Vol. 53, No. 8APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Aug. 1987, p. 1803-18110099-2240/87/081803-09$02.00/0

Infectivity and Antigenicity Reduction Rates of Human RotavirusStrain Wa in Fresh Waters

OSCAR C. PANCORBO,1* BRIAN G. EVANSHEN,' WILLIAM F. CAMPBELL,' STEPHEN LAMBERT,'SHERILL K. CURTIS,4 AND THOMAS W. WOOLLEY5

Environmental Health Science Program, Department of Food Science and Technology, The University of Georgia,Athens, Georgia 30602,1 and Department of Environmental Health, School of Public and Allied Health,2 Departments ofMicrobiology3 and Biophysics4 and Section of Medical Education,5 Quillen-Dishner College of Medicine, East Tennessee

State University, Johnson City, Tennessee 37614

Received 19 May 1986/Accepted 12 May 1987

The rates of inactivation of human rotavirus type 2 (strain Wa) (HRV-Wa) and poliovirus type 1 (strainCHAT) were compared in polluted waters (creek water and secondary effluent before chlorination) andnonpolluted waters (lake water, groundwater, and chlorinated tap water). Viral infectivity titers were

determined by plaque assays, while HRV-Wa antigenicity also was monitored by an enzyme-linked immuno-sorbent assay. Both viruses persisted longest in lake water and shortest in tap water. The actual inactivationtitnes (i.e., times required for two-logl0 reductions of initial viral titers) for the two viruses were significantlydifferent in all waters except tap water. With the exception of the groundwater and secondary effluent results,the HRV-Wa inactivation times in the fresh waters tested were significantly different. Owing perhaps toaggregation, HRV-Wa appeared less susceptible to the effects of chlorine than previously reported for this virusand for the simian rotavirus SAil. HRV-Wa displayed prolonged survival in lake water and groundwaterexceeding that previously reported for the SAil virus. The HRV-Wa infectivity reduction rate (ki) wassignificantly correlated with the water pH (i.e., as pH increased, ki increased). The water pH may haveinfluenced viral aggregation and thereby HRV-Wa susceptibility to other virucidal factors in the water.Enzyme-linked immunosorbent assay results showed similar inactivation patterns with the most significantreduction in HRV-Wa antigenicity occurring in polluted waters and tap water. In all waters, particularly tapwater, infectivity declined at a faster rate than antigenicity. It is proposed that HRV-Wa can be used as a modelfor future studies of rotaviral persistence in the aquatic environment.

Gastroenteritis is the most prevalent waterborne diseasein industrialized nations (14). Reports of gastroenteritisoutbreaks associated with drinking waters (9, 23) and shell-fish consumption (24) have suggested rotaviruses as theetiologic agents. Evidence of this was obtained during acommunity outbreak of waterborne gastroenteritis whichoccurred in Eagle-Vail, Colo., in March 1981, in whichrotavirus was detected in stools from five of seven personsaffected (17). The human rotavirus (HRV) is considered theprimary cause of acute infantile gastroenteritis (19) anddiarrhea-related irnfantile deaths (7).Because of the association of rotaviruses with waterborne

gastroenteritis, the need for research dealing with the fate ofthese pathogens in the aquatic environment has been recog-nized (14). Working with domestic sewage, Steinmann (31)detected rotaviruses in 25% of the samples examined, whileHejkal et al. (16) found the highest levels of rotavirusesduring winter and spring. Most studies, however, have notdealt with indigenous rotaviruses, but rather have focusedon the use of a model rotavirus, namely, the simian rotavirusSAll (14). For example, the SAl1 virus has been used toestablish (i) rotavirus persistence in fresh waters (18) andestuarine waters (18, 27), (ii) rotavirus association withsediments in estuarine water (27), (iii) rotavirus adsorptionto aluminum hydroxide flocs in tap water and activatedsludge flocs (12), and (iv) rotavirus inactivation by chlorine,chlorine dioxide, and monochloramine (3). SAll also wasemployed to develop a method for concentration of rotavi-

* Corresponding author.

ruses from tap water, treated sewage, and raw sewage (30).These studies have indicated that the SAl1 virus can survivein natural waters for prolonged periods compared with thoseexhibited by poliovirus type 1 (14, 18).Although the SAl1 virus is similar morphologically to and

shares cross-reacting antigens with HRV (36), the persis-tence of HRV infectivity in natural waters has not beenestablished (14). This is attributed to the past inability topropagate HRV in vitro. In 1980, however, Wyatt andco-workers (38) reported the cultivation of an attenuatedHRV (strain Wa) in cultures of fetal rhesus monkey kidneycells and the successful adaptation of a plaque assay for thevirus. Recently, Vaughn et al. (32) observed that dispersedparticles of HRV (strain Wa) and SAl1 viruses were bothinactivated rapidly at low chlorine concentrations (s0.3mg/liter). HRV (strain Wa) was used in the research reportedherein to establish base-line data on HRV persistence innatural freshwater samples from polluted and nonpollutedsources. The rates of rotaviral infectivity and antigenicityreductions in these waters were monitored by a plaque assayand an enzyme-linked immunosorbent assay (ELISA), re-spectively. For comparative purposes, reduction of poliovi-rus type 1 (strain CHAT) infectivity in these fresh watersalso was investigated.

MATERIALS AND METHODSViruses and their propagation. Cell culture-adapted strain

Wa of HRV type 2 (HRV-Wa) and poliovirus type 1 (strainCHAT) were used in this study. The HRV-Wa was kindlysupplied by Richard Wyatt (National Institute of Allergy and

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Infectious Diseases) as a lyophilized suspension of infectedMA-104 cells. The lyophilized sample was rehydrated in 1.0ml of sterile distilled water, pretreated with an equal volumeof 20 ,ug of trypsin (type IX; Sigma Chemical Co., St. Louis,Mo.) per ml, and allowed to incubate at 37°C for 45 min. Astock suspension of the virus was prepared by inoculating 2ml of pretreated HRV-Wa onto confluent MA-104 monolay-ers. The MA-104 cells were kindly provided by MonroeVincent (Department of Pediatrics, Uniform Services Uni-versity of Health Sciences, Bethesda, Md.). The MA-104cells were grown in Eagle minimum essential medium(MEM) in Earle balanced salt solution (EBSS) supplementedwith 10% fetal bovine serum (FBS), 2.0 mM L-glutamine,0.22% NaHCO3, 100 U of penicillin per ml, and 100 p.g ofstreptomycin per ml. Cell culture media were obtained fromGIBCO (Grand Island, N.Y.), and FBS was from FlowLaboratories, Inc. (McLean, Va.). Confluent MA-104monolayers were washed three times with Eagle MEM-EBSS containing 0.5% gelatin (DIFCO Laboratories, De-troit, Mich.), 2.0 mM L-glutamine, 0.22% NaHCO3, 100 U ofpenicillin per ml, and 100 Vig of streptomycin per ml (alsoused as the diluent for HRV-Wa throughout this study)before inoculation of HRV-Wa. After virus adsorption for 1h at 37°C, the monolayers were washed once again and thenmaintained in Eagle MEM-EBSS containing 0.5 p.g of tryp-sin per ml, 2.0 mM L-glutamine, 100 U of penicillin per ml,and 100 jig of streptomycin per ml. The cultures wereincubated at 37°C in a 5% CO2 atmosphere. Virus-infectedcells and fluids were harvested after a generalized cytopathiceffect was observed (48 to 72 h). Harvesting of virus frominfected cells was achieved by subjecting the entire fluidvolume to two freeze-thaw cycles followed by centrifugationat 1,000 x g for 10 min to remove cell debris. Eachsupernatant then was dispensed into vials and frozen at-700C.

Poliovirus type 1 (obtained from the American TypeCulture Collection, Rockville, Md.) also was propagated inMA-104 cell cultures grown as described above. Beforepoliovirus inoculation, the cells were washed once withTris-buffered saline containing 100 U of penicillin per ml, 100jig of streptomycin per ml, and 2% FBS (also used as thediluent for poliovirus throughout this study). After virusadsorption, the cell monolayers were maintained in the cellmaintenance medium described above for HRV-Wa exceptthat 2% FBS was incorporated and no trypsin was added.Incubation, examination for cytopathic effect, and harvestwere as described for the HRV-Wa.

Viral plaque assays. Infectivity titers (PFU per milliliter) ofall samples were determined by a plaque assay methodspecific for each virus. A modification of the procedure usedby Wyatt et al. (37) was used to assay HRV-Wa. Pretreat-ment of the HRV-Wa samples with an equal volume oftrypsin (20 ,ug/ml) at 370C for 45 min was performed beforeinoculation of MA-104 monolayers. In 25-cm2 plaque bot-tles, 0.2 ml of inoculum was added to monolayers that hadbeen washed three times as described above. After HRV-Waadsorption for 1 h, the monolayers were washed once asdescribed above. A soft, 0.5% agar overlay containing EagleMEM-EBSS with 0.5 ,ug of trypsin per ml, 100 ,ug ofDEAE-dextran (Pharmacia, Uppsala, Sweden) per ml, 2.0mM L-glutamine, 100 U of penicillin per ml, 100 ig ofstreptomycin per ml, and 1.0 ,ug of amphotericin B(Fungizone) per ml was then added at 45°C. The bottles werecapped loosely and incubated at 37°C in a 5% CO2 atmo-sphere. After 4 to 5 days, the soft agar was easily drainedand the monolayers were stained with a 10% crystal violet

solution to reveal plaques. This procedure provided greatercontrast for plaque enumeration than the commonly usedfirm agar overlay containing neutral red.The poliovirus plaque assay was similar to that described

for HRV-Wa except that the trypsin and DEAE-dextranwere replaced with 2% FBS and 25 mM MgCl2 6H20 in theoverlay and the confluent monolayers were washed oncewith Tris-buffered saline containing 100 U of penicillin perml, 100 ,ug of streptomycin per ml, and 2% FBS before virusinoculation. After a 1-h adsorption period, the monolayerswere washed again and overlaid with soft agar. After incu-bation for 2 days at 37°C in a 5% CO2 atmosphere, theoverlay was easily drained and the monolayers were stainedwith 10% crystal violet to enable plaque enumeration.ELISA for HRV-Wa. The double-antibody, single-

sandwich ELISA (33) was modified to quantitate the HRV-Wa antigenicity in various water samples previously seededwith the virus.

Rabbit anti-HRV immunoglobulin G and peroxidase-conjugated rabbit anti-HRV immunoglobulin G were ob-tained from Accurate Chemical & Scientific Corp. (West-bury, N.Y.). The peroxidase substrate, o-phenylenediaminedihydrochloride, was obtained from Sigma.A 200-,lI sample of anti-HRV diluted 1:200 in 0.05 M

carbonate buffer (pH 9.6) was added to each well of amicrotiter plate (Nunc-Immuno plate; GIBCO) and incu-bated at 4°C for 14 to 24 h. The plate then was washed fivetimes with phosphate-buffered saline (pH 7.4) containing0.05% Tween 20 (Sigma). Twofold serial dilutions of seededwater samples were prepared with phosphate-buffered salinecontaining 0.05% Tween 20 plus 0.5% bovine serum albumin(Sigma). A 200-,u sample of each dilution was added to eachof two microtiter wells. Stool homogenates from childrenshedding rotavirus were included as positive controls. Plateswere incubated at 37°C for 45 min. After being washed asabove, 200 RI of peroxidase-conjugated anti-HRV (diluted1:200 in phosphate-buffered saline-0.05% Tween 20-bovineserum albumin) was added to each well and the plates wereincubated at 37°C for 45 min. After five additional washingswith phosphate-buffered saline-0.05% Tween 20, 100 RI ofo-phenylenediamine was added to each well. The o-phenylenediamine was diluted to 3 mg/ml in citrate-phosphate buffer (pH 6.0) containing 0.02% hydrogen per-oxide and prepared not more than 20 min before use. Plateswere incubated at room temperature for 15 min in the darkuntil the enzyme-substrate reaction was stopped by theaddition of 50 pul of 6 N H2SO4. Absorbance was then readwith a microELISA spectrophotometer (Dynatech Labora-tories, Inc., Alexandria, Va.) at a wavelength of 490 nm.

Electron microscopy of HRV-Wa stock. Stock pools ofHRV-Wa used to seed water samples were centrifuged at1,000 x g for 10 min to remove cell debris. The supernatantswere transferred to polyallomer tubes and centrifuged at80,000 x g for 60 min in a Beckman (Fullerton, Calif.)ultracentrifuge equipped with an SW41 rotor. The superna-tants were discarded, and the virus pellets were suspendedin 50 ,ul of Eagle MEM containing 0.5% gelatin. One drop ofeach virus concentrate was placed on a Formvar-carbon-coated 400-mesh copper grid. The suspensions were allowedto air dry for 15 min and then were negatively stained with 2drops of 2% phosphotungstic acid (pH 6.5). After 1 min, thegrids were removed, blotted dry, and examined with aHitachi-500 (Tokyo, Japan) transmission electron micro-scope.Water samples. Five water samples were chosen as repre-

sentative of polluted and nonpolluted aquatic environments.

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TABLE 1. Characteristics of water samples employed

Chloride Hardness_Solids_(mg/liter)_Fecal_FecalWater sample pH Conductivity (m f TOC HardnesseSolidso(mg/liter)tFecaocFeca(p.S/cm) (mg gof i (mg/literaasTotal Total Volatile ColFor00ms) streptococciCl/liter) CaCOO) Total volatile suspended suspended (CFU/I ml) (CFU/1 ml)

Lake water 6.0 50 <0.5 1 29 92 88 10 8 2 20Creek water 7.8 450 7.7 4 273 236 52 42 17 150 150Groundwater 6.7 600 <0.5 <1 339 360 100 4 4 <1 <1Secondary effluent 6.6 600 16.3 17 223 340 88 38 34 3.4 x 104 1.2 x 104Tap water" 6.1 250 2.3 1 74 180 104 5 5 <1 <1

a Displayed a total chlorine residual of 0.60 mg/liter, 60% chloramines.

A mountain lake water sample represented a relativelypristine surface water which is used for recreation, swim-ming, and drinking. A polluted creek water sample wasobtained from a creek that flowed through relatively popu-lated areas and received mostly domestic sewage effluents.A groundwater sample, a secondary wastewater treatmentplant effluent sampled before chlorination, and a chlorinatedtap water sample also were employed. Two liters of eachwater sample were collected in sterile Nalgene plastic bot-tles. Conductivity and ambient pH were determined for eachwater sample in the field by standard instrumental methods.In the laboratory, the water samples were evaluated imme-diately for solids (i.e., total, total volatile, total suspended,and volatile suspended), chloride content by the mercuricnitrate method, total organic carbon (TOC) with the use of aBeckman 915-B TOC analyzer, water hardness by the EDTAtitrimetric method, and fecal coliforms and fecal strepto-cocci by membrane filtration procedures. All methods wereperformed as previously described (2). The total chlorineresidual of the tap water also was determined by the N,N-diethyl-p-phenylenediamine ferrous titrimetric method aspreviously described (2). The water samples were employedin the viral survival studies immediately upon collection(i.e., fresh samples which had not been stored).

Viral stability studies. Samples (1 ml) of the HRV-Wa pool(1.7 x 105 PFU/ml) were seeded into duplicate 125-ml sterileglass bottles containing 99 ml of each water sample, therebyyielding a virus titer of 1.7 x 103 PFU/ml. The poliovirusseeding was performed in a similar manner with separatebottles. The seeded samples were kept undisturbed at 20°Cand were exposed to regular room light. A 1:400 dilution ofeach seed pool was frozen to be used later as the positivecontrol for plaque assays. This eliminated the need to assayall samples at one time (27). After addition of virus, eachbottle was capped and gently shaken to disperse the virus. A1-ml sample was then transferred immediately from eachbottle into a sterile 5-ml vial and diluted serially 1:2 and 1:4.These samples (designated day zero with the respectivewater type and virus) were frozen and maintained at -70°Cuntil assayed for infectivity or both infectivity and antigeni-city. On subsequent days (1 to 6, 9, 16, and 21), another 1-mlsample was taken from each bottle and processed in thesame manner.

Statistical analyses. All statistical analyses of the datapresented in subsequent sections of this paper were under-taken as previously described (35).

RESULTS

Characterization of water samples. The pH did not varymarkedly among the water samples used, except for creekwater which had the highest pH (7.8) of all water samples(Table 1). The conductivity, hardness, and solids observed

in creek water, groundwater, and secondary effluent indi-cated a greater amount of dissolved constituents than in theother two samples. As expected, the polluted creek waterand secondary effluent contained substantially higher con-centrations of chloride, TOC, suspended solids, fecal coli-forms, and fecal streptococci than did the other watersamples (Table 1).

Electron microscopy of HRV-Wa stock. Concentrations ofHRV-Wa in the seeded water samples used in this studywere too low to be detected by electron microscopy. Evenconcentration of the viruses in these samples by ultracentri-fugation did not yield sufficient virus to be detected byelectron microscopy. Consequently, only original stocks ofHRV-Wa used to seed water samples were used for electronmicroscopic analysis. After ultracentrifugation, these stockscontained approximately 107 PFU per ml. Electron micro-graphs of these stocks (not shown) demonstrated HRV-Waparticles ranging in diameter from 52 to 70 nm, which mayhave reflected a mixture of single-shelled and double-shelledparticles (11, 30). Also evident were slightly altered andseverely damaged capsids (not shown). Other electron mi-crographs, such as the one shown in Fig. 1, presented ample

FIG. 1. Electron micrograph of HRV-Wa stock used to se.edwater samples. Bar, 100 nm.

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0.5 A Sec.Effluent 5.5 -.144 -.971

0 U Groundwater 3.4 -.158 -.962z0 Creek Water 1.8 -.237 -.964

Z 0 A|/ Tapwater 6.7 -.699 -.9930)0

0

r -2

0 2 4 6 9 16Days

FIG. 2. Infectivity reduction patterns of HRV-Wa in fresh wa-ters. N, and No are the viral titers (PFU per milliliter) as determinedby plaque assays at days t and zero, respectively. Shown are theslopes (rates ki) and correlation coefficients of best-fitting lines.

evidence of HRV-Wa aggregation in stocks used to seedwater samples.

Viral infectivity reduction (inactivation). Viral stability inthe different fresh waters was evaluated by comparing theviability ratios at various times after the water samples wereseeded. The viability ratio is log1o N,/NO, where No equalsthe initial infectivity titer and N, equals the titer at elapsedtime t (in days). Shown in Fig. 2 and 3 are the inactivationpatterns in fresh waters for HRV-Wa and poliovirus, respec-tively. Also shown in Fig. 2 and 3 are the computed slopes(ki) of best-fit lines for the inactivation rates. From thesedata, the time required for a 2-log reduction of initial viraltiter was computed from the equation of the best-fitting lineby assigning a y value of -2 and calculating the correspond-ing x value (days). These values can be regarded as thepredicted mean inactivation times of each virus for anyparticular water sample. In Fig. 4, the time required for this2-log reduction of initial viral titer for each experiment isrepresented graphically. The mean inactivation times ofHRV-Wa and poliovirus in each of the five water samples(Fig. 4) were analyzed by the Newman-Keuls multiplecomparisons of the overall mean inactivation times of thetwo viruses. This analysis indicated that there was signifi-cant (P < 0.05) variation between the mean inactivationtimes for the two viruses with respect to water type. Thisalso was true for variations between water types withrespect to virus. An analysis of simple effects of virusillustrated that significant differences (P < 0.05) in meaninactivation times existed between the two viruses for allwater samples except tap water (Fig. 4). The slopes of the

0

z

z -1_0

0

%-2.0

Ut l ~~~~~~~~~AVOtINITIAL SLOPE CORR.: l ~~~~~~~SAMPLE CONC. k; COEFF..- l ( 103~~~~~~~PFU/ml) (day 1)ro * Lake Water 2.7 -.087 -.953

O _U Groundwater 2.7 -.131 -.934

~~~~~ A~~~~~~~~Sec Ettluent 2.2 -.239 -.940

OCreek Water 2.5 -.442 -.940

ATapwater 2.5 -1.131 -.945

0 2 4 6 9 16 21Days

FIG. 3. Infectivity reduction patterns of poliovirus type 1 infresh waters. N, and No are the viral titers (PFU per milliliter) asdetermined by plaque assays at days t and zero, respectively.Shown are the slopes (rates ki) and correlation coefficients ofbest-fitting lines.

24C00m _a, > 20

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o cJ 12

0-

v-8o CCr:n

00CO >

R 4

_-

" LakeWater

ag Ground-

water

bb

f Sec.Effluent

C

e CreekWater

d^ _ d~~~,e Tapwater

_

_-

HRV-Wa Poliovirus

Virus TypeFIG. 4. Days required for two-logl0 reductions of initial infectiv-

ity titers of HRV-Wa and poliovirus type 1 in fresh waters. Pointsdesignated with different letters are significantly different at P < 0.05by analysis of variance.

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1,

E

C:

0

0)

c'

~0

Co

C:

coJ0

111

0 1 2 3 5 9Days

FIG. 5. Antigenicity reduction patterns of HRV-Wa in freshwaters as determined by ELISA. Points designated with differentletters are significantly different at P < 0.05 by analysis ofcovariance. Shown are the slopes (rates kr,) and correlation coeffi-cients of best-fitting lines.

lines produced in Fig. 4 are proportional to the differences ininactivation times for the two viruses in the different watersamples. The analysis of simple effects of water furtherdemonstrated that at least two water types varied signifi-cantly for each of the viruses. These effects can be seen inFig. 4 by noting the disparity of the mean inactivation timesof the five water samples for each virus. Results of Newman-Keuls multiple comparisons are summarized at the ends ofeach line in Fig. 4. Those points with different letters are

significantly different at P < 0.05. Despite these differences,both viruses survived longest in lake water and shortest inthe chlorinated tapwater.HRV-Wa antigenicity reduction. Antigenicity reduction

patterns of HRV-Wa in the five fresh waters as determinedby ELISA are presented in Fig. 5. Regression analysis of theELISA absorbance values for each water sample demon-strated that the rate of decrease in antigenicity (slope ofbest-fit line, kb,) was least for lake water and groundwatersamples, while the greatest rate of decrease was observed insecondary effluent. To determine whether differences in theELISA absorbance values existed among any of the watersamples on day 9 of the study, we evaluated each watersample by individual comparisons after analysis ofcovariance (35) among means for that day. Significant differ-ences (P < 0.05) were exhibited in each pair of watersamples with the exception of (i) lake water-groundwaterand (ii) tap water-creek water. In addition, the magnitude ofHRV-Wa antigenicity reduction over 9 days of the studyperiod for each water type was determined (Fig. 6). Thegreatest reductions in antigenicity were observed in creek

water, secondary effluent, and tap water. Lesser reductionswere found in lake water and groundwater. Different lettersat the top of each bar (Fig. 6) indicate significantly differentantigenicity reductions (P < 0.05). Essentially, the ELISAresults showed that substantial variation in HRV-Wa anti-genic stability existed between the water samples, except for(i) lake water-groundwater and (ii) tap water-creek water.Comparison of HRV-Wa infectivity reduction and antigeni-

city reduction. The relationship between infectivity titer andthe quantity of antigen (determined by ELISA) was exam-ined for fresh waters seeded with HRV-Wa. This wasaccomplished by comparing the infectivity reduction ratio(N,INo) to the antigenicity reduction ratio (Abs,/Abso) over 9days of the study period (Fig. 7). The slopes of each linewere calculated by regression analysis and represent the rateof change (ki,) in the ratio of infectivity reduction to antige-nicity reduction. Therefore, a negative slope (observed forall water samples, Fig. 7) demonstrated that the infectivitydecreased more rapidly than did the quantity of antigen. Inlake water and secondary effluent, lower negative slopesindicated that infectivity titers diminished at only a slightlygreater rate than did the quantities of antigen. In tap water,the infectivity titer of HRV-Wa decreased very rapidly ascompared with the rate of antigen reduction.HRV-Wa reduction rates and water quality parameters.

Correlations between HRV-Wa reduction rates (i.e., ki, k,and kia) and water quality parameters were examined byregression analyses (Table 2). The reduction rates of HRV-Wa were plotted against the measured values of each waterquality parameter. Correlation coefficients (r) associatedwith the best-fit lines for the water quality parameters thenwere calculated, and statistical significance was determined.The tap water data were excluded from this analysis because

C.7

b bE

o -a .60

a.o> -.

o0 a

co .4 -

< 0) -

~j.C

00c .2

0~~~~~~

Lake Creek Ground- Sec. TapwaterWater Water water Effluent

Water TypeFIG. 6. Magnitude of HRV-Wa antigenicity reduction in fresh

waters over 9 days of the study period as determined by ELISA.Bars designated with different letters are significantly different at P< 0.05 by analysis of covariance.

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SAMPLE (dayI COEFF.

.0

Z A Sec. Effluent -.067 -.721

*

0 Creek Water -.171 -.992

O Groundwater -.191 -.9820

A Tapwater -.455 -.947

0c

0

.2

=3

2 >-1

cDc

0

_W 0

cc -2

0 12 3 5 9

Days

FIG. 7. Ratio of infectivity reduction to antigenicity reduction

for HRV-Wa in fresh waters. N, and No are the viral titers (PFU per

milliliter) as determined by plaque assays at days t and zero,

respectively. Abs, and Abso are ELISA absorbances at days t andzero, respectively. Shown are the slopes (rates kia) and correlationcoefficients of best-fitting lines.

viral inactivation rates in this water undoubtedly were

influenced by the presence of chlorine. There was a statisti-cally significant (P < 0.01) positive correlation between thepH of the water and the viral inactivation rate (ki) (Table 2).Thus, with an increase in the natural pH of the water,HRV-Wa infectivity was reduced at a correspondinglygreater rate. Significant positive correlation (Table 2) was

observed between the viral antigenicity reduction rate (k,)and both chloride content (P < 0.01) and TOC concentration(P < 0.05). This indicated that an increase in the concentra-tion of these parameters was associated with a correspond-ing increase in the reduction rate of HRV-Wa antigenicity.Finally, the reduction rate (kia) of the ratio of infectivityreduction to antigenicity reduction was significantly corre-

lated (P < 0.05) with water hardness. This suggested that as

hardness increased, viral infectivity was reduced more rap-

idly than was antigenicity.

DISCUSSION

The survival of enteric viruses in the aquatic environmentis influenced by a variety of factors including temperature,pH, light, biological activity (e.g., presence of proteolyticenzymes), and adsorption to suspended materials (4).Enteroviruses, such as poliovirus type 1, have been shownto survive for prolonged periods in fresh waters underlaboratory (18) and field (25) conditions. Although Hurst andGerba (18) observed that the survival of the simian rotavirus

SAl1 in fresh and estuarine waters generally fell within therange observed for several enteroviruses (i.e., poliovirustype 1, echovirus type 7, and coxsackievirus B3), theseinvestigators reported that SAl1 and enterovirus survivalrates were not always parallel in certain water samples.Moreover, SAl1 inactivation rates varied from 3 to 14 ormore days for a 3-loglo infectivity reduction (20°C) amongthe different freshwater samples (18). Greatest survival wasobserved in heavily polluted fresh water (i.e., 12 to 14 ormore days for a 3-log reduction). In the study reportedherein, the survival times for HRV-Wa and poliovirus type 1in natural fresh waters were compared. Although similarpatterns of infectivity reduction were found for these twoviruses in the fresh waters tested, statistically significant (P< 0.05) mean inactivation times were observed for the twoviruses in all fresh waters, except tap water. Furthermore,HRV-Wa remained infectious significantly longer in the twopolluted water samples (secondary effluent and creek water)than did poliovirus. In nonpolluted lake water and ground-water, the converse was observed, i.e., poliovirus infectivitypersisted substantially longer than did HRV-Wa infectivity.The lack of significant difference in the survival of the twoviruses in tap water could be attributed to the broad virucidalaction of the chlorine (total residual of 0.6 mg/liter, 60%chloramines, at pH 6.1 and 20°C) in the sample (10). Bothviruses were inactivated in the chlorinated tap water fasterthan in any other water samples (2 to 3 days for a 2-logreduction of infectivity titer). However, the observed inac-tivation rates in chlorinated tap water were slower thanthose previously reported for these and other viruses, pos-sibly because of the lower virucidal action of the chlo-ramines (3, 5, 32). Recently, Berman and Hoff (3) reported a99% (2-log) inactivation of the SAl1 rotavirus within 4 min ofexposure to 0.5 mg of chlorine per liter at pH 6 and 5°C. Thisoccurred in purified preparations of single particles or cell-associated (aggregated) virions. These investigators foundthe aggregated viruses to be more resistant to chlorineinactivation than the single-particle preparations of SAil.Vaughn et al. (32) also found that single-particle suspensionsof HRV-Wa and SAl1 virus were rapidly inactivated (5-logreduction in titer within 20 s) at pH levels of 6 to 8 and freechlorine residuals of 0.3 mg/liter or less. Similarily, Sharp etal. (28) demonstrated increased susceptibility of dispersedpoliovirus type 1 (Mahoney) particles to chlorine. In the

TABLE 2. Correlation of HRV-Wa reduction rates in freshwaters with water quality parameters

Correlation (r) with HRV-WaWater quality parameter reduction rates"

ki k,, kij,

pH 0.-ConductivityChloride 0.976bTOC 0.879cHardness 0.783'Total solidsTotal volatile solidsTotal suspended solidsVolatile suspended solidsFecal coliformsFecal streptococci

11 Rates: ki, infectivity reduction; k,,. antigenicity reduction; ki,, reduction inthe ratio of infectivity reduction to antigenicity reduction. -, Lack ofstatistical significance (i.e., P > 0.05); excluding the tap water results.

b Statistically significant at P < 0.01.' Statistically significant at P < 0.05.

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current study, electron micrographs indicated that the HRV-Wa was aggregated before seeding of the freshwater sam-ples. Although we were unable to examine the HRV-Wa inthe water samples by electron microscopy, it is reasonable toassume that some viral aggregation persisted in the watersamples and that this could have influenced virus survival.As in similar studies (18), no attempt was made to ensurethat only single virus particles were present in the watersamples. While some recent studies (3, 32, 34) have indi-cated that virus inactivation in water is measured mostaccurately when using single virus particles, it must beemphasized that indigenous viruses in water often are aggre-gated (5) and that inactivation rates measured with singleparticles may not reflect the inactivation of aggregatedviruses (3). Nonetheless, in the present study, it is mostlikely that the further dilution of each water sample in EagleMEM-EBSS (with gelatin) or in phosphate-buffered saline(with Tween 20 and bovine serum albumin) before HRV-Waplaque assay or ELISA, respectively, resulted in deaggrega-tion of the viral particles. Hamblet et al. (15) demonstrateddeaggregation of poliovirus type 1 particles and a concomi-tant increase in viral titer after dilution of samples in asolution with high organic content (nutrient broth). Simi-larly, other investigators have diluted SAl virus-seededwater samples in organic solutions such as MEM-L15 me-dium (3) and tryptose phosphate broth (18). Possible aggre-gation, the lower virucidal effect of the chloramines, and thechlorine demand of our test system may have accounted forthe slower inactivation rates for HRV-Wa in chlorinatedwater as compared with those for SAl virus as reported byBerman and Hoff (3) and for HRV-Wa and SAl as reportedby Vaughn et al. (32).With regard to persistence of viral infectivity in the five

water samples, both HRV-Wa and poliovirus exhibitedsimilar patterns of survival, remaining infectious in thenonpolluted waters (e.g., lake water) significantly longer (P< 0.05) than in the polluted waters (e.g., creek water andsecondary effluent). A total of 17 and 8 days were requiredfor a 2-log reduction of initial titers of HRV-Wa in the lakewater and creek water; respectively. Similar inactivationrates were reported by Hurst and Gerba (18) for SAll inpolluted waters (i.e., 12 to more than 14 days for a 3-logreduction). In contrast, however, these investigators ob-served that SAl survived longest in polluted fresh waters.Our research has demonstrated prolonged survival of HRV-Wa in lake water and groundwater exceeding that reportedfor SAl in similar nonpolluted waters (18, 21). At present,there is limited information on the fate of viruses in ground-water (20). In our study, poliovirus type 1 was inactivated ingroundwater at a somewhat slower rate (-0.13 day-') thanreported by Yates et al. (39) (mean decay rate in 11 ground-water samples was -0.162 day-1) and by Keswick et al. (21)(-0.21 day-') for the same virus in groundwater. In con-trast, Bitton et al. (6) reported a much slower inactivationrate (-0.0456 day-') for poliovirus type 1 in a Floridagroundwater sample. The data presented herein indicate thatthe HRV-Wa could persist for extended periods in ground-water (approximately 12 days for a 2-log reduction of initialinfectivity titer). Even longer survival times have beenreported by McDaniels et al. (22) for the calf rotavirusseeded in distilled water and wastewater under laboratoryconditions in the absence of light. These investigators re-ported that 7 to 13 days and 80 days were required for a 90%(1-log) reduction in calf rotavirus infectivity at 26 and 8°C,respectively.With the exception of the groundwater and secondary

effluent results, the HRV-Wa inactivation times in the freshwaters were significantly different. Since physical factorsknown to influence virus survival in the aquatic environmentwere held constant (e.g., temperature), it follows that thedifferences in virus inactivation rates could be attributed todifferences in the chemical or biological properties (or both)of the fresh waters employed. For example, the rapidinactivation rate of HRV-Wa in tap water was due mostlikely to the presence of chlorine. The presence of sus-pended sediments in seawater has been shown to prolong thesurvival of the SAl virus, which remained infectious for 19days when adsorbed to sediments in seawater but for only 9days when freely suspended in seawater (27). Farrah et al.(12), however, reported that the SAl virus adsorbs to alesser degree than does poliovirus type 1 to aluminumhydroxide flocs in tap water and activated sludge flocs. Inthe present study, suspended solids in the water samples didnot prolong the survival of HRV-Wa or poliovirus type 1. Infact, the infectivity of both viruses declined fastest in thepolluted creek water which displayed the highest total sus-pended solids (42 mg/liter). Similarly, the bacteriologic con-tent of the water sample, as determined by the concentra-tions of fecal coliforms and streptococci, did not influencesignificantly the inactivation rate of HRV-Wa. In point offact, secondary effluent, which contained the highest con-centrations of indicator bacteria, displayed the second slow-est HRV-Wa inactivation rate. In contrast, Ward et al. (34)found that microorganisms in fresh waters were associateddirectly with virucidal activity directed against echovirustype 12. After regression analyses were performed betweenHRV-Wa inactivation rates (ki) and chemical or biologicalproperties of the fresh waters, only the pH of the watercorrelated significantly with the ki (i.e., as pH increased, kiincreased). Tap water data were ornitted from this analysisbecause of the influence of chlorine. Owing to the moderatepH differences of the water samples (pH 6.0 to 7.8), thereductions in HRV-Wa infectivities probably did not resultfrom the direct virucidal action of pH. Rather, it seems likelythat the pH of water samples may have influenced the extentto which aggregates of stock HRV-Wa were dispersed (13).It is believed that when dispersed, HRV-Wa is more suscep-tible to other virucidal factors in the water since this phe-nomenon has been observed for poliovirus type 1 and SAll(3, 28). It must be emphasized, however, that the pH andionic conditions that induce aggregation or dispersal of theHRV-Wa are not known.

In our study, the fate of HRV-Wa antigenicity in freshwaters was monitored by ELISA. As with infectivity, theantigenic reactivity of HRV-Wa decreased at a faster rate inthe polluted waters (secondary effluent and creek water)than in the nonpolluted waters (lake water and groundwa-ter). The reduction rates of HRV-Wa antigenicity (ka) cor-related directly (P < 0.05) with the chloride content andTOC concentration of the water samples. Tap water datawere omitted from this analysis because of the influence ofchlorine. It appears that the chloride ion and organic carbon(or related, but unknown parameters) affected the viralprotein coat and the associated reduction in antigenic reac-tivity. Increasing concentrations of chloride and TOC couldhave indirectly affected HRV-Wa antigen stability by dis-persing the viral aggregates. When HRV-Wa is dispersed,antigens (i.e., epitopes) are exposed and may become moresusceptible to inactivating factors in the water samples.The rate of inactivation also was compared with the rate of

antigenicity reduction for HRV-Wa seeded in freshwatersamples. In all water samples, the HRV-Wa infectivity

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declined at a faster rate than did the antigenicity. Thereduction rate (kia) in the ratio of infectivity reduction toantigenicity reduction was greatest (-0.455 day-') for theHRV-Wa seeded in the chlorinated tap water. This mayindicate that the loss of HRV-Wa infectivity in the tap waterwas associated primarily with damage to the viral nucleicacid (RNA), while the quantity of antigen remained some-what more stable. In this regard, research with the bacterio-phage f2 has demonstrated that the viral RNA is the principaltarget of chlorine inactivation (5). In fact, the destructiveaction of a given chlorine molecular appears to be related toits ability to penetrate the protein coat (5). Moreover,chlorine concentrations in excess of 1 mg/liter have beenfound to separate the structural components of poliovirustype 1 (Mahoney) and to produce particles lacking RNA (26).However, at low chlorine concentrations (i.e., <0.8 mg/liter), such as that measured in the tap water in our study,the protein coat of poliovirus type 1 (Mahoney) is affectedwithout separation of viral components (1). Similarly, Churnet al. (8) demonstrated that sodium hypochlorite affected theprotein coat of parvovirus type H-1, while the DNA of thevirus remained intact. These investigators found that thechlorine-treated parvovirus particles did not adsorb to hostcells. In the present study, it is conceivable that the chlorinein tap water damaged the protein coat of HRV-Wa whichresulted in a more rapid reduction of infectivity than antige-nicity, the former by inhibiting viral adsorption to host cells.In the other waters, infectivity titers declined only slightlyfaster than did antigenic reactivities where the reductionrates (kia) of their ratios were correlated directly (P < 0.05)with the water hardness. Although increasing hardness hadno significant effect on rates of infectivity or antigenicityreduction when analyzed independently, it was associatedwith a greater reduction rate in viral infectivity as comparedwith that of antigenicity. The mechanism for this effect ofwater hardness (sum of calcium and magnesium ions) is notknown. However, calcium ions (but not magnesium ions)have been shown to maintain HRV integrity by stabilizingthe outer capsid layer and, therefore, viral infectivity (29).Moreover, Yates et al. (39) reported a positive correlationbetween the calcium hardness of various groundwaters andthe decay rates of bacteriophage MS-2 (i.e., the higher thecalcium concentration, the greater the decay rate of thevirus). However, increasing the calcium concentration in a

particular water sample did not affect the MS-2 decay rate.Therefore, these investigators concluded that some unmeas-ured property of water (associated with the calcium concen-

tration) was involved in the observed variation of MS-2decay rates. In the waters of the present study, the loss ofHRV-Wa infectivity could have been associated with dam-age to the viral RNA, although the antigenic integrity of theviral protein also was affected. Similarly, O'Brien and New-man (25) reported damage to viral RNA as the cause of theinactivation of radioactively labeled poliovirus type 1 andcoxsackievirus Bi in membrane dialysis chambers placed inthe Rio Grande in southern New Mexico. Recently, Ward etal. (34) demonstrated that proteolytic bacterial enzymes infresh water inactivate echovirus type 12 by cleaving viralproteins, which is followed by nuclease digestion of theexposed viral RNA. In the present investigation, comparisonof HRV-Wa infectivity and antigenicity reductions estab-lished that different mechanisms may have contributed to theinactivation of the virus in the different waters.The research repoi-ted herein demonstrated that the sur-

vival of HRV-Wa was significantly different from that ofpoliovirus type 1 (CHAT) in all water samples tested (with

the exception of tap water). Moreover, the survival ofHRV-Wa in lake water and groundwater exceeded thatreported for SAl1 in similar waters (18, 21). It is proposedthat HRV-Wa can be used as a model for future studiesdealing with rotaviral persistence in the aquatic environ-ment.

ACKNOWLEDGMENTS

We thank R. Dean Blevins of the Department of Health Sciences,East Tennessee State University, for his assistance with cell cul-tures.

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