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Applied and Environmental Microbiology, January 2000, p. 406-412, Vol. 66, No. 1
0099-2240/0/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Comparison of Animal Infectivity and Nucleic Acid
Staining for Assessment of Cryptosporidium parvum Viability
in Water
Norman F.
Neumann,1
Lyndon L.
Gyürek,1,2
Leslie
Gammie,3
Gordon R.
Finch,2 and
Miodrag
Belosevic1,4,*
Department of Biological
Sciences,1 Department of Civil and
Environmental Engineering,2 and Medical
Microbiology and Immunology,4 University of
Alberta, and EPCOR,3 Edmonton, Alberta
T6G 2E9, Canada
Received 16 June 1999/Accepted 16 September 1999
 |
ABSTRACT |
Cryptosporidium parvum oocysts were stained with the
fluorogenic dyes SYTO-9 and SYTO-59 and sorted by flow cytometry in
order to determine whether the fluorescent staining intensity
correlated with the ability of oocysts to infect neonatal CD-1 mice.
Oocysts that did not fluoresce or that displayed weak fluorescent
intensity when stained with SYTO-9 or SYTO-59 readily established
infections in mice, whereas those oocysts that fluoresced brightly did
not. Although fluorescent staining profiles varied among different batches of oocysts, a relative cutoff in fluorescent staining intensity
that correlated with animal infectivity was observed for all batches.
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TEXT |
Cryptosporidium parvum is
now recognized as a frequent cause of waterborne disease in humans
(1, 10, 16, 17, 20). A primary means of parasite
transmission is via drinking water, through the use of untreated
surface water, contaminated distribution systems, or water treatment
facilities employing only chlorine disinfection protocols. Significant
morbidity and mortality have been associated with outbreaks of this
parasite, particularly in immunocompromised individuals and in children
(10).
An ongoing challenge of detection and disinfection of
Cryptosporidium spp. is the difficulty in determining
whether a parasite is viable. The presence of dead parasites in
finished water is of little concern for disease transmission. Animals
have been used as surrogates for determining the infectious potential
of C. parvum oocysts (13). However, the animal
infectivity method is tedious, difficult, and expensive and is not
readily amenable to normal laboratory analysis in the water industry.
Several methods have been used to estimate the viability of parasites,
including in vitro excystation (1, 6, 7, 25, 26), infection of cell lines (12, 24, 27), parasite morphology by light microscopy, the uptake or exclusion of fluorogenic dyes (4, 5, 8,
9, 23), and animal infectivity (2-5, 15, 18, 19, 21, 22,
29, 30). Other assays that allow determination of viability of
C. parvum oocysts include immunomagnetic capture PCR
(28) and fluorescence in situ hybridization (FISH)
techniques (11, 32, 33). Of all of these methods, only
animal infectivity provides direct information about the ability of the
parasite to cause disease.
In previous studies, we examined the potential use of fluorescent
nucleic acid binding dyes as indicators of C. parvum oocyst viability under different experimental conditions. We reported that the
staining of C. parvum oocysts with the nucleic acid binding dyes SYTO-9 and SYTO-59 correlated with the viability of these organisms, with heat-killed oocyst preparations used as a positive control (4, 5). In the present study, we demonstrate that fluorescence intensity of SYTO-9- and SYTO-59-stained C. parvum oocysts directly correlates with animal infectivity.
Source of C. parvum oocysts.
The strain of
C. parvum used in this study was originally isolated by
Harley Moon (National Animal Disease Center, Ames, Iowa) and is
referred to as the Iowa strain. C. parvum oocysts were isolated from the feces of infected neonatal Holstein calves by methods
described elsewhere (19). Oocysts were used within 90 days
of isolation in all experiments. In our studies, C. parvum oocysts are never exposed to 2.5% potassium dichromate or sodium hypochlorite, a common procedure, in order to minimize oxidative damage
incurred on the oocysts by this treatment.
Determining C. parvum infections in neonatal CD-1
mice.
A neonatal mouse model was used to evaluate infectivity of
C. parvum oocysts (14, 15). Breeding pairs of
outbred CD-1 mice were obtained from Charles River Breeding
Laboratories (St. Constant, Quebec, Canada). The animals were given
food and water ad libitum and were housed in cages with covers fitted
with a 0.22-µm-pore-diameter filter in a specific-pathogen-free (P-2 level) animal facility.
Oocyst doses were prepared from the stock or flow cytometer-sorted
suspensions by serial dilution to obtain the required dose. The actual
dose given to the mice was determined from quadruplicate hemocytometer
counts of the stock suspension. Five-day-old neonatal mice were
inoculated intragastrically with feeding needles containing a known
number of oocysts suspended in 50 µl of deionized water. Two hours
prior to infection, the neonatal mice (5 days old) were taken away from
the mothers to ensure that their stomachs were empty and ready to
receive the intragastric inoculum of C. parvum. In addition,
neonates from multiple litters were pooled and randomly selected for
infection, in order to minimize variability introduced by inherent
resistance or susceptibility of neonatal littermates to infection with
C. parvum. The infectivity of the oocysts was determined 7 days after infection.
Two methods of detection of parasites in exposed mice were employed in
the experiments. The first method was microscopic examination
of mouse
intestinal homogenates, by procedures described elsewhere
(
19). In the second method,
C. parvum
infections were evaluated
by staining mouse intestinal homogenates with
fluorescein-labeled
anti-
C. parvum monoclonal antibody
(Immucell, Portland, Maine)
and by flow cytometry to detect the
presence of fluorescent oocysts
(FACSCalibur; Becton Dickinson).
Briefly, mice were killed by
cervical dislocation, and the lower half
of the small intestine
was removed and placed in 10 ml of deionized
water. The intestines
were homogenized for 45 to 60 s in a Sorvall
Omni-Mixer and washed
by centrifugation at 2,000 ×
g
for 15 min. The pellet was then
washed once with deionized water
containing 0.01% Tween 20 at
2,000 ×
g for 15 min.
The supernatant was discarded, and the cell
pellet was disrupted by
vigorous mixing. Twenty microliters of
the viscous pellet was pipetted
into a 35-µm-pore-diameter sieve
fitted onto a 6-ml flow cytometer
polystyrene tube (Becton Dickinson),
and the sieve was flushed with 450 µl of 1% bovine serum albumin
(BSA; fraction V; Boehringer Mannheim)
in phosphate-buffered saline
(PBS). The strained suspension was
incubated for 15 min at room
temperature in order to block nonspecific
adsorption of monoclonal
antibodies to intestinal contents. Homogenates
were stained with
a 1:2,000 final dilution of fluorescein-labeled
anti-
C. parvum monoclonal antibody (Immucell) at 37°C for
30 min. Detection of
C. parvum oocysts was done with a
FACSCalibur flow cytometer programmed
under the following settings: (i)
forward side scatter photodiode
setting = E00 and amp gain = 4.00; (ii) side scatter photomultiplier
setting = 402, and (iii)
FL1 photomultiplier setting = 470 (Fig.
1). Fifty thousand events were collected
for each sample. A stock
oocyst suspension was used to define a region
based on size (i.e.,
forward light scatter) and internal complexity
(i.e., side scatter)
of
C. parvum oocysts. This defined
region (region 1) was subsequently
used to discriminate potential
oocysts in mouse intestinal homogenates.
An additional criterion (i.e.,
gate) within this region was defined
based on the fluorescent staining
intensity (i.e., FL1) of particles
within this region.
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FIG. 1.
Comparison between SYTO-9 staining and infectivity of
C. parvum oocysts. Oocysts were stained with SYTO-9 and
sorted with the flow cytometer based on their fluorescence intensity
(regions 2 to 8 [R2 to R8, respectively]). Sorted oocysts were used
to infect neonatal CD-1 mice (100 oocysts/mouse). The ability of the
sorted oocysts to establish infections in the neonatal mice is shown in
the table next to the fluorogram. The results are from four independent
sorting trials with four different batches of oocysts done on different
days. The results demonstrate that although staining profiles may vary
from batch to batch, the ability of stained oocysts to cause infections
in neonatal mice is stable.
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Evaluation of nucleic acid staining of C. parvum
oocysts.
Staining of C. parvum oocysts
(107) was done in a total volume of 500 µl of deionized
water containing 10 µM SYTO-9 or 60 µM SYTO-59. All staining
procedures were done with microcentrifuge tubes at 37°C for 30 min
(SYTO-9) or 1 h (SYTO-59). Unbound dye was washed out by
centrifuging the parasite suspension (10,000 × g for
10 min), removing the supernatant, and resuspending the parasite pellet
in 1 ml of deionized water. Parasites were washed twice to remove
unbound dye before oocysts were analyzed by flow cytometry. Analysis of
C. parvum oocysts was done based on three parameters: (i)
forward light scatter = voltage E00, amp gain of 4.00; (ii) side
light scatter = voltage 402, amp gain of 4.00; and (iii)
fluorescent intensity (FL1 for SYTO-9 = voltage of 480, amp gain
of 1.00; or FL3 for SYTO-59 = voltage of 675, Amp gain of 1.00).
To ensure that daily fluctuations in the flow cytometer were
controlled, calibrations on the flow cytometer were done daily
with
Calibrite beads (Becton Dickinson). "Channel targeting" was
used as
a method to ensure that samples obtained on different
days were
analyzed with identical instrument performances. Channel
targeting was
accomplished with fluorescein isothiocyanate (FITC)
or PerCP-labeled
Calibrite beads (Becton Dickinson). Beads were
run on the flow
cytometer and targeted to a mean channel setting
in the appropriate
fluorescence spectrum (e.g., FL1 = 98.93 or
FL3 = 1,267.55).
These mean channel values were used as target
settings for the flow
cytometer. Daily fluctuations in flow cytometer
performance were
corrected by adjusting instrument settings (i.e.,
photodiode and
photomultiplier tube settings), so that similar
target settings were
consistently achieved. Lot-to-lot variation
in Calibrite beads was also
accounted for by comparing channel
target settings between bead lots.
After channel targeting was
done, samples containing oocysts were
subsequently run on the
flow cytometer. Deionized water was used as the
sheath fluid for
experiments involving SYTO-9 or SYTO-59 dye staining.
All samples
were analyzed at a high flow rate through the flow
cytometer (approximately
60 µl/min).
Oocysts stained with SYTO-9 or SYTO-59 displayed heterogeneous staining
patterns, with a spectrum of fluorescent intensities
observed among
individual oocysts in the population (Fig.
1 and
2). The majority of oocysts within a
given preparation remained
unstained or weakly stained (regions 2 and 3 in Fig.
1 and
2);
oocysts were thought to be viable based on our
previous reports
characterizing SYTO dye fluorescent staining
intensities by using
epifluorescence microscopy (
4,
5). The
proportion of unstained
or weakly stained oocysts ranged from 75 to
95%, depending on
the batch of
C. parvum oocysts. Although
fluorescent profiles
were consistent when a single batch of oocysts was
stained (data
not shown), staining profiles varied when different
batches of
oocysts were stained with SYTO dyes (Fig.
1 and
2).
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FIG. 2.
Relationship between SYTO-59 staining and infectivity of
C. parvum oocysts. Oocysts were stained with SYTO-59 and
sorted on the flow cytometer on the basis of their fluorescence
intensity (regions 2 to 8 [R2 to R8, respectively]). Sorted oocysts
were used to infect neonatal CD-1 mice (100 oocysts/mouse). The ability
of the sorted oocysts to establish infections in the neonatal mice is
shown in the table next to the fluorogram. The results are from four
independent sorting trials with four different batches of oocysts done
on different days. The results demonstrate that although staining
profiles may vary from batch to batch, the ability of stained oocysts
to cause infections in neonatal mice is stable.
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Oocysts of various staining intensities were sorted into a cell
concentrator module (Becton Dickinson) containing a 25-mm-diameter
tissue culture insert (pore size, 1 µm; Becton Dickinson). The
tissue
culture insert had been previously incubated with 1% BSA-PBS
for 30 min at room temperature to block adsorption of the parasites
to the
membrane and ensure efficient recovery of the parasites.
Oocysts were
sorted at a low flow rate (~12 µl/min) by using an
exclusion sort
mode. Because sorting accuracy by flow cytometry
approximates 95%, a
reduced infectivity approach was used to identify
a relative cutoff in
fluorescence staining that could classify
a subpopulation of oocysts as
noninfectious or infectious. Theoretically,
a sort of noninfectious
oocysts may contain as many as 5% contaminating
infectious oocysts,
resulting in the occasional mouse becoming
infected. For this reason,
the experimental approach required
that regions in flow cytometry be
defined on the basis of a reduced
infective potential and not absolute
infectivity.
A portion of the stained and flow cytometer-sorted oocysts was used to
inoculate neonatal CD-1 mice for infectivity analysis.
Unstained
oocysts were highly infectious to neonatal CD-1 mice
(region 2, Fig.
1
and
2). As few as 10 weakly stained sorted oocysts
caused infections in
neonatal CD-1 mice (data not shown). Brightly
stained oocysts (region
4, Fig.
1 and
2) were considerably less
infectious than unstained or
weakly stained
oocysts.
Although oocysts that stained brightly with SYTO dyes were not
infectious to neonatal mice, it was possible that binding of
the dyes
to sporozoite DNA (or RNA) impaired the ability of these
parasites to
replicate within host cells. Thus, the relationship
we observed between
infectivity and SYTO dye staining may be an
experimental artifact due
to the mutagenic properties of the dyes.
However, we consistently
observed that highly fluorescent oocysts
were smaller and had greater
internal complexity than unstained
oocysts (Fig.
3). We subsequently sorted unstained
oocysts, based
on their size and internal complexity only, and infected
neonatal
mice. The smaller and more compact oocysts (oocysts that
fluoresce
brightly when stained with SYTO dyes) did not infect neonatal
CD-1 mice (Fig.
3), whereas the larger and less compact oocysts
(oocysts that do not fluoresce when stained with SYTO dyes) readily
established infections in these mice (Fig.
3). These results indicate
that the inability of intensely stained oocysts to infect neonatal
mice
was not a result of SYTO dyes binding to nucleic acids and
impairing
cellular function or parasite replication within host
cells.
Furthermore, it would appear that death of an oocyst is
accompanied by
physiological changes mediating compaction of the
oocyst.
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FIG. 3.
Results showing that the observed relationship between
SYTO-9 staining and infectivity of C. parvum oocysts is not
an indirect result of the binding of SYTO-9 to cellular nucleic acids.
The events in region 9 (R9) shown on the left dot plot, represent
oocysts that fluoresced brightly when stained with SYTO-9 (oocysts
obtained from region 4 [R4], lower fluorescence histogram). The
events in region 10 (R10) represent oocysts that stained weakly with
SYTO-9 (oocysts obtained from region 3 [R3], lower fluorescence
histogram). Regions 9 and 10 were drawn around these foci and
subsequently used as sorting gates for an unstained preparation of
C. parvum oocysts. Oocysts falling into these regions were
sorted, and 100 oocysts were administered to neonatal CD-1 mice. The
infectivity results of these two regions are shown to the right of the
dot plot. FSC-H, forward light scatter (size); SSC-H, side light
scatter (internal complexity).
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These results were consistently observed between different batches of
oocysts (four different batches used in these experiments
[Fig.
1 and
2]). Although flow cytometric profiles of SYTO dye-stained
oocysts
varied from batch to batch, a relative cutoff in fluorescence
intensity
of stained parasites could be used as an objective criterion
for
classifying an oocyst as being infectious or
noninfectious.
Evaluation of nucleic acid staining by using confocal
microscopy.
Confocal microscopy was done with a Molecular Dynamics
2001 confocal microscope (Sunnyvale, Calif.). The instrument settings for all confocal analyses were as follows: ×100 objective lens, excitation wavelength of 488, 565 beamsplitter, 530 DF 30 filter for
FITC detection, photomultiplier gain of 700 V, and pinhole 100 µm in
diameter. The fluorescence intensity was measured with Image Space 3.10 software. The mean fluorescence intensity of oocysts within the sorted
population was determined by overlaying a circle (diameter of 6 µm)
on individual oocysts in confocal images, and their fluorescent
intensity per image pixel was measured. Ten oocysts were randomly
chosen from these images to approximate the mean fluorescent intensity
of oocysts in sorted populations. Confocal microscopy verified the
observation that more intensely stained oocysts were less infectious
than unstained or weakly stained oocysts (Fig.
4). Noninfectious oocysts had bright
internal staining of the sporozoites within the oocysts.
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FIG. 4.
Confocal image of sorted oocysts. SYTO-9-stained
C. parvum oocysts were sorted by flow cytometry, and split
samples were used to infect neonatal CD-1 mice or to visualize
fluorescent staining profiles by confocal microscopy. Panels a to d
represent the different oocyst subpopulations sorted by flow cytometry.
Red circles in panel a indicate where unstained oocysts were located.
The level of staining (relative fluorescence units [RFU]) was
quantified by confocal microscopy. The infective potentials of the
oocysts depicted in the confocal micrographs are listed in the table.
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Application to the water industry.
The direct correlation
between animal infectivity and SYTO dye staining may be useful as a
technique for identifying infectious C. parvum oocysts in
source drinking waters. SYTO-59 may be particularly useful for this
application, since its fluorescence spectrum does not overlap with that
of FITC, and therefore it may be used in conjunction with commercially
available FITC-labeled anti-C. parvum monoclonal antibodies
to detect and determine the viability or infectivity of oocysts in
environmental samples. At present, we are evaluating the staining
characteristics of different strains and genotypes of C. parvum.
Recently, several other methods have been developed to estimate the
viability and/or infectivity of
C. parvum oocysts. In
vitro
excystation is commonly used as an indicator of
C. parvum viability (
1,
7,
23). However, its applicability for
detection
of viable oocysts in natural waters is limited, due to the
abundance
and diversity of microorganisms within natural water samples
and,
conversely, the small numbers of
C. parvum oocysts
found in these
samples. Nevertheless, in vitro excystation is still
used to estimate
the viable numbers of parasites in stock oocyst
preparations and
to determine the efficacy of inactivation of oocysts
after chemical
inactivation (
7,
15,
23,
25). Although in
vitro excystation
is used as a measure of oocyst viability, viability
does not necessarily
equate to infectivity. In fact, in vitro
excystation has been
shown to repeatedly overestimate the infectivity
of
C. parvum oocysts (
7,
15). Moreover, it is
assumed that intact oocysts
that remain after excystation are nonviable
and therefore are
not infectious. Conversely, those oocysts that do
excyst are viable
and therefore capable of establishing infections. We
have recently
demonstrated that intact oocysts isolated by flow
cytometry, after
in vitro excystation, are capable of establishing
infections in
neonatal CD-1 mice (data not
shown).
Because in vitro excystation is primarily used as a measure of
viability and not infectivity, several researchers have used
excystation in conjunction with in vitro infection of immortalized
mammalian cell lines to assess the infectivity of
C. parvum
oocysts.
In these studies, oocysts are excysted and the suspension
containing
infective sporozoites is inoculated into cell culture.
Infection
and replication within cells in vitro can be measured with
different
assays. Slifko et al. (
27) have recently developed
a focus detection
method for determination of the infectivity of
excysted parasites.
In this assay, infected cell cultures are stained
with an anti-
C. parvum sporozoite-merozoite rabbit
polyclonal antibody, and the
number of infectious foci in cell cultures
was determined by using
an indirect fluorescent antibody assay.
Rochelle et al. (
24)
use reverse transcriptase-PCR to detect
the
C. parvum heat shock
protein 70 (hsp70) in infected cell
cultures. Di Giovanni et al.
(
12) used PCR to detect the
hsp70 gene of
C. parvum in infected
cell lines. Although all
of these assays measure the establishment
and infectivity of the
excysted parasites, there are several potential
problems with the in
vitro cell culture infectivity assays. Since
excystation is used as a
method for obtaining infectious sporozoites,
limitations in the
excystation procedure are added to the evaluation
sensitivity of the in
vitro cell culture infectivity assays. Thus,
oocysts that do not
excyst, but are infectious, cannot be detected
by these assays.
Moreover, oocyst preparations require some degree
of activation and
sterilization (i.e., exposure to bleach to prevent
bacterial
contamination of the cell cultures) prior to the addition
of the
parasites to the cell cultures. Arguably, sterilization
procedures may
adversely affect oocyst viability and/or infectivity.
In addition, it
has been demonstrated that various cell lines
display different degrees
of susceptibility to infection with
C. parvum
(
31), and the correlation between in vitro infectivity
and
animal infectivity has yet to be
established.
Dye permeability assays using fluorogenic dyes such as
4'6-diamidino-2-phenylindole (DAPI) and propidium iodide (PI) have
also
been used as indicators of
C. parvum oocyst viability
(
8,
9). Although DAPI-PI staining of
C. parvum
oocysts correlates
with in vitro excystation (
8), a direct
correlation between
animal infectivity and DAPI-PI is yet to be
established.
Recently, a FISH technique has also been developed that shows
considerable promise as an indicator of
C. parvum oocyst
viability
(
11,
32,
33). In these assays, a fluorescent DNA
probe is
targeted to the 18S rRNA of
C. parvum. The basis
for this assay
is the premise that 18S rRNA is usually present in
viable organisms
and is degraded by cellular RNases in dead or dying
cells. A distinct
advantage of the FISH technique over SYTO-9 or
SYTO-59 staining
is the apparent high degree of species specificity:
the ability
to distinguish viable
C. parvum from other
Cryptosporidium species.
However, unlike the SYTO dyes,
existing FISH techniques are limited
to measuring the viability of
C. parvum oocysts and not their
infectivity.
At present, we are evaluating whether SYTO dyes can be used in
conjunction with flow cytometry to determine the levels of
inactivation
of
C. parvum oocysts after exposure to various chemical
disinfectants. Since the precise mechanisms of inactivation of
protozoan cysts or oocysts are not known, it is premature to assume
that nucleic acid dyes may be useful as indicators of oocyst viability
or infectivity following chemical disinfection. Chemical disinfectants
may potentially alter cell wall integrity, mediating changes in
permeability of the dyes into oocysts (i.e., increasing or decreasing
fluorescence intensity). The use of the nucleic acid dyes as indicators
of viability or infectivity for chemically inactivated parasites
will
require more research to determine the effects of each disinfectant
on
oocyst staining
intensity.
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ACKNOWLEDGMENTS |
We thank Cezary Kucharsky and Shannon Lefevbre for technical assistance.
This work was supported by the American Water Works Association
Research Foundation (AWWARF) and by the Natural Sciences and Engineering Council of Canada (NSERC) to M.B. and G.R.F. L.L.G. was supported by an NSERC postdoctoral fellowship, and N.F.N. was
supported by a Province of Alberta doctoral fellowship.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Biological Sciences, CW-405 Biological Sciences Building, University of
Alberta, Edmonton, Alberta T6G 2E9, Canada. Phone: (780) 492-1266. Fax:
(780) 492-9234. E-mail: mike.belosevic{at}ualberta.ca.
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Applied and Environmental Microbiology, January 2000, p. 406-412, Vol. 66, No. 1
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Copyright © 2000, American Society for Microbiology. All rights reserved.
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