Molecular Infectious Diseases Group,
Department of Paediatrics, Institute of Molecular Medicine, John
Radcliffe Hospital, Oxford OX3 9DS, United
Kingdom,1 and Institute of Medical
Microbiology and Immunology, University of Copenhagen,
Copenhagen,2 and Danish Pest Infestation
Laboratory, Lyngby,3 Denmark
 |
INTRODUCTION |
Animal models have been widely used
as a source of Pneumocystis carinii organisms and for
studying many aspects of P. carinii infection. This is
because sustained in vitro cultivation of P. carinii has not
been possible (1, 38), although recently a new method has
been reported which is now being evaluated in a number of centers
(25). The rat model has been particularly useful in studies
of epidemiology (11, 50), drug sensitivity (59),
immunology (51), and the biology of the organism
(42). Rat- and human-derived P. carinii
organisms, however, are known to differ significantly in many respects.
Considerable divergence has been shown between the genes of rat- and
human-derived P. carinii (36, 43), as well as
antigenic (9) and ultrastructural (5) differences.
Two genetically divergent types of P. carinii organisms,
known as Pneumocystis carinii f. sp. carinii and
Pneumocystis carinii f. sp. ratti, have been
found in rat lungs (34). These were originally identified by
differences in electrophoretic karyotype (7) and
subsequently by DNA sequence variation at a number of genes, including
those for the nuclear 26S rRNA (21, 29), the mitochondrial
large-subunit (mt LSU) rRNA (14), the mitochondrial small-subunit rRNA (12), the TATA binding factor
(45), BiP chaperonin (40), thymidylate synthase
(14), and ATPase (24), and the 
subunit of the
G protein (39). The level of genetic divergence between
these two types of rat-derived P. carinii was sufficiently
high for them to be classed as different formae speciales (34,
44). Although eight different electrophoretic karyotypes of
P. carinii f. sp. carinii and two of P. carinii f. sp. ratti have been identified, very little
genetic variation in the form of DNA sequence polymorphisms has been
observed among isolates of either P. carinii f. sp.
carinii or P. carinii f. sp. ratti (6). In contrast, only one forma specialis has been found in human infections, Pneumocystis carinii f. sp.
hominis. However, divergence has been observed within this
forma specialis, detected as DNA polymorphisms at a number of loci,
including the mt LSU rRNA, the mitochondrial small subunit rRNA, the
arom locus and the internal transcribed spacer (ITS) regions
of the nuclear rRNA operon (15, 20, 22, 47, 48, 49). The ITS
regions have been shown to be more informative in distinguishing
between types, and as many as 59 different types have been identified
at this locus (19, 32; R. F. Miller and A. E. Wakefield, unpublished results).
One of the major differences between the human and rat infections is
that human-derived P. carinii samples are obtained from acutely infected individuals who have been exposed to the environment, whereas the rat model uses laboratory-bred animals housed in close proximity and at some level of isolation from the environment. In this
study, we examined P. carinii infection in wild rats. A
previous study of P. carinii infection in wild rats, using
microscopy for detection of P. carinii cysts, suggested that
a significant proportion of rats carried P. carinii, though
these studies lacked the sensitivity and discriminatory power of
contemporary DNA amplification techniques (35). We have used
a PCR technique which can detect P. carinii DNA with great
sensitivity and which allowed us to differentiate between the two known
formae speciales of rat-derived P. carinii (31).
We used amplification at the mt LSU rRNA gene for this study, since all
known P. carinii formae speciales have been analyzed at this
locus (8, 54, 56) and it is known to be a sensitive and
robust target for diagnostic PCR of clinical samples (49,
58), samples of rat lung (50), and environmental samples (52, 53).
In this paper we provide evidence to suggest the presence of five
different formae speciales of P. carinii in lung samples from Danish wild rats. We report a high incidence of P. carinii organisms in these samples and a high frequency of mixed
infections, and we show that only low levels of sequence heterogeneity
were observed in the ITS regions of P. carinii f. sp.
carinii in the samples. We compare the results to those from
human P. carinii infection, where only one forma specialis
has been found, with a large number of polymorphisms in the ITS regions.
| |
MATERIALS AND METHODS |
Samples.
Wild brown rats (Rattus norvegicus) were
live trapped at various domestic locations throughout Denmark. The 51 nonimmunosuppressed rats were then housed singly or in groups of up to
3 per cage in the laboratory and sacrificed after 0 to 78 days. The
animals were all housed initially in a quarantine room and then
transferred to a test room, both of which were under negative pressure.
There were no immunosuppressed animals housed in the same building. No
immunosuppressive drugs were administered, nor were the rats treated
with drugs against bacterial infections or worms. Six wild rats were
trapped, taken to a different animal facility, and immunosuppressed
with corticosteroids. The two rats WR2653 and WR2715 died after 9 and
13 days, but the remaining rats continued on immunosuppressive drugs
until they were sacrificed at between 38 and 48 days.
The laboratory rats were male Sprague Dawley rats weighing 200 to
220 g obtained from Harlan, Leicester, United Kingdom.
Nonimmunosuppressed laboratory rats were not placed in rooms where
immunosuppressed animals were housed, and they were sacrificed
immediately on arrival at the animal facility. Immunosuppressed
laboratory rats were housed in cages with up to six animals and were
treated with dexamethasone (Organon, United Kingdom) and antibiotics as
previously described (55). The rat lungs were recovered at
sacrifice and stored at 
80°C until analysis.
Spore trap samples were collected, and DNA was extracted from them as
previously described (52, 53).
DNA extraction.
Approximately one-fifth of each rat lung was
used for extraction of total DNA. The tissue was minced using sterile
scalpels and digested with 1 mg of proteinase K ml
1 in 10 mM EDTA (pH 8.0) and 0.5% sodium dodecyl sulfate at 50°C overnight,
after which a further 1 mg of proteinase K ml1 was added
and the sample was incubated for a further 24 h. The DNA was
purified by phenol-chloroform extraction, followed by a DNA binding
resin (Wizard DNA Clean-Up system; Promega, Southampton, United
Kingdom). All precautions were taken throughout these procedures to
eliminate the possibility of cross contamination of samples. All
handling of the samples took place in a laminar flow cabinet, and
negative controls were included in the extraction procedure to monitor
for contamination.
DNA amplification.
PCRs were performed using reagents at the
following concentrations: 50 mM KCl, 10 mM Tris (pH 8.0), 0.1% Triton
X-100, 3 mM MgCl2, 0.04 mM (each) deoxynucleoside
triphosphate, 1 µM oligonucleotide primers, and 0.025 U of
Taq polymerase (Promega) ml1. All PCR
experiments were performed on three different dilutions of sample DNA.
For samples which tested negative for all types of P. carinii, DNA was reextracted from the tissue and the PCRs were
repeated. The utmost care was taken at all times to prevent and monitor
for contamination. Negative controls were included for all samples, and
all handling of reagents occurred in a laminar flow cabinet, using
sterile tubes, pipette tips, and aliquoted reagents.
The primers used to amplify a portion of the mt LSU rRNA were as
follows: pAZ102-H and pAZ102-E (57, 58) and pAZ102-X/RI and
pAZ102-Z/RI (53); RC1, RC2, RR1, and RR2 were used to
specifically detect P. carinii f. sp. carinii and
P. carinii f. sp. ratti DNA (31). The
ITS1 and ITS2 regions are flanked by rRNA genes that are highly
conserved between P. carinii formae speciales and other fungi, whereas the ITS regions themselves are highly divergent. Where
the samples were likely to contain DNA from other fungi, it was
necessary to use PCR primers designed for the ITS regions which would
preferentially amplify P. carinii ITS sequences. The ITS
regions from environmental samples were amplified using primers designed specifically for P. carinii f. sp.
carinii in both the first (ITS21/HI and ITS22/HI) and second
(ITS25/HI and ITS26/HI) rounds of PCR in order to amplify only the
desired sequences from this diverse pool of DNA. Primer ITS21/HI
(5'-CGGGATCCACCTGCGGAAGGATCATTAAT-3') was designed to match
the 3' end of the 18S rRNA gene and the first 2 bp of the ITS1 region,
and ITS22/HI (5'-CGGGATCCCTGATTTGAGGTCAAAGGTTC-3') was
designed to match the 5' end of the 26S rRNA gene and the last 6 bp of
the ITS2 region. The nested primers ITS25/HI
(5'-CGGGATCCGAACTAGTTTATCTGGTTCTTG-3') and ITS26/HI
(5'-CGGGATCCTCTAGGAACAATAGACAAACC-3') were designed within
the ITS regions to confer maximum specificity for P. carinii f. sp. carinii. The ITS region was amplified by nested PCR
on all samples from nonimmunosuppressed rats; these specific primers were also used on 10 of the nonimmunosuppressed wild rats. Samples from
six wild rats (one sample was amplified using both protocols) were
amplified using primers designed for the rRNA genes, ITS30 (5'-TTCCGTAGGTGAACCTGCG-3') and NITSR (48) in the
first round followed by primers ITS21/HI and ITS22/HI in the second
round. Primers ITS30 and NITSR were designed for the highly conserved 18S and 26S RNA genes. The ITS regions of heavily infected laboratory rat samples were amplified using a single-round PCR with primers ITS21/HI and ITS22/HI. The ITS regions of the two immunosuppressed wild-rat DNA samples were amplified using a single-round PCR with primers ITS30 and NITSR. In the first round of nested PCRs, the thermal
cycling conditions were 94°C for 1.5 min, 55°C for 1.5 min, and
72°C for 2 min. In PCRs with a single round and in the second round
of a nested PCR, the thermal cycling conditions were 10 cycles at
94°C for 1.5 min, 55°C for 1.5 min, and 72°C for 2 min, followed
by 30 cycles of 94°C for 1.5 min, 63°C for 1.5 min, and 72°C for
2 min.
The PCR products were cloned into pUC18 (Amersham-Pharmacia Biotech) or
into pGEM T-Easy (Promega), and the recombinant DNA was sequenced using
the Sequenase 2.0 kit (Amersham-Pharmacia Biotech) or the dye
terminator kit (Perkin-Elmer) and the ABI Prism 377 DNA sequencer
running Data Collection Software version 2.1 (Perkin-Elmer Applied
Biosystems). Sequence data analysis was performed using Chromas version
1.44 software (C. McCarthy, Griffith University, Brisbane, Australia).
Sequence alignments were performed using the Wisconsin Package version
10 (Genetics Computer Group, Madison, Wis.). Statistical analyses were
performed using SPSS 7.0.
Nucleotide sequence accession numbers.
The GenBank accession
numbers of the DNA sequence of a portion of the gene encoding the mt
LSU rRNA are as follows: P. carinii f. sp.
carinii, U20169; P. carinii f. sp.
ratti, U20173; Pneumocystis carinii f. sp.
rattus-secundi, AF308807; Pneumocystis carinii f.
sp. rattus-tertii, AF308808; and Pneumocystis
carinii f. sp. rattus-quarti, AF308809.
| |
RESULTS |
P. carinii detected in lungs of nonimmunosuppressed
Danish wild rats.
PCR was used to search for P. carinii
DNA in samples extracted from the lungs of 51 wild rats, live trapped
at various locations in Denmark. After a single round of PCR using the
primers pAZ102-H and pAZ102-E, 11 of the 51 samples gave a P. carinii-specific PCR product indicative of the presence of
P. carinii (Table 1). The
first-round PCR products of the 51 samples were used as templates for
three different nested PCRs. Primer pair RC1 and RC2 specifically amplifies P. carinii f. sp. carinii DNA
(31), and nested PCR with these primers showed that 36 of
the 51 samples contained P. carinii f. sp.
carinii DNA. Similarly, the primer pair RR1 and RR2
specifically amplifies P. carinii f. sp. ratti
DNA (31), and this was found to be present in 32 of the 51 samples (Table 1). The primer pair pAZ102-X/RI and pAZ102-Z/RI
amplifies P. carinii DNA from all host species tested to
date (A. E. Wakefield, unpublished results). Using nested PCR with
primers pAZ102-X/RI and pAZ102-Z/RI, P. carinii DNA could be
detected in 47 of the 51 rats. No P. carinii DNA was found
in 4 of the 51 samples (Table 1).
As a control, a group of 12 nonimmunosuppressed laboratory rats were
examined in the same way. The animals were sacrificed immediately on
arrival at the animal facility and before they came into contact with
any immunosuppressed rats. Single-round PCR was carried out with primer
pair pAZ102-H and pAZ102-E, and 2 of the 12 samples were positive for
P. carinii. Using nested PCR with the P. carinii
special form-specific primers, P. carinii f. sp.
carinii DNA alone was detected in 9 of the 12 rats, both P. carinii f. sp. carinii and P. carinii f. sp. ratti DNAs were detected in 1 of the 12, and no P. carinii DNA was found in 2 of the 12 samples
(Table 2).
Evidence for five different formae speciales of P. carinii in wild rats.
A number of the nested PCRs with
primer pair pAZ102-X/RI and pAZ102-Z/RI gave products of unexpected
size, which were cloned and sequenced (Table
3). Five distinct sequence types were
identified (Fig. 1). One of the sequences
corresponded to P. carinii f. sp. carinii, and
another corresponded to P. carinii f. sp. ratti. The three other sequences, which were isolated from six rats and sequenced several times in more than one PCR experiment, have not been
previously reported.
View larger version (59K):
[in this window]
[in a new window]
|
FIG. 1.
Alignment of the DNA sequence of a portion of the
P. carinii gene encoding the mt LSU rRNA isolated from wild
rats showing the sequences from P. carinii f. sp.
carinii (pc-carinii) and P. carinii f. sp.
ratti (pc-ratti) and the new DNA sequences from P. carinii f. sp. rattus-secundi (pc-rat-sec), P. carinii f. sp. rattus-tertii (pc-rat-ter), and P. carinii f. sp. rattus-quarti (pc-rat-qua). Nucleotides
are boxed to highlight regions of sequence identity. Gaps (-) were
introduced to improve the alignment.
|
|
The three novel sequences were aligned with sequences from all other
known P. carinii formae speciales and with other fungi. Each
of the three new types had a higher level of sequence identity to the
corresponding sequence from P. carinii formae speciales than
to those from the other fungi (data not shown). We propose that these
sequences were amplified from three new formae speciales, which we have
provisionally named P. carinii f. sp.
rattus-secundi, P. carinii f. sp.
rattus-tertii, and P. carinii f. sp.
rattus-quarti. These names are in keeping with the
previously published guidelines and were devised in consultation with
the Pneumocystis Taxonomy and Nomenclature Committee
(34, 44). The P. carinii f. sp. rattus-secundi sequence was found to have the highest
identity to P. carinii f. sp. carinii and
P. carinii f. sp. ratti sequences, but it
contained a 100-bp insertion compared with the sequences from known
rat-derived P. carinii special forms. The P. carinii f. sp. rattus-tertii sequence had the highest
identity to the P. carinii f. sp. rattus-quarti
sequence. The P. carinii f. sp. rattus-quarti
sequence contained a 56-bp insertion at the same position as the
P. carinii f. sp. rattus-secundi sequence (Fig. 1). Some minor variations were noted in the repetitive regions of the
sequences; these could have been due to PCR-induced error and the
instability of such repeats in Escherichia coli plasmids (41). At positions 116 to 117 in the P. carinii
f. sp. rattus-secundi sequence, TG, GA, and a 2-bp deletion
were also observed, and at positions 82 to 83 in P. carinii
f. sp. rattus-tertii, TG and a 2-bp deletion were seen. From
positions 91 to 120 in P. carinii f. sp.
rattus-quarti, a variable number (from 10 to 15) of AT repeats were observed (15 AT repeats are shown in Fig. 1).
Distribution of P. carinii in nonimmunosuppressed wild
rats.
The amount of rat-derived P. carinii DNA detected
was stratified into three levels: (i) presence of P. carinii-specific amplification product, visualized on an agarose
gel after single-round PCR; (ii) presence of P. carinii-specific amplification product, visualized on an agarose
gel after nested PCR; (iii) undetectable P. carinii-specific amplification product. When the P. carinii-specific
amplification product could be visualized on an agarose gel after a
single-round PCR, it was taken to indicate the presence of more
P. carinii than if it could only be detected using nested
PCR. It was found that all female rats harbored detectable levels of
P. carinii DNA, and in 9 of 30, P. carinii DNA
could be detected by single-round PCR. In contrast, 4 of 21 male rats
contained no detectable P. carinii DNA, and in only 2 of 21 was P. carinii DNA found at the higher level. This
distribution suggests a relationship between the amount of P. carinii DNA detected and the sex of the rat (G test, P = 0.011; Williams' correction). There were no associations between the types of P. carinii DNA found and the site of
capture, the weight of the rat at capture, or the month of capture.
P. carinii f. sp. rattus-secundi was detected in
three samples, P. carinii f. sp. rattus-tertii
was found in one sample, and P. carinii f. sp.
rattus-quarti was found in two samples (Table 3).
Considering all five types of rat-derived P. carinii, there was evidence for more than one type of P. carinii in 26 of
the 51 rats (Table 1). After capture, 8 of 51 rats were killed
immediately, and the remainder were housed in the animal facility for
up to 78 days before sacrifice. We examined the proportions of the five different formae speciales as a function of the time the animals spent
in captivity after capture. P. carinii f. sp.
rattus-secundi, P. carinii f. sp.
rattus-tertii, and P. carinii f. sp.
rattus-quarti were detected in 6 of 20 rats housed in
captivity for less than 10 days but not in any of the 31 rats housed
for 11 days or more. A significant relationship was detected between
the presence of the three new formae speciales sequences and the amount
of time between capture and sacrifice (P = 0.0027;
2 test; Yates' correction). In contrast, the
proportion of rats in which P. carinii f. sp.
carinii was detected increased from 11 of 20 (55%) in
captivity for less than 10 days to 25 of 31 (81%) in captivity for 11 days or more (P = 0.05; 2 test).
P. carinii in immunosuppressed wild rats.
In order
to obtain larger amounts of P. carinii DNA, six wild rats
were live trapped and immunosuppressed at an animal facility different
from that used in the study of the nonimmunosuppressed wild rats,
whereupon they developed symptoms of P. carinii pneumonia. DNA was extracted and tested using PCR in the same way as for the
nonimmunosuppressed wild rats. After a single round of PCR, four of the
six rats had detectable P. carinii DNA. When nested PCR was
performed, all six rats were positive for both P. carinii f.
sp. carinii and P. carinii f. sp.
ratti DNA (Table 4). None of
the samples showed evidence of the presence of P. carinii f. sp. rattus-secundi, P. carinii f. sp.
rattus-tertii, or P. carinii f. sp.
rattus-quarti DNA.
Low level of diversity in the ITS regions from P. carinii f. sp. carinii.
In order to further examine
diversity within P. carinii f. sp. carinii, we
studied DNA polymorphisms in the ITS regions of the nuclear rRNA
operon. DNA was isolated from a total of 32 samples: (i) the lungs of
10 immunosuppressed laboratory rats spanning a 10-year period, (ii) 15 nonimmunosuppressed wild rats collected throughout Denmark over a
period of 6 months, (iii) 2 wild rats trapped on the Danish island of
Bornholm and then immunosuppressed, and (iv) 5 environmental samples
collected using spore traps at a rural location in the United Kingdom.
Single-round PCR was used to amplify the ITS regions from the
immunosuppressed rats, whereas nested PCR was required to amplify the
same regions from the environmental samples and the nonimmunosuppressed
wild rats. The ITS PCR products were cloned, and the DNA sequences were
compared. In all, 13 clones from 10 immunosuppressed laboratory rats,
35 clones from 15 nonimmunosuppressed wild rats, 10 clones from 2 immunosuppressed wild rats, and 12 clones from five spore trap samples
were analyzed. A very low level of heterogeneity was detected. A number
of single-base polymorphisms were found, but most of these were seen in
only 1 of the 70 clones examined. There were 10 positions within the
sequences of ITS1 and ITS2 at which a single-base polymorphism was
observed in two different samples, and one of these polymorphisms in
the ITS1 region was seen in three samples, an immunosuppressed
laboratory rat, a nonimmunosuppressed wild rat, and an environmental sample.
| |
DISCUSSION |
High incidence of P. carinii infection in rats.
We
found P. carinii DNA in 92% of wild rats, which is higher
than most previously reported estimates of prevalence. Using a single
round of PCR, the incidence of P. carinii was very similar to that found in a previous Danish survey which used the same collection methods (35) and to the incidence of P. carinii in the field vole, Microtus agrestis, and the
common shrew, Sorex araneus, in Finland (16). The
nested-PCR assay detected low numbers of organisms which would not be
seen by histochemical staining or immunocytochemistry (23, 28, 33,
46, 50) and which would not cause pathological changes in the
lung (26, 60). Our data show an association between the
amount of P. carinii DNA detected by PCR and the sex of the
rat, with female rats carrying more P. carinii DNA than
males. This is in agreement with data from laboratory rats in which
corticosteroid-treated female rats acquired severe P. carinii pneumonitis faster than male rats (30).
P. carinii f. sp. carinii and P. carinii f. sp. ratti can coexist in the same laboratory
rat and may compete for resources (6). We found both formae
speciales in nonimmunosuppressed wild rats, in solitary infections and
also in coinfections with other rat-derived P. carinii
types. However, this study cannot conclude that either forma specialis
can exist alone in wild rats. Multiple PCR tests on each sample would
be needed to totally exclude the presence of other types. With over
half of the wild rats carrying at least two formae speciales of
P. carinii, we suggest that infection in the wild rat is a
complex interaction of several distinct types of organisms. Our
findings are consistent with those of another study of the interaction
of populations of P. carinii f. sp. carinii and
P. carinii f. sp. ratti, which suggested that
these two formae speciales may be competing for resources within the
lung (13). It contrasts with human infection, where,
although large numbers of samples have been analyzed, only one special
form, P. carinii f. sp. hominis, has been found
(10, 14, 17, 18, 19, 20, 47, 48, 49).
Our data on prevalence suggest that low-level carriage of P. carinii in wild rats and nonimmunosuppressed laboratory rats is
common. Immunocompetent rats have been shown to be able to clear all
P. carinii organisms if housed under isolator conditions (50), and a model of continual de novo infection via an
airborne route and slow elimination is suggested rather than stable
latency (2, 11, 18, 52, 53). This may also be the case in
human infection, where low levels of P. carinii f. sp.
hominis have recently been reported in some patients who are
not profoundly immunosuppressed, suggestive of short-term, low-level
carriage of P. carinii f. sp. hominis in these
patient groups. These include individuals with other pulmonary disease,
such as chronic lung disease, and also patients with malignant disease
(3, 4, 27, 37).
Evidence for three new formae speciales of P. carinii.
We present data in support of three novel formae speciales of
rat-derived P. carinii. We consider that they are likely to be as-yet-undescribed P. carinii types based on DNA sequence
homology to known P. carinii and fungal mt LSU rRNA
sequences. The majority of other different Pneumocystis
formae speciales were first identified by analysis of the mt LSU rRNA
sequence, and divergence at this sequence has provided a useful
indicator for assigning new formae speciales (8, 32, 36,
56). We consider it highly unlikely that the three new sequences
could have been generated purely by a PCR artifact or the cloning
procedure; none of the sequences is a mosaic of known types, and all
have portions of the sequence which are unique to that sequence. It is
unlikely that the new sequences resulted from some form of
"contamination," since it is difficult to postulate a source of
such contamination. These sequences had never previously been found in
our laboratory; indeed, they have not been described prior to this
study, yet they appear to be of Pneumocystis origin, since
they form a monophyletic group with the other P. carinii
special forms. All of the new sequences were isolated from more than
one PCR, and both P. carinii f. sp. rattus-secundi and P. carinii f. sp.
rattus-quarti sequences were amplified independently from
more than one rat sample. The sequences were isolated using primers
which are capable of amplifying many formae speciales of P. carinii. However, we did not find any evidence for the presence of
non-rat-derived P. carinii in the samples, although they are
present in samples of air spora (52, 53). We therefore
consider it likely that these novel sequences were amplified from
viable P. carinii organisms within the lungs of the feral
rats. We suggest that the sequences detected were amplified from
previously unknown formae speciales of rat-derived P. carinii. While we suggest that these new types make up a part of
the lung fauna of wild rats, there are significant practical problems
in obtaining enough material for further studies. Immunosuppression of
a number of wild rats did not result in the isolation of the new formae speciales.
Low level of diversity in the ITS regions of P. carinii
f. sp. carinii.
Our data on the analysis of the ITS1 and
ITS2 sequences of P. carinii f. sp. carinii
suggested that the level of variation was very low. The lack of
diversity in the ITS regions of P. carinii f. sp.
carinii contrasts strikingly with the extensive diversity found in the ITS regions of P. carinii f. sp.
hominis. To date, we have found 34 types from 55 episodes of
P. carinii pneumonia (A. G. Tsolaki, R. F. Miller,
and A. E. Wakefield, unpublished observations), and Lee et al.
have reported 59 combinations of ITS1 and ITS2 types from 207 clinical
samples (19). The techniques used in the present study were
capable of detecting such heterogeneity if the P. carinii f.
sp. carinii locus had a level of variation similar to that
of P. carinii f. sp. hominis. All our samples came from northern Europe, however, so we cannot exclude the
possibility that other genotypes of P. carinii f. sp.
carinii could be found elsewhere. The primers that were used
to amplify the ITS regions from the spore trap samples and from 18 of
the 45 wild-rat samples were designed for the sequence of the ITS
regions of P. carinii f. sp. carinii. Since the
ITS regions of the special forms of P. carinii are so
divergent, it is conceivable that the primers selectively amplified
sequences identical to the prototype sequence. However, the results
gained from using the primers designed for the sequence of the
conserved rRNA genes were similar. The ITS regions of P. carinii f. sp. carinii are unlikely to form the basis
of a useful typing system like that used in P. carinii f. sp. hominis.
There are a number of explanations to account for the low level of
diversity seen in the ITS regions. One possibility is that the
population of P. carinii f. sp. carinii has
passed through a recent genetic bottleneck; alternatively, the ITS
regions of P. carinii f. sp. carinii may be under
high selective pressure. The genetic-bottleneck hypothesis is
consistent with the natural history of the laboratory animals, since
many rat colonies were derived from the same small populations of
laboratory animals established relatively recently. The lack of
diversity seen in the wild-rat P. carinii population may be
explained by the recent and rapid expansion of the brown rat, R. norvegicus, through Europe in the 18th century from central or
east Asia, displacing the black rat, Rattus rattus, the
carrier of plague (61). P. carinii shows strict
host specificity, and so one would expect the natural history of the
parasite to be greatly influenced by that of the host. This study of
P. carinii infection in wild rats may lead to a better
understanding of the links between the population structures of other
formae speciales of P. carinii and their hosts.
In this study we have shown that P. carinii is frequently
found in the lungs of nonimmunosuppressed rats. Our data support the
notion of short-term carriage of P. carinii f. sp.
hominis in patient groups other than the severely
immunocompromised. However, our results also highlight the differences
between P. carinii infections in rats and humans and
underline the caution needed when using the rat model of infection for
investigating the human disease.
This research was supported by the Medical Research Council
(R.J.P.) and the Royal Society (A.E.W.) and formed a part of the European Concerted Action Biomed 1 "Pneumocystis and
pneumocystosis: impact of the biodiversity of Pneumocystis
carinii on epidemiology, pathology, diagnosis, monitoring and
prevention of pneumocystosis
new therapeutic approaches" PL941118.
| 1.
|
Aliouat, E. M.,
E. Dei Cas,
L. Dujardin,
J. P. Tissier,
P. Billaut, and D. Camus.
1996.
High infectivity of Pneumocystis carinii cultivated on L2 rat alveolar epithelial cells.
J. Eukaryot. Microbiol.
43:22S[Medline].
|
| 2.
|
Bartlett, M. S.,
J. J. Lu,
C. H. Lee,
P. J. Durant,
S. F. Queener, and J. W. Smith.
1996.
Types of Pneumocystis carinii detected in air samples.
J. Eukaryot. Microbiol.
43:44S[Medline].
|
| 3.
|
Calderon, E. J.,
C. Regordan,
F. J. Medrano,
M. Ollero, and J. M. Varela.
1996.
Pneumocystis carinii infection in patients with chronic bronchial disease.
Lancet
347:977[Medline].
|
| 4.
|
Contini, C.,
M. P. Villa,
R. Romani,
R. Merolla,
S. Delia, and R. Ronchetti.
1998.
Detection of Pneumocystis carinii among children with chronic respiratory disorders in the absence of HIV infection and immunodeficiency.
J. Med. Microbiol.
47:329-333[Abstract].
|
| 5.
|
Creusy, C.,
J. Bahon le Capon,
L. Fleurisse,
C. Mullet,
M. Dridba,
J.-C. Cailliez,
M. Antoine,
D. Camus, and E. Dei Cas.
1996.
Pneumocystis carinii pneumonia in four mammal species: histopathology and ultrastructure.
J. Eukaryot. Microbiol.
43:47S-48S[Medline].
|
| 6.
|
Cushion, M. T.
1998.
Genetic heterogeneity of rat-derived Pneumocystis.
FEMS Immunol. Med. Microbiol.
22:51-58[CrossRef][Medline].
|
| 7.
|
Cushion, M. T.,
J. Zhang,
M. Kaselis,
D. Giuntoli,
S. L. Stringer, and J. R. Stringer.
1993.
Evidence for two genetic variants of Pneumocystis carinii coinfecting laboratory rats.
J. Clin. Microbiol.
31:1217-1223[Abstract/Free Full Text].
|
| 8.
|
Durand-Joly, I.,
A. E. Wakefield,
R. J. Palmer,
C. M. Denis,
C. Creusy,
L. Fleurisse,
I. Ricard,
J. P. Gut, and E. Dei-Cas.
2000.
Ultrastructural and molecular characterization of Pneumocystis carinii isolated from a rhesus monkey (Macaca mulatta).
Med. Mycol.
38:61-72[Medline].
|
| 9.
|
Gigliotti, F.
1992.
Host species-specific antigenic variation of a mannosylated surface glycoprotein of Pneumocystis carinii.
J. Infect. Dis.
165:329-336[Medline].
|
| 10.
|
Helweg-Larsen, J.,
A. G. Tsolaki,
R. F. Miller,
B. Lundgren, and A. E. Wakefield.
1999.
Clusters of Pneumocystis carinii pneumonia: analysis of person-to-person transmission by genotyping.
Q. J. Med.
91:813-820[Abstract/Free Full Text].
|
| 11.
|
Hughes, W. T.
1982.
Natural mode of acquisition for de novo infection with Pneumocystis carinii.
J. Infect. Dis.
145:842-848[Medline].
|
| 12.
|
Hunter, J. A., and A. E. Wakefield.
1996.
Genetic divergence at the mitochondrial small subunit ribosomal RNA gene among isolates of Pneumocystis carinii from five mammalian host species.
J. Eukaryot. Microbiol.
43:24S-25S[Medline].
|
| 13.
|
Icenhour, C. R.,
J. Arnold, and M. T. Cushion.
1999.
Interactions of two Pneumocystis carinii populations within rat lungs.
J. Eukaryot. Microbiol.
46:107S-108S[Medline].
|
| 14.
|
Keely, S.,
H. J. Pai,
R. Baughman,
C. Sidman,
S. M. Sunkin,
J. R. Stringer, and S. L. Stringer.
1994.
Pneumocystis species inferred from analysis of multiple genes.
J. Eukaryot. Microbiol.
41:94S[Medline].
|
| 15.
|
Keely, S. P.,
J. R. Stringer,
R. P. Baughman,
M. J. Linke,
P. D. Walzer, and A. G. Smulian.
1995.
Genetic variation among Pneumocystis carinii hominis isolates in recurrent pneumocystosis.
J. Infect. Dis.
172:595-598[Medline].
|
| 16.
|
Laakkonen, J.,
H. Henttonen,
J. Niemimaa, and T. Soveri.
1999.
Seasonal dynamics of Pneumocystis carinii in the field vole, Microtus agrestis, and in the common shrew, Sorex araneus, in Finland.
Parasitology
118:1-5.
|
| 17.
|
Latouche, S.,
E. Ortona,
E. Mazars,
P. Margutti,
E. Tamburrini,
A. Siracusano,
K. Guyot,
M. Nigou, and P. Roux.
1997.
Biodiversity of Pneumocystis carinii hominis: typing with different DNA regions.
J. Clin. Microbiol.
35:383-387[Abstract].
|
| 18.
|
Latouche, S.,
J. L. Poirot,
V. Bertrand, and P. Roux.
1997.
Pneumocystis carinii hominis sequencing for reactivation or de novo contamination and for hypothetic transmission from person to person.
APMIS Suppl.
77:11-13[Medline].
|
| 19.
|
Lee, C. H.,
J. Helweg-Larsen,
X. Tang,
S. Jin,
B. Li,
M. S. Bartlett,
J. J. Lu,
B. Lundgren,
J. D. Lundgren,
M. Olsson,
S. B. Lucas,
P. Roux,
A. Cargnel,
C. Atzori,
O. Matos, and J. W. Smith.
1998.
Update on Pneumocystis carinii f. sp. hominis typing based on nucleotide sequence variations in internal transcribed spacer regions of rRNA genes.
J. Clin. Microbiol.
36:734-741[Abstract/Free Full Text].
|
| 20.
|
Lee, C. H.,
J. J. Lu,
M. S. Bartlett,
M. M. Durkin,
T. H. Liu,
J. Wang,
B. Jiang, and J. W. Smith.
1993.
Nucleotide sequence variation in Pneumocystis carinii strains that infect humans.
J. Clin. Microbiol.
31:754-757[Abstract/Free Full Text].
|
| 21.
|
Liu, Y.,
M. Rocourt,
S. Pan,
C. Liu, and M. J. Leibowitz.
1992.
Sequence and variability of the 5.8S and 26S rRNA genes of Pneumocystis carinii.
Nucleic Acids Res.
20:3763-3772[Abstract/Free Full Text].
|
| 22.
|
Lu, J. J.,
M. S. Bartlett,
M. M. Shaw,
S. F. Queener,
J. W. Smith,
M. Ortiz Rivera,
M. J. Leibowitz, and C. H. Lee.
1994.
Typing of Pneumocystis carinii strains that infect humans based on nucleotide sequence variations of internal transcribed spacers of rRNA genes.
J. Clin. Microbiol.
32:2904-2912[Abstract/Free Full Text].
|
| 23.
|
Lu, J. J.,
C. H. Chen,
M. S. Bartlett,
J. W. Smith, and C. H. Lee.
1995.
Comparison of six different PCR methods for detection of Pneumocystis carinii.
J. Clin. Microbiol.
33:2785-2788[Abstract].
|
| 24.
|
Meade, J. C., and J. R. Stringer.
1995.
Cloning and characterization of an ATPase gene from Pneumocystis carinii which closely resembles fungal H+ ATPases.
J. Eukaryot. Microbiol.
42:298-307[Medline].
|
| 25.
|
Merali, S.,
U. Frevert,
J. H. Williams,
K. Chin,
R. Bryan, and A. B. Clarkson.
1999.
Continuous axenic cultivation of Pneumocystis carinii.
Proc. Natl. Acad. Sci. USA
96:2402-2407[Abstract/Free Full Text].
|
| 26.
|
Millard, P. R.,
A. E. Wakefield, and J. M. Hopkin.
1990.
A sequential ultrastructural study of rat lungs infected with Pneumocystis carinii to investigate the appearances of the organism, its relationships and its effects on pneumocytes.
Int. J. Exp. Pathol.
71:895-904[Medline].
|
| 27.
|
Nevez, G.,
C. Raccurt,
V. Jounieaux,
E. Dei-Cas, and E. Mazars.
1999.
Pneumocystosis versus pulmonary Pneumocystis carinii colonization in HIV-negative and HIV-positive patients.
AIDS
13:535-536[CrossRef][Medline].
|
| 28.
|
O'Leary, T. J.,
M. M. Tsai,
C. F. Wright, and M. T. Cushion.
1995.
Use of semiquantitative PCR to assess onset and treatment of Pneumocystis carinii infection in rat model.
J. Clin. Microbiol.
33:718-724[Abstract].
|
| 29.
|
Ortiz-Rivera, M.,
Y. Liu,
R. Felder, and M. J. Leibowitz.
1995.
Comparison of coding and spacer region sequences of chromosomal rRNA-coding genes of two sequevars of Pneumocystis carinii.
J. Eukaryot. Microbiol.
42:44-49[Medline].
|
| 30.
|
Oz, H. S., and W. T. Hughes.
1996.
Effect of sex and dexamethasone dose on the experimental host for Pneumocystis carinii.
Lab. Anim. Sci.
46:109-110[Medline].
|
| 31.
|
Palmer, R. J.,
M. T. Cushion, and A. E. Wakefield.
1999.
Discrimination of rat-derived Pneumocystis carinii f. sp. carinii and Pneumocystis carinii f. sp. ratti using the polymerase chain reaction.
Mol. Cell. Probes
13:147-155[CrossRef][Medline].
|
| 32.
|
Peters, S. E.,
K. English,
J. Laakkonen, and J. Gurnell.
1994.
DNA analysis of Pneumocystis carinii infecting Finnish and English shrews.
J. Eukaryot. Microbiol.
41:108S[Medline].
|
| 33.
|
Peters, S. E.,
A. E. Wakefield,
S. Banerji, and J. M. Hopkin.
1992.
Quantification of the detection of Pneumocystis carinii by DNA amplification.
Mol. Cell. Probes
6:115-117[CrossRef][Medline].
|
| 34.
|
Pneumocystis Workshop.
1994.
Revised nomenclature for Pneumocystis carinii.
J. Eukaryot. Microbiol.
41:121S-122S[Medline].
|
| 35.
|
Settnes, O. P., and J. Lodal.
1980.
Prevalence of Pneumocystis carinii Delanoë & Delanoë, 1912 in rodents in Denmark.
Nord. Vet. Med.
32:17-27[Medline].
|
| 36.
|
Sinclair, K.,
A. E. Wakefield,
S. Banerji, and J. M. Hopkin.
1991.
Pneumocystis carinii organisms derived from rat and human hosts are genetically distinct.
Mol. Biochem. Parasitol.
45:183-184[CrossRef][Medline].
|
| 37.
|
Sing, A.,
A. Roggenkamp,
I. B. Autenrieth, and J. Heesemann.
1999.
Pneumocystis carinii carriage in immunocompetent patients with primary pulmonary disorders as detected by single or nested PCR.
J. Clin. Microbiol.
37:3409-3410[Abstract/Free Full Text].
|
| 38.
|
Sloand, E.,
B. Laughon,
M. Armstrong,
M. S. Bartlett,
W. Blumenfeld,
M. Cushion,
A. Kalica,
J. A. Kovacs,
W. Martin,
E. Pitt,
E. L. Pesanti,
F. Richards,
R. Rose, and P. Walzer.
1993.
The challenge of Pneumocystis carinii culture.
J. Eukaryot. Microbiol.
40:188-195[Medline].
|
| 39.
|
Smulian, A. G.,
M. Ryan,
C. Staben, and M. Cushion.
1996.
Signal transduction in Pneumocystis carinii: characterization of the genes (pcg1) encoding the alpha subunit of the G protein (PCG1) of Pneumocystis carinii carinii and Pneumocystis carinii ratti.
Infect. Immun.
64:691-701[Abstract].
|
| 40.
|
Stedman, T. T., and G. A. Buck.
1996.
Identification, characterization, and expression of the BiP endoplasmic reticulum resident chaperonins in Pneumocystis carinii.
Infect. Immun.
64:4463-4471[Abstract].
|
| 41.
|
Strauss, B. S.,
D. Sagher, and S. Acharya.
1997.
Role of proofreading and mismatch repair in maintaining the stability of nucleotide repeats in DNA.
Nucleic Acids Res.
25:806-813[Abstract/Free Full Text].
|
| 42.
|
Stringer, J. R.
1996.
Pneumocystis carinii: what is it, exactly?
Clin. Microbiol. Rev.
9:489-498[Abstract].
|
| 43.
|
Stringer, J. R.,
S. L. Stringer,
J. Zhang,
R. Baughman,
A. G. Smulian, and M. T. Cushion.
1993.
Molecular genetic distinction of Pneumocystis carinii from rats and humans.
J. Eukaryot. Microbiol.
40:733-741[Medline].
|
| 44.
|
Stringer, J. R.,
A. E. Wakefield,
M. T. Cushion, and E. Dei Cas.
1997.
Pneumocystis taxonomy and nomenclature: an update.
J. Eukaryot. Microbiol.
44:5S-6S[Medline].
|
| 45.
|
Sunkin, S. M., and J. R. Stringer.
1995.
Transcription factor genes from rat Pneumocystis carinii.
J. Eukaryot. Microbiol.
42:12-19[Medline].
|
| 46.
|
Tamburrini, E.,
P. Mencarini,
A. De Luca,
G. Maiuro,
G. Ventura,
A. Antinori,
A. Ammassari,
E. Visconti,
L. Ortona,
A. Siracusano,
E. Ortona, and G. Vicari.
1993.
Diagnosis of Pneumocystis carinii pneumonia: specificity and sensitivity of polymerase chain reaction in comparison with immunofluorescence in bronchoalveolar lavage specimens.
J. Med. Microbiol.
38:449-453[Abstract].
|
| 47.
|
Tsolaki, A. G.,
P. Beckers, and A. E. Wakefield.
1998.
Pre-AIDS era isolates of Pneumocystis carinii f. sp. hominis: high genotypic similarity with contemporary isolates.
J. Clin. Microbiol.
36:90-93[Abstract/Free Full Text].
|
| 48.
|
Tsolaki, A. G.,
R. F. Miller,
A. P. Underwood,
S. Banerji, and A. E. Wakefield.
1996.
Genetic diversity at the internal transcribed spacer regions of the rRNA operon among isolates of Pneumocystis carinii from AIDS patients with recurrent pneumonia.
J. Infect. Dis.
174:141-156[Medline].
|
| 49.
|
Tsolaki, A. G.,
R. F. Miller, and A. E. Wakefield.
1999.
Oropharyngeal samples for genotyping and monitoring response to treatment in AIDS patients with Pneumocystis carinii pneumonia.
J. Med. Microbiol.
48:897-905[Abstract].
|
| 50.
|
Vargas, S. L.,
W. T. Hughes,
A. E. Wakefield, and H. S. Oz.
1995.
Limited persistence in and subsequent elimination of Pneumocystis carinii from the lungs after P. carinii pneumonia.
J. Infect. Dis.
172:506-510[Medline].
|
| 51.
|
Vasquez, J.,
A. G. Smulian,
M. J. Linke, and M. T. Cushion.
1996.
Antigenic differences associated with genetically distinct Pneumocystis carinii from rats.
Infect. Immun.
64:290-297[Abstract].
|
| 52.
|
Wakefield, A. E.
1994.
Detection of DNA sequences identical to Pneumocystis carinii in samples of ambient air.
J. Eukaryot. Microbiol.
41:116S[Medline].
|
| 53.
|
Wakefield, A. E.
1996.
DNA sequences identical to Pneumocystis carinii f. sp. carinii and Pneumocystis carinii f. sp. hominis in samples of air spora.
J. Clin. Microbiol.
34:1754-1759[Abstract].
|
| 54.
|
Wakefield, A. E.
1998.
Genetic heterogeneity in Pneumocystis carinii: an introduction.
FEMS Immunol. Med. Microbiol.
22:5-13[CrossRef][Medline].
|
| 55.
|
Wakefield, A. E.,
J. M. Hopkin,
J. Burns,
J. B. Hipkiss,
T. J. Stewart, and E. R. Moxon.
1988.
Cloning of DNA from Pneumocystis carinii.
J. Infect. Dis.
158:859-862[Medline].
|
| 56.
|
Wakefield, A. E.,
S. P. Keely,
J. R. Stringer,
C. B. V. Christensen,
P. Ahrens,
S. E. Peters,
V. Bille-Hansen,
S. A. Henriksen,
S. E. Jorsal, and O. P. Settnes.
1997.
Identification of porcine Pneumocystis carinii as a genetically distinct organism by DNA amplification.
APMIS
105:317-321[Medline].
|
| 57.
|
Wakefield, A. E.,
F. J. Pixley,
S. Banerji,
K. Sinclair,
R. F. Miller,
E. R. Moxon, and J. M. Hopkin.
1990.
Amplification of mitochondrial ribosomal RNA sequences from Pneumocystis carinii DNA of rat and human origin.
Mol. Biochem. Parasitol.
43:69-76[CrossRef][Medline].
|
| 58.
|
Wakefield, A. E.,
F. J. Pixley,
S. Banerji,
K. Sinclair,
R. F. Miller,
E. R. Moxon, and J. M. Hopkin.
1990.
Detection of Pneumocystis carinii with DNA amplification.
Lancet
336:451-453[CrossRef][Medline].
|
| 59.
|
Walzer, P. D.,
J. Foy,
P. Steele,
C. K. Kim,
M. White,
R. S. Klein,
B. A. Otter, and C. Allegra.
1992.
Activities of antifolate, antiviral, and other drugs in an immunosuppressed rat model of Pneumocystis carinii pneumonia.
Antimicrob. Agents Chemother.
36:1935-1942[Abstract/Free Full Text].
|
| 60.
|
Weisbroth, S. H.,
J. Geistfeld,
S. P. Weisbroth,
B. Williams,
S. H. Feldman,
M. J. Linke,
S. Orr, and M. T. Cushion.
1999.
Latent Pneumocystis carinii infection in commercial rat colonies: comparison of inductive immunosuppressants plus histopathology, PCR, and serology as detection methods.
J. Clin. Microbiol.
37:1441-1446[Abstract/Free Full Text].
|
| 61.
|
Zissner, H.
1935.
Rats, lice and history.
George Routledge and Sons, London, United Kingdom.
|
![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
Top
Abstract
Introduction
