, HC-resistant lines: JHU.C, JHU.D, JHU.F, and JHU.G (5). Products of -529 nucleotides were amplified from each line but not from sensitive JHU.A or NMRI parasites (Fig. 4c). Since pSM HCR2 was cloned from the resistant JHU.G line and because the sequence of pSM HCR2 showed that the dinucleotide TA located 134 bp downstream from the junction of the 18S gene was the target of the mutation, it appears that the mutant sequence was inserted between the same TA dinucleotide (or adjacent residues) in each of the four HCresistant lines.

In response to exposure to a xenobiotic agent, drug resistance will typically spread gradually through a population of target organisms as the consequence of selection of resistant genotypes present at low frequency. In marked contrast, the observations of Bueding and coworkers (3-5) on resistance in schistosomes to thioxanthenones ostensibly contradict this convention, since resistance appears universally in the first filial progeny of the parasites exposed to the drug. The phenotype of the schistosome population appears to convert in concert from sensitive to resistant. This phenomenon is reminiscent of Lamarckian inheritance. In this paper we (5) characterize the structure of a BamHI RFLP that we had observed previously to accompany the appearance of resistance in the Bueding model. The RFLP represents a mutant 18S ribosomal gene that arises through the insertion of an extra 732-bp sequence into the 5' end of one or a few copies of the tandemly arrayed rRNA genes of this parasite. A sensitive PCR analysis employing ajunctional oligonucleotide unique to the RFLP failed to amplify the target sequence from DNA pooled from a large population of '3000 drug-sensitive JHU.A line schistosomes but amplified a product of the expected size from DNA from their drugresistant progeny. Since resistance is obtained routinely by treatment of mice carrying >






I I I CC C c 7 Cz










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b 1






FIG. 4. PCR analyses of DNAs from drug-resistant vs. drug-sensitive schistosomes primed with oligonucleotides PJB6C and PJB1A. (a) Comparison of the drug-sensitive parent schistosomes (JHU.A) (lane 3), their drug-resistant progeny (line JHU.G) (lane 4), and an unrelated drug-sensitive strain (NMRI.A) (lane 2) for the presence of the mutant 18S ribosomal gene. Lane 1, DNA size standards (in nucleotides); lane 5, 500-nucleotide product from phage A DNA using A-specific oligonucleotide primers, as a control for the PCR. (b) Determination of the sensitivity of the PCR for the junctional mutation. Decreasing amounts of drug-resistant JHU.C DNA were diluted into 100 ng of drug-sensitive JHU.A DNA, and the DNA mixtures were used as templates. A product of -529 nucleotides (arrow) was detected in the mixtures containing 60 pg (lane 2), 40 pg (lane 3), and 20 pg (lane 4) of resistant DNA but not in the sample containing 10 pg (lane 5) of resistant DNA. Lane 1, size standards; lane 6, JHU.A DNA only. (c) Comparison of independently derived drug-resistant lines for the PCR products identified in a. Genomic DNAs from the drug-sensitive parent JHU.A line of schistosomes (lane 5), from the drug-resistant lines JHU.C, JHU.D, JHU.F, and JHU.G (lanes 1-4, respectively), and from drug-sensitive NMRI.A, NMRI.B, and NMRI.C lines (lanes 6-8) were examined. Lane 9, amplification of the phage A control; lane 10, size standards (in nucleotides).


Medical Sciences: Brindley et al.

We speculated previously that genomic rearrangements associated with drug resistance might involve transposition of DNA as a result of environmental stresses, such as exposure to HC (5). However, transposition does not readily explain the appearance of the BamHI RFLP in the progeny of the schistosomes exposed to HC, since the inserted sequence does not exhibit any of the hallmarks of mobile DNA, such as inverted terminal repeats or target site duplication (15). Instead, the mutation resembles an incomplete duplication of the 18S gene. Because the change occurs at the locus of a multiple copy gene, it may arise by unequal crossing-over. Alternatively, schistosomes may possess a site-specific recombination mechanism that is altered as a consequence of exposure to HC. In this regard, it should be noted that HC and its metabolites intercalate DNA (16, 17) and that HC is a frameshift mutagen (18). Only one, or at the most a few, of the 18S genes undergo the mutation that results in the appearance of the BamHI RFLP. Similar mutations in more than just a few of these genes may be lethal through the loss of ribosomes, or, alternatively, only a limited number of the genes may be susceptible to the mutation. If so, sequence microheterogeneity may exist among the copies of the 18S genes, raising the intriguing possibility that different types of 18S ribosomal genes may occur in schistosomes, in a way that is analogous to the situation in malarial parasites where different 18S genes are transcriptionally active during the insect and mammalian developmental forms of the protozoan (14, 19). Length fluctuations in ribosomal genes involving duplication of sequences in expansion segments have been reported in Drosophila, but these fluctuations are reported to reflect slippage-like mechanisms resulting in the duplication or loss of short, simple sequence motifs (20). We are unaware of other examples oflarge insertions within the 18S rRNA genes of other eukaryotes, although insertions occur within the 28S ribosomal genes and within the spacers that flank these genes. Long and Dawid (21) showed that 28S rRNA genes bearing these insertions were not functional or were transcribed only at very low levels. In like fashion, it appears that the mutated 18S gene is not operational since we were unable to detect transcripts of the expected, larger size (data not shown). The absence of transcription from the mutant 18S ribosomal gene in adult, drug-resistant parasites suggests that the interaction of HC with RNA (or ribosomes) is not involved in the mechanism of action of the drug (see refs. 16 and 22). The exact relationship between the internal duplication in rDNA described in this study and the genetic change responsible for drug resistance is not clear. We hypothesize that they arise through related genomic alterations (5) because both phenomena accompany one another and occurred reproducibly in independent experiments where resistance was induced in the JHU strain. In the case of the appearance of the -3.8-kb BamHI RFLP, the change involves a specific sequence in the target rRNA gene. It is unlikely that random mutagenic effects of HC (18) could cause the same rDNA alteration in each of the multiple occasions that resistance to the drug was induced. Similarly, it is also unlikely that the genetic change responsible for resistance to HC results through the mutagenic effects of the drug, unless the target resistance gene is large and unusually sensitive to mutation. An intriguing hypothesis to account for the induction of resistance to HC in schistosomes has been proposed by Cioli

Proc. Nad. Acad. Sci. USA 88 (1991) et al. (23). These authors postulate that HC is converted enzymically in vivo into a reactive ester that alkylates DNA, particularly deoxyguanosine bases (17), leading in due course to the death of drug-sensitive schistosomes through the interruption of nucleic acid synthesis. They argue that the enzyme is absent or defective in drug-resistant schistosomes. If this represents the mode of antischistosomal action of HC, then genomic rearrangements that lead to the dysfunction of the gene encoding the putative HC-esterifying enzyme may have taken place during the production of HC resistance, in a similar manner to the mutation that may have inactivated transcription from the target 18S rRNA gene characterized in this study. We thank Dr. F. Lewis for the schistosomes and Drs. T. Brodin, P. Englund, and E. Pearce and Ms. S. Rathke for helpful comments on this manuscript. This work was supported by the United Nations Development ogramme/World Bank/World Health Organization Special Programme for Research and Training in Tropical Diseases. 1. Cook, J. A. (1987) in Bailliere's Clinical Tropical Medicine and Communicable Diseases: Schistosomiasis, ed. Mahmoud, A. A. F. (Bailliere Tindall, London), Vol. 2, pp. 449-463. 2. Dias, L. C. de S., Pedro, R. de J. & Deberaldini, E. R. (1982) Trans. R. Soc. Trop. Med. Hyg. 76, 652-659. 3. Rogers, S. H. & Bueding, E. (1971) Science 172, 1057-1058. 4. Jansma, W. B., Rogers, S. H., Liu, C. L. & Bueding, E. (1977) Am. J. Trop. Med. Hyg. 26, 926-936. 5. Brindley, P. J., Lewis, F. A., McCutchan, T. F., Bueding, E. & Sher, A. (1989) Mol. Biochem. Parasitol. 36, 234-243. 6. Sabah, A. A., Fletcher, C., Webbe, G. & Doenhoff, M. J. (1986) Exp. Parasitol. 61, 294-303. 7. Beverley, S. M., Coderre, J. A., Santi, D. V. & Schimke, R. T. (1984) Cell 38, 431-439. 8. Stirewalt, M. A. & Uy, A. (1969) Exp. Parasitol. 26, 17-28. 9. Vogelstein, B. & Gillespie, D. (1979) Proc. Natd. Acad. Sci. USA 76, 615-619. 10. Simpson, A. J. G., Sher, A. & McCutchan, T. F. (1982) Mol. Biochem. Parasitol. 6, 125-137. 11. Sanger, F., Nicklen, S. & Coulson, A. R. (1977) Proc. Natl. Acad. Sci. USA 74, 5463-5467. 12. Saiki, R. K., Gelfand, D. H., Stoffel, S., Scharf, S. J., Higushi, R., Horn, G. T., Mullins, K. B. & Erlich, H. A. (1988) Science

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13. Simpson, A. J. G., Dame, J. B., Lewis, F. A. & McCutchan, T. F. (1984) Eur. J. Biochem. 136, 41-45. 14. McCutchan, T. F., de la Cruz, V. F., Lal, A. A., Gunderson, J. H., Elwood, H. J. & Sogin, M. L. (1988) Mol. Biochem. Parasitol. 28, 63-68. 15. Finnegan, D. J. (1989) Trends Genet. 5, 103-107. 16. Pica-Mattoccia, L., Cioli, D. & Archer, S. (1988) Mol. Biochem. Parasitol. 31, 87-96. 17. Archer, S., El-Hamouly, W., Seyed-Mozaffari, A., Butler, R. H., Pica-Mattoccia, L. & Cioli, D. (1990) Mol. Biochem. Parasitol. 43, 89-96. 18. Hartman, P. E., Levine, K., Hartman, Z. & Berger, H. (1971) Science 172, 1058-1060. 19. Gunderson, J. H., Sogin, M. L., Wollett, G., Hollingdale, M., de la Cruz, V. F., Waters, A. P. & McCutchan, T. M. (1987) Science 238, 933-937. 20. Hancock, J. M. & Dover, G. A. (1988) Mol. Biol. Evol. 5, 377-391. 21. Long, E. 0. & Dawid, I. B. (1979) Cell 18, 485-499. 22. De Stasio, E. A., Moazed, D., Noller, H. F. & Dahlberg, A. E. (1989) EMBO J. 8, 1213-1216. 23. Cioli, D., Pica-Mattoccia, L., Rosenburg, S. & Archer, S. (1985) Life Sci. 37, 161-167.

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