Epidemiology of Fasciola hepatica in Florida with emphasis on the population dynamics and infection prevalence of the pr...


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Epidemiology of Fasciola hepatica in Florida with emphasis on the population dynamics and infection prevalence of the primary snail intermediate host, Fossaria cubensis
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ix, 269 leaves : ill. ; 29 cm.
Kaplan, Ray Matthew, 1960-
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Subjects / Keywords:
Research   ( mesh )
Fasciola hepatica -- pathogenicity -- Florida   ( mesh )
Fasciola hepatica -- isolation & purification   ( mesh )
Snails -- pathogenicity -- Florida   ( mesh )
Fascioliasis -- transmission -- Florida   ( mesh )
Fascioliasis -- epidemiology -- Florida   ( mesh )
DNA Probes -- diagnostic use   ( mesh )
Population Dynamics -- Florida   ( mesh )
Prevalence   ( mesh )
Disease Vectors   ( mesh )
Cattle   ( mesh )
Department of Pathobiology thesis Ph.D   ( mesh )
Dissertations, Academic -- College of Veterinary Medicine -- Department of Pathobiology -- UF   ( mesh )
bibliography   ( marcgt )
non-fiction   ( marcgt )


Thesis (Ph.D.)--University of Florida, 1995.
Bibliography: leaves 258-268.
Statement of Responsibility:
by Ray Matthew Kaplan.
General Note:
General Note:

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University of Florida
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oclc - 49847266
notis - ALQ4629
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Many hours of thought, planning and plain old work go

into the creation and completion of a dissertation. I wish to

thank and acknowledge the people who have given me their time

and their hearts in the past four and one half years. Without

their help and support, this project could not have been


First, I would like to thank my committee chair and

mentor, Charles H. Courtney. Charlie has served as an

excellent example of what a professor should be. In addition

to being an excellent teacher and mentor in parasitology, he

has shown great care on a personal level, nurturing my growth

both as a professional and as an individual.

I would also like to thank my committee members, Dr. John

Dame who directed the molecular portion of this project, Dr.

Fred Thompson who provided his expertise in snail biology,

Dr. P.V. Rao who aided in the statistical analysis, and Dr.

Ellis Greiner who guided me through diagnostic parasitology.

Dr. Greiner's warmth, caring nature and sense of humor were

invaluable throughout my four and one half years as a PhD

student. Ellis and Mary Greiner open their hearts and home to

their students. They represent the best in academe.

Special thanks go to Dr. Roman Reddy who served as my

teacher, collaborator and friend. Dr. Reddy was always there

to answer questions, suggest alternate techniques, and

encourage me to run yet anther gel. Dr. Qi Yun Zeng also

deserves special recognition. She has helped me beyond

measure over the past few years both as a teacher and

collaborator. With Qi Yun around, I never had to worry about

not having time to get things done -- she always made the time

to help me no matter what else she had scheduled. Qi Yun's

caring, friendly and cheerful nature makes Dr. Courtney's

laboratory a fun place to work.

I also would like to acknowledge Brian R. Kreitz and

Denette 0. Cooke, veterinary students now beginning their

senior year. The work that both Brian and Denette contributed

to this project were invaluable to me. They also helped to

keep things fun. From lunchtime basketball to thoughtful

discussions ranging from the Gators to Mike to OJ. From the

Angus to the Panda. I wish them both the best of luck as

they begin their new careers as veterinarians.

I also wish to acknowledge Florida Extension Agent Pat

Miller and his family, Susie, Carrie and Becky. Pat was

instrumental in my field work in southern Florida. His

contacts with local ranchers, knowledge of the area and

hospitality were greatly appreciated. The latter included

relaxing with a beer after a day in the hot Florida sun

chasing snails, family barb-b-ques when Holly and Ellie got to


join me on "snail hunting" trips, stops at the Duck Pub for

Duck Burgers, and dips in the pool. Pat's friendship and the

added perks thereof made what could have been repetitious

dreary trips to Okeechobee into fun mini-vacations for myself

and my travel partners.

Finally, I want to thank my family. My parents, Aaron

and Elaine, and sister, Karen, for babysitting, delicious

dinners, and emotional support. Thank you to Ellie, my snail

hunting buddy, for all the times she patiently understood that

Daddy was working and thank you for the times she made me

forget about working. Thank you to Harry for sleeping the

night through, so I was cognizant enough to write the last

chapters. Thank you to Holly for everything. It is to my

family that this dissertation is dedicated.



ABSTRACT . . vii



History and Distribution of Fasciola Hepatica 1
Biology and Life Cycle of Fasciola hepatica .. 2
Biology of Fossaria cubensis . 7
Clinical Signs and Pathology . 9
Host Immunity . .. 12
Economic Importance . .. 14
Diagnosis .. . 18
Seasonal Transmission Dynamics of Fasciola
Hepatica in the United States .. 20
Rationale for Seasonal Control . 22
Current Treatment Recommendations for the Gulf
Coast States . .. .. 25
Current Treatment Recommendations for the
Northwestern States . 26
Epidemiology and Control of Bovine Fascioliasis in
Florida: Review of Literature, Research Needs and
Implications for Improved Control
Recommendations . .. 27


Introduction . ....... 33
Materials and Methods . .... 37
Results . .. 40
Discussion .. . . 42


Introduction . .. 57
Materials and Methods . 61
Results . . 67
Discussion . . 71

ASSAY ............. ... 84

Introduction . . 84
Materials and Methods . 87
Results . .... .. 97
Discussion . . .. 101


Introduction . . .. 120
Material and Methods . .. 124
Results . . 132
Northern Florida . .132
University of Florida Dairy Research Unit
(DRU), Hague, Alachua County, Florida .132
H. E. Wolfe Ranch, St. Augustine, St.
Johns County, Florida .. 137
Central Florida . .. 142
Deseret Ranch, Deer Park, Brevard County,
Florida . . 142
Creek Ranch, Lake Hatchenehaw, Polk County,
Florida . . 149
Southern Florida . 154
Brighton Seminole Indian Reservation, (Glades
County), Florida . .. 154
John Williams Ranch, Okeechobee, Okeechobee
County, Florida . .. 161
Rio Ranch, Okeechobee, Okeechobee County,
Florida . . 166
Water Budgets for Selected Sites in Florida 173
Discussion . . 174


REFERENCE LIST . .. ... 258


Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy



Ray Matthew Kaplan

August 1995

Chairperson: Charles H. Courtney, DVM, PhD
Major Department: Department of Pathobiology

Fascioliasis is one of the most important diseases of

cattle in Florida. Southern Florida, where more than 66% of

beef cows are pastured, is subtropical, and conjectural

evidence suggests that seasonal transmission of Fasciola

hepatica may differ from that in temperate northern Florida.

Current recommendations for control of fascioliasis in cattle

are based on data only from northern Florida. Therefore, it

is extremely important to the cattle industry of southern

Florida to determine the seasonal transmission dynamics of F.

hepatica. In this project, the epidemiology of F. hepatica in

northern, central and southern Florida was determined over 2

years by studying the bionomics of Fossaria cubensis

populations on 7 cattle ranches. Additionally, snail

infection prevalence was determined during year 2 of this


study. Because existing techniques lacked the sensitivity and

specificity needed to obtain accurate infection prevalence

data, a DNA probe was developed. This DNA probe (pFh5)

contains 2 124 bp repeat sequences belonging to a large family

of 124 bp repeats constituting approximately 15% of the F.

hepatica genome. pFh5 has excellent sensitivity and

specificity; a single miracidia is detected and it does not

cross hybridize with DNA of Fascioloides magna, Paramphistomum

liorchis or Heterobilharzia americana, trematodes that share

the same intermediate host and enzootic range as F. hepatica.

Using this DNA probe, over 5,000 snails from 6 ranches were

assayed. Infection prevalence for individual ranches varied

from 0.1% to 3.1%. In a separate experiment, it was

demonstrated that eggs of F. hepatica can survive and develop

on pasture during the summer with greater than 80% survival

after 28 days.

Results of this study demonstrated that seasonal

transmission of liver flukes is virtually the same for all of

Florida. Transmission occurs predominantly in winter and

spring. Wet summers followed by cool, wet weather in early

autumn could result in significant levels of transmission

occurring by December. Summer transmission will rarely occur

in Florida. Although snail habitats remain wet throughout

summers of most years, ecological conditions of Fossaria

habitats are not conducive to snail activity. Therefore,

current control recommendations remain unchanged except that


annual treatment should be given in late summer, 1 to 2 months

sooner than recommended previously.


History and Distribution of Fasciola hepatica

Fasciola hepatica, the common liver fluke, belongs to the

phylum Platyhelminthes, class Trematoda, order Digenea, family

Fasciolidae. Domestic ruminants are the definitive hosts,

however numerous other mammals, including man, may become

infected (Malek, 1980). The earliest known reference to the

liver fluke (and liver rot) was made in 1379 by Jean de Brie

in France, in his treatise on the proper management of sheep.

Sir Anthony Fitzherbert published the first recognizable

description of the liver fluke in 1523, and Francesco Redi in

1668 published the first illustration. A major breakthrough

in the understanding of trematode biology came in 1842, when

J. Steenstrup published his work on the theory of "alternation

of generations". This work drew wide attention and paved the

way for the independent discovery of the complete extra-

vertebrate life cycle of F. hepatica by A.P. Thomas in England

and R. Leuckart in Germany in 1882. The discovery of the

migratory path taken by larval flukes in their vertebrate

hosts was made by D.F. Sinitsin in 1914 in Russia, thus

completing the knowledge of the liver fluke life cycle

(Reinhard, 1957).


Fascioliasis, caused by infection with the liver fluke,

Fasciola hepatica, remains one of the most important diseases

of grazing ruminants throughout much of the world (Malek,

1980). The transmission of F. hepatica is dependant upon the

presence of its lymnaeid snail intermediate hosts, therefore

the distribution of the parasite is limited to those

geographic areas where the appropriate snail species are

present. In the United States, F. hepatica is enzootic

primarily in the Gulf coast and western states, where high

annual rainfall, large areas of poorly drained pasture and

certain soil types provide suitable lymnaeid snail habitats

(Malone, Loyacano, Armstrong, & Archbald, 1982a).

Additionally, the distribution and prevalence of liver flukes

may increase in areas where irrigated pastures are used. This

is particularly important in the western United States

(Malczewski, Wescott, Spratling, & Gorham, 1975). In Florida,

F. hepatica is almost exclusively restricted to the peninsula

south and east of the Suwannee River (Shearer, Courtney, &

Richey, 1986) where 92% of Florida's 1.1 million beef brood

cows are pastured (Florida Department of Agriculture and

Consumer Services, 1994).

Biology and Life Cycle of Fasciola hepatica

Adult F. hepatica reside in the intrahepatic biliary ducts

of host animals and eggs are carried with the bile into the

bowel lumen and then passed in the feces. Each of these eggs


contains a fertilized ovum which develops into a ciliated

larva called a miracidium. The rate of miracidial development

is dependent on temperature and oxygen availability and can

take from 10 days to several months. The critical temperature

below which egg development ceases is 9.5C and the thermal

constant for the optimal temperature range for egg development

is between 200 and 220 day-degrees, the values obtained when

eggs are incubated at 23"C and 180C respectively. The maximum

temperature that eggs can be exposed to and still survive is

not known although temperatures above 300C increasingly

inhibit development. Additional requirements for successful

development and hatching of eggs include separation from the

feces, and a film of moisture on the egg surface during the

entire developmental period from deposition onto pasture until

hatching of the free-swimming, ciliated miracidia (Rowcliffe

& Ollerenshaw, 1960). Once the miracidium is fully developed,

exposure to sunlight stimulates it to hatch, although the

precise mechanism of hatching has not been proven. Roberts

(1950) demonstrated that violet and blue wavelengths of the

light spectrum are an essential part of the light stimulus.

Rowan (1956) suggested that the operculum is cemented to the

shell and that the miracidium releases an enzyme at the time

of hatching that digests this cementing substance. Wilson

(1968) refuted this explanation and suggested that the

miracidium causes an alteration of the permeability of the

membrane surrounding the viscous cushion. This change in


permeability causes an expansion of the cushion which

compresses the miracidium, leading to an increase in the

internal pressure within the egg until the operculum finally

ruptures. Both studies demonstrated that the escape of the

miracidium from the open egg is due primarily to the

hypertonicity of the egg contents which forcefully expels the

miracidium, and only secondarily to muscular activity.

After hatching, the miracidium has only a few hours in

which to find and penetrate a suitable snail host. During the

process of penetration, the miracidium loses its ciliated

covering and becomes a sporocyst. The sporocyst, which has a

rudimentary digestive tract and is filled with germinal cells,

is usually found in the wall of the mantle cavity, the wall of

the pulmonary chamber, or in the tissue around the esophagus

(Kendall, 1965). The germinal cells within the sporocyst

develop into rediae, the next larval stage. Within 2 to 3

weeks the mature rediae leave the sporocyst by rupturing the

wall and migrate to the digestive gland (hepato-pancreas).

Rediae may then produce either a daughter generation of rediae

or cercariae depending upon various environmental stimuli.

Stresses such as snail host starvation and high or low

temperature tend to stimulate daughter redial production

(Wilson & Draskau, 1976). The tadpole-like cercariae leave

the rediae through a birth pore and enter into the

perivisceral space before emerging from the snail. Cercarial

emergence is a mostly passive process caused by an increase in


pressure in the perivisceral spaces. This rise in pressure

results from contractions of the walls of the mantle cavity

which occur with increased snail activity. Immersion of the

snail in fresh water appears to be the principal factor

governing this process (Kendall and McCullough, 1951). This

has important epidemiologic implications since cercariae will

not be shed if the environment is dry when dispersion and

survival of the cercariae would be unlikely. Cercariae have

been observed to emerge in large numbers following rain that

was preceded by a dry period (Boray, Happich, & Andrews,

1969). Infection of a snail with a single miracidium can

result in the production of hundreds of cercariae and this

reproductive process greatly contributes to the success of

this parasite. The rate of development of these

intramolluscan stages is dependent upon temperature and the

nutritional state of the snail (Kendall, 1965). Under optimal

conditions, parasite maturation within the snail takes

approximately 5 to 7 weeks.

The free-swimming cercariae loses its tail after contacting

vegetation, secretes a protective cyst covering, and completes

its development during the first 2 or 3 days after encystment

(Malek, 1980). This stage, the metacercariae, contains a

fully developed immature fluke and is the infective stage of

the parasite. Ruminant hosts become infected primarily by

ingesting the metacercarial cysts on forage, but they also can

become infected by ingesting cysts suspended on soil and


detritus while drinking contaminated water. The length of

time that metacercariae survive on pasture is dependant on

environmental factors. Moisture is the principal factor

controlling the length of life of metacercariae, with a

minimum of 70% relative humidity considered necessary for

prolonged survival. Metacercariae were killed within two days

of exposure to temperatures of 98.6 to 105F when exposed in

air to direct sunlight, however when placed in water kept at

room temperatures ranging from 71.6 to 80.6F they survived

for 4 months. In Great Britain, metacercariae shed onto

pasture in the autumn remained infective for periods of

between 270 to 340 days and metacercariae shed onto pasture

during the summer remained viable for up to 180 days (Kendall,

1965). Under the hot and dry pasture conditions of coastal

Texas during the summer, metacercariae were rapidly killed

(Olsen, 1947), however under conditions of high humidity such

as exist in Louisiana during the summer, metacercariae may

survive for extended periods (Malek, 1980). Once ingested by

a ruminant host, the metacercariae excysts releasing a

juvenile fluke. Excystment is an active process that occurs

in 2 phases: (1) the metacercariae becomes activated in the

rumen and; (2) after passing into the small intestine and

contacting bile, the juvenile fluke escapes through a small

hole on the ventral surface of the cyst wall (Dixon, 1966).

The juvenile fluke penetrates the wall of the small intestine,

migrates through the peritoneal cavity over a week's time, and


then penetrates through the liver capsule. Juvenile flukes

migrate through the hepatic parenchyma for about 6 to 8 weeks

before entering the bile ducts where they mature. Egg

production can begin as early as 8 weeks post-infection (de

Leon, Quifones, & Hillyer, 1981), however most infections do

not become patent until after about 11 to 12 weeks (Ross,

Todd, & Dow, 1966). Thus, completion of the entire parasite

life cycle, from the time an egg is shed onto pasture until a

newly infected animal reinfects the pasture with the next

generation of fluke eggs, requires 16 to 24 weeks.

Biology of Fossaria cubensis

Since the population dynamics of the lymnaeid snail

intermediate host is fundamental to fluke transmission,

understanding the biology of the snail is also important. In

the United States, Fossaria cubensis and Fossaria bulimoides

are the intermediate hosts of primary importance (Malone,

1986), however several other lymnaeid snail species including

Pseudosuccinea columella and Fossaria modicella may also serve

as hosts (Krull, 1934). In Europe and throughout much of the

rest of the temperate enzootic range of F. hepatica, Lymnaea

truncatula, is the principal snail intermediate host. Several

other lynmaeid snail species of similar morphology and biology

to Fossaria spp. and L. truncatula also serve as intermediate

hosts for F. hepatica in various regions of the world (Malek,

1980). These snails are small (<10mm), "right handed" snails


that can be recognized by the dextral whorl of the shell and

are semi-aquatic or amphibious in nature, being found most

often in shallow water during the cooler months and on wet mud

during the warmer periods of the year. These snails prefer

water that is medium in chlorides (60 ppm), medium in calcium

(21 ppm) and has a pH of 7.5 to 8.5 (Batte & Swanson, 1951).

Lymnaeid snails have the ability to survive adverse times

of the year by hibernation during the winter in cold climates

or by a similar process called aestivation during hot and/or

dry periods. At such times they burrow into the mud and enter

into a state of reduced metabolic activity. Snails that

survive the adverse times of the year via these processes,

reemerge when environmental conditions improve (spring in cold

climates, fall in warm climates) forming the beginnings of the

next generation. If snails are infected with F. hepatica when

they enter into the hibernation or aestivation period, the

parasite will also cease development until the snail

reemerges. These infected snails thus provide a limited means

of seasonal carryover of fluke infection.

Lymnaeid snails are hermaphroditic, therefore the entire

sexually mature population is capable of producing fertile

eggs (Olsen, 1944). Fossaria bulimoides attain sexual

maturity when the shell length reaches 4.5 mm, and under

optimal temperature and nutritional conditions, snails can

grow to this size as soon as 14 days after hatching. These

snails can produce from 2 to 3 egg masses per day containing


on the average 17 eggs. Once snails begin to lay eggs they

can continue to do so for 3 to 7 months, which is probably

close to their entire life span under natural conditions.

Four laboratory raised snails produced an average of 5,112

eggs each over their lifetimes (Olsen, 1944). These features

allow for tremendous increases in the density of snail

populations over relatively short periods of time when

environmental conditions are favorable. Snails are also

capable of colonizing new areas by passive lateral migration

from runoff of high water following periods of heavy or

prolonged rainfall. Under such flooded conditions, snails

often rise to the top and cling to the underside of the water


Clinical Signs and Pathology

There are many similarities between cattle and sheep in the

pathophysiology of fascioliasis but there are also some

important differences. Acute disease is much more common in

sheep than cattle and results from the extensive damage to the

hepatic parenchyma caused by the migrating juvenile flukes.

Chronic disease on the other hand results from a combination

of the partially resolved hepatic damage that follows the

acute phase and the blood sucking activities of the adult

flukes within the bile ducts. In both species, chronic

fascioliasis is the more common form. Black disease

(infectious necrotic hepatitis) and bacillary hemoglobinuria


due to Clostridium novyi and C. hemolyticum, respectively,

also can be complications of fascioliasis. The distribution

of these two diseases parallels the distribution of F.

hepatica (Jubb & Kennedy, 1970).

In cattle, infection with liver flukes usually causes no

clinical signs and in general can be considered a subclinical

disease. However, where extremely high fluke burdens rapidly

accumulate and/or nutritional stress or other concurrent

disease complicate otherwise subclinical infections, outbreaks

of acute or subacute bovine fascioliasis can occur (Malone,

Smith, Loyacano, Hembry, & Brock, 1982b; Reid, Doyle, Armour,

& Jennings, 1972; Ross & Dow, 1966). Clinical disease

resulting from chronic infection is most frequently seen in

cattle under 2 years old and is characterized by weight loss,

anemia, hypoproteinemia, eosinophilia, general depression and

occasionally death (Armour, 1975). Concurrent infections with

Ostertagia ostertagi can complicate the clinical presentation.

During the parenchymal phase of fluke migration in cattle,

tissue regeneration is minimal and fibrosis is marked (Ross et

al., 1966). In response to the traumatic injury caused by the

flukes, tracts of coagulative necrosis develop which result in

a diffusely fibrotic hepatic parenchyma containing hemorrhagic

streaks and foci. In the bile ducts, the adult flukes produce

a mechanical irritation which causes cholangiohepatitis. This

leads to dilation, thickening and extensive fibrosis of the

duct wall resulting in stenosis and calcification. This


biliary response is believed to be at least partially

responsible for the short life span (less than 1 year) of the

liver fluke in cattle.

In sheep, fascioliasis can occur either as an acute or

chronic disease depending upon the pattern and level of

infection. Acute disease occurs when large numbers of flukes

are acquired over a short period of time. Unlike the response

in cattle, during the parenchymal phase of the juvenile fluke

migration in sheep, liver reaction is partially regenerative,

fibrosis is minimal and parenchymal destruction is extensive.

When large numbers of parasites are present, a critical period

develops in which extensive hemorrhage occurs, causing the

death of the host from acute fascioliasis (Dow, Ross, & Todd,

1968). Clinical signs include anorexia, depression and

weakness. Sudden death may occur within 48 h of the

appearance of signs and deaths may continue for 2 to 3 weeks

(Wescott & Foreyt, 1986).

Chronic fascioliasis in sheep presents with similar but

usually more pronounced clinical symptoms as those seen in

cattle. Some deaths may occur but most animals will survive,

albeit, in poor condition. The biliary reaction to the fluke

is minimal and ducts expand to contain the parasites.

Although some fibrosis of the duct wall does occur, the duct

wall remains fairly pliable and calcification is not a

feature. In this biliary environment flukes can survive for

prolonged periods (Dow et al., 1968).


Host Immunity

Sheep acquire very little immunity to infection with F.

hepatica evidenced by the fact that sheep develop only minimal

fibrosis and no calcification of their biliary ducts and

flukes have been reported to survive for up to 11 years. As

a result, liver fluke infections are persistent, additive and

essentially unimpeded by host response. This is largely

responsible for the extreme pathogenicity of F. hepatica in

this species (Boray, 1969). Cattle on the other hand develop

a moderate degree of resistance to this parasite. The life

span of flukes in cattle is quite variable and related to the

pattern and intensity of infection (Ross, 1968). In low level

single experimental infections (200 metacercariae), 75% of

flukes that became established were lost between the 5th and

21st month after infection. In high level experimental

infections (2,500 to 15,000 metacercariae), the life span was

reduced to 6 to 7 months. Where challenge infections were

involved the life span was reduced to 4 to 5 months and an

"acquired self cure" was observed in that there was a turnover

of the adult population. Ross (1968) concluded that the life

span of the parasite in the bile ducts of cattle is directly

related to the number of parasites present at any one time and

to the intensity of the reaction that they produce in the bile


Doyle (1971) also studied the immune response of cattle to

infection with F. hepatica but in greater detail. He found


that calves eliminated 85% of flukes derived from an initial

experimental infection of 750 metecercariae between 16 and 30

weeks after infection. Additionally, a high level of

resistance was seen in challenge infections where only 16% of

the fluke burden found in control calves was recovered from

calves which had been reinfected. Of the 2 groups of calves

that were administered a challenge infection, 1 group was

first treated with flukicide to remove the initial infection

and the other was left untreated. Although the total number

of flukes recovered from these 2 groups was similar, in the

untreated group the flukes were largely derived from the

initial infection indicating that these calves had resisted

the establishment of the new population more successfully than

did the treated group. In both of these studies an "acquired

self cure" was demonstrated beginning approximately 4 months

following initial infection. However, results of the

challenge experiments in these 2 studies contradict each

other. Whereas Ross reported that a new population of flukes

replaced the old following challenge, Doyle reported that the

old flukes persisted and an acquired immunity prevented the

new population from becoming established. It is likely that

differences in experimental design and the age and breeds of

calves used were responsible for these differences.

From a practical standpoint, the fact that cattle exhibit

a partially protective immune response is far more important

than the particular details and differences of these studies.


It is clear that a dynamic relationship exists between the

host animal and the parasite. The interaction of factors such

as the age of the host, the innate resistance of the host, the

previous exposure of the host and the present level of

parasite exposure will determine the level of immunity, the

degree of parasite establishment and the pathologic impact of

the infection. Older cattle with previous exposure will have

a greater resistance to infection than young parasite naive

calves. Additionally, flukes are gradually eliminated over

time so that most flukes acquired during the major

transmission period of one year will be lost prior to the same

time the following year. Fluke burdens in cattle are

therefore not cumulative as they are in sheep. These issues

should be considered when designing control programs for


Economic Importance

Economic losses from Fasciola hepatica result directly from

increased death losses and liver condemnations at slaughter

and indirectly from decreased livestock productivity.

Although direct losses are easier to measure, indirect losses

are considered to be far more economically important (American

Association of Veterinary Parasitologists, 1983). Beef

producers are affected by increased culling of cows, reduced

sale weights of culled cows, lowered reproductive performance

in the brood cow herd, and reduced calf weaning weights.

Economic losses in feedlots result from reduced feed-

conversion ratios and lowered average daily gains, and fluke-

infected dairy cows produce less milk (Hope Cawdery,

Strickland, Conway, & Crowe, 1977; Randell & Bradley 1980;

Malone, 1982; Simpson, Kunkle, Courtney, & Shearer, 1985).

Precise economic benefits of liver fluke control are

difficult to quantify due to the interactions of physiologic,

nutritional, associated disease, and climatic/geographic

factors (Simpson et al., 1985). The interaction of these

factors causes tremendous variation in pasture infectivity and

the physiologic consequences of infection from year to year

and ranch to ranch. As a result, there is little consistent

and well-documented data on the benefits of liver fluke

control. However, by piecing together data from several

studies it is clear that the economic benefits of controlling

liver flukes are substantial.

Significant production losses may occur in cattle herds

when the prevalence of infection is greater than 25% (Malone,

1986). Subclinical infections averaging 54 flukes per calf (8

to 9 months old at the time of infection) reduced weight gains

by 8% over the first 6 months of infection. Higher levels of

infection (average of 140 flukes per calf, 14 to 15 months

old) reduced weight gain by 28% and caused the appearance of

clinical signs in some animals. Although impaired weight

gains were demonstrated for up to 6 months, the majority of

losses occurred during the first 4 months. Significant

production losses did not occur in infections of greater than

6 months duration (Hope Cawdery et al., 1977). In cow-calf

studies in Louisiana, calves from groups of cows that were

treated for liver flukes approximately 6 and 12 months earlier

had a 10.1 kg advantage in 205-day adjusted weaning weight as

compared to calves from groups of untreated cows.

Additionally, several studies on the performance of fluke

infected feedlot calves demonstrated that average daily gains

were increased by an average of 9.5% by flukicidal treatment

(Malone et al., 1982a). Dairy cows treated for liver flukes

had a significant increase in average daily milk yield (+ 4.2

kg/day, 90-120 days after treatment) over daily yields of the

previous lactation. Nontreated control and uninfected treated

cows did not show significant gains over yields of the

previous lactation (Randell & Bradley, 1980).

Data obtained from a large survey of cattle producers in

Florida recently showed that specific benefits from control of

liver flukes through appropriate treatment of cattle included

18-22 lb. heavier cull cows, 1-3 percent more calves, and 30-

45 lb. heavier calves at weaning, yielding a net return to the

producer of $15.19-$31.03 per brood cow, depending upon the

size of the calf crop and calf prices (Simpson & Courtney,

1990). Based upon liver condemnations at slaughter, approxi-

mately 64% of beef cows in peninsular Florida are infected

with liver flukes, thus proper fluke control applied to these

one million plus beef cows has the potential to increase net


income to the cattle industry in Florida by 10 to 20 million

dollars annually. A separate economic analysis of liver fluke

control in Florida reported benefit-cost ratios ranging from

4:1 to 16:1 depending upon the number of treatments given and

the range of estimates included in the analysis of production

parameters (Simpson et al., 1985). A more recent study

performed in central Florida on a large cow-calf herd reported

that treatment with clorsulon showed a positive net return

regardless of the price of calves and cows (Simpson, Greiner,

& Richey, 1989). Cows treated with clorsulon had a net

positive return that ranged from $46.96 for calf and cow

prices of $0.60 and $0.35 per pound respectively, to $72.84

for calf and cow prices of $0.90 and $0.50 per pound

respectively. Although Florida may have a more serious

problem with liver flukes than most other states, these

figures give an estimation of the economic impact which liver

flukes can have on the cattle industry in highly enzootic



Fecal sedimentation remains the standard method for the

diagnosis of fluke infections in individual animals. However,

this technique is labor intensive (requiring 20 to 30 minutes

per sample) and it can have poor sensitivity. Problems with

sensitivity are due to: (1) the low egg output which is

typical of fluke infected adult cattle; and (2) excessive


fecal debris in samples which may obscure the eggs and prevent

detection. A modification of the sedimentation technique

(Flukefinder*, Visual Difference, Moscow, ID) considerably

improves both the sample processing speed and the sensitivity

of egg detection. The technique utilizes a two sieve system

which filters most of the fecal debris leaving a relatively

clean sample which can then be sedimented and read. When

examining fecal samples it is important to differentiate eggs

of liver flukes (F. hepatica) from those of rumen flukes

(Paramphistomum spp.). Rumen flukes are not affected by

available flukicides and therefore eggs of these flukes are

frequently present in greater numbers than those of liver

flukes. Infections with the deer fluke, Fascioloides magna,

do not become patent in cattle, and therefore cannot be

diagnosed by fecal examination.

Several factors need to be recognized when interpreting

fecal sedimentation results. As occurs with most parasitic

diseases, flukes are over-dispersed within the cattle

population, resulting in a situation where a small percentage

of animals will carry the greatest fluke burdens and therefore

shed the most eggs. This causes a large variation in the

number of eggs shed by different animals grazing the same

pasture. Additionally, since most fluke infected cattle shed

relatively few eggs (less than 5 per gram even in heavily

infected herds) (Malone & Craig, 1990), a minimum of 10

samples should be examined before clinical impressions of herd


prevalence are made. Other factors that must be considered

are the long prepatent period of liver flukes and the

variation in seasonal egg production that occurs depending

upon transmission patterns and the duration of the infection.

Enzyme-linked immunosorbent assays (ELISA) have been

developed for the serologic diagnosis of liver flukes, but

none are commercially available presently. Diagnostic

laboratories at veterinary colleges in Texas, Louisiana,

Oregon and Washington have used several versions of these

tests on trial bases but none proved sensitive or specific

enough to improve on diagnosis by fecal sedimentation in

individual patients (Malone & Craig, 1990). Although these

tests have not been reliable for determining the infection

status of individual cattle, they have potential usefulness in

screening groups of animals such as stocker calves arriving at

feedlots or young cattle on pasture suspected of harboring

immature infections. Improvements in these tests continue to

occur so it is likely that new versions could become

commercially available in the future.

Seasonal Transmission Dynamics of Fasciola Hepatica in the
United States

The seasonal transmission profile for liver flukes in any

given locale depends upon a series of interactions between the

biology of the parasite and the environment. Within this

framework of interactions are two essential requirements for


the completion of the fluke life-cycle: (1) adequate moisture

to sustain populations of the semi-aquatic lymnaeid snail

intermediate host for prolonged periods; and (2) temperatures

above 10C, since below this temperature both parasite and

snail development ceases (Ollerenshaw & Rowlands, 1959). The

upper limits of temperature in which the parasite and snail

can survive has not been well established. Since conditions

necessary for completion of the fluke life cycle occur at

different times of the year in each climatic/geographic

region, the seasonal transmission patterns will also be


Transmission of F. hepatica in the Gulf Coast states occurs

primarily during the winter and spring. In Louisiana, most

fluke transmission occurs between the months of February and

July (Malone, Loyacano, Hugh-Jones, & Corkum, 1984/85). A

very similar pattern probably occurs in Texas as well (Craig

& Bell, 1978). In north central Florida, fluke transmission

occurred almost exclusively between the months of December and

June, with the peak months being February to April (Boyce &

Courtney, 1990). In Louisiana, this transmission pattern is

directly related to yearly temperature and precipitation

patterns. Soil moisture recharge begins in the autumn and

snail populations increase throughout the late autumn and

winter. By February, infected snails are shedding large

numbers of cercariae. So long as rainfall is sufficient to

maintain surplus soil moisture, snails will remain and cattle


will become infected with flukes. Fluke transmission ceases,

however, with the first sustained drought of the summer since

snails die or aestivate and metacercariae are rapidly killed

(Malone et al., 1984). Water deficits tend to persist

throughout the summer and early autumn, therefore snails

remain in estivation until the autumn rains begin, usually in

November. Similar seasonal climatic patterns are probably

responsible for the transmission profile seen in Texas (Olsen,

1947; Craig & Bell, 1978). Climatic parameters responsible

for the transmission profile demonstrated for northern Florida

(Boyce & Courtney, 1990) have not been determined.

In southern Idaho, where the cold climate limits the

pasture grazing season from April to November, fluke

transmission occurred from June to November (Hoover, Lincoln,

Hall, & Wescott, 1984). This transmission profile is similar

to that of northern Europe (Armour, 1975; Ross, 1977; Shaka &

Nansen, 1979). Summers in Idaho are semi-arid, therefore

flood irrigation at regular intervals is routinely performed.

This practice provides sufficient moisture for transmission to

occur and paradoxically, it is the shallow irrigation ditches

which provide the best snail habitat. It is likely that other

areas of the cold temperate northwest United States have a

similar transmission profile, however there are no other

published reports to confirm this. More importantly, there

are no published reports on seasonal transmission from any of

the warmer areas of the western United States.


Rationale for Seasonal Control

To develop a rational seasonal control program for liver

flukes in a given area, it is important to understand local

transmission dynamics, fluke biology, seasonal pasture forage

availability, local ranch management practices and the

variable efficacy of available flukicidal drugs against the

different age classes (juvenile, immature, mature) of F.

hepatica. Currently, only two drugs are available in the

United States for the treatment of liver flukes: clorsulon

(Curatrem, Ivomec-F; Merck Sharp & Dohme Research

Laboratories); and albendazole (Valbazen; SmithKline Beecham

Animal Health).

At the labeled dosage (7 mg/kg), Curatrem, which is

administered as an oral drench, has the best efficacy of these

three products against adult stages (>99%) and will also kill

immature bile duct stages (8-12 weeks) with fairly high

efficacy (85 to 95%) (Kilgore, Williams, Benz, & Gross, 1985;

Malone, Ramsey & Loyacano,1984; Malone, Williams, Lutz, Fagan,

Jacocks, Jones, Marbury, & Willis, 1990). This product will

only kill flukes, however, and has no effect on other

parasites. Ivomec-F, which is administered by subcutaneous

injection, is a combination product which is formulated to

deliver a full therapeutic dose of Ivomec (200 ug/kg) plus a

reduced dose (2 mg/kg) of clorsulon. The reduced subcutaneous

dose of clorsulon in Ivomec-F has the approximate biological

equivalence of a 3.5 mg/kg oral dose of Curatrem. This dose


has about 97 to 99% efficacy against adult flukes (Kilgore et

al., 1985; Wyckoff & Bradley, 1983; Zimmerman, Wallace,

Schons, & Hoberg, 1986) but is not very effective against

immature stages (Malone et al., 1984). The addition of

Ivomece makes this a broad spectrum product that will kill

nematodes and arthropod parasites in addition to flukes.

Valbazene is a broad spectrum product, administered by oral

drench, which kills flukes, nematodes and cestodes. At the

labeled dosage (10 mg/kg), the efficacy of albendazole against

adult flukes has been variable (76 to 92%) and the efficacy

against immature stages is poor (Craig, Qureshi, Miller, Wade,

& Rogers, 1992; Kilgore et al., 1985; Malone et al., 1982b).

However, at higher doses (:15 mg/kg), the efficacy of

albendazole approaches that of clorsulon (Ronald, Craig, &

Bell, 1979). None of these three flukicidal products will

kill the migrating juvenile stage (<8 weeks), (Richards,

Bowen, Essenwein, Steiger, & Buscher, 1990) therefore the

limited efficacy of these drugs against juvenile and immature

stages of F. hepatica must be taken into account when making

decisions regarding the timing of treatments.

Flukicidal treatment is most effective if given when

pastures are devoid of metacercariae and snails, and flukes

infecting cattle are mature. This occurs during the late

winter-early spring in northern climates and during the late

summer-early autumn in southern climates. At these times

virtually all the flukes in the pasture biota are concentrated


in the ruminant hosts as mature stages. Therefore, flukicidal

treatment will not only kill most of the flukes in the cattle

but will also reduce the potential for fluke transmission to

snails when they emerge from hibernation or estivation.

Decisions regarding treatment of liver flukes must be made

as part of an overall herd health management program and

cannot realistically be done without accounting for other

management factors such as stocking rate, pasture management

and forage availability. Although animals will usually

benefit from treatment regardless of other factors, treatment

is most valuable in situations where cattle are also suffering

from nutritional stress or some other production limiting

disease. In the gulf coast states, most cattle are maintained

year round on extensively managed open pastures. During the

winter months, forage quality and availability is poor and

most cows will lose substantial weight during this period even

with supplementation. This is also the time of year when

parasitic gastroenteritis can be an important health problem.

Under these conditions fluke infections will have their

greatest impact, thus treatment programs should aim to reduce

fluke burdens during this time.

Current Treatment Recommendations for the Gulf Coast States

Fluke transmission in the gulf coast region ceases during

summer because snail intermediate hosts burrow into the mud

and aestivate, thus contamination of the pasture with


metacercariae ceases. Fluke control recommendations for beef

cattle in the Gulf coast states are currently predicated on

the occurrence of this summer gap in fluke transmission.

Cattle on ranches having known endemic fluke infections should

be treated with a flukicide once annually in the early autumn

with an optional spring treatment based on local parameters of

risk. The rationale behind this treatment scheme is as

follows. Flukes are presumed to survive the summer only as

adult flukes in the livers of cattle. Fluke eggs shed in the

manure of cattle during the summer will be killed by the

persistently high summer temperatures. Pastures will be

cleansed of metacercariae shed in the spring by summer's heat

and metacercariae are not replaced after snails enter summer

aestivation. Finally, very few fluke infected snails are

presumed to survive aestivation due to the combined stresses

of both aestivation and fluke infection. Flukicidal treatment

of cattle in the early autumn will prevent the shedding of

large numbers of fluke eggs onto pasture when the essentially

fluke-free snails emerge from summer aestivation with the

onset of cooler weather. Since newly emerged snails and their

subsequent offspring are not likely to become infected if

cattle have been treated, fluke transmission to cattle is

greatly reduced during the following winter and spring. As an

added benefit, virtually all flukes in cattle in the autumn

will be mature adults and therefore fully susceptible to



The optional spring treatment is primarily palliative

rather than curative, because many flukes would be immature at

that time of year and thus not susceptible to flukicides. Its

purpose is to reduce the fluke burdens carried over the summer

by cattle during high risk years or on high risk properties

(Malone, Williams, Muller, Geaghan, & Loyacano, 1987). Since

some flukes will survive spring treatment, this treatment

should be given in addition to, but never in place of an

autumn treatment.

Current Treatment Recommendations for the Northwestern States

Fluke transmission in the northwestern U.S. ceases in

December because temperatures are too low for parasite

development and cattle are removed from pasture. Since most

fluke transmission occurs from September to November, a single

treatment should be given in the late winter or early spring.

At this time all flukes will be mature and fully susceptible

to flukicide treatment. By eliminating flukes from the cattle

at this time, contamination of the pasture with eggs will be

minimized in the spring when cattle are returned to pasture

and snail populations return. Few infected snails,

metacercariae, or eggs will survive the winter and do not seem

to have an important epidemiological role in fluke

transmission in this region. Autumn treatment is likely to be

of little benefit because most flukes will be immature at this

time and not susceptible to flukicides. Therefore, a single

annual treatment should never be given only in the autumn.

Although a twice yearly (early spring/late autumn) treatment

program could be used, because the fluke transmission cycle in

cold climates is relatively short, a single late winter-early

spring treatment should be satisfactory in most cases.

Epidemiology and Control of Bovine Fascioliasis in Florida:
Review of Literature. Research Needs and Implications for
Improved Control Recommendations

Swanson (1949) reported that the lymnaeid snails

Pseudosuccinea columella Say and Fossaria cubensis Pfeiffer

were the snail intermediate hosts of F. hepatica in Florida.

These same snails were also reported to serve as intermediate

hosts for the rumen flukes Paramphistomum cervi Schrank and

Cotylophoron cotylophorum Fish which are commonly found in

cattle raised in F. hepatica enzootic areas. In 1947 and

1948, economic losses resulting from condemned fluke-infected

livers at the large slaughterhouses in Florida amounted to

over $100,000. This value did not include losses of condemned

livers in many small local butcher establishments throughout

the state, nor did it include death or production losses

(Swanson, Batte, & Dennis, 1952).

In 1952 liver fluke disease was known to be present in 25

of the 67 counties of the state. River tributaries, lakes and

ponds were noted as constant sources of infection, especially

if lime rock or marl-type soils were present. Seeping banks,

springheads, bay heads and lands that were heavily limed also

were areas where snail intermediate hosts could be found.

Cypress lowlands and piney woods areas were noted to be too

acidic for snail development and these areas were therefore

not important in fluke transmission (Swanson et al., 1952).

Water from artesian wells was noted to create ideal breeding

grounds for snails. These free-flowing wells also provide

snails with areas of refuge where they can survive during

periods of drought (Batte & Swanson, 1951).

Recommendations for control of fascioliasis were based on

a program which included snail eradication through the use of

pasture management and moluscicides, and treatment of cattle

with hexachlorethane. Ranchers were advised to shut off

artesian wells; drain low-lying wet areas using a V-type ditch

cut; and where it was impractical to drain ponds, swamps or

other wet areas, then those areas should be fenced off from

cattle. Copper sulfate (bluestone, blue vitriol) was

recommended as the moluscicide of choice to be applied at the

rate of 20 pounds per acre if spread over pasture or 24 pounds

per cubic foot per second of water flow if treating streams.

After establishing a program of snail eradication, ranchers

were advised to treat all cattle over 4 months of age with

hexachlorethane at a dosage of 10 g/100 lbs. body weight.

This treatment was to be repeated in 21 days to kill the

flukes that were immature at the time of the first treatment

and therefore unaffected by the drug (Swanson et al., 1952).

Proper timing for this treatment or for the application of

moluscicide was not addressed. Apparently, no research into

the seasonal transmission dynamics of liver flukes in Florida

was performed and therefore the most beneficial time for

performing these treatments was not known.

From June 1984 to June 1987, 37 groups of four fluke-free

ewes each grazed in monthly succession a 0.25 ha pasture in

order to determine the seasonal transmission dynamics of F.

hepatica in north central Florida (Boyce & Courtney, 1990).

The pasture, located on the University of Florida campus in

Gainesville, was known to support populations of both P.

columella and F. cubensis. At the end of each month, that

month's group of ewes was held in a fluke-free pen for 2

months, necropsied, and the flukes recovered and counted.

Fluke transmission was found to occur from December to June

with February, March and April as the peak months. This

seasonal pattern was repeated with only minor variation over

3 years. No summer transmission occurred in any of the 3

years: in 1986 and 1987 transmission ceased in May and in 1984

and 1985 transmission ceased in June. Fluke transmission

resumed during the late autumn or early winter: November

(1986), December (1987), or January (1985).

This pattern of transmission was very similar to that

reported in Louisiana and Texas (Craig & Bell, 1978; Malone et

al., 1984/85), the other states of the Gulf coast where F.

hepatica is highly enzootic. This gave the researchers

confidence that this was the true transmission pattern for


north central Florida, although it was noted that data

collected from a single study site may not be typical of an

entire region. This may be particularly true for Florida,

since northern Florida is in a warm temperate climatic zone

whereas southern Florida is in a subtropical climatic zone.

However, with no other data to base decisions on,

veterinarians at the University of Florida have made fluke

treatment and control recommendations for Florida cattle

predicated on the absence of summer and early autumn

transmission (Courtney, Shearer, & Plue, 1985; Shearer et al.,

1986). It was acknowledged, however, that information on

liver fluke transmission dynamics was desperately needed for

southern Florida where 66% of Florida's beef cows are


Recently, a climate forecast model was developed for

prediction of relative fluke risk in different years, and thus

whether there is a need to treat cattle once or twice annually

(Malone et al., 1987; Malone, Williams, Loyacano, & Muller,

1989). This forecast model also predicts the seasonality of

fluke transmission for a given region. Although not fully

verified for Florida, this forecast model correctly predicted

in a retrospective study (Malone & Courtney, unpublished data)

the transmission profile (i.e., the seasonality and intensity

of fluke transmission) that had been reported by Boyce and

Courtney (1990) in north central Florida. This same model,

however, predicts that there may be two transmission seasons

(winter and summer) in subtropical southern Florida (Malone &

Craig, 1990).

If fluke transmission in subtropical southern Florida

occurs mostly in the winter and summer then treatment of

cattle in early autumn and spring would be ill advised.

Therefore, it is of extreme importance to the cattle industry

of southern Florida to determine the seasonal transmission

dynamics of F. hepatica in this region. Additionally, the

seasonal transmission dynamics should be studied in central

Florida to ascertain any differences that may exist in this

transitional area.

The seasonal transmission dynamics of fascioliasis is often

established for a particular area through the monthly

slaughter of sentinel animals grazing known fluke-infected

pastures, but this is an expensive process. Since transmission

of F. hepatica closely follows the availability of its snail

intermediate hosts, it is also possible to measure seasonal

transmission of F. hepatica indirectly by studying the

population dynamics these snails. The latter approach was

taken in this project because it has the advantage of allowing

the investigation of more sites at a far lower cost than can

be done using sentinel animals. In addition to studying snail

population dynamics, a DNA probe assay was developed to

determine the infection prevalence of snail intermediate hosts

with larval stages of F. hepatica. To determine whether

runoff from an artesian well would alter the snail population


dynamics as compared to other snail habitats, one pasture was

selected for study which had a free-flowing artesian well.

Fluke egg survival during the summer was also studied, since

summer transmission could only occur if eggs were able to

survive and hatch during periods of extreme heat.



Climate is the primary factor affecting seasonality of

transmission for F. hepatica (Ollerenshaw & Rowlands, 1959).

In cool-temperate areas of the world such as the northwestern

United States, Ireland and Great Britain, liver fluke

transmission occurs mostly during the summer and autumn

(Armour, 1975; Hoover et al., 1984; Ross, 1977), whereas in

warmer areas such as the Gulf coast region of the United

States, transmission occurs primarily during the winter and

spring (Boyce & Courtney, 1990; Craig & Bell, 1978; Malone et

al., 1982a). Liver flukes in cool and warm climates will

therefore face disparate environmental pressures for survival

and successful transmission and natural selection will favor

those parasites that successfully adapt to the set of

environmental parameters which they face. Therefore, it is

possible that data on transmission dynamics and bionomics of

F. hepatica from a cool temperate climate may not accurately

reflect the transmission dynamics and bionomics that exist in

a tropical or subtropical climate.


Rowcliffe and Ollerenshaw (1960), determined the conditions

necessary for the survival, development and hatching of eggs

of F. hepatica in a series of experiments using eggs recovered

from fluke-infected sheep in Great Britain. Great Britain has

a cool temperate climate with maximum temperatures rarely

exceeding 25C. On the contrary, the areas of Florida where

the majority of cattle are pastured and where F. hepatica is

highly endemic is subtropical. In this region, daily high

temperatures in the summer months average 33 to 34C, with

daily mean temperatures of 27 to 28"C. Rowcliffe and

Ollerenshaw (1960) found that when eggs were stored in water

free of fecal matter above 30C, temperature increasingly

inhibited development and mortality reached 100% when eggs

were stored at 370C for 24 days. In experiments where eggs

were incubated in the presence of sheep feces at 100% humidity

at temperatures of 18 or 270C, mortality reached 100% by 24

and 13 days respectively. Eggs in sheep feces exposed to

normal fluctuations of rain and drying during the summer

reached greater then 90% mortality within 17 days and 100%

mortality by 35 days.

Internal temperatures of cattle fecal pats deposited on

open areas of pasture rise and fall in a diurnal pattern in

accordance with air temperature and amount of solar radiation.

In the early morning, air and fecal pat temperatures are the

same, however, by mid-afternoon, fecal pat temperatures exceed

air temperatures. In pilot studies, we frequently recorded


fecal pat temperatures in mid-afternoon of greater than 37C

with temperatures exceeding 40C on some days. Although these

high temperatures were not sustained for long periods, clearly

the eggs in these fecal pats would frequently experience very

hot temperatures (>35C) for several hours and moderately warm

temperatures (>270C) for long periods of each day.

If eggs of F. hepatica in Florida have similar temperature

limitations for survival and development as those in Great

Britain, it seems highly unlikely that eggs on pasture would

survive long enough to develop and hatch during the summer.

However, if eggs of F. hepatica have adapted to the hot

climate and can survive and hatch during the summer, then this

would have important epidemiological implications. We have

determined that the snail intermediate host for F. hepatica in

Florida, Fossaria cubensis, is rarely present on pasture

during the summer months in Florida. A dry period in the late

spring (April and May) of most years usually causes snail

habitats to dry and snails to aestivate. Snail populations do

not reappear until the autumn even though snail habitats

become wet once again when summer rains begin in June.

In most years, therefore, liver fluke transmission ceases

during the in Florida summer because no new metacercariae are

shed onto pasture once snails aestivate and metacercariae are

rapidly killed under hot and dry conditions (Olsen, 1947).

Fluke control recommendations for beef cattle in the Gulf

coast states are currently predicated on the occurrence of

this summer gap in fluke transmission (Kaplan, 1994; Malone et

al., 1982a). Flukicidal treatment of cattle in the early

autumn will kill virtually all of the adult flukes thus

preventing the shedding of large numbers of fluke eggs onto

pasture when snails emerge from summer aestivation with the

onset of cooler weather. Since newly emerged snails and their

subsequent offspring are not likely to become infected if

cattle have been treated, fluke transmission to cattle is

greatly reduced during the following winter and spring. If

eggs can survive on pasture for any length of time, however,

flukicide would need to be given far enough in advance of the

return of snails to gain the full benefit of this treatment.

Current recommendations regarding the timing of autumn

treatment do not take the possibility of egg refugia on

pasture into account.

Alternatively, we have observed instances where small

numbers of snails were present throughout the summer if snail

habitats remained wet during the late spring and summer.

Therefore in years with above average spring rainfall and

normal or above average summer rainfall, it is possible that

snail populations would be present on pastures in sufficient

numbers to transmit liver flukes provided eggs survive long

enough to infect them. Therefore, as part of a larger study

to determine the seasonal transmission dynamics of F. hepatica

in Florida it seemed desirable to determine the ability of

eggs to survive, develop and hatch on pasture during the

summer months.

Materials and Methods

Collection of eggs. Eggs of F. hepatica were recovered

from the bile of gall bladders collected from infected bovine

livers at a local abattoir. Eggs were cleaned by rinsing the

bile through two screens using tap water. The first screen

(No. 60, 250g mesh) kept large particulate matter from passage

while allowing eggs of F. hepatica to pass through. The

second screen (No. 400, 38A mesh) allowed passage of clean

bile, while trapping the eggs. The eggs were then stored in

tap water at 4C in darkness to inhibit development of the


Placement of egg-containing fecal pats on pasture. At the

University of Florida's Animal Research Facility, Gainesville

Florida, a section of unshaded pasture was mowed and a 0.3 m

x 1.5 m x 3.0 m protective barrier of wooden stakes and

translucent sheet plastic was constructed to protect the

experiment from disruption. The barrier was sectioned in

half, leaving two 1.5 m2 plots. On 2 separate occasions

(experiment 1, 21 June; experiment 2, 09 August) fresh feces

was obtained from fluke-free calves at Sandhill Research Unit,

University of Florida, Gainesville Florida. Six kg of feces

was thoroughly mixed with the eggs of F. hepatica to a

concentration of 100 eggs per gram of feces. The feces was


then divided into 2 even portions and replicate fecal pats

were prepared by dropping the feces from a height of 1.25 m

into each of the 1.5 m2 plots. A sample of the feces was

retained as a control to confirm the concentration of eggs in

the feces and to determine egg development time and viability

under controlled conditions.

Collection and processing of samples. A 4-gram sample was

taken from each of the 2 fecal pats on: days 0, 1, 2, 4, 7,

15, 21, 28 and 35 in experiment 1; and days 0, 3, 7, 14, 21,

29 and 35 in experiment 2. Samples were collected during the

mid-afternoon in the warmest portion of the day. All samples

were taken from a moist portion of the interior of the fecal

pat after a portion of the outer crust was removed. Following

sampling, the crust was replaced to reform the pat and the

sample was taken to the laboratory for processing. Sampling

of fecal pats was ended when feces could no longer be

collected due to disintegration of the pat. Two gram samples

were processed using a modified sedimentation technique

(Flukefinder, Visual Difference, Moscow,ID). After filtering

samples through the Flukefinder, they were sedimented in 50

ml plastic conical test tubes for 3 minute intervals until the

sample was clean. In most cases 2 to 3 sedimentations per

sample was adequate. In samples with substantially more

debris, an additional sedimentation was performed in a 15 ml

test tube for 2 minutes. The concentrated suspension of eggs

was then transferred by pipette into 4 wells of a 24-well

tissue culture plate and placed in darkness at 27"C.

Examination of eggs. Eggs were examined under an inverted

microscope at weekly or biweekly intervals until fully

developed miracidia that were near the hatching stage were

noted. Thereafter eggs were examined daily. Prior to

examination, plates were placed under artificial light for at

least five minutes to induce hatching of mature miracidia.

Daily examination was continued until at least 50% of all eggs

had hatched or greater than 50% of eggs were dead. A

classification scheme was established to follow the

progressive development of the eggs as follows: undeveloped,

developed, and hatched. An undeveloped egg had an interior

that was completely filled with cells, appeared dense and

granular and was golden brown in color. A developed egg had

several stages including Stage 1+, Dead, and Stage 2+. Stage

1+ eggs lost their granular appearance and appeared coarsely

cellular with or without empty space within the egg. A Dead

egg was one that appeared either dead or dying; these eggs

looked relatively acellular and were somewhat lighter in

color. A Stage 2+ egg contained a formed miracidia. A

hatched egg was empty and had an open operculum. All eggs

present in the sample up to 100 were classified. The day of

hatching was defined as that on which fifty percent of viable

eggs had hatched (Rowcliffe & Ollerenshaw, 1960). Duplicate

samples collected from the 2 fecal pats were processed and


examined separately. Results are reported as an average of

the values obtained from the 2 replicate samples.

Weather information. Rainfall and temperature data were

obtained from a local weather station at the Tobacco Unit,

University of Florida, Gainesville, Florida. Additionally,

during a pilot experiment (07-Jun to 18-Jun), daily air

temperatures and fecal pat temperatures were recorded in the

morning, early afternoon, and evening. Throughout experiments

1 and 2, air and fecal pat temperatures were recorded only at

the time of sample collection.


Experiment 1 (21 June to 26 July). Eggs from a control

sample unexposed to pasture (day 0) required 15 days at 27C

to reach 50% hatching. Eggs from samples collected on days 1

and 2 hatched in the same amount of time as the control (14.5

and 15 days respectively), however, egg development and

hatching was inhibited in samples taken on days 4,7, and 15

(Figure 2-1). Eggs from feces exposed to pasture for 4 days

required 21.5 days to hatch whereas eggs exposed for 7 and 15

days required 18 and 20 days respectively. Eggs from samples

collected on days 21 and 28 required less time in the

incubator to hatch than the control eggs (11 and 10 days

respectively), indicating that some degree of development had

occurred within the fecal pat. Regardless of developmental

inhibition or enhancement within the fecal pat, the longer the


eggs were contained within the fecal pat on pasture, the

longer the total time needed for the eggs to hatch (time on

pasture plus time at 270C) (Figure 2-2). Throughout the

sampling period egg mortality was minimal with greater than

97% of eggs still viable after 21 days on pasture and 80%

still viable after 28 days. By day 35 the fecal pat was

fairly well disintegrated from rainfall and the activities of

dung disrupting invetebrates and an adequate sample could not

be obtained.

During the first 2 weeks of this experiment (21 June to 04

July) there was a considerable amount of rain that helped keep

the interior of the fecal pats wet (Figure 2-3, Table 2-1).

Heavy and frequent rainfall continued over the next 3 weeks of

the study causing fecal pats to disintegrate to a point where

an adequate sample of feces could not be obtained by day 35.

Fecal pat temperatures recorded at the time of sampling were

50C to 100C (average = 6.60C) higher than air temperatures

recorded at the same time and frequently reached 400C or more

(Table 2-2).

Experiment 2 (09 August to 13 September). Eggs from a

control sample unexposed to pasture (day 0) required 16 days

in the incubator at 27C to reach 50% hatching. Eggs from

samples collected on day 3 also required 16 days to hatch.

Beginning with samples collected on day 7, the time required

for eggs to hatch when transferred to 270C decreased with each

successive sample indicating that the eggs were developing

within the fecal pat (Figure 2-1). On the final sampling (day

35), greater than 50% of eggs recovered had already hatched by

the time the eggs were first examined after being isolated

from the feces. Although there was continuous development

within the fecal pat, the total length of time required for

the eggs to hatch increased with longer pasture exposure time

(Figure 2-2). Throughout the sampling period egg mortality

was minimal with 87% of eggs still viable after 21 days on

pasture and 84% still viable after 35 days.

Only a small amount of rain fell during the first 2 weeks

of experiment 2 (09 August to 13 September), however, the

interior of fecal pats remained moist (Figure 2-3, Table 2-1).

The frequency of rainfall increased over the next 3 weeks of

the study which helped maintain adequate fecal moisture in the

interior of the pat. Fecal pats disintegrated slower than in

experiment 1 and adequate fecal samples were still obtained on

day 35. However, by day 35 the integrity of the pat was

severely disturbed and no further samples were taken. Fecal

pat temperatures were 0 to 70C (average = 2.80C) higher than

air temperatures recorded at the same time and on one occasion

reached 410C (Table 2-2).


Our results on survival, development and hatching of eggs

of F. hepatica exposed to summer environmental conditions

differ substantially from those of Rowcliffe and Ollerenshaw


(1960). In studies on the bionomics of eggs of F. hepatica,

they demonstrated that eggs will not develop in sheep feces or

in concentrated fecal suspensions and development is inhibited

by high temperatures (>300C). Although both of these

phenomena were demonstrated to some extent in experiment 1, in

both experiments our data also contradict those findings. In

our study, eggs of F. hepatica contained within feces for

extended periods of time were able to survive and develop even

though temperatures in the fecal pats frequently exceeded


After being deposited onto pasture, fecal pats of cattle

quickly form a dry outer crust helping to maintain a moist

interior. In these experiments, the interior of the fecal

pats remained moist for the duration of each experiment when

disintegration of the pat finally occurred between days 28 to

35. In experiment 1, inhibition of egg development was

observed in eggs from feces collected on days 4, 7 and 14,

however, in experiment 2 inhibition was not observed.

Apparently some combination of factors that inhibited

development were present in the first experiment, but not the

second. Based on the findings of Rowcliffe & Ollerenshaw

(1960), one might expect higher temperatures early in

experiment 1 to cause this difference. However, during the

first 14 days of experiment 1 (21-Jun to 04-Jul), the average

of mean daily and high temperatures were lower than those

during the first 14 days of experiment 2 (09-Aug to 13-Sep)

(Table 2-2). Whether or not this small difference in air

temperatures is important is debatable, however, since it is

fecal pat and not air temperatures that will affect the eggs.

Probably more important than air temperature is the amount of

daily solar radiation. Although solar radiation was not

measured in this study, average monthly values for solar

radiation (available from published tables; ASCE, 1990) are

greatest during June and July. The mean of average solar

radiation values for June and July in Gainesville is 23,575

cal/cm2, whereas the mean of average solar radiation values

for August and September is 20,474 cal/cm2. This difference

may be important since fecal pat temperatures averaged 6.6C

higher than air temperature in experiment 1 whereas the

difference in experiment 2 averaged only 2.80C.

Differences in rainfall also may have contributed to the

differences in egg development seen in these 2 experiments

(Table 2-2). During the first 10 days of experiment 1 there

were 9 days in which there was 2 0.1 cm of rain and total

rainfall was 12.2 cm. The frequent and heavy rainfall during

this period kept the fecal pats very wet. During the first 10

days of experiment 2 there were only 2 days in which there was

k 0.1 cm of rain and total rainfall was 0.6 cm. Although the

interior of the fecal pat remained moist during this time, the

fecal pats were noticeably dryer than in experiment 1.

We hypothesize that the differences in egg development seen

between the 2 experiments are due to differences both in


fecal pat temperature and moisture content. Physical

properties of a wet fecal pat are very similar to those of a

concentrated fecal suspension, therefore it can be expected

that egg development will not occur. However, when the outer

crust of a fecal pat becomes dry it begins to crack and

crumble allowing the interior to become increasingly aerated.

Additionally, insect activity in and on the fecal pats causes

numerous small hollows and cavities to form which further

disrupts the integrity of the pat. Eggs would still be able

to survive under the moist, high humidity conditions within

the interior of the pat, but the increasingly oxygenated fecal

environment would no longer resemble the more anaerobic

environment of a concentrated fecal suspension. This would

probably allow egg development to take place and would be a

much different environment than that which exists in sheep

feces. A combination of high fecal pat temperatures with a

wet fecal suspension-like pat probably resulted in the

inhibition of development that was observed in experiment 1.

The dryer and cooler conditions of the fecal pat in experiment

2 enabled development to take place.

The maximum temperature that eggs could be exposed to and

still survive was not determined. However, on 12 July a fecal

pat temperature of 44'C was recorded and egg mortality was not

increased over the previous collection. The threshold

temperature for survival is therefore greater than 44C, a

temperature that would only occasionally be reached in fecal


pats on pasture. It is more likely that sustained high

temperatures for long periods of time would kill the eggs, but

this does not occur under natural conditions because of

diurnal temperature variation. A pilot study was performed in

early June for the purpose of optimizing the methods used in

the actual study. The detailed results are not presented

because of deficiencies in the methods and controls used in

the pilot, but some of the results are noteworthy nonetheless.

Most eggs from these fecal pats did not develop and egg

mortality was very high after only 3 days on pasture (first

collection). In one fecal pat no egg development was noted

even after 41 days at 27C. The other fecal pat had a few

eggs hatch at day 14 (3%) and by day 41 only 14% had hatched

with dead and undeveloped eggs accounting for 84% of the

total. The period of time in which these fecal pats were on

pasture was the hottest of the summer. During the 4 days that

the fecal pats were on pasture (days 0-3), the average daily

high temperature was 36.8 and the average daily mean

temperature was 28.0C with no rain and little cloud cover.

Therefore, either the lethal threshold temperature is slightly

greater than 440C and that temperature was reached (44C was

the maximum fecal pat temperature recorded), or daily

sustained temperatures in the fecal pats of >40C for several

hours were sufficient to kill most of the eggs. These results

suggest that eggs deposited during a short, very hot period

with day-long direct solar radiation will be killed. However,

because in Florida solar radiation reaches its greatest

intensity in June and decreases thereafter, it is unlikely

that fecal pats will reach temperatures sufficiently hot with

high enough frequency to have a major impact on overall egg

survival during the summer season.

Rowcliffe and Ollerenshaw (1960) determined that 9.5C was

the critical temperature below which egg development would

cease. Using this value, they concluded that the thermal

constant for the optimal temperature range for egg development

was between 200 and 220 day degrees, the values obtained when

eggs were incubated at 23 and 18 degrees respectively.

Interpolation of graphs illustrating the number of degree days

and the actual number of days required for egg hatching at

different incubation temperatures (Rowcliffe & Ollerenshaw,

1960) gave a value of approximately 190 or approximately 11

days for 27C. The eggs used in our experiments required 15

days to hatch when incubated at 270C. Using the same critical

temperature of 9.5C, the number of day degrees necessary for

hatching of the eggs used in these experiments is 263.

Clearly, the eggs used in this study obtained from fluke-

infected Florida cattle, developed much more slowly than the

eggs obtained from fluke-infected sheep in Great Britain.

Whether this difference in rate of development is due to a

difference in the minimum critical temperature (which would

affect the calculation of day degrees) or is due to an actual

difference in development rate per degree above 9.50C was not

determined in this study. It can be postulated, however, that

the warm year-round temperatures of south-central Florida

(average daily low temperature in January = 9.50C, average

daily mean temperature = 160C) would minimize the selective

pressure on eggs to develop at low temperatures and could

therefore shift the thermal constant for the optimal

temperature range upward.

The study of Rowcliffe and Ollerenshaw (1960) represents

the only comprehensive examination of egg bionomics of F.

hepatica that has been published. Our study was not meant to

re-examine their results, but designed only to determine if

eggs of F. hepatica could survive on pasture during the summer

in Florida. The differences between the results of the 2

studies suggest that there is a distinct biological variation

between liver flukes of Florida and Great Britain, but some of

the differences also can be explained by dissimilarities in

design. Sheep feces are deposited as small pellets and

therefore will heat and desiccate much more rapidly than will

cattle feces. Since cattle feces remain moist for long

periods of time and become increasingly aerated as they dry

and crack open, it is probably a much better environment for

egg development than is sheep feces under drying conditions.

However, the differences between the studies in egg

development rate and survival at high temperatures suggest

that the temperature requirements and limitations are

different for F. hepatica in Florida than for F. hepatica in

Great Britain. Therefore, major differences in climate appear

to have selected for parasites better able to survive under

the subtropical conditions of Florida. It would be

interesting to compare side by side using the same methods,

the bionomics of eggs obtained from F. hepatica in a cool

temperate climate (Great Britain, northwestern United States),

a subtropical climate (Florida), and a tropical climate

(Caribbean, Central America).

This study was performed under controlled conditions and

only partially resembles the situation that would exist in the

areas of a cattle pasture where fluke transmission is

occurring. Eggs of F. hepatica require three essential

factors to hatch. (1) They must be separated from the feces

and later exposed to light, (2) the temperature must be

adequate for survival, and (3) there must be a film of

moisture on their surface during the entire developmental

period from deposition onto pasture until release of the free-

swimming, ciliated miracidia (Rowcliffe and Ollerenshaw,

1960). Furthermore, transmission of F. hepatica occurs only

in and around the habitats of its snail intermediate hosts

because only those eggs deposited in the wet snail habitat

will ever be able to hatch, infect snails and ultimately

infect their ruminant hosts. Eggs deposited elsewhere on

pasture will almost never have the opportunity to infect a

snail no matter how long the egg might survive. Also, water

has a moderating effect on surface temperature so eggs

deposited in water will not be exposed to the same high

temperatures as those deposited on dry ground. In a 2-year

study of snail intermediate host bionomics in Florida where

surface soil and water temperatures were recorded at 3-week

intervals, surface soil and ground water temperatures reached

400C only once, occasionally reached 37 to 38C, and usually

were less than 350C. Taken together, the results of this

study strongly suggest that eggs of F. hepatica can survive,

hatch and potentially infect snails during the hot summer

season in Florida. This conclusion has 2 important

implications in regard to liver fluke epidemiology and


First, the timing of autumn treatment with flukicide must

take into account the ability of eggs to survive on pasture

during the late summer. Current recommendations for early

autumn (September to October) treatment as the most

advantageous time to treat cattle for flukes in Florida

(Courtney et al., 1985; Kaplan, 1994; Shearer et al., 1986)

does not take egg survival into consideration. This is

especially important because fluke transmission in Florida is

greatest during the spring months (Boyce & Courtney, 1990;

Kaplan, 1994) and egg production peaks 16 to 20 weeks after

infection (Doyle, 1971; de Le6n et al, 1981). Therefore,

flukes acquired during the spring will reach maximum egg

production levels in the late summer. In Florida, emergence

of snails from aestivation usually occurs in late September or


October, thus a large refugia of eggs will accumulate in snail

habitats just before snails emerge from aestivation and begin

to repopulate these areas. Treatment in late September or

October is therefore too late to prevent pasture contamination

with eggs capable of infecting the new generation of snails.

The precise amount of time following treatment that is

necessary for eggs previously deposited in snail habitats to

hatch and die cannot be determined from the results of this

study. However, it is likely that treatment approximately 4

weeks prior to the emergence of snails from aestivation should

be sufficient time to allow eggs already on pasture to hatch

and miracidia to die. Treatment for flukes in Florida should

therefore be given in the late summer, mid-August to early

September, to obtain maximum benefit.

The second important epidemiologic implication of egg

survival during the summer is that in years when snail

intermediate hosts are present on pasture during the summer,

even in low numbers, it is very likely that they will become

infected with liver flukes. Although the level of fluke

transmission resulting from this situation will probably be

low, flukes accumulated during the summer will not be killed

with high efficacy by treatment with flukicide in the late

summer or early fall because many flukes will still be

immature and not fully susceptible to available flukicidal

drugs. Surviving flukes will reach maximum egg output in the

late autumn which is the time of year in Florida when snail


populations frequently reach their greatest levels. This

would greatly decrease the benefits of late summer/early

autumn treatment and would favor high levels of fluke

transmission during the following winter and spring.

Table 2-1.

Summary of precipitation and temperature data
while fecal pats were on pasture.

No. of rain days

No. of rain days
(2 0.1 cm/day)

Total rainfall (cm)

High temperature (C)

Average of daily
high temperature (C)

Average of daily
mean temperature (C)

Experiment 1
6/21-7/04 7/05-7/26









Experiment 2
8/09-8/22 8/23-9/13









Table 2-2. Temperatures (oC) on sample collection dates,
experiments 1 and 2.

Date Weather station Pasture site* Fecal pat*
(daily high)

Experiment 1
22-Jun 35.3 33 38
23-Jun 35.1 33 40
25-Jun 34.9 33 40
28-Jun 31.7 30 35
06-Jul 37.6 35 42
12-Jul 35.6 34 44
19-Jul 35.8 ND ND
26-Jul 36.5 34 39

Experiment 2
12-Aug 36.2 34 41
16-Aug 34.8 32 35
23-Aug 36.4 32 35
30-Aug 33.0 33 33
07-Sep 33.9 29 31
13-Sep 33.2 33 36

* Temperatures recorded at time of sample collection in mid-aftemoon
ND Not Determined






5 10 15 20 25 30 35
Number of Days on Pasture

-M- 21June to 19-July

Figure 2-1.

o 09-August to 13-September

Number of days of incubation at 270C required
to reach 50% hatching vs. number of days that
eggs were in fecal pats on pasture prior to



0 20


Number of Days on Pasture

Figure 2-2.

0 09-August to 13-September

Total number of days (days on pasture plus
days in incubator at 27C) required for 50% of
eggs to hatch vs. the total number of days
that eggs were in fecal pats on pasture prior
to collection. Linear regression was
performed to obtain a best fit line for the
data. The R2 values for experiment 1 and
experiment 2 were 0.85 and 0.99 respectively.

M 21-June to 19-July

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'0 10 so (E N

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1 14



Fasciola hepatica, the common bile duct fluke of cattle

and sheep has a worldwide distribution and is a major cause of

liver condemnation, mortality and decreased productivity of

livestock (Foreyt & Todd, 1976a; Malone et al., 1982a; Price,

1953). Transmission of F. hepatica is dependant on the

presence of its lymnaeid snail intermediate hosts, thus data

on the bionomics and infection rates of these snails can be

used to evaluate changes in seasonal pasture contamination

levels and the ensuing risk to grazing livestock. In the

United States, Pseudosuccinea columella, Fossaria cubensis and

Fossaria bulimoides are known to serve as the primary

intermediate hosts for F. hepatica. However, Fossaria spp.

are the more prevalent in enzootic areas and are the

intermediate hosts of greatest importance (Malone, 1986).

Investigations into the epizootiology of F. hepatica

frequently include the monitoring of snail infection rates.

Historically, three techniques have been used to diagnose

trematode infections of snails. These are (1) observation for

cercarial shedding, (2) microscopic dissection, and (3)


crushing followed by microscopic examination. In the first

method, individual snails are examined for the shedding of

cercariae by placing them in glass tubes containing a small

volume of water. This method has been used routinely in the

study of schistosome infection rates in field populations of

snails (Sturrock & Karamsadkar, 1979) but it has been used

only rarely in published studies on snail infection rates of

F. hepatica (Olsen, 1944).

In the second method, individual snails are carefully

dissected under low and/or medium power microscopy. This

technique has been used frequently for studying infection

rates of snails with F. hepatica (Boray, Happich, & Andrews,

1969; Olsen, 1944; Smith, 1981), but there are several

problems with it. First, snail dissection is tedious and time

consuming. Second, intramolluscan stages of different

trematodes are difficult to distinguish from one another prior

to cercarial development, thus histological examination of

stained tissue sections is often required to confirm the

identification. The ruminant trematodes Fascioloides magna

(giant liver fluke) and Paramphistomum spp. (rumen flukes of

cattle and deer) share the same intermediate host and have the

same enzootic range as F. hepatica (Castro-Trejo, Garcia-

Vasquez, & Casildo-Nieto, 1990; Soulsby, 1982; Wescott &

Foreyt, 1986) thereby confounding the identification of

intramolluscan stages of trematodes recovered from snails.

Third, prior to the release of rediae from the sporocyst and


their subsequent migration through the tissue of the snail

(around day 21 post-infection), sensitivity of detection is

poor even with careful dissection. Thus, dissection suffers

from inadequate sensitivity in early stages of infection and

in those few snails where early intramolluscan stages are

found, specificity is problematic.

In the third method, snails are crushed and immediately

examined under low power microscopy for the presence of rediae

or cercariae. This technique has been used frequently in the

study of snail infection rates with schistosomes (Chernin &

Dunavan, 1962; Sturrock & Karamsadkar, 1979). A faster and

less sensitive modification of this technique has been used

recently in several large studies of the epizootiology of F.

hepatica (Malone et al., 1984/85; Khallaayoune, Stromberg,

Dakkak, & Malone, 1991). Although quick and simple, this

modified technique fails to detect prepatent infections.

Because all three of these techniques have problems of

sensitivity and/or specificity, the development of a highly

sensitive and specific method for detecting infected snails

would greatly aid in the study of the epizootiology of F.

hepatica by enabling the collection of complete and accurate

data on infection rates in field populations of snails. The

use of a DNA probe capable of sensitive and specific detection

of snails infected with F. hepatica could solve these

problems. Highly repetitive DNA constitutes a large

percentage of the eucaryotic genome (Britten & Kohne, 1986).

Since these sequences are present in multiple copies, the

relative abundance of this repetitive DNA enables very

sensitive detection. Additionally, these sequences evolve

more rapidly than gene coding sequences making them excellent

candidates for species-specific probes for eucaryotic

organisms (McLaughlin, Collins, & Campbell, 1987). DNA

probes using highly repetitive sequences have been developed

for identification of several different parasite species:

Plasmodium falciparum, (Barker, Suebsaeng, Rooney, Alecrim,

Dourado & Wirth, 1986); Brugia malayi, (Sim, Piessens & Wirth,

1986); Trichinella spiralis, (Dame, Murrell, Worley, & Schad,

1987); Onchocerca volvulus, (Shah, Karam, Piessens, & Wirth,

1987); Babesia equi, (Posnett & Ambrosio, 1989); Babesia

bigemina, (Buening, Barbet, Myler, Mahan, Nene, & McGuire,

1990); and Opisthorchis viverrini (Sirisinha,

Chawengkirttikul, Sermswan, Amornpant, Mongkolsuk, & Panyim,


For a nucleic acid probe to be useful as a diagnostic

tool in a large epidemiologic study, not only must the probe

be highly sensitive and specific, but sample preparation must

be relatively quick and efficient. Without efficient sample

processing, a diagnostic probe cannot realistically be used in

a study where thousands of samples need to be processed at a

relatively low cost. We report here the development of both

a DNA probe for the sensitive and specific detection of F.

hepatica infected snails and a quick and inexpensive DNA

extraction protocol for use in field collected snails.

Materials and Methods

Trematodes and snails. Live adult F. hepatica obtained

from condemned bovine livers at an abattoir in Florida, were

incubated for 4 h in 0.85% NaCl at room temperature to remove

adherent host cells and to empty intestinal cecae. Flukes

were then rinsed once with DNA extraction buffer (50 mM

Tris-HCl pH 8.0, 100 mM NaCl, 50 mM EDTA) and frozen with an

equal volume of extraction buffer in an alcohol bath at -700C.

Adult F. hepatica from Montana also were obtained from

condemned bovine livers, but these were frozen in liquid

nitrogen following saline incubation. Eggs of F. hepatica

were recovered from the bile of gall bladders collected from

infected bovine livers and stored in the dark at 4*C. Rediae

of F. hepatica were obtained by dissection of experimentally

infected P. columella 28 days post-infection (p.i.), rinsed in

DNA extraction buffer and quick frozen in 200 Ml of DNA

extraction buffer. Adults of Fascioloides magna and

Paramphistomum liorchis were obtained from the liver and rumen

respectively of white-tailed deer (Odocoileus virginianus)

brought to a hunting check station in Florida. Fascioloides

magna were incubated in 0.85% NaCl for 4 h and were quick

frozen in liquid nitrogen. Paramphistomum liorchis were

washed with 0.85% NaC1 to remove contaminating rumen contents,


incubated in 0.85% NaCl for 4 h, rinsed once with DNA

extraction buffer, and quick frozen with an equal volume of

DNA extraction buffer in a liquid nitrogen bath. All

trematodes were stored at -70C until subsequent use in DNA

preparation. Pseudosuccinea columella were obtained from

established cultures in our laboratory (Boyce, Courtney &

Thibideau, 1986) and F. cubensis were collected from a fenced

field in Florida devoid of domestic ruminants.

Laboratory infections of snails. Eggs of F. hepatica

were removed from the refrigerator and incubated in darkness

at room temperature for two weeks to allow miracidial

development. Following incubation, eggs were exposed to light

to stimulate hatching. Pseudosuccinea columella or F.

cubensis were individually placed in wells of a 24 well tissue

culture plate along with 2 3 freshly hatched miracidia and

200 300 pl of tap water. After 3 4 h the snails were

removed from the wells and reared in culture.

Isolation of genomic DNA. Frozen flukes were ground to

a fine powder in liquid nitrogen using a pre-chilled mortar

and pestle. Genomic DNA was extracted from the powdered

flukes by SDS-Proteinase K digestion followed by

phenol/chloroform extraction (Sambrook, Fritsch & Maniatis,

1989). The recovered DNA was dissolved in 10mM Tris-HCl-lmM

EDTA, pH 7.6 (TE) buffer, and contaminating RNA was removed by

incubation with RNAse A for 60 min at 370C followed by a

second phenol/chloroform extraction and ethanol precipitation.

Uninfected snails were kept without food in distilled water

containing 100 Ag ml-' ampicillin for 48 h to clear their

digestive system and minimize bacterial contamination prior to

the extraction procedure (Strahan, Kane, & Rollinson, 1991),

rinsed in distilled water, and their shells crushed and

removed. Intact tissue from four snails was rinsed once in

DNA extraction buffer, placed in a microcentrifuge tube with

400 Al of extraction buffer and crushed with a polypropylene

pestle (Kontes Chemistry & Life Science Products, Langhorne

PA). Genomic DNA from snails was then extracted essentially

as described above.

Construction of F. hepatica genomic DNA library. Genomic

DNA of F. hepatica was digested to completion with the

restriction endonuclease Sau 3A (BRL, Gaithersburg MD),

purified by phenol/chloroform extraction and recovered by

ethanol precipitation. Digested DNA was ligated into the Bam

HI site of the plasmid Bluescript SK+ (Stratagene, La Jolla,

CA) using T4 DNA ligase. Ligation products were transformed

into Escherichia coli, strain XL1-Blue and were grown

overnight at 37C on Luria Bertaini (LB) plates containing

ampicillin, tetracycline, X-gal (5-bromo-4-chloro-3-indolyl-P-

D-galactoside) and IPTG (isopropylthio-P-D-galactoside).

Selection of clones containing repetitive sequences.

Ninety six recombinant clones were selected at random,

transferred to a 96-well microplate and grown overnight in LB

media at 370C. Replicates of the clones were made on


nitrocellulose filters (Schleicher & Schuell, Keene, NH)

placed on LB plates, grown overnight at 37C, treated to

release the DNA from E. coli (Thayer, 1979) and baked for 2 h

in a vacuum oven at 80C. The filters were incubated for 2 h

at 65C in a prehybridization solution (6X sodium chloride,

sodium citrate pH 7.5 (SSC), 1.0 mM EDTA, 10X Denhardts, 0.5%

SDS, 10 mM NaPO4, and 100 Ag ml-' denatured salmon sperm DNA).

A radiolabeled probe was prepared using genomic DNA of F.

hepatica and a random primer labeling kit (Prime-It*,

Stratagene, La Jolla, CA) with a32P-dATP (111 TBq/mMol, E.

I.DuPont De Nemours & Co. Inc., Wilmington, DE). Labelled

probe was boiled for 5 min with 1.0 mg of sheared salmon sperm

DNA and added directly to the filters in prehybridization

solution for overnight hybridization (16 h). The filters were

washed once in 6X SSC, 0.1% SDS at room temperature for 10

min, twice in 2X SSC, 0.1% SDS at 50C for 30 min, and once

with 0.2% SSC, 0.1% SDS at 500C for 30 min. Hybridizing clones

were identified by autoradiography using X-OMAT-AR film

(Kodak, Rochester, NY) and an intensifying screen. Positive

clones were grown in LB broth at 370C overnight, and plasmid

DNA was isolated by the boiling prep method (Holmes & Quigley,

1981). Insert DNA was excised by digestion with Eco RI and

Xba I and DNA fragments were separated by agarose gel

electrophoresis. DNA fragments were transferred from the gel

to a nylon membrane (Southern, 1975) and probed with

radiolabeled genomic DNA of F. hepatica as described above.


Sensitivity and specificity of selected clones.

Replicate slot blots were made onto nylon membrane (Hybond

N+*, Amersham, Arlington Heights, IL) using a vacuum

filtration apparatus (Minifold II Slot Blot System -

Schleicher & Schuell, Keene, NH). Blots were made with 1.0 Mg

each of genomic DNA from adults of F. magna, P. liorchis and

F. cubensis, and 1.0 Ag, 100 ng, 10 ng, 1.0 ng, and 0.1 ng of

genomic DNA from F. hepatica. DNA was crosslinked to the

membranes by exposure to UV light (Stratalinker, Stratagene,

La Jolla CA) prior to prehybridization. Each of the selected

clones was radiolabeled for use as a probe and was hybridized

overnight with one of the replicated slot blots.

Hybridization and washing conditions were the same as

described for the genomic DNA library screen for clones with

repetitive sequences.

Determination of fragment size and insert number in F.

hepatica specific clones. Insert DNA of F. hepatica specific

clones was PCR amplified in a Perkin Elmer Cetus DNA thermal

cycler using a standard reaction mixture (AmpliTaq*, Perkin

Elmer Cetus, Norwalk CT) with KS (GCTATGGCAGCTGGAGC) and SK

(TCTAGAACTAGTGGATC) primers. PCR products were digested with

Sau 3A for 1 h at 370C and fragments were separated by

electrophoresis on a 1.5% agarose gel.

Sensitivity and specificity of selected probe. Genomic

DNA from Florida and Montana isolates of F. hepatica, genomic

DNA of F. magna and P. liorchis, and genomic DNA from the


Florida isolate of F. hepatica mixed with 10 gg of P.

columella DNA was spotted in 10X dilutions from 1.0 Lg down to

10 pg onto nylon membrane. DNA of rediae dissected free of

experimentally infected snails, and DNA of individual fluke-

infected snails (28 days p.i.) also were spotted onto the

membrane. Hybridization and washing conditions were the same

as described for the genomic DNA library screen.

Minimum biologic sensitivity of the probe. Laboratory

raised P. columella were infected with miracidia of F.

hepatica. Groups of 4 snails were removed from culture tanks

on days 0 (4 h post-infection), 1, 4, 7, 14, 21, and 28 p.i.,

crushed in 200 Ml lysis buffer (8% triton-X 100, 250 mM

sucrose, 50 mM TE pH 7.6) using plastic pestles, and frozen at

-20C prior to extraction. DNA was extracted using a protocol

which was modified from that used by Strahan et al. (1991).

Snail lysates were thawed, 20 A1 of freshly prepared

Proteinase K (10 mg ml-') was added, and samples were incubated

in a waterbath at 680C for 2 h. Following incubation, samples

were centrifuged for 1 min at 12,000 x g and the supernatant

was transferred to a clean tube. Nucleic acids in the lysate

were precipitated by the addition of 800 Ml of warm 2%

hexadecyltrimethyl ammonium bromide solution (CTAB) and tubes

were inverted 10 times to mix. The precipitate was pelleted

by centrifugation for 1 min at 12,000 x g. Supernatant was

aspirated and 200 pl of 2.5 M NaCI containing 10mM EDTA was

added to dissolve the pellet. To facilitate dissolution the


samples were placed back into a 680C waterbath over night. A

single chloroform extraction was performed using 400 Al of

chloroform/isoamyl alcohol (24:1) and the aqueous phase was

transferred to a clean tube. Twenty Al of 2N NaOH was added

to denature the DNA and the sample was applied onto nylon

membrane using a dot blot manifold (Schleicher & Schuell,

Keene, NH). DNA was crosslinked to the membrane using UV

light and was hybridized with a32P-dATP labeled probe overnight

at 650C as described above. The hybridized membrane was

washed twice for 15 min in 2X SSC, 0.1% SDS and once in 0.2X

SSC, 0.1% SDS for 15 min at 550C.


Screening of F. hepatica genomic library for highly

repetitive sequences. Thirteen out of 96 recombinant clones

from a genomic DNA library of F. hepatica hybridized strongly

with radiolabeled genomic DNA of F. hepatica. All clones

contained small inserts, ranging in size from approximately

190bp to 900bp. Ten of the 13 clones were selected for

further analysis. All 10 clones displayed strong

hybridization signals when probed with total genomic DNA of F.

hepatica, suggesting that they contained highly repetitive


Characterization of selected clones. Radiolabelled

probes were prepared from each of the 10 clones to identify

those which hybridize specifically with F. hepatica DNA but

not with DNA of F. cubensis, F. magna or P. liorchis. Four

clones cross hybridized with DNA of F. magna and one clone

cross hybridized with DNA of P. liorchis. Most clones showed

a slight non-specific cross hybridization with DNA of F.

cubensis. This slight non-specific signal was not seen in

later experiments with DNA of F. cubensis or P. columella

confirming the specificity of these clones against DNA of the

snail intermediate hosts. Each of the 5 clones specific for

F. hepatica detected as little as 1.0 ng of F. hepatica

genomic DNA (Figure 3-1).

Insert DNA of the 5 F. hepatica specific clones ranged in

size from 190bp to 520bp. Inserts from these clones were

amplified by PCR using vector derived primers


Sau 3A. A 125bp fragment was seen in all 5 of the clones.

Since all 5 clones contained a similar sized DNA fragment,

cross-hybridization experiments were performed to determine

the relationship of the 5 clones. Results (not shown)

revealed that insert DNA of all 5 clones hybridized with

varying intensity to probes prepared by radiolabelling each of

the 5 cloned inserts.

pFh5 and pFh76 contained cloned inserts of approximately

250bp and were selected for sequencing since this suggested

that each clone may contain two-125bp repeats. Clone pFh5

contained two 124bp repeats (pFh5rl & pFh5r2) and clone pFh76

contained one 124bp repeat (pFh76rl) together with a short

unrelated sequence. The sequences of the 3 repeats differed

only slightly from each other. Repeats pFh5rl and pFh5r2 had

11 base mismatches, pFh5rl and pFh76rl had 14 base mismatches,

and pFh5r2 and pFh76rl had 15 base mismatches. Thus, the 3

repeats have a 91%, 89%, and an 88% sequence identity

respectively (Figure 3-2). A search of the Genbank database

showed that no other previously reported sequences were

homologous to those reported here.

The abundance of this 124bp repeat within the F. hepatica

genome was determined by quantitative dot-blot analysis

(Figure 3-3). Phosphor image analysis of hybridization

signals demonstrated that this 124bp repeat constitutes 15%

of the F. hepatica genome. Assuming that the genome of F.

hepatica is similar in size to that of Schistosoma mansoni,

i.e. 2.7 x 108 (Simpson, Sher & McCutchan, 1982) there are

greater than 300,000 copies of this 124 bp repeat present in

the genome of F. hepatica.

Sensitivity and specificity of the selected probe, pFh5.

1.0 ng of genomic DNA of F. hepatica was detected easily with

the probe and this level of sensitivity was not diminished by

the presence of 10 pg of snail DNA. The probe showed similar

hybridization intensities to genomic DNA of the Montana and

Florida isolates of F. hepatica verifying the reliability of

the probe against another isolate from a geographically

distant locale (data not shown). The probe did not cross

hybridize with DNA of the trematodes F. magna or P. liorchis,


or with DNA of the snail intermediate hosts, F. cubensis and

P. columella. Additionally, DNA from a field-collected snail

(F. cubensis) infected with schistosome-like cercariae

presumptively diagnosed as Heterobilharzia americana did not

hybridize with pFh5. Heterobilharzia americana is the only

other digenetic trematode other than F. hepatica, F. magna and

Paramphistomum spp. known to use Fossaria spp. as intermediate

hosts (Malek, 1980).

Since the purpose of this probe is to detect all infected

snails, the goal set for sensitivity was the detection of the

minimum biologic unit of the parasite. Thus, if a single

miracidium following penetration of the snail can be detected,

no further sensitivity is required. The probe developed in

this study is capable of this level of sensitivity (Figure

3-4). This probe detected the parasite DNA in all 4 snails

whose DNA was extracted less than 4 h following miracidial

penetration. The amount of parasite DNA remained about the

same on day 1 p.i., but by day 4 considerable parasite

development had occurred and this development continued in a

logarithmic fashion throughout the 28 day time period included

in the experiment (Figure 3-5, Table 3-1). One infection was

not detected on day 14, and this was most likely due to the

snail never having become infected or the snail having

rejected its infection, although laboratory error during the

extraction and detection process is also possible. These

snails were each exposed to 2-3 miracidia to ensure that most


snails would become infected, however, in a separate

experiment, DNA from a single miracidia extracted together

with an uninfected snail was easily detected with the probe

(data not shown). Data from the phosphor image analysis of

blots containing known dilutions of genomic DNA of F. hepatica

and the same dilutions of DNA of F. hepatica but added to

snails and extracted, demonstrated that DNA recoveries using

the CTAB based extraction protocol average 50-60% regardless

of the concentration of the added DNA of F. hepatica.


The DNA probe and snail extraction protocol reported here

solve the problems of sensitivity, specificity and efficiency

which are inherent in the visual examination of snails for the

purpose of detecting F. hepatica infection. Infected snails

can be detected immediately following miracidial penetration,

thus a sensitivity equivalent to the minimum biologic unit of

the parasite is achieved. Additionally, signal intensity

increases in a logarithmic fashion over time, therefore,

signal intensity can be used to estimate the age of the

infection giving further information on the bionomics of the

infected snail populations on pasture. However, for a nucleic

acid probe to be useful as a diagnostic tool, not only must it

be sensitive, but it must also be specific for the organism of

interest. The screening and selection process used in this

study ensured that the probe would be both highly sensitive



and specific. pFh5 was selected from among the 5 candidate

probes based upon sequencing results which showed that it

contained two repetitive sequences but no other unrelated DNA.

This probe did not cross hybridize with 1.0 jg of genomic DNA

of F. magna or P. liorchis, trematodes which share the same

enzootic range and same intermediate hosts as F. hepatica.

All 5 of the clones which were specific for F. hepatica

contained a fragment of DNA approximately 124 bp in size. In

experiments to determine if these fragments were related, all

5 cross-hybridized with each other to varying degrees

suggesting that they all were similar but not identical in

sequence. This conclusion was further supported when sequence

data from clones pFh5 and pFh76 demonstrated that the 3 repeat

sequences contained in these clones shared only about 90%

identity. This level of sequence variation between repeats is

characteristic of interspersed repetitive sequences (Jelinek

& Schmid, 1982), although the arrangement of this repeat in

the genome has not been determined. It therefore appears that

the F. hepatica genome contains a large family of 124 bp

repeats that are not identical but share a high level of

sequence identity. Quantitative dot-blot analysis revealed

that this 124 bp repeat accounts for approximately 15% of the

entire F. hepatica genome. This level of abundance is similar

to that reported for a 121 bp repeat (12%) in S. mansoni

(Hamburger, Turetski, Kapeller & Deresiewicz, 1991). It is

interesting that a search of the Genbank data base revealed no


homology between the 124 bp repeat sequences reported here and

the 121 bp repeat sequence reported for S. mansoni. The 124

bp repeats of F. hepatica and the 121 bp repeat of S. mansoni

were cloned from genomic DNA libraries made from DNA digested

with Sau 3A. It would be interesting to see if other

trematode species also contain abundant repeats of similar

length as this would be an excellent means of quickly

isolating species specific DNA probes.

Before a nucleic acid probe can be used in a large

epidemiologic survey, sample processing must be made both time

and cost efficient. Phenol/chloroform extraction was used in

the development and initial testing of the probe but problems

associated with this technique excluded it from practical

consideration as a method for large scale snail DNA

preparations. Hamburger, Weil, & Pollack (1987), reported a

NaOH-based technique for quick DNA extraction of Schistosoma

mansoni infected snails (Biomphalaria glabrata). However, in

our laboratory using different snail species (F. cubensis, P.

columella), poor results were obtained using this technique.

We have developed a DNA extraction protocol which solves

all the major problems associated with both of these

techniques. The protocol described here utilizes the cationic

quaternary ammonium compound, CTAB, which selectively

precipitates nucleic acids in preference to proteins and

polysacharides at low salt concentrations (Yap & Thompson,

1987). DNA recoveries using this procedure average 50-60%,

___ __


and the extract is easily filterable through a nylon membrane.

When extracting large numbers of snails with this technique,

total processing time per sample, excluding an overnight

incubation, is only about two minutes. Total cost per snail

for the entire assay including all miscellaneous supplies,

solutions, reagents, enzymes, and disposables (i.e. tubes,

pipette tips, etc.) is only about US $0.33. This sample

processing speed enables extraction of hundreds of snails over

a 2 day period and the low cost minimizes economic

considerations when determining the number of snails that must

be assayed (thousands) to obtain statistically meaningful

data. This protocol therefore possesses all of the desirable

qualities of a DNA extraction process for use in a large scale

epidemiologic study. It is quick, inexpensive, safe, and

gives both excellent DNA recovery and an extract that is

easily filterable.

Recently, nucleic acid probes for the detection of F.

hepatica infected snails have been published in two separate

reports (Shubkin, White, Abrahamsen, Rognlie, & Knapp, 1992;

Heussler, Kaufman, Strahm, Liz, & Dobbelaere, 1993). Shubkin

et al., (1992) reported the development of a nucleic acid

probe using rRNA sequences for the detection of F. hepatica

infected snails, however, the probe was not tested for

specificity against any other trematodes. Fascioloides magna

and F. hepatica both belong to the family Fasciolididae.

Therefore, it is very likely that the DNA of these organisms


have a high degree of sequence homology, especially in

conserved genes such as those for rRNA. In addition,

Paramphistomum spp. and F. magna both can infect the same

snail intermediate hosts as F. hepatica and frequently infect

livestock in the same areas enzootic with F. hepatica. Thus,

before any nucleic acid probe can be used as an epizootiologic

tool in the study of F. hepatica, it must be demonstrated that

it does not cross hybridize with nucleic acid of either of

these trematodes. Additionally, this report did not address

the question of sample preparation efficiency and cost, both

of which will ultimately determine whether a nucleic acid

probe becomes a useful research tool or an academic exercise.

Heussler et al. (1993) also failed to demonstrate the

practical specificity of the DNA probes they developed. They

screened their probe for specificity against 3 different

trematodes: Dicrocoelium dendriticum, Xiphidiocercaria spp.,

and Diplostoma spp. All of these trematodes are

phylogenetically divergent from F. hepatica, use different

snail intermediate hosts, are found in different ecological

niches than F. hepatica, and only D. dendriticum is a parasite

of ruminants. Additionally, sensitivity of these probes was

not tested against purified genomic DNA of F. hepatica and

when squash blots of individual F. hepatica rediae or

cercariae were probed, only a weak signal was detected. This

is in contrast to the sensitivity of the probe assay reported

here in which the DNA of a single miracidia can be detected

__ __


and the DNA of a single rediae gave a very strong

hybridization signal that was similar to the signal strength

seen in 14-21 day old infections.

The DNA probe assay reported here will enable researchers

to obtain truly accurate infection rate data in field-

collected snails for the first time. This should lead to a

better understanding of the epizootiology of F. hepatica. By

monitoring the population density and infection rates of

snails over time, patterns of seasonal pasture infectivity

with metacercariae can be determined. Knowing this, the risk

of infection to grazing ruminants can be predicted. Because

numerous snail habitats in a given region can be studied at a

relatively low cost using this approach, the chances of

geographic bias are reduced greatly. This is extremely

important for any disease in which transmission is dependant

upon an invertebrate animal whose population density is so

easily affected by minor differences in local physiographic

parameters and precipitation levels.

Monitoring of schistosome infection rates in snails

frequently has been used to evaluate the success of control

measures and to better understand transmission dynamics

(Barnish, 1982; Chernin & Dunaven, 1962; Christie & Upatham,

1977). This has not been attempted for F. hepatica, however.

By monitoring snail infection rates with F. hepatica, the long

term effects of currently recommended control measures on

liver fluke transmission dynamics can be better evaluated.

This may permit the adjustment of fluke risk prediction models

(Malone et al., 1987) to compensate for chemotherapeutic

intervention, thus making improved control recommendations


Additionally, this DNA probe assay can be used to

increase our basic understanding of the biology of the snail-

fluke relationship. This includes the ability to rapidly

determine snail host suitability for F. hepatica in areas of

the world where it is not yet known. Finally, it is very

likely that the DNA probe assay reported here for the

detection of F. hepatica infected snails can be modified and

used for the study of other helminth parasites which utilize

an invertebrate animal during a portion of their life-cycle.

pFh5 pFhlO pFhl5 pFhl7

1.0 pg Fh
0.1 pg Fh
10.0 ng Fh
1.0 ng Fh
0.1 ng Fh
1.0 pg PI
1.0 pg Fm
1.0 pg Fc

pFh56 pFh67


pFh69 pFh71

_ -

Figure 3-1.

Screening of clones containing repetitive
sequences for sensitivity and specificity.
Each clone was radiolabeled with a3P-dATP and
used to probe replicate slot blots containing
DNA of F. hepatica (Fh) in 10X dilutions from
1.0 Pg down to 0.1 ng, and DNA of F. magna
(Fm), P. liorchis (P1) and F.cubensis (Fc) in
1.0 Ag quantities.




1.0 pg Fh
0.1 pg Fh
10.0 ng Fh
1.0 ng Fh
0.1 ng Fh
1.0 pg PI
1.0 pg Fm
1.0 pg Fc







51 100




Figure 3-2.

DNA sequences of repeats in clones pFh5 and
pFh76. Clones pFh5 and pFh76 were sequenced
in both strands using the PrismTm ready
reaction dye primer cycle sequencing kits,
containing M13 forward and reverse primers
(Applied Biosystems, Foster City CA) and an
ABI 373a automated DNA sequencer. Nucleotide
sequence data have been submitted to the
GenBank database and assigned the accession
numbers: PFH5R1: U11819; PFH5R2: U11818;
PFH76R1: U11817.



c d e f g h

F. hepatica


Figure 3-3.


Quantitative dot-blot analysis for estimation
of the abundance of the 124 bp repeat in the
genome of F. hepatica. Genomic DNA of F.
hepatica and PCR amplified insert DNA of pFh5
was blotted onto nylon membrane in doubling
dilutions from (a) 100 ng to (h) 0.78 ng for
F. hepatica and from (a) 1.0 ng to (h) 7.8 pg
for pFh5 and hybridized with "P-labeled pFh5
insert. Intensity of the hybridization signal
was measured by storage phosphor imaging
(PhosphorImager 400-S, Molecular Dynamics,
Sunnyvale, CA) and the image was quantitated
using image analysis software (ImageQuant
Version 3.22, Molecular Dynamics).
Hybridization and washing conditions were as
in Figure 3-4. under materials and methods.
Autoradiographic exposure was for 5 h.

a b

C1C2-1 0 1 4 7 142128

B 9
C *

Figure 3-4.

Detection of snails infected with F. hepatica
using probe pFh5. DNA of 4 uninfected snails
(-1) and 4 infected snails on days 0, 1, 4, 7,
14, 21 and 28 p.i. (rows A-D). To enable
quantitation of hybridization signal of
infected snails and calculation of DNA
recovery per cent during extraction, genomic
DNA of F. hepatica in 10X dilutions from 1.0
Ag to 0.1 ng was blotted directly (lane C1,
rows A-E) or added to individual uninfected
snails (lane C2, rows A-E). DNA of all snails
was extracted, blotted onto nylon membrane,
and hybridized overnight with a32P-dATP
labeled probe. Autoradiographic exposure was
for 12 h.

-- Average of 4 Snails

0 7.. .. .
0 7

14 21 28

Days Post Infection

Figure 3-5.

Graphical representation of data from Table
3-1. Increase in the total genomic DNA of
intramolluscan stages of F. hepatica over the
first 28 days of infection. Line is through a
point representing the average of the 4
individual data points (4 snails).

1800 -

1500 -

1200 -

900 -

600 -

300 -


__ 1_ 1

TABLE 3-1.

F. hepatica
snails as
analysis of

DNA recoveries (ng) from infected
determined by phosphor image
blot shown in Figure 3-4*.

Days Post Infection

Snail 1
Snail 2
Snail 3
Snail 4


0.20 0.55 2.76 23.18 241.22 376.0 881.56
0.77 0.87 1.31 3.30 ND 401.91 1645.03
0.74 0.23 2.30 9.26 97.89 234.01 692.10
1.71 0.93 2.47 47.86 79.28 434.34 1519.44



2.21 20.90 139.46 361.56 1184.53

ND = Not Determined
* Storage phosphor imaging was performed by exposing the
hybridized membrane to an imaging screen (Molecular Dynamics,
Sunnyvale, CA) for 24 hr. at room temperature. The screen was
then scanned using the Molecular Dynamics PhosphorImager* 400-
S and the image was quantitated using image analysis software
(ImageQuante Version 3.22, Molecular Dynamics). Background
counts were subtracted from all samples and linear regression
was performed using standard DNA concentrations vs. observed
counts. Results of linear regression were used to calculate
actual amounts of F. hepatica DNA recovered from each infected
snail. True amount of F. hepatica DNA would be approximately
67-100% greater than the values reported here since extraction
recoveries average 50-60%.



Fasciola hepatica, the common liver fluke, is enzootic

throughout most of the major livestock producing areas of the

world. Control programs aimed at minimizing economic losses

from F. hepatica are usually based on the properly timed

treatment of infected livestock with flukicidal drugs (Armour,

1975; Malone et al., 1982a; Shearer et al., 1986). Because of

regional and local differences in seasonal transmission

dynamics, the proper timing of these treatments differs for

each geographic area. Therefore, before appropriate treatment

strategies for the control of liver flukes can be developed

for a particular region, the seasonal transmission dynamics

must be first determined.

Studies designed to determine seasonal transmission

dynamics of F. hepatica typically use worm-free sentinel

animals, "tracers", that are placed on pasture at 4 or 6 week

intervals and necropsied after 6-8 weeks of confinement

(Armour, Urquhart, Jennings, & Reid, 1970; Boyce & Courtney,

1990; Craig & Bell, 1978; Hoover et al., 1984). Although

tracer studies are often regarded as the most accurate means

for determining seasonal transmission profiles (Armour et al.,

1970) they are also extremely expensive to conduct. For

example, at the University of Florida, a 2-year single site

tracer study with 6 calves placed on pasture at 6 week

intervals would cost approximately $75,000. The high cost

associated with tracer studies frequently limits them to 1 or

2 years at 1 or 2 sites and can therefore lead to studies

where data is incomplete and/or subject to geographic and

yearly climatic biases. The great expense of tracer studies

also limits the number of investigations that are conducted.

This has caused a dearth of knowledge regarding F. hepatica

transmission dynamics in many areas of the United States and

around the world. This is especially important in developing

countries where a lack of information regarding parasite

transmission prevents the design of rational cost effective

control programs.

Tracer studies also frequently include the determination

of snail intermediate host bionomics and snail infection

prevalences (Boray et al., 1969; Khallaayoune et al., 1991;

Malone et al. 1984/85; Ross, 1977). This data is helpful in

interpreting the worm acquisition data and enables the

researcher to better explain the seasonal transmission

patterns that are seen. Unfortunately, existing microscopic

techniques lack the sensitivity, specificity and efficiency

necessary to obtain accurate and meaningful infection

prevalence data in most studies. As a result, the effort to


determine snail infection prevalence is often wasted because

the numbers of infected snails detected are too low to have

any epidemiologic value (Khallaayoune et al., 1991; Lindsay,

1976; Malone et al. 1984/85; Wilson & Samson, 1971). However,

accurate snail infection prevalence data has the potential to

be extremely useful in determining seasonal transmission

dynamics of trematodes (Christie & Upatham, 1977; Sturrock,

1973). Patterns of F. hepatica transmission are the direct

result of interactions between the snail intermediate host,

the parasite and the environment (Ollerenshaw, 1959).

Therefore, data on the bionomics and infection prevalences of

these snails can be used to predict changes in seasonal

pasture contamination levels and the ensuing risk to grazing

livestock. Thus, it is possible to determine seasonal

transmission dynamics without using expensive tracer animals,

particularly if a method for rapidly testing large numbers of

snails for infection is available.

We recently reported the development of a highly

sensitive and specific DNA probe for the detection of F.

hepatica infected snails together with an efficient DNA

extraction protocol suitable for large scale testing of field-

collected snails (Kaplan et al., 1995). Here we report a

modification of that protocol that improves the assay

efficiency and makes use of chemiluminescent detection.

Using this DNA probe assay we monitored the infection

prevalence of more than 5,000 snails on 6 cattle ranches in


Florida during the second year of a 2-year study on snail

intermediate host bionomics. Additionally, the infection

prevalence of snails on 1 ranch was determined for the first

year using the same DNA probe but a different assay system.

This report represents the first published use of a nucleic

acid probe in the study of F. hepatica transmission dynamics

and demonstrates the usefulness of this alternate approach.

Materials and Methods

Snail collection and storage (field-collected snails).

Fossaria cubensis were collected at 3 week intervals from 12

May 1993 to 26 April 1994 using the methods described in

chapter 5. Snails were collected from pastures of 6 cattle

ranches in Florida: H. E. Wolfe Ranch, St. Johns County;

Deseret Ranch, Brevard County; Creek Ranch, Polk County; Rio

Ranch, Okeechobee County; John Williams Ranch, Okeechobee

County; and Brighton Seminole Indian Reservation (Glades

County). At the time of collection, snails were placed in

small (-50 ml) plastic jars filled with water obtained at the

snail collection site. Jars with snails were then stored on

ice in a chest-cooler until brought back to the laboratory.

Once at the laboratory, snails were poured into a strainer

covered with 2 layers of cheese cloth and rinsed well with tap

water to remove as much mud and debris as possible. Snails

were then placed back into clean containers with dechlorinated

tap water and stored in the refrigerator at 40C for 1 to 7 days.


Snail processing prior to DNA extraction. Snails were

removed from the refrigerator, rinsed again with tap water

through cheese cloth and were then placed in a 100 x 15 mm

petri dish containing distilled water. Snails were

individually removed from the petri dish, measured to the

nearest millimeter (rounding upward), and placed on a 15 x 20

cm piece of clean lab-bench protector paper. All snails (or

184 snails, whichever was less) from an individual ranch were

measured and counted except for two collections (20 Jan 1994,

06 April 1994) when greater than 184 snails were assayed from

the Brighton Seminole Indian Reservation. Microcentrifuge

tubes were labelled according to snail size (1 mm to 10 mm)

and 200 pl of lysis buffer (8% triton-X 100, 250 mM sucrose,

50 mM TE pH 7.6) for snails : 7 mm or 300 pl for snails k 8 mm

was added. Individual snails were then placed in tubes and

were either immediately crushed using plastic pestles or were

stored in the refrigerator for 1 to 4 days prior to being

crushed. Snails from a single ranch were processed as a unit

and were kept at room temperature while being crushed.

Crushed snails were then stored in the refrigerator at 4C if

DNA extraction was planned for the same week, otherwise

crushed snails were stored in the freezer at either -200C or -

700C depending upon available storage space.

DNA extraction procedure. Twenty microliters of freshly

prepared Proteinase K (10 mg ml-1) was added to crushed snails

using a motorized microliter pipet (EDP-Plus, Rainin


Instrument Co. Inc, Woburn, MA) and samples were incubated in

a waterbath at 68C for 2 to 3 h. For snails 1 8 mm, 40 Al of

Proteinase K was added and incubations were done for 3 to 4 h.

After approximately 1 h of incubation, samples were mixed by

gently inverting the tube racks several times. Following

incubation, samples were centrifuged for 30 sec at 12,000 x g

to pellet sample debris (shell plus undigested snail tissue)

and to draw the lysate down and off the tube-lid to prevent

sample cross-contamination when the tubes were opened.

Lysates of snails k 4 mm were transferred to clean tubes,

however, the amount of debris in snails : 3 mm was minimal and

these lysates were not transferred. Nucleic acids in the

lysates were precipitated by the addition of 800 Al of warm 2%

hexadecyltrimethyl ammonium bromide solution (CTAB). Tubes

were inverted 10 times to mix, the nucleic acid precipitate

was pelleted by centrifugation for 30 sec at 12,000 x g, and

the essentially nucleic acid-free supernatant was aspirated

using a glass pasteur pipet attached to a vacuum apparatus.

200 jl of 2.5 M NaC1 containing 10 mM EDTA was added to

dissolve the DNA-CTAB pellets and the samples were placed back

into a 68C waterbath over night to facilitate dissolution.

The next morning, racks with tubes were shaken for 10 sec and

tubes were centrifuged for 5 sec to draw sample droplets off

the lid. A single organic extraction was performed to

separate the CTAB from the DNA using 500 ul of

chloroform/isoamyl alcohol (24:1) which was stained pink by


the addition of Sudan III (.025 % w/v) (Sigma Chemical Co.,

St. Louis MO). One hundred and eighty microliters of the

aqueous phase (-90% of the sample) was transferred to a well

of a 96-well microplate and microplates containing the DNA

samples were stored in the refrigerator at 4C (1 to 7 days)

until they were dot-blotted onto nylon membrane. Each

microplate contained 92 test snails, one positive snail

control, one negative snail control and 2 empty wells for

hybridization controls. All solutions used in the assay in

large volumes (2 200 Al; lysis buffer, CTAB, NaCl/EDTA,

Chloroform) were dispensed using repeat dispensers (Repipet*,

Labindustries, Berkeley, CA).

Preparation of snail controls for DNA extraction

procedure. Uninfected laboratory raised Pseudosuccinea

columella were removed from culture tanks and placed in a 100

x 15 mm petri dish containing distilled water for -1 h.

Snails were dried on clean lab-bench protector paper and added

to tubes containing 200 A1 of lysis buffer. Ten nanograms of

F. hepatica genomic DNA was added to positive control snails

and both positive and negative control snails were crushed and

stored at -700C until needed. Control snails were thawed as

needed for each round of snail DNA extractions and were

handled exactly the same as sample snails throughout the

extraction process.

DNA dot blots. Microplates containing the snail DNA

extracts were removed from the refrigerator. Each microplate

had 2 wells that were left empty for use as a positive and

negative control for the blotting, hybridization and detection

process. 180 gl of 2.5 M NaC1 containing 10 mM EDTA (aqueous

solution of snail DNA extracts) was added to each control well

and the positive control well also had 10 ng of genomic DNA of

F. hepatica added. Twenty microliters of 2N NaOH (0.2N final

concentration) was then added to each well using a motorized

microliter pipet (EDP-Plus*, Rainin Instrument Co. Inc,

Woburn, MA) and the microplate was placed in a 370C incubator

for 15 min to 3 h to denature the DNA. 100 gl of each sample

(one half of volume) was transferred to a nylon membrane

contained within a dot blot manifold (Schleicher & Schuell,

Keene, NH) using a multichannel pipettor (12-Pette, Costar

Corporation, Cambridge MA). The remaining 100 pl were stored

at -70C. HybondTM N+ (Amersham Corporation, Arlington

Heights, IL) nylon membrane was used when hybridization was

with 3P-labeled probe and positively charged nylon membrane

(Boehringer Manheim Corporation, Indianapolis, IA) was used

when hybridization was with Digoxigenin-labeled probe.

Membranes were air-dryed and DNA was crosslinked to the

membrane by UV light (Stratalinker, Stratagene, La Jolla CA).

Membranes were stored in plastic zip-lock bags until used for


Probe preparation and labelling. Probe was PCR amplified

from clone pFh5 in a Perkin Elmer Cetus DNA thermal cycler

using a standard reaction mixture (AmpliTaqO, Perkin Elmer

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