J. Mol. Biol. (1997) 269, 203±213
Regulation and Trafficking of Three Distinct 18 S
Ribosomal RNAs During Development of the
Malaria Parasite
Jun Li1, Robin R. Gutell2,3, Simon H. Damberger2, Robert A. Wirtz4
Jessica C. Kissinger1, M. John Rogers1, Jetsumon Sattabongkot5
and Thomas F. McCutchan1*
1
Growth and Development
Section, Laboratory of Parasitic
Diseases, National Institute of
Allergy and Infectious Diseases
National Institutes of Health
Bethesda, MD 20892-0425
USA
2
Department of MCD Biology
Campus Box 347, University of
Colorado, Boulder, CO 803090347, USA
3
Department of Chemistry and
Biochemistry, Campus Box 215
University of Colorado, Boulder
CO 80309-0215, USA
4
Entomology Branch, Division
of Parasitic Diseases-NCID
Centers for Disease Control and
Prevention, 4770 Buford
Highway NE, Atlanta
GA 30341-3724, USA
5
The human malaria parasite Plasmodium vivax has been shown to regulate
the transcription of two distinct 18 RNAs during development. Here we
show a third and distinctive type of ribosome that is present shortly after
zygote formation, a transcriptional pattern of ribosome types that relates
closely to the developmental state of the parasite and a phenomenon that
separates ribosomal types at a critical phase of maturation. The A-type
ribosome is predominantly found in infected erythrocytes of the vertebrate and the mosquito blood meal. Transcripts from the A gene are
replaced by transcripts from another locus, the O gene, shortly after fertilization and increase in number as the parasite develops on the mosquito
midgut. Transcripts from another locus, the S gene, begins as the oocyst
form of the parasite matures. RNA transcripts from the S gene are preferentially included in sporozoites that bud off from the oocyst and migrate
to the salivary gland while the O gene transcripts are left within the
oocyst. Although all three genes are typically eukaryotic in structure,
the O gene transcript, described here, varies from the other two in core
regions of the rRNA that are involved in mRNA decoding and translational termination. We now can correlate developmental progression of
the parasite with changes in regions of rRNA sequence that are broadly
conserved, where sequence alterations have been related to function in
other systems and whose effects can be studied outside of Plasmodium.
This should allow assessment of the role of translational control in parasite development.
# 1997 Academic Press Limited
Entomology Department, US
Army Medical Component
Bangkok, Thailand
*Corresponding author
Keywords: small subunit (SSU) rRNA; ribosome; development; malaria;
translational control
Introduction
The Apicomplexa comprises a group of parasitic
protozoa with increasing medical importance,
which includes Cryptosporidium, Toxoplasma, Theileria, Babesia and Plasmodium. The control of development of these parasitic protozoa is unusual and
appears to involve a broad control mechanism
lying at the center of translational machinery, ribosomal RNA. At least in the malaria parasite, Plasmodium species, structurally distinct ribosomes are
Abbreviations used: ITS1, 30 -internal transcribed
spacer.
0022±2836/97/220203±11 $25.00/0/mb971038
active during different stages of development and
differentiation (McCutchan et al., 1995). In most
eukaryotic organisms there is a correlation between
proliferative states of development and enhanced
rRNA transcription and ribosome production
(Sollner-Webb & Tower, 1986). The increase in ribosome production is occasionally accompanied by
modi®cation to the ribosome (Etter et al., 1994).
Plasmodium appears to respond to developmental
stimuli in a similar fashion except that points of developmental commitment and cell proliferation are
accompanied not only by increased ribosome production but also by changes in the probable catalytic moiety of the ribosome, the rRNA. Malaria
# 1997 Academic Press Limited
204
Ribosomal RNAs During Development of the Malaria Parasite
parasites maintain a life cycle which includes three
different asexual multiplicative stages and one sexual reproductive stage, all with different rates of
replication in the vertebrate and mosquito host
(Figure 1). Asexual reproduction takes place in the
liver (exoerythrocytic schizogony) and the blood of
the vertebrate host (erythrocytic schizogony), and
also on the mosquito midgut (sporogony). Sexual
reproduction initiates in the vertebrate blood
(gametogenesis) and ®nishes by fertilization of the
female macrogamete with the ex¯agellated male
microgamete in the mosquito midgut after engorgement of a blood meal. In previous studies, two
distinct 18 S rRNA (small subunit rRNA) genes
were identi®ed from Plasmodium vivax (Li et al.,
1994b). Transcripts from the type A gene are dominant in erythrocytic stages and gametocytes, while
the type S gene seems to be transcribed in the sporozoite (Figure 1). However, neither of these rRNA
types could be detected in the infected mosquito
during the early stage of P. vivax development (Li
et al., 1994b). The gap between termination of A
gene transcripts and expression of the S gene
prompted a more exhaustive investigation for a
correlation between development and the expression of rRNAs. Here we describe a third type
of 18 S rRNA (type O) which is associated with oocyst development in infected mosquitoes (Figure 1)
Figure 1. Schematic representation of malaria life cycle and stage-speci®c ribosomal RNAs. Human infection by Plasmodium begins when an infected female anopheline mosquito inoculates sporozoites into the bloodstream during
feeding. The sporozoites invade liver cells and transform into trophozoites. In six to eight days one mature schizont
will release thousands of liver-stage merozoites into the bloodstream (exoerythrocytic schizogony). The second
asexual proliferative stage (erythrocytic schizogony) is initiated when the liver-stage merozoites invade the erythrocytes giving rise to blood stage trophozoites. About 14 to 16 erythrocytic merozoites are generated in a 48-hour cycle
for re-infection. The merozoites may alternately differentiate into single gametocytes, the initial stage of the sexual
reproduction (gametogenesis). Mosquito infection begins when the gametocytes are drawn in the blood meal, and the
male microgametocyte ex¯agellates into individual microgametes and fertilizes the female macrogamete. The resulting zygote transforms into a motile ookinete, which penetrates the mosquito midgut and rounds up as an oocyst on
the external surface. After a period of 9 to 14 days, thousands of sporozoites are differentiated in the mature oocyst
(sporogony), the only multiplicative stage in the mosquito.
Ribosomal RNAs During Development of the Malaria Parasite
and show the existence of a mechanism for sorting
ribosomes in the oocyst which selects the type S ribosome for inclusion into the maturing sporozoite,
with the concomitant exclusion of the type O ribosome.
Exploring the multi-rRNA system in Plasmodium
species may both elucidate a basic mechanism of
developmental control and reveal targets for metabolic intervention of parasite growth. For example,
it has been shown that the ribosomal GTPase center, a site of antibiotic interaction, produced in the
asexual blood stage is different from that found in
the sporozoite stage (Rogers et al.,1996). One must
now also entertain the idea that there is a difference in af®nity of the type O ribosome for a
speci®c subset of mRNAs. Finally the addition of
the third type of rRNA enhances the possibility
that the direction of events leading to the evolution
of this multi-gene family, perhaps even the parasite, can be determined.
Results
Three rRNA genes are associated with
P. vivax infection
We have monitored Plasmodium ribosomal RNA
sequences from the blood of malaria patients as an
indicator of the species of infecting parasite (Li
et al., 1997). Investigation of patients in Thailand,
who were initially diagnosed by microscopy as
having P. vivax infection, revealed three distinct
types of P. vivax 18 S rRNA genes associated with
the infection. Partial 18 S rRNA genes were ampli®ed from DNA isolated from infected blood using
PCR primers that are conserved in the genus Plasmodium (Li et al., 1995). Sequence analysis of the
products revealed three genes. Two of these genes
have been reported as the A and S genes of P.
vivax (Li et al., 1994a). The third type of gene is referred to as type O for reasons that will be described below. Sequence information from a
fragment of this gene allowed the design of O
gene-speci®c oligonucleotides (Figure 2) for further
investigation, as the PCR from patients' blood was
designed with primers to conserved regions which
presumably amplify all the P. vivax 18 S rRNA
genes.
To con®rm association of the O gene with P.
vivax, we collected P. vivax-infected blood from
patients in different locations of central Thailand.
Ampli®cation by RT/PCR of RNA from patients
yielded products that hybridized to the probes
speci®c to the type A gene of P. vivax but not to
those designed to detect the other human malaria
parasites, including P. falciparum, P. malariae and P.
ovale (Li et al., 1995). Volunteer patients also allowed laboratory-reared mosquitoes, Anopheles
dirus, to feed on their arms before they were treated for malaria. The species of parasite maturing in
the mosquitoes was also determined using species
speci®c monoclonal antibodies directed to the P.
205
vivax circumsporozoite protein, which is the major
surface protein expressed in sporozoites (Wirtz
et al.,1991). The four samples collected from
patients did not have mixed infections as determined to the limits of our detection techniques
(data not shown). To further substantiate the association of the O gene with P. vivax, the complete
18 S rRNA O gene was isolated from a laboratorymaintained strain of P. vivax, Sal-1, originally isolated in El Salvador. The O gene was produced in
two overlapping fragments (1.6 kb and 1.9 kb) ampli®ed with speci®c primer pairs, 705/683 and
706/573 (Figure 2). Hybridization shows that both
fragments are recognized by an O gene speci®c
oligo-probe (741) targeted to the central overlapping regions (Figure 2). The cloned fragments contain the entire 18 S rRNA genes and the ITS1
region.
The type O transcript appears soon after
fertilization in developing ookinetes and
oocysts but is segregated from
budding sporozoites
The temporal and developmental pattern of
rRNA expression was determined for the period of
parasite development within the mosquito
(Figure 3a). For this purpose, mosquitoes were collected at different time intervals after the infectious
blood meal and total RNA was isolated from the
separated thorax and abdomen. The origin of the
parasite RNA could be determined, since mature
sporozoites migrate from developed oocysts on the
midgut wall to the salivary glands of the thorax
(Figure 3a). RNA originating from mature sporozoites in salivary gland was distinguished from
that found in the oocyst by severing the mosquito
at the junction separating the abdomen and the
thorax. Only RNA from mature sporozoites is
found in the thorax sample while the abdomen
sample contains RNA from both parasites in the
blood meal and those proceeding through development to the mature oocyst stage. The RNA from
the thorax and abdomen was then analyzed for expression of the distinct genes during the time
course of development in the mosquito (Figure 3b).
The results indicate that the A-type transcript was
the dominant rRNA detected in infected blood and
in the gut of engorged mosquitoes. The blood
stage rRNA decreased rapidly and was detected
only in low levels in the mosquito gut at 24 hours
after feeding (Figure 3b). The novel type of rRNA,
which we present here, was identi®ed after the disappearance of the A gene transcript and remained
throughout oocyst development (Figure 3b). By
convention with nomenclature, the third type of
gene is referred to as the O gene as it re¯ects development starting in the ookinete stage and continuing through the oocyst stage (Figure 1). Type S
rRNA was not detected until day 6 (Figure 3b),
when the oocyst is close to mature and sporozoites
are differentiated (Li et al., 1994b). The increasing
206
Ribosomal RNAs During Development of the Malaria Parasite
signal after this point may relate to an increase in
the mass and number of parasites as they develop into mature sporozoites. Clearly, the type O
and S genes are transcribed during the later
period of oocyst development but they are distin-
guished by the pattern and location of expression. The type O gene is transcribed earlier
and limited to the abdomen of the infected mosquito (Figure 3b), and is not transferred to the
thorax portion of the mosquito. The type S rRNA
Figure 2. Sequence alignment of three 18 S rRNA genes from P. vivax Sal-1 strain. The coding region for the mature
rRNAs starts from the 50 end and is followed at the 30 end by the internal transcribed spacer ITS1 (open box) and the
50 end of the 5.8 S rRNA gene (shaded box). The upper line shows the O gene sequence; the middle and lower lines
represent the S and A gene, respectively, and are shown only where the sequences differ from the O gene. Dashes
represent gaps where the sequences cannot be directly aligned. Oligonucleotides complementary to the sequences
used for cloning and analysis of the gene expression are indicated by shaded boxes, with numbers and direction. The
GenBank accession numbers for the P. vivax 18 S rRNA gene sequences are: type A U07367, type O U93095 and type
S (previously C) U07368.
Ribosomal RNAs During Development of the Malaria Parasite
peaks around day 10 to day 12 when rapid
nuclear division and differentiation of sporozoites
in oocysts occur. On day 14 S-type rRNA was
detected in the thorax, suggesting that it is the
207
dominant type of rRNA in sporozoites that migrate from the maturing oocyst on the midgut in
the abdomen to the salivary glands in the thorax.
This association was also supported by the fact
Figure 3. Developmentally regulated transcription of the rRNA genes in P. vivax. a, Schematic view of parasite development in mosquitoes following an infectious blood meal (in days, where D 0 ˆ day 0, etc.). The progressive course
of differentiation and growth stages is drawn appropriate to days following the blood meal. R, ring form; T, trophozoite; Sc, schizont; M, merozoite; G, gametocytes (male and female); Z, zygote; K, ookinete; O, oocyst and S, sporozoites. Stages are colored for ease of identi®cation, and a line indicates separation of thorax and abdomen RNA
samples. b, Autoradiograph of a Northern blot analysis of total RNA prepared from infected mosquitoes at different
days after an infectious blood meal. The RNA, from either thorax or abdomen, was ®rst separated by electrophoresis
on an agarose gel and immobilized on a nylon membrane. Then, triplicates of the blots were individually probed
with 32P-labeled oligonucleotides 741, 742 and 743, which are speci®c for type A, type O and type S rRNAs, respectively. Each slot represents the average signal of RNA from one tenth of a ten-mosquito pool collected at each time
point. The sample from day 0 (D0) was collected two hours after the mosquito fed on the infected patient. Lanes B
and M represent RNA from P. vivax infected patient blood and uninfected mosquito, respectively. The middle panel
was autoradiographed for 12 hours while the top and lower panels were autoradiographed for three hours.
208
Ribosomal RNAs During Development of the Malaria Parasite
that the S-type rRNA is the only type detected in
sporozoites puri®ed from the salivary glands
(data not shown).
The secondary structure of the type O rRNA
differs from the other 18 S rRNAs in regions
associated with translational function
Sequence alignment of the three P. vivax rRNA
genes (Figure 2) indicates that the O gene is different from the other two genes, although they share
signi®cant homology. Comparison of the ITS1 sequences shows little similarity among the three
genes (Figure 2), further indicating three distinct
transcription units. The secondary structures for
the three 18 S rRNAs were derived by the comparative method. By searching for positional covariances secondary structure models for the type
O, A and S rRNAs were determined, based on the
existing Eukarya 18 S rRNA and Eubacterial 16 S
rRNA models (R.R.G., data not shown; available
from http://pundit.colorado.edu:8080/root.html).
Analysis of the O gene sequence shows that it is,
overall, very similar in sequence and structure to
the A and S genes (Figure 4). The structural analysis shows that it forms all of the expected domains
found in the Eukarya 18 S rRNA consensus,
although, as discussed below, there are anomalous
insertions and deletions. The secondary structure
of the O gene has many compensatory base-pair
changes from the A and S genes and the overall
conservation with the Eukarya, together with its
stage speci®c expression (Figure 3), strongly
suggests an active and functional role. In sharp
contrast, there are about 36 positions where the O
gene is different from the Apicomplexa and Eukarya consensus. There are an additional 18 positions
where the O gene differs from the Apicomplexa
consensus. Overall, about 12 positions are insertions in the O gene, with three occurring in highly
variable regions and probably not signi®cant (denoted in green, Figure 4). Nine insertions are prominent as they occur at positions well conserved in
the Eukarya, and in many cases these are conserved in all rRNAs. The size of the insertion
ranges from a single nucleotide to more than 20
nucleotides in the O gene (Figure 4). There are also
about nine signi®cant deletions in the O gene, relative to the Eukarya consensus, which are at several
highly conserved regions of the 16 S-like rRNA
(Figure 4). It is noteworthy that the sequence of the
O gene obtained from ampli®cation of genomic
DNA was identical to that derived from rRNA
transcripts (data not shown). Northern blots were
also probed with oligonucleotides complementary
to the unique insertions in the O gene to con®rm
expression of these regions in the mosquito stages
and, hence, these sequences are present in the mature O gene product (Figure 3b). The very long
``inserted'' variable region (approximately 1135 in
Escherichia coli numbering) is unusual in the O
gene. It is longer than the equivalent region in the
A and S genes and in the O gene forms a very
long, extended helix with few internal loops and
mismatches. In contrast, the equivalent helix contains several internal loops in the A and S genes,
making their helices more irregular (R.R.G., data
not shown).
Of the anomalous differences in the O gene, the
most signi®cant are discussed here. There are two
regions in the O-type rRNA that vary at positions
that have been shown in other studies to be involved in the decoding step in protein synthesis
(Figure 4); these are unique variations in the O
gene as the A and S genes resemble all other Eukarya. One of these regions is near the 30 -end,
shown in red (Figure 4; 1400 region in E. coli numbering). This area is usually termed the decoding
site as it is in close proximity to tRNA at the A, P
and E sites in the ribosome and also associates
with the mRNA (Zimmerman, 1996). The Sal-1 O
gene has both a 22 nt insertion and a deletion of a
core conserved base-pair (Figure 4) (corresponding
to the conserved 1399 1504 base-pair in E. coli)
which disrupts the O gene transcript's secondary
structure in this region. Another region altered in
the O-type rRNA corresponds to helix 34
(nucleotides 1046 to 1067 and 1189 to 1211 in E. coli
numbering). This helix is also in intimate contact
during protein synthesis as it is in close proximity
with mRNA and tRNA (Zimmerman, 1996). A deletion at position 1054 was originally isolated as a
suppresser of UGA termination codons (Murgola
et al., 1988), although subsequent analysis has
shown a wider effect on translational accuracy in
E. coli and Saccharomyces cerevisiae (Chernoff et al.,
1996; Goringer et al., 1991; Moine & Dahlberg,
1994). The type O SSU rRNA in this region has
three insertions in the 30 -half of helix 34, one of
which corresponds to an insertion at position 1200
(Figure 4). Mutation at this position in E. coli has
signi®cant effects on translational accuracy, causing read-through and frameshifting (Moine &
Dahlberg, 1994). The insertions in the 30 -side of
helix 34 in the type O rRNA also correspond to a
site of rRNA cleavage of the P. falciparum type A
rRNA that degrades the transcript during early development in the mosquito (Waters et al., 1989),
and the possible signi®cance of this is discussed
below. Another insertion occurs at positions corresponding to the 912 region in E. coli. Mutation at
this position also affects translational accuracy in
E. coli and S. cerevisiae (Liebman et al.,1995;
Lodmell et al.,1995). Hence it is likely that the O
gene transcripts have an altered speci®city in translation or decay of certain mRNA species. These
mRNAs are likely to be critical for development of
the parasite. The O gene has other ``non-intron''
and anomalous insertions that occur in highly conserved regions of this rRNA have, to date, only
been documented in this O gene sequence (shown
in red; Figure 4). All of these insertions should be
considered signi®cant and most unusual. There are
a few insertions that occur in less conserved regions of this rRNA (green boxes; Figure 4). These
Ribosomal RNAs During Development of the Malaria Parasite
occur in the O gene relative to the A and S genes.
In summary, there are nine signi®cant (red) insertions and three less important (green) insertions. In
209
addition, there are several nucleotides deleted in the
O gene, relative to the Eukarya consensus. These
positions are denoted with a large red dot (Figure 4).
Figure 4. Comparison of the secondary structures of P. vivax 18 S rRNAs: type O compared with type A and S. The
helices are represented according to Watson-Crick, wobble (G*U) and unusual (G*A) base-pairing. Regions of variable sequence are left mostly as nucleotides in line format. Insertions and missing nucleotides relative to the A and S
genes are color coded, as discussed in the text, and insertions in variable and conserved regions in the O gene are
color shaded. Compensatory base-pairs and differences in the O gene sequences are indicated by the symbols (!*;
see the legend on the Figure). Secondary structures for the P. vivax rRNAs are available from the World Wide Web
site for RNA secondary structures (http://pundit.colorado.edu:8080)
210
Ribosomal RNAs During Development of the Malaria Parasite
These all occur in conserved regions of the 18 S
rRNA and should be considered signi®cant.
Phylogenetic analysis suggests an order of
divergence of the three rRNA genes
Plasmodium are members of the Apicomplexa,
with Dino¯agellates as the likely sister group
(Allsopp et al., 1994; Escalante & Ayala, 1995). The
relationship of the three P. vivax rRNA genes to
each other and other relevant Apicomplexa was estimated by maximum parsimony analysis
(Swofford, 1993). The sequences used for analysis
are conserved regions that can be unambiguously
aligned among all sequences compared (R.R. Gutell, unpublished). The results (Figure 5) show that
Plasmodium species are well separated from the
other Apicomplexans but that the relationships of
the different Apicomplexa lineages to each other
are less certain, with no bootstrap value over 60%.
The rRNA genes of other members of the Apicomplexa are more distantly related (Sarcocystis, Toxoplasma and Cryptosporidium). As expected,
Dino¯agellata form the sister group (Prorocentrum
and Symbiodinium), and Ciliophora (Paramecium
and Oxytrichia) are outgroups in the phylogram
(Figure 5). Among the P. vivax rRNA genes there
have been two independent gene duplication
events; the ®rst leading to the O gene and the antecedent of the A/S lineage. The second duplication
event occurred later and lead to the A and S genes.
The duplication and divergence leading to the A
and S genes is not a recent event because when the
A-type genes from the monkey malarias (P. fragile,
P. knowlesi, P. simium) and the human parasite P.
malariae are added to the data set, they form a
monophyletic group. This demonstrates that the A
and S genes diverged before speciation. The O
gene has been detected by expression in at least
two other Plasmodium species (J. L. et al., unpublished), and is therefore likely to be a common or
perhaps universal feature of the genus.
Discussion
Plasmodium species exhibit remarkable control
over ribosome production during parasite development. The rRNA genes are few in number, between four and eight rDNA units (McCutchan et al.,
1995), and are genetically unlinked, being dispersed throughout the genome (Wellems et al.,
1987). The genes appear to accumulate mutations
independently of each other and hence sequence
differences between the units do not occur at the
same rate as in other organisms (Rogers et al.,
1995). The presence of structurally distinct rRNA
genes and the physical separation of the transcription units on different chromosomes may be involved in selectively accessing and expressing
stage-speci®c rRNA during different periods of the
parasite development. The controlled switching of
rRNA gene transcription in Plasmodium was
suggested in earlier studies (Gunderson et al.,1987;
McCutchan, 1986; McCutchan et al.,1988; Waters
et al., 1989). The exact timing of the transcriptional
switches varies somewhat with the species of parasite, and even the temperature at which the parasite is being maintained, but the sequence of events
is thought to be uniform. The type A rRNA is the
dominant transcript in erythrocytic stages includ-
Figure 5. Phylogram of 18 rRNA genes. Horizontal branch lengths between nodes correspond to the number of
shared derived changes. Bootstrap percentages are indicated above each branch. Analysis parameters are de®ned in
Materials and Methods. For ease of interpretation, the Phyla are differentiated by the intensity of background
shading.
Ribosomal RNAs During Development of the Malaria Parasite
ing gametocytes, the initial form of sexual stages.
Here we show that after zygote formation in the
mosquito midgut, the type A rRNA is replaced by
the type O rRNA. Transcription of the O gene continues in the oocyst through the entire development of the parasite in the mosquito. The S type
rRNA is initiated about a week after the infectious
blood meal, when differentiation of sporozoites begins in maturing oocysts. The rapid increase of the
S-type rRNA corresponds with the period for
differentiation of sporozoites, which can migrate
from the mature oocyst to the salivary gland,
ready for infection of humans. The S type rRNA is
replaced by the type A rRNA when merozoites
differentiate in maturing liver stage schizonts (Li
et al, 1991). The merozoites released from mature
schizonts will initiate the erythrocytic stage of the
parasites in which the A gene dominates. Thus, the
switch of rRNA types correlates more with the progression of distinct stages of development than
with the presence of the parasite in one host or the
other (summarized in Figure 1), suggesting that
regulated transcription of stage-speci®c rRNA
could be an integral part of the developmental control of the Plasmodium species.
The possible functional signi®cance of maintaining alternative forms of rRNA is indicated by the
developmental regulation of their expression and
by structural differences in core regions known to
be associated with biological function in other organisms (Figure 4). The three distinct rRNA genes
identi®ed from P. vivax each correspond to a discrete proliferative stage during parasite development. The O gene is signi®cantly different in the
core regions which are associated with the accuracy center in the ribosome (Liebman et al., 1995).
Changes in sites universally identi®ed with mRNA
association and decoding suggest that differences
in ribosome af®nity for subsets of mRNA or suppression of translation termination are involved in
the developmental progress. There also appear to
be active mechanisms involved in the turnover
from one type of ribosome to another. The ®rst indication that a distinction was being made between
ribosomal types co-existing within the developing
parasites in the mosquito came from a study showing a preferential breakdown of type A transcripts
in the zygote (Waters et al., 1989). This was initially
dif®cult to understand, since the point of cleavage
within the type A transcript occurs in a core region
of the RNA that is common to eukaryotic small
subunit rRNAs. We present two possible explanations for this ®nding. The O and A gene transcripts vary in sequence at the processing site and
speci®c cleavage could be responsible for the stability of the O ribosome in the presence of degrading A ribosomes. Physical separation of the
ribosomal types may also account for the selective
stability of the O type ribosome. Localization of
transcripts during the period of oocyst development indicates that there are mechanisms involved
in physically separating one type of ribosome from
the other. This occurs despite the fact that the cyto-
211
plasm of the oocyst is known to be open to traf®cking of organelles to the maturing sporozoites until
the point when the sporozoite buds off into the
hemocoel. Here we show that although the cytoplasm is open to traf®cking, the type O ribosomes
are found only in oocysts and do not go into the
sporozoite. Hence there is a mechanism keeping
the newly forming translational system separate
from the existing one.
Having three distinct types of rRNAs associated
with a single organism permits one to study the
progression of events associated with the evolution
of a new ribosome and the global effects associated
with such a change, which is unprecedented. A
number of initial conclusions can be reached with
these data. The multiple ribosome system represents a relatively recent adaptation after the
divergence of Plasmodium from all other Apicomplexa that have been analyzed to date, since
other members of the genus maintain complex life
cycles and do not have a multiple ribosome system. We show that the genes themselves appear to
have evolved via two gene duplication events. The
O gene, speci®c for oocyst development, diverges
before the other two genes in P. vivax (A and S),
when other Apicomplexa are used to root the phylogenetic tree. Either the O gene or the A/S progenitor most closely resembles the ancestral state.
The most recent duplication and divergence gave
rise to the A and S genes. Neither of these duplications occurs very deep on the Plasmodium
branch, suggesting that their origin is fairly recent.
There are numerous examples of genes originating by duplication of a parent gene, sequence drift
and co-option (Li & Graur, 1991). This scenario for
the generation of a multiple ribosome system, involving changes in core regions of rRNA, would
indicate a more complicated process. This would
include global changes involving compensatory alteration of molecules associated with the ribosome
complex (e.g. ribosomal proteins and mRNAs). The
other scenario which has resulted in different cytoplasmic ribosomes being expressed in a single cell
is secondary endosymbiosis, where one eukaryote
engulfs another, permanently retaining a secondary nucleus, the nucleomorph, surrounded by its
own cytoplasm (Palmer & Delwiche, 1996). Endosymbionts are essentially a cell within a cell which
maintain separated cytoplasmic compartments
with different ribosomes. The possibility that Plasmodium species are derived from an endosymbiotic
event has been indicated (Wilson et al., 1994). The
work described here does not indicate cytoplasmic
compartmentalization in the oocyst, as there is no
indication of this from ultrastructural studies
(Sinden & Strong, 1978). The presence of segregated ribosomes may prompt re-investigation of
the question and presents the tools for such inquiry. This presents a challenge to the molecular
phylogeneticist, who, when other related genes are
available, should be able to relate ribosomal
change to the highly successful adaptation of this
parasite to its present niche. For those who study
212
Ribosomal RNAs During Development of the Malaria Parasite
the machinery involved in protein synthesis the
opportunity is presented to understand the adaptation of an essential and complex system in the
presence of an already successful one.
Materials and Methods
Amplification of parasite rDNA
Oligonucleotides used for ampli®cation and detection
of DNA fragments are designated by number and
shown in Figure 2. Genomic DNA of P. vivax was prepared from four Thai isolates, which were con®rmed by
the CS protein based-ELISA test (Wirtz et al., 1991), and
one laboratory strain Sal-1 (El Salvador strain) as described previously (Li et al., 1994b). The complete 18 S
rRNA gene, including the 30 internal transcribed spacer
(ITS1) region, was ampli®ed by polymerase chain reaction (PCR) from the DNA of the Sal-1 strain with oligonucleotides 705 and 573 as the 50 and 30 -end primers,
respectively. Partial 18 S rDNA fragments were produced from the four Thai isolates with a pair of primers
566 and 570, approximately 140 and 40 nucleotides
shorter from the 50 and 30 -ends of the coding region. The
reaction was in a volume of 100 ml containing 20 to 50 ng
DNA, 200 mM of each dNTP, 50 mM KCl, 10 mM TrisHCl (pH 8.3), 2 mM MgCl2, and 2.5 units Taq DNA polymerase (Perkin Elmer Cetus; Norwalk, CT) in a Perkin
Elmer DNA Thermal Cycler with the following parameters: 94 C/one minute, 55 C/one minute, 72 C/two
to three minutes and a total of 30 cycles.
Cloning and sequence analysis
The procedure for cloning ampli®ed rRNA genes has
been described (Li et al., 1994b). The complete sequences
of the three 18 S rRNA genes and the ITS regions were
determined in both directions from the Sal-1 strain and
Thai isolates. Sequence alignment was performed with
Lasergene software (DNASTAR Corp., Madison, WI)
and AE2 software (T. Macke, Scripps Clinic; available
from Ribosome Database Project at http://rdp.life.uiuc.edu). Secondary structure models were inferred from
comparative sequence analysis. The sequences of P. vivax
rRNAs were aligned with those of the available 18 S
rRNAs, and the models were constructed according to
maximum similarity in both primary and secondary
structures (Gutell et al., 1994), since identical structure
can be folded from the same type of rRNA with different
primary sequences. Our comparative studies have also
identi®ed tertiary interactions (Gautheret et al., 1995).
The structures were drawn with the computer program
XRNA developed by B. Weiser and H. Noller (University
of California, Santa Cruz; available at ftp://fangio.ucsc.edu/pub/XRNA).
Isolation of RNA from infected mosquitoes
Laboratory-reared mosquitoes, Anopheles dirus, were
fed on P. vivax-infected Thai patients. The engorged mosquitoes were held at 26 C and ten mosquitoes were removed and frozen at ÿ70 C at intervals beginning two
hours after the blood meal and thereafter every two days
except day 4. The development of the parasite in the
mosquito was followed by microscopic examination of
midguts for oocysts and salivary glands for sporozoites.
Preparation and analysis of total RNA from the P. vivax
infected blood and mosquitoes were based on the meth-
od previously described (Chomczynski & Sacchi, 1987;
Li et al., 1994b).
Phylogenetic analysis
18 RNA sequences were aligned based on the conserved sequences and corresponding secondary structures (Gutell et al ., 1994). Regions of the genes that were
not unambiguously aligned were not used in the analysis. Sequences were analyzed by the parsimony method
(Swofford, 1993) using 100 heuristic random addition replicatives and 100 bootstrap replicates. Uninformative
characters were ignored. Genbank accession numbers for
the sequences used in this study are: P. vivax (Sal-1)
U07367, U03768 and U93095 (this study); P. vivax (Thai)
U93233, U93234 and U93235 (this study); Plasmodium cynomolgi L08241 and L08242; Babesia equi Z15105; Theileria
buffeli Z15106; Sarcocystis gigantia L24384; Toxoplasma
gondii M97703; Cryptosporidium wrairi U11440; Prorocentrum micans M14649; Symbiodinium microadriaticum
M88521; Paramecium tetraurelia X03772; Oxytricha nova
M14601.
Nucleotide sequence data reported in this paper have
been reported to GenBank with the accession numbers
U93233, U93234, U93235 and U93095.
Acknowledgements
Procedures for drawing blood and feeding mosquitoes
on patients were approved by the human use committees of the Ministry of Public Health, Thailand, and US
Army Medical Research and Development Command.
The work was supported by a World Health Organization grant, 890093, and by NIH grant GM48207
(awarded to R.R.G.) and the NSF/Sloan Foundation
(J.C.K.). R.R.G. and S.H.D. thank the W.M. Keck for
their generous support of RNA Science on the Boulder
Campus.
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