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Applied and Environmental Microbiology, November 1998, p. 4439-4445, Vol. 64, No. 11
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Influence of Pretreatment and Experimental
Conditions on Electrophoretic Mobility and Hydrophobicity of
Cryptosporidium parvum Oocysts
Charles F.
Brush,1,*
Michael F.
Walter,1
Lynne J.
Anguish,2,
and
William C.
Ghiorse2
Department of Agricultural and Biological
Engineering1 and
Section of
Microbiology,2 Cornell University, Ithaca,
New York 14853
Received 26 March 1998/Accepted 10 July 1998
 |
ABSTRACT |
Surface properties of Cryptosporidium parvum oocysts
were investigated by using electrophoretic mobility and hydrophobicity measurements. Oocysts purified from calf feces by several sucrose flotation steps and deionized water (DI) washes (DIS method) had an
electrophoretic mobility (neutral surface charge) near 0.0 m2 V
1 s1 over a pH range of 2 to 10. The mean electrophoretic mobility of oocysts stored in DI
containing a mixture of antibiotics had a lower standard deviation (
= 0.36) than that of oocysts stored in DI without antibiotics ( = 0.53); their electrophoretic mobility remained unchanged up to 121 days
after collection. The electrophoretic mobility of oocysts purified on a
cold Percoll-sucrose gradient after the feces was defatted with ethyl
acetate (EAPS method) varied linearly with pH from 0.0 m2 V1 s1 at pH 2.4 to 
3.2 × 108 m2 V1 s1 at
pH 10 ( = 0.52), thus displaying the negative surface charge at
neutral pH observed by other researchers. The hydrophobicity of oocysts
and two types of polystyrene beads was measured as a function of ionic
strength by adhesion to polystyrene. Oocysts were purified by the DIS
method. The ionic strength of the suspending solution was varied from 0 to 95 mmol liter1. Two-week-old oocysts exhibited strong
adhesion (~85%) at ionic strengths of 0 to 10 mmol
liter1 and moderate adhesion (~20%) at ionic strengths
of 20 to 95 mmol liter1. Two-month-old oocysts exhibited
high adhesion (~60 to 80%) at all ionic strengths. These results
show that adhesion properties governed by the electrophoretic mobility
of purified C. parvum oocysts can be altered by the method
of purification and that hydrophobicity can change as oocysts age.
| |
INTRODUCTION |
The protozoan parasite
Cryptosporidium parvum has been responsible for several
recent waterborne disease outbreaks in the United States (26,
31). This gastrointestinal illness is transmitted by an
environmentally durable oocyst (15). C. parvum
oocysts have been identified in significant amounts in surface waters throughout the United States and Canada (27, 34, 38). Public drinking water supplies derived from filtered surface waters were implicated in all U.S. waterborne cryptosporidiosis outbreaks between
1984 and 1993 (10). Filtration is an important barrier in
drinking water purification, because C. parvum oocysts are highly resistant to disinfection with chlorine (25).
Recent studies suggest that the surface properties of C. parvum oocysts may differ from those of bacteria and other
microbes. Results reported by Fogel et al. (16) suggest that
significant numbers of C. parvum oocysts bypassed a
filtration plant that retained smaller coliform bacteria, indicating
that oocysts may not adhere to filter media as readily as other
microbes. In a series of microscope studies, Anguish and Ghiorse
(2) reported that C. parvum oocysts seeded into
soil samples and suspended in deionized water (DI), phosphate-buffered
saline (PBS), or 0.1% sodium pyrophosphate did not closely associate
with inorganic or organic soil particles.
The surface properties of the oocyst wall affect the interactions of
the oocysts with filter media and with environmental chemicals and
surfaces. Changes in the oocyst wall as they age may affect the
adhesion and transport properties of oocysts in natural environments.
Changes in the oocyst wall may also affect oocyst survival. For
example, Robertson et al. (37) noted that when oocysts were
stored in fecal material, the oocyst wall permeability of potentially
viable oocysts decreased over time.
Net surface charge and hydrophobicity are important factors mediating
microbial adhesion to surfaces (21, 46). Understanding the
surface charge and hydrophobicity of C. parvum oocysts will aid the development of optimum filtration media and coagulants to
remove oocysts from drinking water and sewage in treatment plants. Such
basic knowledge will also help clarify the microscale processes
involved in sorption of oocysts onto particle surfaces in natural
waters. The identification of noninfective surrogates with similar
surface properties will also help facilitate development of treatment
strategies and laboratory transport experiments.
Surface charge measurements for C. parvum oocysts have
recently been reported by Ongerth and Pecoraro (33), Drozd
and Schwartzbrod (14), and Rice et al. (36). Each
of these studies utilized different oocyst sources, purification
methods, storage solutions, and suspending media, and the reported
results varied widely. Some chemicals used for oocyst purification in
these studies may damage the oocysts (9) and change oocyst
surface properties, including surface charge. A survey of the
literature reveals few if any studies of electrophoretic mobilities for
C. parvum oocysts in which oocysts were purified and stored
under controlled conditions with concern about the use of
surface-active chemicals.
Microbial adhesion to hydrophobic surfaces such as polystyrene can be
used as a surrogate measurement of microbial adhesion to organic
material in the soil. We developed a method for estimating oocyst
hydrophobicity that relied on microscopic direct counting of suspended
oocyst concentrations after adhesion to a standard polystyrene surface
(40, 45, 46). This method was used to measure oocyst
hydrophobicity as a function of the ionic strength of the suspending
solution. Polystyrene was an ideal substrate for these tests because it
is very hydrophobic (1), and the percentage of particles
adhering to the polystyrene substrate under the mixing action of a
micropipettor provided a reliable qualitative measure of the
particle-surface adhesion energy (43, 48).
The objectives of this study were to estimate the electrophoretic
mobility of C. parvum oocysts and to determine the effects of purification method and presence of antibiotics on the
electrophoretic mobility. We also measured the effects of solution
ionic strength on the hydrophobicity of oocysts and polystyrene beads
and determined how the electrophoretic mobility and hydrophobic
properties of oocysts change as they age.
| |
MATERIALS AND METHODS |
Oocyst purification.
C. parvum oocysts were purified
by two methods (described below), referred to here as the DIS method
and the EAPS method. All oocysts were obtained from fresh neonatal
Holstein calf feces which was stored at 4°C until use. Storage time
varied with treatment from 12 days to 6 months.
DIS method.
Dwight Bowman (Department of Microbiology and
Immunology, Cornell University) utilized a modification of the
continuous-flow flotation method of Vetterling (47) to
purify oocysts from calf feces within 48 h of collection. A sample
of fresh neonatal calf feces (less than 24 h old) was screened
through a U.S. Standard Mesh sieve of mesh size 20, diluted
approximately 1:10 with DI, and mixed with sugar solution (specific
gravity [s.g.] of 1.3) to produce a mixture with an s.g. of
approximately 1.12. This material was then introduced into a
continuous-flow centrifuge (model V; International Equipment
Corporation, Boston, Mass.) with a nonperforated basket and centrifuged
at 800 × g. The outflow was collected, diluted with DI
to an s.g. equal to 1.05 to 1.09, and recentrifuged. The sediment was
diluted to the same s.g. and concentrated again by a third passage
through the centrifuge under the same conditions. The resulting
sediment was further purified by using several sucrose step gradient
centrifugations around an s.g. equal to 1.08; each centrifugation was
performed in an ultracentrifuge (model L8M; Beckman Instruments, Palo
Alto, Calif.) with a swinging bucket rotor for 15 min at 8°C and at
full speed. After the oocyst-containing layers from the step gradients
were collected by aspiration, the oocysts were concentrated and washed three times by dilution in DI with centrifugation at 800 × g. After the last wash, the number of oocysts per milliliter was determined by using a hemacytometer and was adjusted to 1.0 × 107 oocysts ml1 by the addition of DI.
Samples stored with antibiotics received 100 U of penicillin G sodium
ml1, 100 µg of streptomycin sulfate ml1,
and 0.25 µg of amphoteracin B ml1. After purification,
all oocysts were stored at 4°C until use. Electrophoretic mobility
measurements were conducted on DIS-purified oocysts stored with
antibiotics at 4, 7, 31, 94, and 121 days after feces collection.
Electrophoretic mobility measurements were conducted on DIS-purified
oocysts stored without antibiotics at 4 to 7 days after feces
collection. Hydrophobicity measurements were conducted on DIS-purified
oocysts at 12 to 17 days and 60 to 68 days after feces collection.
EAPS method.
C. parvum oocysts were purified from calf
feces by the method described by Despommier et al. (13).
Briefly, this method involved sieving the fecal sample, washing with
DI, fixing with formalin, extracting with ethyl acetate (EA), and
floating on a cold Percoll-sucrose (PS) cushion (s.g., 1.18), followed
by three or four DI washes. EAPS-purified oocysts were suspended in DI
with antibiotics added as described above and stored at 4°C until
use. For one fecal sample, purification was completed 35 days after
collection, and electrophoretic mobility measurements were completed
within 5 days of purification. The collection date for the other batch
was not recorded; electrophoretic mobility measurements were completed
within 3 weeks of purification and perhaps as much as 6 months after
collection. These two samples were used to gauge the general effect of
the EAPS method on oocyst electrophoretic mobility for comparison with
literature values.
Electrophoretic mobility measurements.
The electrophoretic
mobility of C. parvum oocysts was measured with a Lazer Zee
meter model 501 (Pen Kem, Bedford Hills, N.Y.) at 150 V. Oocysts were
suspended at approximately 106 ml1 in 0.010 M
KNO3. This solution is easily prepared, it has a
conductivity low enough that the solution conductivity does not
interfere with the measurement of electrophoretic mobilities, and
K+ and NO3 are less likely than
Na+ and Cl to react with ionogenic compounds
on the oocyst surface (17). The pH was adjusted by addition
of 0.1 or 0.01 N HNO3 or KOH. Prior to measurement of
oocyst electrophoretic mobility, the Lazer Zee meter was calibrated
with a standard charged polymer solution.
The initial electrophoretic mobility measurement was taken at the pH of
the prepared oocyst suspension (generally between 7.0 and 7.4). Each
electrophoretic mobility measurement represents the average for a cloud
of 10 to 20 oocysts visible in the microscopic field of view of the
Lazer Zee meter chamber (18). After each measurement, the
chamber was emptied and the solution pH and temperature were recorded.
The pH was then adjusted downward approximately 1 pH unit by addition
of HNO3 and returned to the electrophoresis chamber for the
next measurement. This was continued until an electrophoretic mobility
measurement was recorded at a pH of below 3.0. The pH was then raised
approximately 1 pH unit at a time by addition of KOH, and measurements
were continued until an electrophoretic mobility measurement was
recorded at a pH of above 8 and up to 10, after which the oocyst
suspension was discarded. Readings were recorded as electrophoretic
mobilities, rather than zeta potentials, according to the
recommendations of van Loosdrecht et al. (46). All
electrophoretic mobility measurements were corrected to 20°C by the
relationship µcorr = µmeas × [1 0.02 × (T 20)], where µcorr = electrophoretic mobility corrected to 20°C and µmeas = electrophoretic mobility measured at T°C (35).
Hydrophobicity measurements.
Adhesion experiments were
performed with C. parvum oocysts and two types of
polystyrene microspheres. C. parvum oocysts were purified
from calf feces by the DIS method described above. Each purified sample
received 100 U of penicillin G sodium ml1, 100 µg of
streptomycin sulfate ml1, and 0.25 µg of amphotericin B
ml1. Experiments were conducted with purified oocysts at
12 to 17 days and 60 to 68 days after excretion. Stock suspensions of
4.5-µm-diameter Polybead uncharged polystyrene microspheres (catalog
no. 17135; Polysciences, Warrentown, Pa.) and 6.0-µm-diameter
Fluoresbrite carboxylated polystyrene microspheres (catalog no. 16392;
Polysciences) were prepared by diluting to approximately
103 microspheres ml1 in DI. All particle
suspensions were stored at 4°C until use.
Particle hydrophobicity as a function of ionic strength was assessed by
adhesion to polystyrene (
40,
45,
46). Polystyrene
microtiter
plates (Falcon 3915; Becton Dickinson Labware, Franklin
Lanes, N.J.)
from a single production batch were used as the standard
surface for
adhesion measurements. Each microtiter plate contained
96 400-µl
wells. Experimental treatments consisted of 20 to 40
different ionic
strengths, ranging from 0 to 95 mmol liter
1, with two
replicates. For each experiment, treatments were randomly
distributed
across a single microtiter
plate.
Particle suspensions of the appropriate ionic strengths were prepared
in the microtiter wells. Appropriate quantities of DI
and PBS
(containing 2.31 µM NaH
2PO
4 · H
2O, 5.93 µM Na
2HPO
4
[anhydrous],
120.00 µM NaCl, and 12.69 µM NaN
3, with
the pH adjusted to 7.0
± 0.1 with HCl) were first placed in the
microtiter wells. A pipettor
was used to add 20 µl of vortexed oocyst
or bead suspension to
each well, yielding a 240-µl suspension at the
desired ionic strength.
The particle concentration in the stock
solution was measured
with a hemacytometer (
7) immediately
prior to filling the wells.
Due to the complex ionic makeup of the PBS
solution, the ionic
strength,
I (moles
liter
1), was calculated from solution electrical
conductivity, EC (decisiemens
meter
1), by using the
relationship
I = 0.0135 × EC (
50).
Microtiter plates were covered and incubated at 4°C. After 24 h,
the contents of each microtiter well were mixed with a pipettor
by
three rapid extractions and reinjections of 10 µl of liquid.
The
pipettor tip was inclined at a 45° angle, with the tip of
the
pipettor in the center of the well and approximately 1 mm
above the
bottom of the well. The suspended-particle concentration
in each well
was then measured with a hemacytometer, with two
to four counts per
microtiter well. Hydrophobicity, measured as
percent adhesion, was
calculated as [(
C0 Ci)/
C0] × 100, where
C0 is the mean number of particles measured for
the control and
Ci is the mean number of
particles measured for a given
treatment.
Due to a high degree of variability in the data set, data were
transformed into Lowess plots by using Minitab (version 10.5
Xtra
Power; Minitab Inc., State College, Pa.). Lowess plots use
a robust
locally weighted regression method to fit a smooth curve
through a
scatterplot, helping to reveal trends in the data (
11,
12).
| |
RESULTS |
Electrophoretic mobility.
Oocysts purified by the DIS method,
with and without antibiotics, exhibited an electrophoretic mobility of
between approximately +108 and 108 m2 V1 s1 and a linear
regression slope of near zero over the entire pH range investigated
(Fig. 1a and 1b). Since the regression
lines for these data did not cross the x axis, there are no
clear isoelectric points for these measurements. For the oocyst sample
stored in antibiotics, there was no difference in electrophoretic
mobility between oocysts tested at 4, 7, 37, 94, and 121 days after
collection. The presence of antibiotics in the DIS-purified oocyst
suspensions did not significantly change the average electrophoretic
mobility but did reduce the variation in electrophoretic mobility
within the sample. This was reflected in a 31% drop in the standard
deviation (Table 1). Again these data
show no clear isoelectric point. Oocysts purified by the EAPS method
exhibited a strongly linear relationship between electrophoretic
mobility and pH, with a distinct isoelectric point at pH 2.37 (Fig.
1c).
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FIG. 1.
Electrophoretic mobility versus pH for DIS-purified
C. parvum oocysts without antibiotics (a), DIS-purified
C. parvum oocysts with antibiotics (b), and EAPS-purified
C. parvum oocysts with antibiotics (c). The squares
represent individual electrophoretic mobility measurements, and the
dashed lines represent the least-squares regression lines through the
data.
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TABLE 1.
Summary of experimental measurements and literature data
on electrophoretic mobility of C. parvum oocysts
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Hydrophobicity.
The hydrophobicity method developed for this
study utilized microtiter plates to measure adhesion to polystyrene.
This method allowed a large number of adhesion tests to be conducted in
a short time. For a given ionic strength, adhesion results had a high
variability. Lowess plots revealed trends in the different oocyst and
microsphere treatments.
Two-week-old and 2-month-old
C. parvum oocysts obtained from
the same calf exhibited markedly different adhesion characteristics
(Fig.
2). Adhesion characteristics of
2-week-old oocysts were
strongly dependent on the ionic strength of the
suspending solution.
Over 80% of the 2-week-old oocysts adhered at an
ionic strength
of 0 mmol liter
1. The percent adhesion
dropped nearly linearly as the ionic strength
rose from 0 to
approximately 20 mmol liter
1 and then remained between 10 and 20% as the ionic strength rose
to 95 mmol liter
1.
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FIG. 2.
Data and Lowess plots for adhesion of 12- to 17-day-old
DIS-purified C. parvum oocysts (squares and solid line) and
>60-day-old DIS-purified C. parvum oocysts (circles and
dashed line) to polystyrene at different ionic strengths.
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|
The adhesion characteristics of 2-month-old
C. parvum
oocysts showed less dependence on the ionic strength of the suspending
solution (Fig.
2). Approximately 70% of 2-month-old oocysts adhered
to
polystyrene at an ionic strength of 0 mmol liter
1. As the
ionic strength increased, the percent adhesion dropped
to approximately
62% at approximately 15 mmol liter
1 and then rose again
to remain between approximately 70 and 80%
at ionic strengths of 30 to
95 mmol liter
1.
The adhesion characteristics of uncharged polystyrene microspheres
showed a moderate dependence on solution ionic strength
(Fig.
3). Approximately 60% of the uncharged
microspheres adhered
to the polystyrene substrate at an ionic strength
of 0 mmol liter
1. As the ionic strength increased to
approximately 40 mmol liter
1, the percent adhesion rose
to approximately 85%. As the ionic
strength increased further,
adhesion decreased nearly linearly
to approximately 75% at an ionic
strength of 95 mmol liter
1.
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FIG. 3.
Adhesion of 6.0-µm-diameter uncharged polystyrene
microspheres (electrophoretic mobility = 0.0 m2 V1 cm2) to polystyrene at different ionic
strengths.
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|
The adhesion characteristics of carboxylated polystyrene microspheres
also showed a moderate dependence on the ionic strength
of the
suspending solution (Fig.
4). Over 70%
of the carboxylated
microspheres adhered to the polystyrene substrate
at an ionic
strength of 0 mmol liter
1. As the ionic
strength increased to 20 mmol liter
1, adhesion decreased
to approximately 60%. As the ionic strength
continued to increase to
95 mmol liter
1, adhesion to polystyrene increased nearly
linearly to approximately
70%.
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FIG. 4.
Adhesion of 4.5-µm-diameter carboxylated polystyrene
microspheres (electrophoretic mobility = 68 m2 V1 cm2) to polystyrene at different ionic
strengths.
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| |
DISCUSSION |
Electrophoretic mobility.
Oocysts purified by the DIS method
exhibited an electrophoretic mobility of very close to zero throughout
the pH range examined. These oocysts are hypothesized to be similar to
oocysts as they occur in natural waters, because they have had little
contact with chemicals that would change their surface properties. The DIS method utilized in this study employed only sucrose and DI in the
purification steps, followed by several DI washes with very large
dilution ratios and storage in DI with and without antibiotics.
The presence of antibiotics had little if any effect on electrophoretic
mobility. However, electrophoretic mobility measurements
for oocysts
stored in the presence of antibiotics had a lower
standard deviation
than measurements for oocysts stored without
antibiotics. While the
exact reasons for this effect are unclear,
they could include buffering
by the antibiotics, binding of the
antibiotic molecules to reactive
groups on the oocyst surface,
or interactions between ionic species in
the antibiotic solution
and ionogenic groups on the oocyst surface.
Further studies under
controlled conditions with simpler buffering
solutions are needed
to clarify this
effect.
Oocysts purified by the EAPS method exhibited an electrophoretic
mobility strongly dependent on pH. These oocysts had come
in contact
with a protein cross-linking agent (formalin), a defatting
agent (ethyl
acetate), and a polyvinylpyrrolidone (PVP)-containing
reagent (Percoll)
in addition to DI and sucrose during purification.
We hypothesize that
the EAPS-purified oocysts had their surface
chemistry, including
charged ionic groups, altered to a greater
degree during purification
than oocysts purified by the DIS method.
Our results clearly show that
oocysts purified by the EAPS method
were significantly different in
their surface charge from oocysts
purified by the DIS
method.
Of the three most active ingredients used in the EAPS method (formalin,
ethyl acetate, and PVP), ethyl acetate, which is commonly
used to
extract fats from fecal suspensions, may have caused the
most
significant changes in the oocyst surface chemistry. Indeed,
the
corrosive action of this solvent on polytetrafluoroethylene
and
polystyrene test tubes necessitates the use of glass test
tubes. Other
studies in our laboratory indicate that the permeability
of the oocyst
wall increased significantly after contact with
ethyl acetate
(
3), indicating that this solvent may have removed
lipids or
other ethyl acetate-soluble compounds and thus modified
the oocyst
surface. Formalin would be expected to cross-link glycoproteins
but not
to have major effects on surface charge. The effects of
PVP on the
surface chemistry of oocysts are not known, but this
reagent can be
used to purify macromolecules such as DNA contaminated
with humic acids
(
5,
51). We recommend that ethyl acetate
not be used to
purify oocysts for use in studies of their surface
properties and that
reagents such as formalin, potassium dichromate,
and Percoll be avoided
or used with
caution.
Previous studies of oocyst surface charge were carried out by Rice et
al. (
36), Drozd and Schwartzbrod (
14), and
Ongerth
and Pecoraro (
33). In general, the results of these
three studies
are difficult to compare with ours, because each study
utilized
different, often not-well-described purification methods, as
well
as different suspending solutions and techniques for surface
charge
measurements. Rice et al. (
36) and Drozd and
Schwartzbrod (
14)
reported zeta potential rather than
electrophoretic mobility,
while Ongerth and Pecoraro (
33)
reported both zeta potential
and electrophoretic mobility. van
Loosdrecht et al. (
46) recommend
reporting results as
electrophoretic mobilities, the measured
variable, rather than
converting to zeta potential, since the
conversion to zeta potential
relies on the unproven assumption
that the oocyst surface is
impermeable to
ions.
Ongerth and Pecoraro (
33) and Drozd and Schwartzbrod
(
14) used DI as their suspending solution. We decided to use
0.01
M KNO
3 rather than DI as the suspending medium for
electrophoretic
mobility measurements for several reasons. Busscher et
al. (
8)
found irreproducible results when they measured
electrophoretic
mobilities in DI but stable and reproducible results on
addition
of small amounts of HNO
3 or NaOH. These workers
reasoned that
this result was probably due to low ionization of surface
groups
in DI. In addition, if the Smoluchowski approximation is to be
used to convert electrophoretic mobility measurements to zeta
potential
(
= 14.1µ, where
= zeta potential and µ = electrophoretic
mobility), then the ionic strength must be adjusted, since the
approximation is valid only for suspending mediums with ionic
strengths
of greater than 0.001 mol liter
1 (
24).
Rice et al. (
36) used filtered natural lake water as their
suspending medium. While surface charge measurements in natural
waters
may provide guidance for specific filtration systems, the
results
obtained with these media may be variable and therefore
difficult to
compare and interpret. For example, the complex ionic
makeup, the
presence of unknown chemical species and buffering
capacity, or the
presence of chemically reactive ionic species
all can affect
electrophoretic mobility measurements. Furthermore,
the chemical makeup
in natural waters may change dramatically
over time, especially in
river waters. Nonetheless, Rice et al.
(
36) do show a zeta
potential for oocysts that is very similar
to our results (Table
1).
Electrophoretic mobilities for oocysts purified by the DIS method in
this study were different from those reported by Ongerth
and Pecoraro
(
33), who reported a regression slope of 0.471
and an
isoelectric point of 3.90 (Table
1). The purification
method used by
Ongerth and Pecoraro (
33) differs from our DIS
method
primarily in that they used Percoll in their purification
protocol.
Also, their washing and flotation steps were carried
out in centrifuge
tubes (
20,
34), whereas we used a continuous-flow
centrifuge, which may have provided purer oocyst suspensions.
We are
currently testing these parameters to determine their effects
on oocyst
electrophoretic
mobility.
Electrophoretic mobilities for the oocysts purified by our EAPS method
were similar to those reported by Drozd and Schwartzbrod
(
14), who also used formalin-treated oocysts. Their oocysts
also exhibited a strong inverse relationship between surface charge
and
pH, with an isoelectric point of 2.18 (Table
1).
Hydrophobicity.
The data for adhesion to polystyrene indicate
that except at extremely low ionic strengths, 2-week-old oocysts are
less strongly attracted to negatively charged surfaces than 2-month-old
oocysts (Fig. 2). For ionic strengths of over 20 mol
liter1, approximately 20% of the 2-week-old oocysts
adhered to the substrate, compared to over 60% of the 2-month-old
oocysts. These results suggest that the hydrophobicity of the oocyst
surface changes as the oocysts age after they are excreted. The reasons
for such changes are not known, but in general they may be attributed
to changes in the physical and chemical properties of the oocyst surface.
As physical objects, like viruses and bacteria (
6,
46),
oocysts can be considered colloidal particles and modeled as
such. The
theory of Derjaguin, Landau, Verwey, and Overbeek (DLVO
theory)
provides a good description of the long-range forces involved
in
temporary particle-surface adhesion (
42) and has been
successfully
used to describe the adhesion properties of many bacteria
and
viruses (see, e.g., references
6,
23,
39,
41, and
45).
According to the DLVO theory, the net interaction energy
(
VT)
between a spherical particle (e.g., an
oocyst or polystyrene microsphere)
and a charged plate (e.g.,
polystyrene) is the sum of the attractive
van der Waals energy
(
VA) and the repulsive electrostatic energy
(
VE) (
23,
41,
42):
|
(1)
|
The value of the long-range energy balance
VT determines whether temporary adhesion occurs
and whether the opportunity for
permanent adhesion exists. Hogg et al.
(
22) and Visser (
48)
describe the van der Waals
and electrostatic energies of interaction
between a plate and spherical
particle as follows:
|
(2)
|
|
(3)
|
where
C1 = 1 +
exp(
h),
C2 = 1
exp(
h),
C3 = 1
exp(
2
h),
A is the system Hamaker coefficient (ergs),
R is the
particle
radius (meters),
h is the distance between the
plate and particle
(meters),
is the dielectric constant of the
suspending medium,
and
P and
S (volts) are the surface charges of the
plate and
sphere, respectively.
(meters
1), referred
to as the double-layer thickness, is a function of
the ionic strength
of the suspending medium (
49).
For
C. parvum oocysts interacting with a flat polystyrene
surface, many of the variables in equations 2 and 3 are fairly easy
to
estimate. Our electrophoretic mobility measurements showed
that
S for both 2-week-old and 2-month-old oocysts
was near 0.0
mV (Table
1). The electrostatic repulsion term (equation
3) then
reduces to
VE =
RPln[1
exp(
2
h)]/4. For
a given plate surface
potential and solution ionic strength,
VE reduces to a function
of the separation
distance
h. For oocysts, the value of
A in
equation
2 is the only remaining
unknown.
Hamaker (
19) devised a continuum approach using the
coefficient
A to approximate the total van der Waals
interaction between
two bodies, where the value of
A depends
on the chemical compositions
of their surfaces. While their complex
chemical makeup makes it
difficult to determine an exact
A
for biological particles, the
value of
A may be approximated
from the composition of the particle
surface (
32).
Unfortunately the lack of knowledge regarding
the surface composition
of
C. parvum oocysts (
4,
30,
44)
precludes
estimation of
A for oocysts at this time. However, something
can be said about how a change in the oocyst surface composition
would
affect oocyst hydrophobicity. For example, the replacement
of lipids in
the surface coat with sugars or glycoproteins would
result in a
significant increase in
A (
32). This rise in
A would
cause a corresponding rise in the attractive energy
VA (equation
2). Since the surface charge (and
thus
VE) remained constant as
the oocysts aged,
we hypothesize that the observed increase in
adhesion over time
resulted from changes in the chemical makeup
of the oocyst surface as
they
aged.
Oocysts stored in feces or in another environment may behave
differently than the oocysts used in this experiment, which underwent
some processing and were stored at 4°C in DI containing antibiotics.
McEldowney and Fletcher (
29) noted that both pH and
temperature
affected bacterial adhesion to polystyrene. Loeb et al.
(
28)
also suggested that the ionic makeup of the suspending
solution
will affect the values of
P and
S. It is unclear how specific
ions (e.g.,
Na
+ and Cl
) in the suspending solution
interact with ionogenic compounds
on the oocyst surface, influencing
the oocyst surface charge.
Future studies should explore the
sensitivity of oocyst hydrophobicity
to changes in temperature and in
the ionic makeup and pH of the
suspending solution and how storage in
fecal suspensions, river
water, and other environments affects oocyst
hydrophobicity.
| |
ACKNOWLEDGMENTS |
This research was supported in part by grants from the New York
City Department of Environmental Conservation and the Edna Bailey
Sussman Fund.
We thank Dwight Bowman for supplying C. parvum oocysts and
Leonard Lion for the use of his Pen Kem Lazer Zee meter. We also thank
Juliet Bryant, Michael Jenkins, Charles McCulloch, Kelvin Minniefield,
and Mark J. Walker for their assistance and Kathleen Buckley, Nancy
Doon, Hilary Grimes, and Sharon Guest-Tagliavento for laboratory assistance.
| |
FOOTNOTES |
*
Corresponding author. Present address: Department of
Geology and Geophysics, University of Hawaii at Manoa, Honolulu, HI
96822. Phone: (808) 956-7865. Fax: (808) 956-5154. E-mail:
cbrush{at}akule.soest.hawaii.edu.
Present address: New York State College of Veterinary Medicine,
Ithaca, NY 14853.
| |
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Abstract
Introduction
