AN ABSTRACT OF THE THESIS OF
Claire A. Fuller for the degree of Doctor of Philosophy
presented on October 25, 1994. Title: Host Parasite
Relationships Between Deer Mice (Peromyscus maniculatus) and
Their Eimerian Parasites (Protozoa).
Redacted for Privacy
Abstract approved:
Andrew R. Blaustein
Hosts and their parasites can affect each other's
population dynamics in numerous ways. Parasites may affect
host reproduction or survival, and they may alter host
behavior in such a way as to increase transmission from one
host to the next. Hosts can affect parasite population
dynamics if they develop immunity to their parasites or
avoid contracting infections through behavioral mechanisms.
Here,
I examine the population dynamics of the parasites,
Eimeria arizonensis and E. delicata and their deer mouse
host, Peromyscus maniculatus.
Deer mice shed up to 106 E. delicata oocysts over the
course of a 9 day infection. They rapidly developed immunity
to E. delicata under laboratory conditions. In addition,
free-living adult deer mice were infected less frequently
than free-living juveniles. This pattern suggests that deer
mice also developed immunity to E. delicata in the field.
The prevalence of E. delicata was not correlated with
temperature or rainfall during its free-living stage.
Deer mice shed up to 107 E. arizonensis oocysts over
the course of a 10-12 day infection. They developed immunity
to E. arizonensis under laboratory conditions, although not
as rapidly as to E. delicata. However, several lines of
evidence suggest that acquired immunity to E. arizonensis
was more variable in the field: 1) the prevalence did not
decrease with deer mouse age, 2) animals frequently became
reinfected and the intensity of infections (number of
oocysts shed) decreased with subsequent infections only in
some years and 3) acquired immunity to experimental
infections in enclosures and the field was more variable
than under laboratory conditions. The prevalence of E.
arizonensis was strongly correlated with temperature during
its free-living stage, suggesting that this factor may be
more important than immunity in its population dynamics.
Eimeria arizonensis affected both deer mouse recruitment
(females) and over-winter survival (males).
Eimeria arizonensis and E.delicata population dynamics
were primarily affected by different factors, environmental
temperature and immunity, respectively. Because deer mice
rapidly developed immunity to E. delicata, adults were
probably able to escape adverse effects. In contrast, many
animals did not develop immunity to E. arizonensis and it
negatively affected their survival.
Host Parasite Relationships Between Deer Mice
(Peromyscus maniculatus) and Their Eimerian Parasites
(Protozoa)
by
Claire A. Fuller
A THESIS
submitted to
Oregon State University
in partial fulfillment of
the requirements for the
degree of
Doctor of Philosophy
Completed October 25, 1994
Commencement June 1995
Doctor of Philosophy thesis of Claire A. Fuller presented on
October 25, 1994
APPROVED:
Redacted for Privacy
/
MaDor Professor, representing Zoology-­
Redacted for Privacy
Chair of Department 6f Zoology
Redacted for Privacy
Dean of Gr
I understand that my thesis will become part of the
permanent collection of Oregon State University libraries.
My signature below authorizes release of my thesis to any
reader upon request.
Redacted for Privacy
Claire A. Fuller, Author
ACKNOWLEDGEMENTS
I am greatly indebted to my graduate committee, Andrew
R. Blaustein (chair), who was supportive throughout this
venture, Robert G. Anthony, Mark A. Hixon, Philippe A.
Rossignol, David McIntire (graduate representative, retired)
and Karen Timm (grad. rep.) for many helpful discussions and
for reading many drafts of my manuscripts. Sarah Fryer
served as an unofficial committee member and had all of my
ideas bounced off her.
This work could not have been completed without the
contribution of many people. First and foremost are Dan Edge
and Jerry Wolff. They not only allowed me to use their
enclosures at a critical juncture in my research, they also
were a source of field assistants and equipment. In
addition,
I had numerous undergraduate assistants without
whom the work would not have been possible, or nearly as
fun. Two of these became co-authors on chapter 2. Two people
assisted with species identification: Martin Adamson and Don
Duszynski. Don also helped me throughout this project by
reading manuscripts and allowing me to work in his
laboratory. The work was funded by the OSU Zoology Dept.,
OSU IACUC, Sigma Xi and the American Society of
Mammalogists. Friday Harbor Laboratories provided space and
peace of mind for writing.
My friends at OSU made my time there great fun and helped in
many aspects of the research. I'd especially like to thank Dave
Booth, Sarah Fryer, Maureen Larson, Dede Olson, Diane Rowe, Paul
Sikkel, Brian Tissot, Liz Walsh, Sedonia Washington, Jerry Wolff
and Ulrike Zelck. The Blaustein lab was a great place for ideas
to grow: discussions with Andy Blaustein, Grant Hokit, Joe
Kiesicker, Hugh Lefcort, Rick O'Hara, Dede Olson, Jeff Peterson
and Susan Walls were always interesting.
Finally,
I would like to thank my family for always
supporting me in my academic endeavors. Special thanks go to
Jeff, Joanne and Richard Fuller, Nancy Hayward, John Nelson, and
my husband, friend and colleague Paul Sikkel. My son Niko has
been the joy in my life for the past 17 months and has renewed my
belief that learning keeps the child in us alive.
CONTRIBUTION OF AUTHORS
Jerry Hefner and Elizabeth Wrosch were involved in the
design, sample collection and writing of chapter 2.
TABLE OF CONTENTS
Page
CHAPTER 1: GENERAL INTRODUCTION
1
Abstract
2
Introduction
2
Theoretical Studies
4
Empirical Studies (Level of the Individual Host)
7
Reproductive Success
Survival
Other Measures of Mortality
Transmission
Immunity
8
9
9
15
16
Population Level Empirical Studies
18
Conclusions
20
Overview of Thesis
22
Study Sites
Study Animal
CHAPTER 2: PARASITE LIFE HISTORIES
23
25
27
Abstract
28
Introduction
28
Methods
30
Results
33
Daily Oocyst Output
Diurnal Periodicity in Oocyst Output
Fecal Oocyst Concentration
Effect of Inoculurn Period and Level
Immunity
Discussion
Daily Oocyst Output
33
35
35
39
39
43
43
TABLE OF CONTENTS (Continued)
Page
Diurnal Periodicity in Oocyst Output and Fecal
Oocyst Concentration
Immunity
CHAPTER 3: POPULATION DYNAMICS OF E. ARIZONENSIS and
E. DELICATA
45
47
49
Abstract
50
Introduction
50
Methods
53
Data Collection
53
Statistical Analysis
55
Host Factors
Environmental Factors
Results
Overall Prevalence and Intensity
Hypothesis 1: Host. Factors
55
56
57
57
59
Prediction 1: Occurrence of Double Infections
59
Prediction 2a: Variability Among Sub-Populations
59
of Deer Mice, Age and Sex Differences
Prediction 2b: Differences Among Reproductive
64
Classes
Hypothesis 2: Environmental Factors
Discussion
Factors Influencing E. arizonensis
Factors Influencing E delicata
CHAPTER 4: THE EFFECT OF E. ARIZONENSIS ON DEER MOUSE
SURVIVAL
64
67
67
68
72
Abstract
73
Introduction
73
TABLE OF CONTENTS (Continued)
Page
Methods
Observations
Experiments
Eimeria arizonensis Cultures
Field Experiment
Enclosure Experiment
Statistical Analysis
Results
Prediction 1: Establishment of Residency
Prediction 2: Overwinter Survival
Field Observations
Enclosure Experiment
Prediction 3: Weight Changes
Field Observations
Field Experiment
Enclosure Experiments
Discussion
CHAPTER 5: VARIABLE LEVELS OF IMMUNITY TO E.
ARIZONENSIS
76
76
77
77
78
80
80
82
82
82
82
84
87
87
87
87
90
95
Abstract
96
Introduction
97
Methods: Field Observations
101
Results: Field Observations
102
Prediction 1, Complete Immunity
Prediction 2, Partial Immunity
Methods: Immunity Experiments
102
104
104
TABLE OF CONTENTS (Continued)
page
Laboratory Experiment
Field Experiment
Enclosure Experiment
Statistical Analyses
Results: Immunity Experiments
Effectiveness of inoculations
Laboratory Experiment
Comparisons Between Laboratory, Field and
Enclosure Results
Discussion
BIBLIOGRAPHY
106
106
108
109
111
111
111
114
114
121
LIST OF FIGURES
Figure
Page
1.1
Parameters Affecting Subpopulations of Deer Mice
2.1
Oocyst Output of E. arizonensis and E. delicata
34
2.2
Percent Daily Oocyst. Output
36
2.3
Total Oocyst Output Over a 20 Day Infection
41
3.1
The Prevalence of E. arizonensis and E. delicata
58
3.2
The Intensity of E. arizonensis and E. delicata
60
4.1
Over-Winter survival of Free-Living Deer Mice
85
5
4.2 Over-Winter Survival of Enclosure Deer Mice
86
4.3 Weight Change of Enclosure Deer Mice
89
5.1 Number of Times Individual, Free-Living Deer Mice Were
Infected With Naturally Occurring Eimeria arizonensis
Each Year
103
5.2 Change in Oocyst Output with Successive Infections.105
5.3 Experimental Infections in Naive Animals
5.4
Immunity to Experimental Infections in the Field
112
115
LIST OF TABLES
Table
1.1
1.2
Page
Studies Addressing Parasite-Mediated Population
Regulation in Vertebrates
10
Studies by Host and Parasite Taxa
21
2.1 Oocysts Per Fecal Pellet
37
2.2 ANOVA of the Number of Oocysts Excreted
40
2.3 ANOVA of the Reproductive Index
40
2.4 Immunity to E. arizonensis
42
2.5 A Comparison Between E. arizonensis and E. delicata
Life History Parameters
3.1 Frequency of Double Infections
48
61
3.2 Percent infected with E. arizonensis in different
sex and age classes of deer mice
62
3.3 Percent infected with E. delicata in different sex
and age classes of deer mice
63
3.4 Percent infected with E. arizonensis in different
reproductive classes of adult deer mice
65
3.5 Percent infected with E. delicata in different
reproductive classes of adult deer mice
66
4.1 Protocol for Field Experiment
79
4.2 Predictions and Experimental Paradigms
81
4.3 E. arizonensis and Deer Mouse Recruitment
83
4.4 Weight Change in Free-Living Deer Mice
88
5.1 Protocol for Laboratory and Field Experiments
107
5.2 Sample Sizes in Enclosure Experiments
110
5.3
Infections Among Sham Inoculated Animals
113
Host Parasite Relationship Between Deer Mice
(Peromyscus maniculatus) and their Eimerian Parasites
(Protozoa)
Chapter 1
General Introduction
2
Abstract
Parasites may regulate the population dynamics of their
hosts. In this review, I summarize mathematical models
addressing this topic and the parameters of the most basic
model are outlined. I then review empirical studies which
address model parameters at the level of the individual
host.
I next summarize studies which directly examine
whether parasites regulate host populations (ie., at the
population level). All empirical studies included in this
review examine macroparasite (Protozoa, helminths and
arthropods)
vertebrate systems and were conducted under
natural or semi-natural conditions. This review illuminates
several areas for further study including 1) experimental
studies at the population level, 2) experimental studies at
the level of the individual host and 3) studies which
include a greater spectrum of host and parasite taxa.
Introduction
The relative importance of factors regulating animal
populations is one of the most controversial and widely
discussed topics in ecology (Hestbeck 1987). Much of the
controversy has focused on emigration, competition and
predation. For example McMahon & Tash (1988) demonstrated
that emigration maintains desert pupfish population density
and Buskirk and Smith (1991) showed that intraspecific
3
competition was density dependent and negatively affected
salamander populations.
The potential for parasites to regulate their host
populations has also been established through mathematical
models (Anderson & May 1979, 1986, May & Anderson 1979, Aron
& May 1982). Moreover, many laboratory studies have
documented negative effects of parasites on individual hosts
and a few studies have shown that parasites can regulate
host populations under controlled laboratory conditions (see
reviews by Gregory & Keymer, 1989, Keymer & Read 1991, Scott
& Dobson 1989, Scott & Lewis 1987). However, many factors
differ between the laboratory and the field including food
levels, the presence of other parasites, predators and
competitors. Thus, while it is clear that parasites can
control host populations under laboratory conditions, it is
unclear from models and laboratory studies how important
parasites are in the presence of other factors such as food
limitation and predation.
Until recently, emprical studies of host-parasite
relationships conducted under field conditions have largely
been lacking. While a few long-term field experiments have
been conducted at the population level (see below), the
majority of recent studies have focused on specific aspects
of host-parasite interactions such as parasite-induced
mortality or host immune response. In this review,
I will
briefly outline models of animal population regulation by
parasites. I will then summarize studies of parasites in
4
vertebrate hosts conducted under natural or semi-natural
(ie., field enclosures) conditions which examine specific
parameters of the models at the level of the individual
host. Finally,
I will discuss studies conducted at the
population level under natural and semi-natural conditions.
Theoretical Studies
Until 1979, epidemiological models of host-parasite
systems were static, examining one point in time (Anderson &
May 1979) . Anderson & May (1979) and May & Anderson (1979)
modelled changes in host and parasite populations over time.
Figure 1.1 (modified from Anderson & May 1979) shows the
parameters of the basic model (Anderson & May 1979). Animals
susceptible to, infected with and immune to parasites are
represented by X,
Y and Z, respectively (a = intrinsic host
birth rate and b = the intrinsic host death rate. Alpha =
parasite induced death rate). Beta, v and y represent the
rate at which hosts move between states X, Y and Z by
transmission (B), recovery (v) and loss of immunity (y).
There are several assumptions of the basic model which have
been addressed in subsequent models. For example, X,
Y and Z
are assumed to have the same birth rate; Dobson (1988)
incorporated parasite induced reduction in host reproductive
success. Another assumption is that all susceptible hosts(X)
are equally susceptible to parasitism; Anderson & May (1985)
5
.___._._.____...------T----­
births (a)
deaths
(b + alpha)
deaths (b)
(B)
X
Y
(susceptible)
(infected)
(v)
(y)
Z
(immune)
deaths (b)
Figure 1.1. Parameters affecting subpopulations of deer
mice. Parameters from the basic model of Anderson and May
(1979) on parasite-mediated population regulation. B, v and
y represent transmission, recovery and loss of immunity,
respectively and are the rates at which animals move between
subpopulations X, Y and Z. Arrows leading from the circle
represent host deaths. Susceptible and immune hosts die at
some intrinsic rate (b). Infected hosts have an additional,
parasite-induced death rate (alpha). All host subpopulations
are assumed to have the same intrinsic birth rate (a) and
offspring are assumed to be susceptible (ie., no vertical
transmission of parasites or immunity).
6
incorporated genetic heterogeneity in host
susceptibility.
Anderson & May (1979) also discussed two fundamentally
different types of parasites. Microparasites (viruses &
bacteria) reproduce within the host, recovered hosts become
completely immune and immunity can last the host's lifetime.
Thus, the prevalence (% infected) of microparasites is often
used in epidemiology studies (Aron & May 1982).
Macroparasites (helminths and arthropods) generally do not
complete a reproductive cycle within a host, immunity is
density dependent (ie., immunity increases with increasing
exposure) and is generally lost at some rate (Anderson
1985). Thus, because macroparasite infections are not all­
or-nothing, the intensity of infection (# parasites per
host) is thought to be a better measure in epidemiological
studies (Anderson 1986). Protozoans reproduce like
microparasites but generally elicit density-dependent
immunity, thus, they do not fit neatly into either category.
Although Anderson and May (1979) originally included
protozoans as microparasites, subsequent work has generally
treated them as macroparasites (Anderson 1991, Aron & May
1982, Nowell & Higgs 1985).
Several conclusions can be drawn from Anderson and
May's model: 1) there is a certain baseline host population
density below which a parasite cannot persist, 2) this level
decreases with increasing transmission and 3)
increases with
increasing pathogenicity. Finally, for a parasite to
7
regulate its host population, parasites must be
overdispersed such that only a few hosts carry the majority
of parasites, parasite induced mortality must increase with
parasite burden and parasite survival/reproduction must be
density dependent within individual hosts (Gregory & Keymer
1989)
.
Empirical Studies (Level of Individual Host)
I have limited the scope of the studies reviewed to
vertebrate-macroparasite systems, because 1) acquired
immunity will play a role in these systems and 2)
vertebrate-microparasite studies have been summarized
elsewhere (Minchella & Scott 1991, Scott 1988). In addition,
to avoid anecdotal accounts, I have only included
observational studies which examine at least 25 hosts.
Finally, a great deal of recent work has addressed the
effect of parasites on host behavior, especially on sexual
selection. For example, parasites affect behavior of the
hosts such that transmission is facilitated or they weaken
hosts such that sexual displays are affected. In addition,
females may avoid mating with parasitized males. These
studies have been reviewed recently (Combes 1991, Dobson
1988, Hart 1990, Moore 1987) and are not included here.
Examination of Anderson & Mays' models suggests that
there are 5 basic parameters which could affect parasites'
ability to determine host population size. The parameters
8
can be divided into pathogenicity factors (effects on
reproduction or survival) and susceptibility factors (rate
of parasite transmission [B], recovery [v] and loss [y] of
immunity. Table 1.1 summarizes studies that address these
parameters by host and parasite taxa.
Reproductive Success
Most studies have examined the effect of parasites on
correlates of reproductive success such as pregnancy rate
(Boonstra et al. 1980, Theis & Schwab 1992), clutch size
(Clinchy & Barker 1994, Dobson et al. 1992, McPhail &
Peacock 1983, Poiani 1993, Schall 1983), testis size
(Boonstra et al. 1980, Timm & Cook 1979), dominance (Schall
& Dearing 1987), or nestling weight (de Lope et al. 1993, de
Lope & Moller 1993, Moore & Bell 1983). A few studies have
documented parasite effects on fledging success (Howe 1992,
Hudson 1986, de Lope et al. 1993, de Lope & Moller 1993,
Moller 1993). Four of the studies on reproductive success
have included field manipulations of nest mite (de Lope et
al. 1993, de Lope & Moller 1993, Moller 1993) or nematode
(Hudson 1986) levels and all 4 of these studies showed
significant negative effects of parasites on fledging
success (all in birds).
9
Survival
Several studies have measured the effect of parasites
on adult survival using recapture data (Boonstra 1977,
Boonstra et al. 1980, Gulland 1992, Hudson et al. 1992 a, b,
Keith et al. 1986, Lehmann 1992, Miller & Getz 1969, Moss et
al. 1990, Samson et al. 1987, Weatherhead & Bennett 1992).
Other studies have used statistical methods comparing
expected and observed distribution of parasites in host
populations (Adjei et al. 1986, Dobson et al. 1992, Gordon &
Rau 1982; method sensu Croften 1971, Pacala & Dobson 1988).
Temple (1987) showed that a predator (red-tailed hawk)
captured highly parasitized prey in prey species which were
difficult to capture but not in species that were easy to
capture (based on number of strikes to capture). Only 3 of
the studies included in this survey manipulated parasite
burdens to determine whether there is a causal relationship
between parasites and host mortality (Lehmann 1992, Hudson
et al. 1992b, Samson et al. 1987). All three studies showed
that arthropod or nematode parasites decreased host
survival.
Other Measures of Mortality
Several authors have examined mechanisms through which
parasites might influence host survival or reproduction
including:
1) weight change associated with parasitism
(Bennett et al. 1988, Booth et al. 1993, Boonstra et al.
10
Table 1.1. Studies addressing parasite-mediated population
regulation in vertebrates (sensu Anderson & May 1979).
Parasites taxa include Protozoa (P), Acanthocephala (Ac),
Cestoda (C), Nematoda (N), Trematoda (T) and Arthropoda
(Ar). 0 indicates observational studies and E indicates
experimental studies. Under headings reproduction,survival
and other, (-) indicates a negative effect of the parasite,
0 indicates no effect. Under transmission, parameters
measured are indicated by B = behavioral, H = habitat
differences, R = reinfestation time, S = seasonal variation,
Sury = free-living parasite survival,T = temperature
variation, V = vertical transmission. Under immunity, A-I
and A-P = age-intensity or age-prevalence, spleen = spleen
size, RE = changes in immunity with reproductive effort.
Table 1.1
11
Host
Parasite
Repro
Sury
er
Trans
Immunity
References
Fish
Gasterosteus aculeatus
(3-spine stickleback)
Schistocephalus
solidus (C)
Culaea inconstans
(brook stickleback)
Apatemon qracilis
Saurida tumbil
S. undosquamis
(lizard fish)
Callitetrarhynchus
qracilis (Ac?)
(
0,
McPhail & Peacock 1983
0,
Gordon & Rau 1982
0,­
0,-
Adjei et al. 1986
)
Amphibians
Scaphiopus couchii
(spade-foot toad)
Pseudodiplorchis
americanus (T)
0, T
0,Y A-I
Tinsley 1990, Tocque 1993,
Tocque & Tinsley 1991
Reptiles
Sceloporus occidentalis
(western fence lizard)
Plasmodium
mexicanum (P)
Anolis lineatopus
Centrorhynchus
spinosus (Ac)
Mesocoelium
danforthi (T)
Cyrtosumum
sceloperi (N)
Thelandros
cubensis (N)
(anole lizard)
Anolis spp.
various helminths
o,
Schall 1983, Schall & Dearing
1987, Schall et al. 1983
0,N A-P/I
Vogel & Bundy 1')87
0,N A-P/I
0,N A-P/I
0,N A-P/I
o,
o,
0, H
Dobson et al. 1992
Birds
Various passerines
Hematozoa (P)
Oreosoptes montanus
(sage-thrasher)
Protocalliphora
braueri (Ar)
Ficedula hypoleuca
(pied flycatcher)
Ceratophyllus
qallinae (Ar)
Delichon urbica
Oeciarus
hirundinis (Ar)
E,
Plaqiorhynchus
cylindraceus (Ac)
0,
(house martin)
Sturnus vulqaris
(starling)
Troglodytes aedon
(house wren)
Protocalliphora
pavarom (Ar)
Bennett et al. 1988
0, -/0
Howe 1992
E, B
Mappes et al. 1994
de Lope et al. 1993
Moore & Bell 1983
Johnson et al. 1991
Table 1.1, cont.
12
Host
Parasite
Agelaius phoeniceus
(red-winged blackbird)
Hematozoa (P)
Molothrus ater
(brown-headed cowbirds)
Parus major
Hematozoa (P)
Repro
Trans
Sury
0,N A-P
0,0
0,0
(swallow)
Hematozoa (P)
0,Y RE
Oeciacus
hirundinis (Ar)
Ornthonyssus bursa
References
Weatherhead & Bennett 1991
Weatherhead & Bennett 1992
(great tit)
Hirundo rustica
Immunity
E,
Norris et al. 1994
de Lope & Moller 1993, Moller
1993
(Ar)
Columba livia
(rock dove)
Ischnocera sp.
Booth et al. 1993
(Ar)
Anser caerulesceus
(lesser snowgoose)
various (N,T,P)
Laqopus laqopus
Trichostronqylus
tenius (N)
(red grouse)
0,0
Clinchy & Barker 1994
0,0
0,-/E,­
0,SAT
0,N A -P
0,-/E,­
Moss et al 1990,1993, Shaw &
Moss 1989, Wilson 1983, Hudson
1986, Hudson et al. 1992 a,b
Mammals
Microtus montanus
(montane vole)
M. townsendii
(Townsend's vole)
Babesia microti
0,Y Spleen Watkins et al. 1991
(P)
0,
V.,31fartia vigil
(Ar)
Cuterebra qrisea
0,
0,­
Boonstra 1977
0,­
Boonstra et al. 1980
(Ar)
M. aqrestis
(short-tailed vole)
Clethrionomys qlareolus
Babesia (P)
0,N A-P
Hepatozoa (P)
0,S
(bank vole)
C. q. skomerensis
(island bank vole)
C. rutilis
(red vole)
Trypanosoma (P)
Healing 1981,
Haukisalmi et al. 1988
0,Y A -P
various helminths
Apodemus sylvaticus
(wood mouse)
Eimeria spp.
0,S
(P)
Heliqmosomoides
polyqyrus (N)
Peromyscus leucopus
(white-footed mouse)
Cuterebra sp.
Hymenolepis
citelli (C)
(Ar)
0,Y A-I
Ball & Lewis 1984
Gregory 1992, Gregory et al.
1992, Quinnell 1992, Montgomery
& Montgomery 1988
O,
0,0
0,
0,Y Spleen Miller & Getz 1969, Munger &
Karasov 1989, Payne et al. 1965,
Timm & Cook 1979
Munger & Karasov 1994
Table 1.1, cont.
13
Host
P. maniculatus
(deer mouse)
Parasite
Taenia
taeniaeformis (C)
Capillaria
hepatica (T)
Ixodes dammini
Repro
Sury
Ot
0,0
(Ar)
ectoparasites
Rattus norveqicus
(brown rat)
Siqmodon hispidus
(cotton rat)
Gerbillus andersoni
Toxoplasma gondii
(P)
Trans
ectoparasites (Ar)
0,N A-P
Theis & Schwab 1992
0, S
0,N A-P
Herman 1981
0, H
Adler et al. 1992
E,R
Glicken & Schwab 1980
0, V
Webster 1994
Spermophilus elegans
(Wyoming ground squirrel)
Eimeria spp.
S. townsendii
(Townsend's ground
squirrel)
Eimeria spp.
Tamius striatus
(eastern chipmunk)
Sylvilaqus floridanus
(cottontail rabbit)
Sciurus carolinensis
total parasite
burden
0,varies
E,-
References
0, S
blood parasites
(gerbil)
Immunity
E,'
Lochmiller et al. 1994
Lehmann 1992
0, H
0,N A-P
Stanton et al. 1992
0, S
0,N A-P
Wilber et al. 1994
0, var
Temple 1987
(gray squirrel)
Myocaster coypus
Eimeria spp.
0, S
(nutria)
Lepus americanus
(snowshoe hare)
Obeliscoides
cuniculi (T)
Nematodirus
trianqularis (N)
Trichuris leporis
0,­
0,0
(C)
0,
0,0
0,
Eimeria spp.
Ovis aries
(Soay sheep)
(P)
Ostertraqia
circumcincta (N)
Trichostronqvlus
sp.
0,0
0,0
o,
Protostronqvlus
stilesi (N)
P. sp.
0,Y A-P
0, S
0,Y A-P
0,Y A-P
0
0,Y A-P
0, S
0, S
(N)
0,Y A-P
Gulland 1992, Gulland & Fox 1992
0,Y A-P
E,Sury
0, S
(?)
(bighorn sheep)
Keith et al. 1985, 1986
E, Sury
Cooperia carticii
O. canadensis
0,N A-P
0,Y A-P
0,0
(N)
Protostronqvlus
boughtoni (N)
Taenia pisiformis
0, S
Ball & Lewis 1984
0,Y A -P
E,sury
E,
0, S
0,Y A-I
Arnett et al. 1993 Samson et al.
0,Y RE
Festa-Bianchet 1989
1987
Table 1.1, cont.
14
Host
Odocoileus virqinianus
(white-tailed deer)
Parasite
Repro
Su
Other
Trans
Immunity
References
Fascioloides magna
(T)
Eimeria mccordocki
0,0
Mulvey et al. 1994
(2)
Ranqifer tarandus
(reindeer)
Alopex laqopus
(arctic fox)
Homo sapiens
(humans)
Hypoderma tarandi
0,Y A-P
Samuel & Trainer 1971
E,Y Sex
Folstad et al. 1989
0,Y A-P
Prestrud et al. 1993
0,Y A-I
Fulford et al. 1992
0,Y A-P
Woolhouse et al. 1991
(Ar)
Trichinella sp.
(N)
Schistosoma
mansoni (T)
S. haemotobium (T)
0,0
15
1980, Keith et al 1986, Lehmann 1992, Mulvey et al. 1994,
Munger & Karasov 1994, Prestrud et al. 1993, Weatherhead &
Bennett 1992), 2)
changes in metabolic rate (Booth et al.
1993, Munger & Karasov 1989, 1994),
3) changes in the
relative abundance of plasma proteins (Payne et al. 1965),
4) anemia (Lehmann 1992, Schall et al. 1982, Tocque 1993)
and 5) changes in conception date (Mulvey et al. 1994).
Three studies included in this survey were experimental
(Booth et al. 1993, Lehmann 1992, Munger & Karasov 1989)
with variable results (see Table 1.1).
In general, one problem with these studies is that it
is often unclear whether parasite induced changes are
actually associated with mortality and to what degree
(Munger & Karasov 1989). Thus, unless some measure of
survivorship is also included, the studies are difficult to
interpret.
Transmission
Transmission of parasites from one host to another is
especially difficult to measure in the field because one
must know how many of the available parasites are actually
encountered and subsequently recruit into the host. In some
cases, vertical transmission (when a parasite passes to
offspring through the placenta) may also occur (Webster
1994). Where the parasite's life-cycle includes free-living
stages, it is also relevant to know the survival time of the
16
free-living parasite under various environmental conditions.
Most field studies which touch on the subject of
transmission document the association between the prevalence
or intensity of infections and temporal changes (Ball &
Lewis 1984, Gregory 1992, Haukisalmi et al. 1988, Moss et
al. 1993), differences in habitats (eg., xeric vs. mesic;
Adler et al. 1992, Dobson et al. 1992, Stanton et al. 1992)
or weather variables (Moss et al. 1993, Tocque & Tinsley
1991). A few studies have examined survival of free-living
stages (Levine 1980, Gulland & Fox 1992) and reinfestation
after removal (Glicken & Schwab 1980). Hosts may also avoid
(Mappes et al. 1994) or be manipulated by parasites (Moore &
Bell 1983). Behavioral mechanisms of parasite avoidance and
parasite manipulation of hosts to increase transmission have
been reviewed by Combes (1991), Dobson (1988), Hart (1990)
and Moore (1987).
Immunity
Vertebrate hosts clearly become immune to a wide variety
of parasites under laboratory conditions (Gregory, Keymer &
Clarke 1990, Higgs & Nowell 1988, Lillehoj 1988, Wakelin
1978, 1987, Wassom et al. 1986). However, it is
less clear
how frequently immunity occurs under field conditions. Both
genetic (Wakelin 1992) and environmental (McCallum 1990)
factors may modify immunocompetence. Several methods have
been used to detect immunity including serological measures
17
(Elliott et al. 1994, Lochmiller et al. 1994), increases in
spleen size (Timm & Cook 1979, Watkins et al. 1991) and the
presence of convex age-intensity/age-prevalence curves
(Arnett et al. 1993, Ball & Lewis 1984, Fulford et al. 1992,
Gulland & Fox 1992, Haukisalmi et al. 1988, Healing 1981,
Hermann 1981, Keith et al. 1986, Quinnell 1992, Prestrud et
al. 1993, Samuel & Trainer 1971, Shaw & Moss 1989, Stanton
et al. 1992, Tinsley 1990, Vogel & Bundy 1987, Watkins et
al. 1991, Weatherhead & Bennett 1991, Wilber et al. 1994).
The first 2 methods only detect previous exposure or general
immune readiness, rather than current immune response to a
specific pathogen. The last method assumes that exposure
increases with age after an initial period of low juvenile
exposure. Thus, as animals first become exposed, parasite
burden increases; as immunity develops, parasite burden
decreases. This method is widely employed (used by 20 of 24
authors included here), but convex age-intensity curves can
be caused by other factors (Bundy 1988, Fulford et. al 1992,
Pacala & Dobson 1988).
Two studies of immunity included here were
experimental. However, there were problems with both.
Folstad et al.
(1989) examined differences in intensity
among animals with different hormonal levels. Thus,
behavioral differences among age or sex classes could
explain differences in parasite burdens. Quinnell (1992)
documented immunity in enclosure populations of wood mice
(Apodemus sylvaticus). However, except for natural
18
temperature and lighting, enclosures did not mimic natural
environments. In addition, challenge infections (a second
experimental infection given to the same animal designed to
test the immune response to the first infection) were
conducted in the laboratory. Thus, harsh environmental
conditions which could compromise the immune response
probably were not adequately imitated.
Population Level Empirical Studies
Three studies have examined host population regulation
by macroparasites in vertebrate hosts at the population
level. All examined mammal-nematode systems; 1 used
comparative data and the other 2 were experiments conducted
in field enclosures.
The nematode Parelaphostronqylus tenius causes
neurological damage in moose but not in deer. It has been
hypothesized that moose decline in the presence of deer and
that this decline is caused by increased transmission of P.
tenius from deer to moose with increasing deer population
density (Whitlaw & Lankester 1994). Whitlaw and Lankester
used reports of deer and moose densities and moose
neurological disease on 6 sites over an 80-year period to
test this hypothesis. They found that moose and deer
densities were negatively correlated but there was no
significant correlation between moose declines and reports
of P. tenius related disease. They critique the quality of
19
their data set and conclude that "historical data are
unlikely to provide further insight" in this system.
Barker et al.
(1991) and Gregory (1991) used enclosures
to examine population regulation in the house mouse (Mus
musculus)-Capillaria hepatica system and the wood mouse
(Apodemus sylvaticus)-Heliqmosomoides polygyrus system,
respectively. Barker et al.
(1991) found little evidence of
population regulation by C. hepatica; control (n=3) and
treatment (n=3) populations fluctuated synchronously and
overall population densities, reproduction and survival did
not differ between control and treatment populations over
the 1.5 year study. They suggested that food quality or
intraspecific competition may have regulated mouse
population density and masked any effect of C. hepatica.
Gregory (1991) found that control populations (n=3) grew
faster and had significantly greater host densities at the
end of the 2-year study than treatment (n=3) populations.
Reproduction was not affected, rather treatment animals did
not survive as well after weaning as control animals.
Although these studies were closer to field conditions than
laboratory studies, in many ways the enclosures were closer
to the laboratory than the field. Enclosures were small
(Barker et al. = 15 x 15 m; Gregory = 9 x 4 m), food and
water were supplied ad libitum and predators and
interspecific competitors were excluded. While Gregory
(1991) demonstrated that H. polyqyrus can regulate wood
mouse populations in the absence of other factors, he does
20
not provide information about the relative importance of
these parasites in population regulation.
Conclusions
1) Future studies should include a wider variety of
host and parasite taxa. The majority of field work has
focused on avian and mammalian hosts (Table 1.2). Even
within these groups, only a few host orders have been
examined. The majority of studies in mammals have been
conducted on rodents and ungulates; in birds most hosts have
been passerines. The parasites examined have also had a
relatively narrow focus in birds: most studies have been on
nest mites and hematozoa (Protozoa). In mammals the scope of
the parasites studied is substantially broader and includes
protozoans, cestodes, trematodes, nematodes and arthropods.
The taxonomic scope of experimental studies is particularly
limited (Table 1.2).
2) More studies should address all individual
parameters of population regulation but especially
transmission and immunity. The aspects of population
regulation that have been addressed include all model
parameters, however, the majority of work has focused on
parasite-induced host mortality. Other than documenting
seasonal variation in parasites and age-intensity curves,
very few studies have examined transmission dynamics or
acquired immunity.
21
Table 1.2. Studies by host and parasite taxa. The number of
observational (0) and experimental (E) studies addressing
various model parameters in host and parasite species. Repro
= parasite effects on host reproduction, Sury = parasite
effects on host survival, Other = parasite effects on other
aspects of host fitness (eg., weight loss), Trans =
transmission and Immun = host immune response.
Repro
Other
Sury
Trans
Immun
Taxonomic group
0
E
1
0
E
0
0
E
0
E
0
E
1
0
1
0
1
0
Host Taxa
Fish
3
0
Amphibians
Reptiles
2
0
1
0
1
0
1
0
1
0
Birds
4
3
3
1
3
1
2
1
2
0
Mammals
3
0
6
2
5
2
10
1
14
2
Protozoa
2
0
2
0
4
0
6
0
8
0
Acanthocephala
1
0
1
0
Cestoda
2
0
1
0
1
1
1
0
2
0
Trematoda
1
0
2
0
3
1
2
0
5
0
Nematoda
2
1
5
3
4
0
6
0
11
1
Arthropoda
3
2
3
1
3
2
1
1
1
1
Parasite Taxa
22
3) Field experiments are critical. Keymer and Read
(1991) called for field experiments to test the causal
relationship between parasites and their hosts. Although
most of the 52 studies reviewed here did not incorporate
field manipulations, those that did were relatively recent
(10 of 12 were published after 1988,
8 after 1991). Thus,
experimental evidence about the ability of parasites to
regulate host populations is beginning to accumulate. Within
the experimental studies, only 3 of 12 examined transmission
or immunity. All others examined various aspects of
mortality.
4) Long-term experiments at the population level are
needed. Two of the 3 studies of parasite-mediated population
regulation were inconclusive (Barker et al. 1991, Whitlaw &
Lankester 1994). The third provided evidence that parasites
can regulate hosts but did not address the conditions under
which this might occur (Gregory 1991). In addition, studies
should follow several host generations to adequately assess
the impact of parasites on population dynamics.
Overview of Thesis
I examined the host-parasite interactions of deer mice
(Peromyscus maniculatus) and its intestinal protozoans
(Eimeria arizonensis and E. delicata). My dissertation
provides new information in several of the areas addressed
above:
23
1) Most studies of protozoans have focused on blood
parasites. Little is known about other groups in this
Kingdom.
2)
I have examined several of the individual parameters
including parasite-induced host mortality, transmission
dynamics and acquired immunity.
3) The second chapter provides information on parasite life-
history and was conducted in the laboratory. All other
chapters were conducted in the field.
4) Chapters 4 and 5 include field experiments on survival
and acquired immunity, respectively. These represent the
first experimental field work on protozoan parasites. To my
knowledge, the immunity experiments are the first to address
whether individual hosts acquire immunity under natural and
semi-natural conditions in any host or parasite taxa.
Study Sites
Observational and experimental studies on free-living
animals were conducted in MacDonald Forest, Oregon State
University's experimental forest (approx. 15 km NW,
Corvallis, OR). The elevation at the trapping sites ranged
from 500-1500 ft. All areas trapped were chosen to be
similar in the composition of dominant species. Dominant
trees included Douglas fir (Psuedotsuqa menziesii), grand
fir (Abies qrandis), big leaf maple (Acer macrophyllum) and
vine maple (Acer circinatum). Shrubs included Oregon grape
24
(Berberis nervosa) and salal (Gaultheria shallon); other
common understory plants included poison oak (Rhus
diversiloba), sword fern (Polystichum munitum) and moss.
Many other species of small mammals co-occurred with
deer mice. Common species (captures at least 1 in 200 trap
nights) included voles (Microtus oregoni and Clethrionomys
californicus), Townsend's chipmunk (Tamias townsendii),
flying squirrels (Glaucomys sabrinus) shrews (Sorex vaqrans
and S. trowbridqii) and shrew-moles (Neurotrichus qibsii).
Species caught infrequently (< 1 in 1000 trap nights)
included red tree voles (Phenacomys lonqicaudus), bushy-
tailed woodrats (Neotoma cinerea), Pacific jumping mice
(Zapus trinotatus), Douglas' squirrels (Tamiasciurus
douqlasii), coast moles (Scapanus orarius) and ermine
(Mustela erminea).
Deer mouse predators known to occur in this area
include weasels (M. erminea and M. frenata), skunks
(Mephitis mephitis), raccoons (Procyon lotor), coyotes
(Canis latrans), bobcats (Felis rufus), garter snakes
(Thamnophis sertalis and T. eleqans), gopher snakes
(Pituophis melanoleucus) and spotted, screech, long-eared
and great horned owls (Strix occidentalis, Glaucidium qroma,
Otus asio, Asio otus and Bubo virqinianus, respectively).
The research site for the enclosure studies was located
at the Hyslop Experimental Farm of Oregon State University,
approx. 10 km north of Corvallis, OR. Twenty-four 0.2 ha
(0.5 acre) enclosures were constructed at the research site;
25
4 were used in my research. Each enclosure was 45 x 45 m and
was constructed of galvanized sheet metal approx. 90 cm high
and 90 cm deep to prevent escape of or entry by burrowing
animals. Each enclosure was planted with alfalfa in summer
1991. During the course of my study, vegetation was
maintained below a stand height of 0.4 m. A 1 m wide strip
along the inside fence of each enclosure was kept bare or
mowed short to reduce usage and therefore minimize escape
rates of small mammals.
One hundred trap stations set in a 10 x 10 array at 5 m
intervals were established in each enclosure. Each station
was marked by a 70 cm high wire flag. One Sherman live trap
was placed at each trapping station for a total of 100
traps/enclosure.
Study Animal
The biology of Peromyscus, including deer mice (P.
maniculatus) has been reviewed by King (1968) and Kirkland
and Layne (1989). Deer mice are new world rodents in the
Family Muridae, Subfamily Sigmodontinae (Carleton 1989).
Gestation lasts 24 days and litters range between 2-8 pups
(mean = 4.5; Millar 1989). Laboratory-reared pups from the
MacDonald Forest population could be weaned as early as 18
days and both sexes can become reproductively mature as
early as 60 days (pers. obs.). Typically, juvenile females
remain on their natal home range and juvenile males disperse
26
before they become reproductively mature (Wolff 1989).
Breeding tends to be seasonal with the majority of
reproduction occurring in spring and fall, however, a small
percentage of animals continue to breed throughout the year
in many years. Typically, females born in the spring begin
to reproduce in the fall but females born in the fall do not
reproduce until the following spring (pers. obs.). Mating
systems are generally promiscuous, ie., both sexes mate with
several different partners (Wolff 1989).
Deer mice live in a wide variety of habitats including
meadows, clear-cuts, riparian stream beds, and in old growth
(MacMillen & Garland 1989). At most densities they have
overlapping home ranges of up to 750 m2. However, aggression
increases with density and at high densities during the
breeding season, they exhibit territoriality (Wolff 1989).
During colder weather, non-reproductive animals may nest in
groups of up to 4. They primarily eat seeds and arthropods,
but will also eat conspecific young (pers. obs.).
Deer mouse populations exhibit annual density cycles
with peaks occurring in the spring. Several factors are
thought to affect population density including food
availability, intra- and interspecific competition and
predation (Kaufman & Kaufman 1989).
27
Chapter 2
Parasite Life Histories
C.A. Fuller, J. Hefner and E. Wrosch
In Press, Journal of Parasitology
28
Abstract
We compared the life histories of 2 eimerians (Eimeria
arizonensis and E. delicata) which co-occur in the deer
mouse (Peromyscus maniculatus). Laboratory-reared deer mice
were given 103, 104
(E. delicata), or 103,
104, or 105
(E.
arizonensis) oocysts by stomach intubation. Eimeria
arizonensis infections lasted longer (11-13 days) than E.
delicata infections (9-10 days). Eimeria arizonensis
infections also produced more oocysts for each oocyst
ingested at both levels compared. Both parasites exhibited
periodicity in oocyst output at all levels compared.
However, peaks in E. arizonensis output occurred at
approximately 20-24 hr intervals whereas peaks in E.
delicata output occurred at 12-16 hr intervals. Finally,
deer mice developed immunity to both eimerians after only 1
inoculation at all levels tested. Based on these parameters,
we expect E. arizonensis to have a greater reproductive
potential than E. delicata in free-living deer mice.
Introduction
Many aspects of host-parasite relationships are
receiving increasing attention from ecologists including
population regulation of animals by their parasites (Scott
and Dobson, 1989; Gregory, 1991; Holt, 1993), host-parasite
coevolution (Reduker et al., 1987; Gardner, 1991) and the
effect of parasites on host behavior (Schall and Dearing,
29
1987; Dobson, 1988; Brodeur and McNeil, 1989; Moore et al.,
1994). To conduct basic ecological research in these areas,
it is vital to obtain information about parasite life
histories and the interaction between the host environment
and the parasite. For example, the age and immune status of
a host population may affect a parasite's ability to invade
those hosts and thus will influence the population dynamics
of both the host and/or parasite (Anderson, 1991).
Eimerians (Apicomplexa: Coccidia) are excellent
parasites to use in ecological studies in the laboratory and
in the field. They are easily maintained and experimental
infections are easily produced. Because they are generally
parasites of the gastrointestinal tract which shed oocysts
in their host's feces, long-term, non-invasive monitoring of
infections can be conducted. Finally, infections generally
are of short duration (usually < 20 days), so it is possible
to follow repeated infections in individual hosts.
Eimerian parasites of domestic animals, e.g., poultry
and livestock, have received a great deal of attention
(Catchpole, et. al 1976; Rose, 1987; Augustine, 1988). In
these systems, much is known about the effect of host age
and immune response, inoculum level and periodicity. Much
less is known about coccidians which occur exclusively in
natural populations. For example, hundreds of rodent
coccidians have been described (Levine and Ivens, 1989);
however, only a few studies have documented aspects of their
30
life histories (Tilahun and Stockdale, 1981; Higgs and
Nowell, 1988) .
Here, we present a comparison of the infection patterns
caused by Eimeria arizonensis and E. delicata in the deer
mouse, Peromyscus maniculatus. The prepatent (the period
between ingestion of infective oocysts and shedding of
oocysts) and patent (period during which oocysts are shed)
periods, periodicity of oocyst output and the effect of a
prior infection were investigated for both eimerians.
Methods
Oocysts of Eimeria arizonensis were isolated from the
feces of 2 P. maniculatus live-trapped from MacDonald
Douglas Forest (14 km NW, Corvallis, Benton County OR) in
May 1990. Oocysts
of Eimeria delicata were isolated from 8
P. maniculatus from the same population in May-September
1991. Sporulated oocysts of both species were maintained at
4 C in 2.5% (w/v) aqueous potassium dichromate (K2Cr2O7)
solution and new oocysts were generated periodically in
captive P. maniculatus (see below). Oocysts were no more
than 7-wk-old at the time of inoculation.
Ten male and 10 female P. maniculatus collected from
MacDonald Douglas Forest in the fall of 1989 were used to
start a coccidian-free colony. Animals from the colony were
never observed to shed oocysts and all hosts used in this
study were 3rd-5th generation laboratory-reared. Deer mice
31
were verified to be coccidian-free by microscopic
examination of feces several times before inoculation. All
deer mice were housed in hanging wire mesh cages, provided
with nesting material and Purina Mouse Breeder Chow and
water ad lib. and a pan was suspended from each cage to
collect fecal samples. The room was on a 8L:16D light-dark
cycle; the temperature was maintained between 24-28 C. These
conditions approximate those found May
September in
western Oregon.
Twenty-five, 9-wk-old P. maniculatus were randomly
divided into 5 groups, anesthetized with metafane and
inoculated by stomach intubation with either 103,
104, or 105
sporulated E. arizonensis oocysts, or with 103 or 104
sporulated E. delicata oocysts (N = 5 in all groups). Feces
were collected from each individual deer mouse for 20 days
PI in 24 + 0.25 hr periods. Samples were stored in 2.5%
(w/v) K2Cr207 solution at 4 C until examined.
Feces were collected in 4 hr sampling periods on days
5-7 PI to determine whether periodicity in oocyst output
occurred. These samples were collected in dry pans and
weighed before suspension in K2Cr207 solution.
Fecal samples were homogenized and filtered through a
285 um mesh screen, allowed to settle and then resuspended
in a total volume of 10 or 50 ml water (for 4 and 24 hr
samples, respectively). The oocysts in 2 subsamples were
counted using MacMaster's counting chambers using sugar for
flotation (Dunn & Keymer, 1986). Samples from each
32
consecutive day were examined until no oocysts were seen for
3 days and every 2-3 days thereafter until day 20 PI. If no
oocysts were detected in 4 hr samples, the entire sample was
examined using Sheather's sugar flotation method (Ash &
Orihel 1987).
On day 21 PI all animals (except 1 animal originally
inoculated with 104 E. arizonensis oocysts which died during
the challenge inoculation) were reinoculated with the same
number and species of oocysts as in the initial inoculation.
In addition, 18 naive control animals were similarly
inoculated (N: E. arizonensis, 103=4,
104=5, 105=5; E.
delicata, 103=4). Feces from individual animals were
collected for 20 days PI. Because these samples were very
large, they were filtered through a graded series of brass
mesh filters (30 and 60 mesh) and stored in 2.5% (w/v)
K2Cr2O7 solution until examined. The filtrate was suspended
in 100-500 ml water and 4-5 subsamples were examined using
MacMaster's chambers. If no oocysts were detected, 2,
5 ml
samples were examined by Sheather's sugar flotation method.
There were clear differences in the number of oocysts
shed by naive treatment animals during the first inoculation
and naive control animals during the challenge inoculation.
Thus, inoculation period (first inoculation or challenge
inoculation) was included as a factor in all analyses of
total oocyst output and reproductive index. The oocyst
output of E. arizonensis was compared across treatment
levels by 2-Way ANOVA. Levels of the treatment were compared
33
using the Least Significant Difference test. T-tests were
used to compare output of E. delicata oocysts between
animals inoculated with 103 oocysts (first and challenge
inoculations) and between animals inoculated with 104
oocysts (first inoculation only). Where necessary, data were
arcsin and log transformed to meet the assumptions of
parametric analyses. In all cases, alpha of 0.05 was
considered significant.
Results
Daily Oocyst Output
The prepatent period of E. arizonensis was 4 days.
Animals began to shed oocysts day 4 PI with a peak in oocyst
output generally on day 5-8 PI (Figure 2.1 A). After 10 days
animals inoculated with 104 and 105 oocysts no longer shed
oocysts. Oocysts were detected in feces of animals
inoculated with 103 oocysts until day 12 PI.
The prepatent period of E. delicata was 6 days and the
patent period was 3 days for animals inoculated with 104 and
4 days for animals inoculated with 103 oocysts (Figure 2.1
B). The peak in oocyst output occurred on day 7 PI. In
addition, animals shed small (<1000) numbers of oocysts
intermittently until day 20 PI (when animals were
reinoculated).
34
105 OOCYSTS
04 OOCYSTS
103 OOCYSTS
107
A
106
106
104
103
102
10'
10°
106
105
104 H
103 7
102 H
10'
10°
2
3
4
5
6
7
8
9
10
11
12
13
14
DAYS POST-INOCULATION
Figure 2.1. Oocyst output of E. arizonensis and E. delicata.
Total daily oocyst output in deer mouse feces from days 2-14
post-inoculation for Eimeria arizonensis (A) and E. delicata
(B). Animals inoculated with E. delicata shed small (< 1000)
numbers of oocysts intermittently until reinoculated (not
shown). Animals were inoculated with 103, 104 or 105 oocysts
on day 0. Values given at each inoculum level are means
(+SE) of five animals.
35
Diurnal Periodicity in Oocyst Output
Both E. arizonensis and E. delicata exhibited a clear
cycle in oocyst output (Figure 2.2 A,B). For E. arizonensis
peaks in oocyst output occurred during the dark phase and
were most apparent after day 5 PI. Eimeria delicata showed 4
peaks in 2 days at 12
16 hr intervals. Thus, periodicity
occurred in this species, but not a diurnal cycle.
Fecal Oocyst Concentration
Individual deer mouse fecal pellets weighed
0.02 +
0.0013 g. This value was used to estimate the concentration
of oocysts per fecal pellet during light and dark periods.
Oocyst concentration of E. arizonensis ranged from 1.5-14.6
x 104 oocysts per fecal pellet and pellets produced during
the light period had a 1.3-2 greater concentration of
oocysts than pellets produced during the dark period (Table
2.1).
Oocyst concentration per pellet of E. delicata was much
lower than for E. arizonensis; however, pellets produced
during the light period also contained a higher
concentration of oocysts than those produced during the dark
period (Table 2.1).
36
105 00CYSTS
104 00CYSTS
103 00CYSTS
A
40
30
20
10­
DARK
LIGHT
DARK
LIGHT
DARK
LIGHT
0
DARK
8
7
6
5
DARK
LIGHT
LIGHT
B
50-1
40-?
30
20 -1
10 ­
0
7
8
9
DAYS POST-INOCULATION
Figure 2.2. Percent daily oocyst output. Percent of total
daily oocyst output shed in deer mouse feces during four hr
sampling periods for Eimeria arizonensis (A) and E. delicata
(B) during days of peak oocyst output. Animals were on a
16L:8D light-dark cycle. Symbols represent the end of each
sampling period.
37
Table 2.1. Oocysts per fecal pellet. The number of oocysts
per Peromyscus maniculatus fecal pellet (0.02g feces) during
the day and night on various days post-inoculation (PI).
Values represent means of 5 animals in all cases; It = light
period = 16 hrs. dk = dark period = 8 hrs.
Table 2.1
Oocysts of
Eimeria delicata (x 102)
Oocysts of
Eimeria arizonensis (x 104)
Day 6 PI
dk
It
Inoculum
level
Day 5 PI
It
dk
103
6.3
4.9
14.6
7.3
104
9.2
6.7
7.3
4.0
105
3.3
1.9
2.3
1.5
Day 7 PI
dk
It
4
-
Day 7 PI
dk
It
Day 8 PI
dk
It
2.6
21.4
2.6
3.8
2.3
-
18.4
4.4
4.9
2.2
-
-
-
-
­
39
Effect of Inoculum Period and Level
Both inoculum period and level of E. arizonensis
significantly affected total oocyst output of naive animals
(Table 2.2). Total oocyst output increased with inoculum
period and decreased with the number of oocysts inoculated
(Fig. 2.3 A). In addition, there was a significant
interaction between period and level (Table 2.2), indicating
that the effect of period depended on the number of oocysts
ingested. The reproductive index (RI = the number of oocysts
shed per oocyst ingested) also increased with period and
decreased with level (Table 2.3).
For E. delicata there was no significant difference in
total oocyst output or RI of naive animals between the first
and challenge inoculations at 103 oocysts (t = 0.08, P >
0.05; Figure3B) or in total oocyst output at different
inoculum levels (t = 2.287, P > 0.05). However, RI for
animals inoculated with 103 oocysts was higher than for
animals inoculated with 10' oocysts (Table 2.4; t = 3.611, P
< 0.01).
Immunity
All animals shed fewer E. arizonensis oocysts during
challenge infections than when initially inoculated (Table
2.4). Only 1 of 14 animals showed < 90% decrease in oocyst
output compared with the output of controls inoculated with
the same number of oocysts. This animal shed 34% of the
40
Table 2.2. ANOVA of the number of oocysts excreted.
The factor "level" is the number of oocysts inoculated
and "period" is whether animals were in the first or
second experimental group.
Factor
df
F
P
Level
2
20.4
0.001
Period
2
18.0
0.001
Interaction
4
3.7
Error
0.04
34
Table 2.3. ANOVA of reproductive index (number of oocysts
excreted for each oocyst ingested). The factor "level"
is the number of oocysts inoculated and "period" is
whether animals were in the first or second experimental
group.
Factor
df
F
P
Level
2
131.36
0.001
Period
2
58.08
0.001
Interaction
4
2.69
Error
34
0.05
41
30
A
Naive Treatment Animals
c(4)
Naive Control Animals
25
0
Co
w
20
C
VJ
>­
15
0
0
--I
10
I­
b
0
10,000
1000
100,000
20
B
d
1000
10,000
INOCULUM LEVEL
Figure 2.3. Total oocyst output over a 20 day infection
(+SE). Deer mice were naive (not previously infected) when
inoculated with Eimeria arizonensis (A) and E. delicata (B).
The sample size was 5 except where shown by parentheses. The
same letters indicate grou7s that did not differ
significantly at the 0.05 significance level (Least
Significant Difference Test).
42
Table 2.4. Immunity to E. arizonensis. The effect of
challenge infection on Eimeria arizonensis oocyst output in
deer mice.
Inoculum
level
Mean oocyst output x 106 (+SE)
of animals at 1st & 2nd
%
Reduction in
inoculation
oocyst output*
Naive control
103
10.15 (1.84)
Challenge
0.54
(0.17)
97.5
104
8.38
(2.33)
0.39
(0.23)
97.9
105
3.09
(0.67)
0.39
(0.24)
95.5
* compared to naive control animals
** excluding one outlier (see text)
43
expected value, probably did not become immune and was
excluded from further analysis. Table 2.4 shows that the
mean percent decrease in oocyst output was at least 90%.
Three animals were challenged with E. delicata at each
inoculum level. None of these animals shed fecal oocysts for
at least 20 days PI.
Discussion
In this study we examined several parameters of the
life history of 2 eimerians. Eimeria arizonensis has been
reported previously in P. maniculatus, P. eremicus, P.
leucopus, Reithrodontomys fulvescans, R. meqalotis and R.
montanus (Reduker et al., 1985, Upton et al., 1992); E.
delicata has only been reported in P. maniculatus (Levine
and Ivens, 1960; Reduker et al., 1985). However, with the
exception of prepatent and patentperiods for E. arizonensis,
little is known about the life history of these 2 species.
Daily Oocyst Output
The prepatent and patent periods reported herein for E.
arizonensis are similar to those reported previously.
Reduker et. al (1985) found a prepatent period of 3-4 days
and a patent period of 9-11 days in deer mice. Upton et. al
(1992) reported a patent period of 4-14 days in the western
harvest mouse (Reithrodontomys megalotis) and a patent
period of 4-16 days in the pinyon mouse (P. truei). Eimeria
44
delicata had a longer prepatent and shorter patent period
than E. arizonensis (Table 2.5). Interestingly, the patent
period in deer mice inoculated with 103 oocysts of either
species was longer (+2 days for E. arizonensis; +1 day for
E. delicata) than in animals inoculated with higher oocyst
levels. This suggests that the patent period depends on
initial dose. Oocysts of E. delicata also reappeared
intermittently after the majority of oocysts were shed but
E. arizonensis oocysts did not. Although it is possible that
animals became reinfected with E. delicata by cage
contamination, despite precautions taken (frequent cage
changes, wire cages), it seems unlikely. When the patent
period and sporulation time (minimum of 5 days; pers. obs.)
are taken into account, animals that were reinfected from
contaminated cages would not begin to shed oocysts until at
least day 16 PI, but E. delicata oocysts reappeared as early
as day 12 PI.
We found that the total oocyst output (E. arizonensis)
and the reproductive index (both E. arizonensis and E.
delicata) declined with increasing inoculum. This pattern of
negative density dependence has been observed in many other
eimerians (Hall, 1935; Brackett and Bliznick, 1952; Tilahun
and Stockdale, 1981; Higgs and Nowell, 1988; Lillehoj,
1988). In general, these studies report an optimum inoculum
which produces the largest reproductive index. Below this
level, the parasites may be too sparse to reproduce
efficiently during the sexual phase. Above the optimum
45
level, host cells/nutrients may be limited (see Fernando
[1982] for a review of potential mechanisms). We did not
find a lower limit for either species; however, our lowest
dose (103) may have already been higher than the optimum.
It is unclear why output of oocysts increased in the
second group of naive animals we inoculated. Oocyst
viability may have changed slightly in the 3 weeks between
inoculations although it seems unlikely that enough oocysts
would die to create the differences seen here. Alternately,
the deer mice may have undergone sexual maturation during
this time (Millar, 1989). Several studies have shown effects
of age and reproductive hormones on parasites (Augustine,
1988; Higgs and Nowwell, 1988; Harder et. al, 1992).
Diurnal Periodicity in Oocyst Output and Fecal Oocyst
Concentration
Although periodicity has been documented in many
helminth parasites (Hawking, 1975), there have only been a
few studies of this phenomenon in coccidial parasites
(genera Isospora and Eimeria) and, to our knowledge, all of
these have been in avian hosts (Boughton, 1988). These
studies showed that peak oocyst output occurred shortly
before or during the beginning of the dark period. If the
light-dark cycle was changed, oocyst output also changed
such that peak oocyst output still occurred at the beginning
of the dark period. Parasite invasion of host tissue
occurred at the same time relative to the light period
46
regardless of when the inoculation occurred (Boughton,
1988). Apparently periodicity is determined by environmental
factors and is not intrinsic to the parasite or the host.
The cycle of oocyst output in E. arizonensis was similar to
those reported in previous studies in that peak oocyst
output occurred at night, despite the fact that, in contrast
to the birds used in Boughton's studies, deer mice are
nocturnal. Interestingly, E. delicata showed roughly 2 peaks
in oocyst output per day rather than 1. Boughton (1988)
suggested that there are 2 waves of cellular invasion 24 hr
apart and each reproductive cycle lasts 48 hr. Thus, peaks
occur at 24 hr intervals. Eimeria delicata may be unusual in
having accelerated development.
More total oocysts were shed at night; however, because
less feces was produced during the day, each daytime pellet
contained a higher concentration of oocysts than those
produced at night (Table 2.1). This may have ramifications
for parasite transmission. The oocysts shed at night will be
in relatively low concentrations and spread widely
throughout the environment. However, there will be a very
concentrated oocyst source inside of deer mouse nests.
Animals from the population from which our laboratory stock
originated have been observed to use latrine sites below
bedding in artificial nest boxes and to share nests with up
to 3 adults (C.A. Fuller, pers. obs.). Nests may provide a
sufficiently warm and moist environment to facilitate oocyst
sporulation. Thus, deer mice may be exposed to high
47
concentrations of oocysts in their nests and this may be the
primary site of transmission.
Immunity
Both species examined in this study elicited
immunity
(>90% reduction in oocyst output) after 1 large, inoculating
dose. However, E. arizonensis elicited partial immunity
while E. delicata seemed to elicit complete immunity at both
levels tested. Other studies have reported a large decrease
in oocyst output after 1 large exposure (Hall, 1935;
Augustine, 1988; Lillehoj, 1988) or after several smaller,
e.g., 102, doses (Higgs and Nowell, 1988, Fitz-Coy and
Edgar, 1989). Both cell mediated and antibody response are
thought to be involved in the immune response to eimerians
(Rose, 1987) .
Although E. arizonensis and E. delicata co-occur in the
deer mouse, they vary widely with respect to most life-cycle
parameters examined (Table 2.5). Eimeria arizonensis
infections produced a much larger number of oocysts over a
slightly longer patent period and did not elicit complete
immunity after 1 exposure as did E. delicata infections. In
addition, the number of oocysts per fecal pellet was much
(3000-5000 x) greater for E. arizonensis. These findings
suggest that E. arizonensis has a much greater reproductive
potential in wild deer mice than E. delicata.
48
Table 2.5. A comparison between Eimeria arizonensis and
Eimeria delicata life history parameters.
Parameter
Length in days:
Prepatent period
Patent period
Total oocyst output
Eimeria arizonensis
E. delicata
3
6
7-9
3-4
> 106
« 106
Development of
immunity
partial
complete
Periodicity in
oocyst output
1 peak/day
2 peaks/day
Oocysts/fecal pellet
> 1.5 x 104
< 3000
49
Chapter 3
Population Dynamics of E. arizonensis and E. delicata
50
Abstract
Because many parasites have a free-living stage, their
survival is affected by 2 sets of factors: 1) host factors
and 2) abiotic factors in the external environment.
I
examined the population dynamics of 2 deer mouse eimerians
(Protozoa) under field conditions to determine which factors
were more important. The prevalence of Eimeria arizonensis
was not affected by host sex or reproductive condition and
increased with host age. However, E. arizonensis prevalence
was strongly correlated with environmental temperature. In
contrast, E. delicata had a significantly higher prevalence
in juvenile than in adult deer mice, but was not related to
any other factor measured. In addition, there was no
evidence of competition between the 2 eimerians.
These data
suggest that host factors were probably less important than
abiotic factors in the population dynamics of E. arizonensis
and more important than abiotic factors in E. delicata.
Introduction
By definition, parasites have a negative impact on
their host. Thus, parasites may be important in host
population regulation (Anderson & May 1979, May & Anderson
1979). Parasites may also cause or exacerbate the decline of
threatened or endangered host species (Scott 1988).
Understanding the basic population dynamics of parasites can
give us insights into which individual hosts will be
51
affected (eg., male or female, old or young) and whether the
effect will occur at the population level (Anderson 1986,
Scott 1988). Because parasites generally live in at least 2
environments (within/attached to the host and free-living
outside of the host), 2 sets of factors may affect their
survival. Within host factors include host immune status and
resource competition among parasites (Holmes et al. 1977).
Factors external to the host include abiotic parameters such
as temperature, moisture and uv radiation (Levine 1980).
Two different categories of parasites are recognized:
1) microparasites (viruses, bacteria and protozoans) go
through multiple reproductive cycles within the host and
usually induce complete immunity after only one infection
(Anderson & May 1979); 2) macroparasites (helminths and
arthropods) shed eggs or larvae from the host and larvae
must infect a new host before reproduction occurs. Acquired
immunity is often partial and the degree of immunity may be
dose (density) dependent (May & Anderson 1979). In addition,
acquired immunity to macroparasites may be lost if enough
time passes between infections.
Several studies have documented population dynamics of
natural macroparasite infections in free-living hosts
(Arnett et al. 1993, Gregory 1992, Gulland & Fox 1992,
Prestrud et al. 1993, Shaw & Moss 1989, Theis & Schwab 1992,
Wilson 1983). Less attention has focused on the population
dynamics of microparasite infection (Ball & Lewis 1984,
Healing 1981, Stanton et al. 1992, Weatherhead & Bennett
52
1991, 1992, Wilber et al. 1994).
I compared the population
dynamics of 2 deer mouse protozoans, Eimeria arizonensis
(Ea) and E. delicata (Ed; Apicomplexa; Eimeriidae). These
parasites have only one host, are transmitted by ingestion
of infective oocysts (the form excreted) and generally live
in the hosts' intestinal epithelium. The infections are
usually of short duration (< 20 days) and, because
reproduction occurs by multiple fission within the host, as
many as 106 oocysts can be produced for each oocyst
ingested. Oocysts are shed directly into the lumen of the
intestine and can be detected by fecal examination. Oocysts
are shed in an uninfective form and require approx. 1 week
at room temperature (pers. obs.) to become infective
(sporulate). During this time, extreme temperatures and
humidity have been shown to affect oocyst survival and
sporulation rate (Graat et al. 1994, Marquardt et al. 1960).
Although eimerians are generally categorized as
microparasites because of their mode of reproduction, they
are often more similar to macroparasites with respect to
acquired immunity (Healing & Nowell 1985, Quinnell & Keymer
1990)
.
In this study, I compared the population dynamics of E.
arizonensis and E. delicata using naturally-infected free-
ranging deer mice. I used these data to determine whether
host factors,
environmental factors or a combination of the
two are important determinants of parasite abundance in this
system. If host factors are the primary determinant of
53
parasite population dynamics, several predictions can be
made. First, if competitive exclusion among parasites is
important, E. arizonensis and E. delicata will be found
together less frequently than expected by chance (Reduker &
Duszynski 1985, Lafferty et al. in press). Second, if
immunity is important, young animals should be parasitized
more frequently than older animals due to exposure
(Prediction 2a). Alternatively, because reproductive
hormones often lead to changes in immunocompetence
(Alexander & Stimson 1988, Bundy 1988), there may be
differences in parasite prevalence among males and females
or reproductive and non-reproductive animals (Prediction
2b). Third, if environmental factors are most important in
determining parasite population dynamics, temperature and/or
moisture during oocyst sporulation will be correlated with
parasite prevalence.
Methods
Data Collection
Two groups of deer mice were captured in Sherman live-
traps in MacDonald Forest (15 km NW Corvallis, OR). The
first group was part of a mark-recapture study. These
animals were individually marked and released at their point
of capture. Only the first capture of each animal was used
in the current study. The second group of animals was
collected by live-trapping at other sites around MacDonald
54
Forest. These sites were not trapped more than once and were
at least 500 m from the mark-recapture grid. Both groups
were captured between 15 April
Nov 1991 and 7 March
17 Nov 1990, 30 March
17
12 July 1992. Animals from both
sources were processed in the same manner and assumed to be
comparable with respect to Eimeria infections. Animals were
assessed for age (juvenile or adult, based on pelage) and
reproductive status (females: reproductive = pregnant and/or
lactating, non-reproductive = no visible signs of
reproduction; males: reproductive = scrotal testes, nonreproductive = abdominal testes). All feces were collected
from traps, weighed and incubated in a 2.5% w/v potassium
dichromate
(K2Cr2O7)
solution at room temperature for one
week to allow any coccidia present to sporulate (this is
necessary to identify oocysts to species). Fecal samples
were macerated and examined for the presence (prevalence) of
oocysts by Sheather's sugar flotation (Ash & Orihel 1987).
Oocysts can only be detected in patent infections. However,
the length of the prepatent period (the time from ingestion
of infective oocysts to the beginning of oocyst output) was
obtained from laboratory experiments (prepatent to patent
ratio: Ea = 3:9 days, Ed = 6:4 days; Chapter 2) and used to
estimate the total number of animals infected. In addition,
the number of oocysts per gram of feces was estimated as 0,
1-9, 10-99, 100-999, etc., to compare the severity of
infection between Eimeria species.
55
Temperature and rainfall data were obtained from a
weather station at Hyslop Farm, approx. 2.5 km east of
MacDonald Forest.
Statistical Analyses
In all cases, alpha of 0.05 was considered
statistically significant.
Host Factors
Data were pooled by month because trapping occurred at
variable intervals. Due to the high occurrence of 0
prevalence in monthly samples, transformation did little to
increase the normality of the data set. Thus, nonparametric
statistics were used in analyses comparing infection
prevalence among various sex, age and reproductive classes
of deer mice. To account for seasonal variability,
prevalence data were compared with either Wilcoxon Sign-Rank
Test or Friedman's 2-way ANOVA (Zar 1984). These analyses
compared the data set by month either pairwise (Wilcoxon
Test) or by groups of > 2
(Friedman's ANOVA). In both
analyses, any month with missing values or tied ranks was
excluded. If Friedman's ANOVA was significant, Wilcoxon Test
was used as a post-hoc test to determine which groups
differed. Where predictions were tested, 1-tailed alpha
levels are given; other alpha levels are 2-tailed.
56
To determine whether E. arizonensis and E. delicata
occurred together more or less frequently than expected by
chance, data were divided into 4 time periods (late 1990,
early 1991, late 1991 and early 1992). The probability of
having both infections was calculated as the product of the
proportion of animals infected with either eimerian:
PEaxPEd.
The occurrence of observed and expected double infections
within each time period were compared by X2 analysis.
Environmental Factors
In deermice, the prepatent period of E. arizonensis and
E. delicata is 3 and 6 days respectively. Oocysts are shed
for up to 9 (Ea) or 4
(Ed) days and sporulation takes 4-7
days under laboratory conditions (Chapter 2). If
environmental factors such as rainfall or temperature affect
prevalence of infection, they should have the greatest
impact during oocyst sporulation
(see above) because this is
the primary time when oocysts are exposed to the external
environment. This period occurs 7-13 days before the actual
observed infection. Thus, I used mean high and low
temperature and total rainfall for the 2-week period prior
to the
capture period in analyses.
I used stepwise multiple regression to compare the
relative strengths of environmental factors (Wilkinson
1990). All variables were examined for normality and
57
transformed as necessary prior to analysis. The residual
errors from the final model were examined graphically to
insure they met the assumptions of linear regression.
Results
Overall Prevalence and Intensity
432 deer mice were captured in 21 months of the study.
Of these, all were assessed for the presence of E.
arizonensis and 348 were assessed for the presence of E.
delicata (E. delicata was incorporated into the study in
August 1990). Over the course of the study, the prevalence
of E. arizonensis ranged from 0-80% and of E. delicata from
0-53% (Figure 3.1 A). Peaks in E. arizonensis prevalence
occurred in spring (June, April and May in 1990, 1991 and
1992) and spring/summer for E. delicata (July and May, 1991
and 1992).
There were no significant differences in monthly
prevalence of patent infections between E. arizonensis and
E. delicata infections (Wilcoxon Sign-Rank Test, z=0.3341,
N=16 months). However, when the estimated number of animals
with prepatent infections (see above) was added to animals
shedding oocysts, significantly fewer animals were infected
with E. arizonensis than with E. delicata (z=2.43, P=0.015,
N=16; Figure 3.1 B).
Oocyst output per gram feces was quantified in 94 E.
arizonensis and 94 E. delicata infections. Infected deer
58
1.0
1990
A
1992
1991
B
0.0
Ma
Ju
Se
Ma Ma
Ju
Se
No
Ap
Ju
Figure 3.1. The prevalence of E.
arizonensis and E.
delicata. The proportion of deer mice shedding oocysts (ie,
with patent infections) of Eimeria arizonensis (solid lines)
and E. delicata (dotted lines)
(A) and the estimated
proportion of animals with prepatent and patent infections
(see text) (B). The proportion of animals with patent and
prepatent infections combined was greater for E. delicata
but there were no differences between species for the number
of animals with patent infections alone. See tables 3.2 and
3.3 for sample sizes.
59
mice shed significantly greater numbers of E. arizonensis
than E. delicata oocysts (Figure 3.2; X2=36.15, df=4, P <
0.001) .
Hypothesis 1: Host Factors
Prediction 1: Occurrence of Double Infections
Eimeria arizonensis and E. delicata were not found
together significantly more or less frequently than expected
by chance in any of the 4 time periods examined (Table 3.1).
Prediction 2a: Variability Among Sub-Populations of Deer
Mice, Age and Sex Differences
Friedman's analysis indicated that there were
significant differences in prevalence among different
classes of animals with naturally occurring E. arizonensis
infections (Fr=8.61, df=3, P < 0.02; Table 3.2). Further
analysis showed that juveniles were infected less frequently
than adults, the opposite of predicted (z=2.86, N=14, P <
0.005). In addition, males were infected more frequently
than females; however, this difference was not significant
(z=1.76, N=17, P=0.08).
There were also significant differences in prevalence
among E. delicata infections (Fr=6.69, df=3, P < 0.04; Table
3.3). As predicted, juveniles were infected more frequently
than adults (z=2.05, N=11, P < 0.02). There were no
60
40
E. arizonensis
E. delicata
30
10­
0
1
2
3
4
5
6
7
Oocysts Per Gram Feces
Figure 3.2. The intensity of E.
arizonensis and E. delicata.
The number of animals with various levels of oocysts in
99, etc.). The
their feces (1 = 1-9 oocysts/gram, 2 = 10
animals
in
which
data were combined for all infected
3
years
of
the study.
oocysts/gram was assessed across the
Animals infected with Eimeria arizonensis shed significantly
more oocysts than those infected with E. delicata.
61
Table 3.1. Frequency of double infections. The number of
deermice infected with separate Eimeria arizonensis and E.
delicata infections, infected with both or neither. E.
arizonensis and E. delicata did not appear together more or
less frequently than expected by chance in any of the 4 time
periods. This suggests that facilitation or cross-immunity
are unlikely. Numbers in parentheses indicate the number of
months included in each time period. Df = 1 in all analyses.
Time period
1990-late (3)
1991-early (4)
1991-late (5)
1992-early (5)
Ea
Ed
Both
8
13
3
31
16
5
24
20
18
Neither
X2
56
0.29
41
0.24
5
50
0.75
2
39
2.31
17
62
Table 3.2. Percent infected (sample size) with Eimeria
arizonensis in different sex and age classes of deer mice.
Adults
Males
1990
April
May
June
July
Aug
Sept
Oct
12
20
33
(16)
0
(9)
(9)
(5)
(6)
Females
18
0
100
Males
(11)
0
(1)
(1)
20
25
(8)
0
(8)
0
(13)
12
12
18
(11)
12
(1)
0
(2)
67
83
(6)
(4)
50
38
33
(22)
(21)
75
75
42
25
(24)
(12)
(6)
0
(3)
0
(9)
(6)
25
(8)
(3)
0
(5)
(2)
0
(2)
(0)
0
20
(5)
(4)
(3)
25
(8)
(2)
0
(15)
6
(18)
25
(4)
0
(2)
0
11
38
0
(4)
(5)
(4)
(3)
(7)
(8)
(0)
Juveniles
Females
25
0
0
(4)
(2)
(5)
(0)
0
(9)
8
(12)
(0)
1991
March
April
May
June
July
Aug
Sept
Oct
Nov
0
20
0
(6)
0
(4)
33
29
0
(0)
(0)
(0)
(3)
(7)
(2)
(3)
(5)
(5)
0
0
11
0
20
(0)
(0)
(0)
(3)
(1)
(9)
(1)
0
(5)
(2)
0
(1)
40
(5)
(0)
(1)
(2)
1992
March
April
May
June
July
20
75
33
40
50
50
(0)
(0)
(1)
(0)
(4)
(4)
0
0
63
Table 3.3. Percent infected (sample size) with Eimeria
delicata in different sex and age classes of deer mice.
Adults
Males
Females
Males
Juveniles
Females
1990
Aug
Sept
Oct
11
31
(13)
0
(1)
33
33
27
38
33
44
67
20
(6)
(9)
22
50
(9)
(8)
0
0
(11)
37
0
(2)
(7)
(8)
(0)
50
25
29
(4)
(4)
(0)
(0)
(0)
(0)
(0)
0
(12)
(0)
1991
March
April
May
June
July
Aug
Sept
Oct
Nov
(6)
8
(24)
(12)
67
(3)
25
(8)
0
(3)
(2)
0
(6)
(5)
(2)
(0)
40
(5)
50
(8)
0
(4)
(3)
0
(2)
(15)
22
(18)
(4)
0
(2)
(22)
(21)
(6)
(9)
0
(0)
(3)
67
(3)
(7)
0
(1)
56
60
(2)
(3)
(5)
0
(5)
0
(9)
(1)
(5)
(2)
(0)
(1)
(0)
(4)
100
20
(4)
0
67
71
50
0
0
0
1992
March
April
May
June
July
33
27
25
0
(0)
50
50
0
(1)
(5)
(0)
(1)
(2)
64
significant differences in prevalence between males and
females (z=1.29, N=14) .
Prediction 2b: Differences Among Reproductive Classes
There were no significant differences in prevalence of
naturally-occurring E. arizonensis infections among
reproductive and nonreproductive males (Table 3.4; Z=0,
N=11). However, reproductive females were infected
significantly more frequently than nonreproductive females
(Table 3.4; z=2.07, N=9, P < 0.04).
There were no significant differences in the prevalence
of naturally-occurring E. delicata infections among
reproductive and nonreproductive males or females (Table
3.5; z=0.153, N=10 and z=0.398, N=7 for males and females,
respectively).
Hypothesis 2: Environmental Factors
The prevalence of E. arizonensis was negatively
correlated with mean high temperature (r = -0.725, N = 24, P
< 0.001), mean low temperature (r = -0.583, P < 0.005). The
correlation between prevalence and total rainfall was not
significant (r = 0.382, P < 0.10). In stepwise regression
analysis, only mean high temperature explained a significant
amount of variance (prevEa = 103.4 + -1.067temp, F = 24.37,
P < 0.001, R2 = 0.526) .
65
Table 3.4. Percent infected (sample size) with Eimeria
arizonensis in different reproductive classes of adult deer
mice. R = reproductively active, NR = not reproductively
active.
Females
Males
1990
April
May
June
July
Aug
Sept
Oct
33
33
20
0
17
0
(3)
(3)
(5)
(7)
(6)
0
100
0
NR
R
NR
R
(2)
(0)
(1)
(0)
(2)
50
100
14
14
(8)
(0)
25
100
(2)
(0)
50
(3)
100
75
75
50
(3)
(5)
(6)
(3)
(0)
(0)
0
0
(1)
(3)
(3)
(4)
(1)
(3)
(4)
0
(2)
(2)
(0)
(1)
(7)
(7)
0
(4)
(0)
14
(3)
(0)
(0)
(0)
(1)
(7)
0
(1)
(1)
(0)
(7)
67
67
47
(3)
(3)
(7)
0
(2)
(7)
(1)
(0)
0
0
1991
March
April
May
June
July
Aug
Sept
Oct
33
100
42
33
40
0
0
Nov
1992
March
April
May
June
July
(1)
(17)
(15)
33
75
33
47
(15)
100
(2)
0
0
0
(4)
(4)
(6)
43
43
0
29
0
(0)
(0)
0
(1)
0
0
(0)
(5)
(1)
(1)
(2)
(2)
(0)
20
(5)
(0)
(0)
33
50
(3)
(2)
9
(11)
0
(0)
0
(0)
(5)
(2)
(0)
(3)
0
(15)
66
Table 3.5. Percent infected (sample size) with Eimeria
delicata in different reproductive classes of adult deer
mice. R = reproductively active, NR = not reproductively
active.
Females
Males
1990
Aug
Sept
Oct
17
0
NR
R
NR
R
(6)
0
(2)
0
(2)
(0)
37
(8)
(0)
0
(3)
(1)
33
25
(3)
(4)
0
0
(4)
29
83
(6)
0
(1)
(7)
(4)
0
(0)
0
(1)
67
33
27
20
0
(1)
(7)
(1)
1991
March
April
May
June
July
Aug
Sept
Oct
33
0
35
20
40
50
100
Nov
(5)
0
(6)
(3)
(3)
(0)
33
33
25
100
29
(3)
0
(4)
(0)
(7)
(7)
(2)
(7)
(1)
(0)
(0)
0
(1)
(0)
(3)
(4)
(3)
100
(2)
(0)
(17)
(15)
(3)
(3)
(15)
(5)
0
(1)
0
(1)
0
(2)
(2)
0
(0)
1992
March
April
May
June
July
0
0
33
27
50
(0)
(0)
(15)
(2)
0
(1)
60
(5)
(0)
33
0
(0)
27
(11)
20
(0)
0
(3)
(2)
(0)
(5)
(2)
67
The prevalence of E. delicata was not significantly
correlated with any environmental factor
rlowtemp =
0.014,
(rhightemp
0 . 1 36,
rrain = 0.301, N = 20) and no factor
explained a significant proportion of the variance in
regression analysis.
The prevalence of E. arizonensis and E. delicata were
not significantly correlated (r = 0.157, N = 20, P > 0.05).
Discussion
Factors Influencing E. arizonensis
There was no evidence of competitive exclusion between
E. arizonensis and E. delicata (Prediction 1). There was
also little evidence that deer mice became immune to E.
arizonensis as measured by a decrease in prevalence with
host age (Prediction 2a). In fact, adults were significantly
more likely to be infected than juveniles. This is
consistent with other studies. Although we showed that deer
mice can become immune under laboratory conditions (Chapter
2), experiments conducted under natural and semi-natural
conditions suggest that these animals do not appear to
exhibit the same degree of immunocompetence as in the
laboratory. GThis pattern is presumably because free-living
animals are exposed to more immunosuppressing factors
(Chapter 5).
There were differences in E. arizonensis prevalence
among sex and reproductive classes of deer mice. Males were
68
slightly more likely to be infected than females and
reproductively active adult females were more likely to be
infected than nonreproductive adult females. Although the
sex ratio varied over the course of the study,
it was not
significantly correlated to E. arizonensis prevalence (R =
0.041, N = 20). In addition, only 62 of 432 animals (14.3%)
were reproductive females. Thus, neither host sex or
reproductive class seemed likely to affect E. arizonensis
population dynamics (Prediction 2b).
On the other hand, mean high environmental temperature
during the period that oocysts were on the substrate, was
strongly correlated with E. arizonensis prevalence. This
pattern suggests that environmental factors were most
important in determining E. arizonensis population dynamics
(Prediction 3)
.
Factors Influencing E. delicata
The opposite seems to be true for E. delicata. Juvenile
deer mice were far more likely to be infected than adults,
suggesting that immunity to E. delicata is common
(Prediction 2a). Evidence from the laboratory also suggests
that E. delicata is more immunogenic than E. arizonensis;
laboratory reared animals developed complete immunity after
only one exposure (Chapter 2). Unfortunately, field
experiments with E. delicata were not conducted because of
logistical constraints.
69
No differences in E. delicata prevalence were seen
among sex or reproductive classes of deer mice, suggesting
that reproductive hormones are not important in
immunocompetence to this species of parasite (Prediction
2b). In addition, E. delicata prevalence was not
significantly correlated with any of the environmental
factors measured (Prediction 3). Thus, it appears that the
population dynamics of E. delicata, in contrast to E.
arizonensis, were primarily controlled by host immunity.
Reduker & Duszynski (1985; after I calculated X2 from
data given) and Stanton et al. (1990) also found little
evidence of competitive exclusion among eimerians. There are
several mechanisms through which co-occurring eimerians can
avoid competition. Eimerians inhabiting the same host may
partition the cells they use. Kheysin (1972) found that
species of co-occurring eimerians often utilize different
areas of the digestive tract even in the absence of other
species. Alternatively, even if > 1 species of parasite is
in the same place at the same time, rapid host cell turnover
and/or low parasite abundance may prevent competition. In
the present study, E. arizonensis and E. delicata may not
co-occur in the same host individual; they may avoid
competition simply because one primarily occurs in adults
and the other in juveniles.
Results are more variable for blood microparasites.
Turner & Cox (1985) also found little evidence of species
interactions in rodents, Schall & Bromwich (1994) found
70
positive associations between 2 species of lizard malaria
and Forbes et al.
(1994) found positive, negative and random
associations among blue grouse hematozoans. Thus, further
studies among a wider variety of hosts may uncover evidence
of non-random associations among eimerians.
Several studies have found evidence of acquired
immunity to microparasites among free-living birds and
mammals, either by examining age-prevalence relationships
(Ball & Lewis 1984, Healing 1981
in 1 of 4 Hematozoans,
Samuel & Trainer 1971, Weatherhead & Bennett 1991
males
only) or differences in prevalence among sex/reproductive
groups (Ball & Lewis 1984, Keith et al. 1986, Weatherhead &
Bennett 1991
in 1 of 4 hematozoa). However, other studies
have found no evidence of immunity (age-prevalence: Healing
1981
in 3 of 4 Hematozoa, Keith et al. 1986, Stanton et
al. 1992, Wilber et al. 1994; sex/reproductive status: Ball
& Lewis 1984). Thus, while acquired immunity may be
important in some host-parasite systems, it seems to be
unimportant in others. In addition, the proportion of
animals in a population exhibiting acquired immunity to a
single parasite species may vary over time due to host or
parasite genetic factors (Wakelin 1992) or immunosuppression
(Chapter 5).
Most studies examining the effect of climate on
parasites have found that parasites were more abundant in
mesic than in xeric habitats (Dobson et al. 1992, Ford et
al. 1990, Shults et al. 1990). This is consistent with
71
results for E. arizonensis but not E. delicata in the
present study. Interestingly, none of the earlier studies
examined the effect of weather within a habitat and all pool
several species of parasite. Thus, it is possible that
differences between parasite species and within habitats
were obscured.
In my system, both parasites were exposed to identical
environmental conditions during sporulation; however, only
E. arizonensis seems to be primarily controlled by this
factor. This pattern suggests that acquired immunity when
present, supersedes environmental factors in determining
parasite population dynamics, at least within a single
habitat. However, even in the presence of acquired immunity,
habitat differences may be more important.
Deer mice rapidly develop immunity to E. delicata, thus
it seems unlikely that this parasite will influence host
population dynamics unless it affects juvenile survivorship.
In contrast, immunity does not play as important a role in
E. arizonensis population dynamics. Deer mice become
reinfected frequently and E. arizonensis does affect host
survival in the field (Chapter 4). Eimeria arizonensis and
E. delicata are closely related and have very similar life
histories, but apparently their populations are limited by
different mechanisms. Thus, it is clear that each parasite
must be examined individually to determine its effect on its
host.
72
Chapter 4
The Effect of E. arizonensis on Deer Mouse Survival
C.A. Fuller and A.R. Blaustein
73
Abstract
By definition, parasites negatively affect their hosts.
However, few experimental studies have documented this effect
under natural or semi-natural conditions.
We studied the
affect of Eimeria arizonensis on deer mouse recruitment, over­
winter
survival
and
weight.
In
field
observations,
E.
arizonensis was negatively related with recruitment success in
females
and
over-winter
survival
in
males.
Experimental
manipulation of E. arizonensis in large field enclosures also
showed that it negatively affects male over-winter survival.
There was
no relationship between deer mouse weight
and
infections, thus, the mechanism through which E. arizonensis
affects survival remains unclear.
Introduction
A major goal of ecology is to understand the factors
which affect animal populations. Predation, emigration and
competition have been the focus of most of the empirical
research on this topic (see review in Hestbeck 1987).
However, Anderson and May demonstrated mathematically that
parasites could regulate the population density of their
hosts (Anderson & May 1979, May & Anderson 1979). Since the
publication of their seminal papers, a few studies have
directly examined the potential for parasites to regulate
host populations in the laboratory (Scott & Anderson 1984,
74
Scott 1987) and in field enclosures (Barker et al. 1991,
Gregory 1991).
Although studies of host population regulation by
parasites should ideally be conducted at the population
level, this is often impractical in natural free-living
animals. Thus, most field work has focused on the effect of
parasites on the individual host (but see Washburn et al.
1991). If a parasite regulates its host's population
density, it must negatively affect the survival and/or
reproduction of the host in a density dependent manner.
There is now ample correlative evidence from the field that
parasites in many taxa (e.g., protozoans, nematodes,
trematodes and arthropods) have a negative impact on
reproduction and/or survival in a variety of hosts,
including mollusks, insects, fish, reptiles, birds and
mammals (Arnqvist & Maki 1990, Burgett et al. 1990, Festa-
Bianchett 1989, Goater et al. 1989, Johnson et al. 1991, Lim
& Green 1990, McPhail & Peacock 1983, Schall 1983). Ideally,
controlled field experiments should be conducted to examine
associations found in field observations. However,
experimental manipulation of parasite levels under field
conditions has been accomplished in only a few host-parasite
systems (Hudson 1986, Hudson et al. 1992a,b, Lehman 1993).
Scott & Dobson (1989) suggested that parasites which
are directly transmitted (i.e., complete their life cycle in
only one host) and have known pathology would be the most
likely to have a detectable effect on their host's
75
population density. Many eimerians (intestinal protozoans in
the class Coccidia) meet both of these criteria. They are
transmitted directly through ingestion of contaminated feces
and many are known to be highly pathogenic (see Long 1982).
We studied the effect of Eimeria arizonensis on deer mouse
(Peromyscus maniculatus) survival using both field
observations and field and enclosure experiments. Eimeria
arizonensis infections last 10-12 days and up to 107 oocysts
are shed in the feces during this period (Chapter 2).
Because infections are of relatively short duration, are
easy to detect using non-invasive methods and individual
hosts become infected repeatedly, this system is ideal for
examining the dynamics of infections and the effect of
infections on individual hosts. Deer mice are excellent
subjects because they readily enter live-traps. In addition,
they can be recaptured and sampled repeatedly in the field.
We tested predictions of the hypothesis that E.
arizonensis has a negative impact on deer mouse survival and
thus may limit deer mouse populations. Although actual
survival is difficult to measure under field conditions,
length of residency in a population has frequently been used
as a measure of survivorship (e.g. Boonstra et al.
1980,
Lehmann 1992). First, we predicted that animals which are
infected when they first enter a population are less likely
to become residents than animals which are uninfected when
they enter a population. Second, animals that do not
reappear in spring (and presumably die over the course of
76
the winter) were infected more frequently the previous year
than animals that do reappear in spring. Third, E.
arizonensis infections are associated with weight loss in
deer mice.
Methods
Observations
A 10 x 12 trapping grid was established in MacDonald
Forest (15 km NW Corvallis, OR) in March 1990. Sherman live-
traps were spaced 10 m apart in 1990. Traps were spaced 15 m
apart in 1991 and 1992 to increase sample size without
increasing trapping effort. Traps were set for 2-3
consecutive nights at 2-3 week intervals from 15 April
Nov, 1990, 30 March
17 Nov, 1991 and 7 March
17
12 July,
1992. All deer mice were either ear-tagged or toe-clipped
for individual identification at first capture. All animals
were released each morning at their point of capture.
Animals were sexed, weighed and adult females were assessed
for reproductive condition. A second group of animals was
trapped in MacDonald forest using the same methods as above.
However, after collection these animals were killed for use
in another study. Data from these animals were combined with
the first capture of each mark-recapture animal in analyses
concerning the effect of E. arizonensis on deer mouse
weight.
77
Feces were collected from traps, weighed and incubated
in K2Cr2O7 solution (2.5% w/v) at room temperature for one
week. This procedure facilitates the development of oocysts
into the infective or sporulated stage and is necessary for
identification to species. Soiled traps were washed before
they were returned to the field.
Fecal samples were macerated and examined by Sheather's
sugar flotation (Ash & Orihel 1987). The number of oocysts
per gram in fecal samples was estimated by either counting
all oocysts present or averaging counts from 10
representative microscope fields and extrapolating to the
entire sample. Thus, counts were estimated as levels 0-6
where 0 = no oocysts detected, 1 = < 10 oocysts/gram, 2 =
10-99 oocysts/gram, 3 = 100-999 oocysts/gram, etc. For
analyses of over-winter survival, we distinguished new
infections from continued infections. If an animal which was
shedding < 1000 oocysts/gram had been shedding oocysts
during the previous trapping session, the second infection
was assumed to be a continuation of the first (Chapter 2).
Experiments
Eimeria arizonensis Cultures
The E. arizonensis oocysts used in these experiments
were obtained from the feces of two deer mice from the
MacDonald Forest population in Spring 1990. The oocysts were
stored at 4°C and fresh oocysts were generated periodically
78
using laboratory born, Eimeria free deer mice which also
originated from the MacDonald forest population. The oocysts
used in the experiments were generated 6 days
4 months
before the initial inoculations, sporulated at room
temperature and refrigerated until used.
Field Experiment
The animals from the mark-recapture population were
used after the observational study had ended. In addition, a
second mark-recapture grid was established approx. 1 km from
the first. The protocol for the field experiment is
summarized in Table 4.1. Animals were lightly anaesthetized
and inoculated by stomach intubation with 20,000 sporulated
oocysts 2 times at 3-week intervals (Oct. and Nov. 1992) or
once (Nov. 1992). Animals were released within 1 hr of
inoculations and were free-living throughout the experiment.
Traps were set overnight on days 5 or 6 post-inoculation
([PI] these are the days of peak oocyst output for E.
arizonensis, Chapter 2)
and animals were released the
following morning. Feces were collected out of traps,
weighed to the nearest 0.01 g and examined by MacMaster's
technique (Dunn & Keymer 1986) to verify success of
infections.
79
Table 4.1. Protocol for field experiment.
Inoculum (N)
Event
Days PI*
Treatment
Inoculation
Feces collection
Challenge
Feces collection
0
2 x 104
(9)
2 x 104
(9)
Control
water (11)
5 or 6
20 or 21
25 or 26
(5-6 PC)**
* PI = post-inoculation
** PC = post-challenge
2 x 104
(12)
80
Enclosure Experiment
Free-living deer mice were live-trapped and placed in
4,
50 m x 50 m (2 treatment and 2 control enclosures) field
enclosures in the fall of 1993. There were 100 live-traps
(10 x 10 grid with 5 m spacing) in each enclosure and each
enclosure was surrounded by a 1 m high fence above ground
and a 0.5 m fence below ground. Thus, animals were exposed
to aerial but not terrestrial predators and could not
disperse from the enclosures. The dominant vegetation was
alfalfa; deer mice were not given water or additional food
but were provided with nest boxes (20/enclosure). In this
experiment animals in treatment enclosures were inoculated
with 5 x 105 oocysts and animals in control enclosures were
inoculated with water 1
3 times at 5-week intervals. The
majority of animals were first inoculated in Sept. 1993,
given a second infection in Oct. 1993 and a third infection
in Nov. 1993 at 5 week intervals. A second group of animals
was first inoculated in Oct. 1993 and given a second
infection in Nov. Animals were trapped as in field studies
and fecal samples were collected directly from the animals.
Fecal samples were processed as in the field experiment.
Statistical Analyses
Table 4.2 shows the predictions, protocols which
address them and statistical methods for each. In the
analysis of prediction 1
(recruitment), we used a three-way
81
Table 4.2. Predictions and experimental protocols.
Predictions 1 & 2 are from the hypothesis that Eimeria
arizonensis negatively affects deer mouse survival by
negatively affecting recruitment (21) and over-winter
survival (P2). P3 predicts that deer mice will lose weight
during infections. a = 3-way X2 contingency analysis, b =
Mann-Whitney U test, c= Paired t-test, d= Student's t-test
and e = ANOVA.
Predictions
Study
Field Observations
1
2
3
Xe
Xb
Xc
Field Experiment
Enclosure Experiment
Xd
Xe
Xd
82
Chi-square analysis (factors = sex, infection status and
year) to factor out the effect of year on infection rate.
Because there was no evidence of variability within
treatments, we pooled data within treatments for analyses of
weight change. In tests of all predictions, 1-tailed
probabilities are given unless stated otherwise and alpha of
0.05 was considered significant.
Results
Prediction 1: Establishment of Residency
Deer mice were considered to be residents if they were
present in at least four trapping sessions (a minimum of 6
weeks). Males that became residents in the population were
infected during their first captures as frequently as males
that did not become residents (Table 4.3; X2=1.841, df=2,
N=79, P>0.2). In contrast, females that were infected when
they first appeared, were significantly less likely to
become residents than females that were not infected (Table
4.3; X2=3.566, df=2, N=63, P<0.05) .
Prediction 2: Over-Winter Survival
Field Observations
Only residents (see above) which were present in one of
the last two trapping sessions of each year were included in
this analysis. Males that remained in the population after
83
Table 4.3. E. arizonensis and deer mouse recruitment. Does
the frequency of infection predict the likelihood of
establishing residency? Values given are percent (sample
size) of animals infected in at least one of first two
captures.
* Females which were infected with E. arizonensis in at
least one of their first two captures were significantly
less likely to become residents than females which were
uninfected during their first two captures (3-way X2
contingency analysis, P < 0.05).
Became residents?
Males
Yes
No
Yes
Females*
No
Year:
1990
32
(22)
10
(19)
19
(16)
40
(10)
1991
69
(13)
60
(10)
21
(14)
50
(8)
1992
73
(11)
75
(4)
54
(11)
75
(4)
Total
52
(46)
33
(33)
29
(41)
50
(22)
84
the winter had been infected significantly less frequently
the previous year than males that did not remain in the
population. This pattern was clear in the spring of both
1991 and 1992 (Figure 4.1 A; 1990-91: Mann-Whitney U=36.5,
N=14, P<0.05; 1991-92: Mann-Whitney U=12, N=7, P>0.02).
Females that remained in the population after the winter of
1990 were not infected less frequently than females that did
not return (Figure 4.1 B; Mann-Whitney U=17, N=12, P>0.2).
There were not enough resident females present in the
population during the last two trapping sessions of 1991 to
conduct an appropriate statistical analysis (N=4). However,
the mean proportion of infections is presented in Figure 4.1
B for comparison.
Enclosure Experiments
As in field observations, males in treatment
populations were significantly less likely to survive the
winter than males in the control populations (ANOVA: F =
40.65, P < 0.025). There were no significant differences
between control and treatment survival for females
0.004; Figure 4.2).
(F =
85
0.6
0.5
0.4
0.3
0.2
0.1 H
0.0
0.6
b
0.5
0.4
0.3
0.2 ­
0.1 ­
0.0
MALES
FEMALES
Figure 4.1. Over-winter survival of free-living deer mice.
Proportion (SE) of captures free-living deer mice were
infected with naturally-occurring Eimeria arizonensis in
1990 (a) or 1991 (b) vs whether they survived the following
winter (open bars represent animals which survived, shaded
bars represent animals which did not survive). Males which
did not survive the winter had been infected significantly
more frequently (* P < 0.05) than males which survived in
both years. There were no significant differences for
females in either year. Numbers in columns are sample sizes.
86
I
CONTROL POPULATIONS
TREATMENT POPULATIONS
100
80
To
E
(/)
60
4a.
szlj
40
O
2
a)
0_
20
MALES
FEMALES
Figure 4.2. Over-winter survival of enclosure deer mice.
Percent of population which survived the winter in control
and treatment enclosure populations. Control deer mice were
sham inoculated and treatment deer mice were inoculated with
5 x 105 sporulated oocysts 1-3 times in the fall. Treatment
males were significantly less likely to survive the winter
than control males (* P < 0.05) but there were no
significant differences in female survival.
87
Prediction 3: Weight Changes
Field Observations
There were no significant differences between infected
and uninfected adults. However, infected juveniles were
significantly heavier than uninfected juveniles (Table 4.4).
Field Experiment
Only males were used in this experiment. There were no
significant differences in weight change from inoculation to
Day 5/6 PI between control and treatment animals in Oct
(t =
-0.49, df = 20) or in Nov (t = -1.50, df = 21) 1992 (Table
4.4)
.
Enclosure Experiments
There were no significant differences in weight change
for adult males or for non-reproductive adult females in the
enclosure experiment whether weights were analyzed
separately or pooled by month (ie., ignoring the number of
times animals had been inoculated; Figure 4.3 A,B). Other
age/sex classes were not present in sufficient numbers to
warrant analysis.
88
Table 4.4. Weight change in free-living deer mice. Weights
(g) of infected and uninfected deer mice in field
observations and experiments: mean + SE. Numbers in
parentheses are # of months (observations) and # of animals
(experiments)
.
* P < 0.025, Mann-Whitney U Test.
Observations: Data were pooled by month within each group
and the mean weight of infected and uninfected animals was
compared by paired t-tests for months in which both infected
and uninfected animals were present. This was done to
correct for seasonal weight changes.
Experiments: Weight change from Day 0
Day 5 post­
inoculation (PI) of free-living deer mice inoculated with
water (control) or 2 x 105 sporulated Eimeria arizonensis
oocysts (treatment) .
Infected
Uninfected
11.8+0.5
12.8+0.6
17.0+0.3
16.9+0.4
20.7+0.7
18.3+1.4
Lactating (5)
17.8+0.6
16.5+0.7
Non-reproductive (6)
14.9+0.6
15.1+0.7
17.8+0.7
16.6+0.6
Observations
Juvenile*
(9)
Adult male (15)
Adult female:
Pregnant
(6)
Total females (17)
Experiments (Adult males only)
October 1992
-0.15+0.4
(10)
-0.37+0.2
(12)
November 1992
-2.18+0.4
(19)
-0.75+0.6
(4)
89
1
Control Animals
1
Sept, 0
Oct, 0
Sept, 0
Treatment Animals
IIIN
Oct, 1
Oct, 1
1
I
Nov, 1
Nov, 2
Nov, 3
Month and Number of Previous Inoculations
Figure 4.3. Weight change of enclosure deer mice. Weight
Day 5 post-inoculation
change was measured from from Day 0
(PI) in animals inoculated with water (control) or 5 x 105
sporulated Eimeria arizonensis oocysts (treatment) 1-3
times. There were no significant differences in weight
change for either males (a) or females (b) in any month and
regardless of the number of times inoculated. Numbers at the
base of columns are sample sizes. Bars are standard errors.
90
Discussion
We tested three predictions of the hypothesis that E.
arizonensis has a negative impact on deer mouse survival. We
found that, as predicted, the frequency of infection was
negatively associated with two measures of survivorship,
establishment of residency (Prediction 1)
and overwinter
survival (Prediction 2). However, males and females were not
affected in the same way. Females infected with naturally-
occurring E. arizonensis when they first appeared in the
population were less likely to become residents than females
that were uninfected, but there was no similar trend for
males. On the other hand, once a male became a resident, the
frequency with which he was infected over the remainder of
the year was negatively associated with overwinter survival
both in naturally-occurring and experimental infections. In
this case, no similar trend was seen for females. In
addition, the fact that the differences in over-winter
survival are significant in all years/studies despite
relatively small samples, suggests that the effect is
especially strong.
Although it is common for males and females of a host
species to have different rates of infection (Alexander &
Stimson 1988, Bundy 1988), it is unclear why E. arizonensis
would affect different aspects of deer mouse life history.
In the study of recruitment, females which did not persist
were infected as frequently as males, whether the males
91
persisted or not. Females which did persist were infected
less frequently than other animals. This pattern suggests
that female immigrants may be more affected by infections
than male immigrants. In contrast, in observations of
overwinter survival, males which did not reappear in spring
were infected more frequently than all other animals.
Although males and females in the enclosure experiment were
inoculated with equal frequency, it is not known how often
they became reinfected between Nov.
(the last experimental
inoculation) and March (collection). Thus, the probability
of surviving the winter may be a function of the number of
times animals were infected.
Several studies have found negative associations
between host survivorship and parasites in naturally-
occurring infections (Arnqvist & Maki 1990, Boonstra et al.
1980, Burgett et al. 1990, Hudson et al. 1992a, Gulland &
Fox 1992, Lehmann 1992, Ross et al. 1989 and Royce &
Rossignol 1990). However, other observational studies did
not find any correlation between parasitism and host
survivorship (Goater et al. 1989, Johnson et al. 1991, Moss
et al. 1990). Keith et al.
(1986) found evidence that only
one of six parasites examined affected snowshoe hare (Lepus
americanus) survival. Only a few studies have used
controlled field manipulations to directly test the effect
of parasites on host survival. In some cases, experiments
have confirmed trends seen in observations (Hudson et al.
1992a, Lehman 1992, Trout et al. 1992); in at least one
92
case, results of experiments have contradicted observations
(Samson 1987).
Because E. arizonensis does not cause obvious pathology
in deer mice, we also examined weight loss as an indication
of a subtle mechanism through which E. arizonensis might
influence deer mouse survival (Prediction 3). Results of
naturally-occurring and experimental infections did not
support the prediction that E. arizonensis infections cause
weight loss in deer mice. Our results indicated no
significant weight loss associated with infections in
adults. However, naturally-infected, free-ranging juveniles
weighed significantly more than uninfected juveniles, the
opposite of prediction 3. Although juveniles may have gained
more weight while infected, it may also be that heavier
juveniles were older. Older juveniles may be more active
outside the nest, increasing their encounter rate with E.
arizonensis; they may also have less maternal immunity,
leaving them more susceptible to infections. Unfortunately,
we were unable to include juveniles in either experiment.
A few other authors have examined weight loss as a
possible mechanism for parasite induced reduction in
survival and/or reproductive success in birds and mammals
(Booth et al. 1993, Howe 1992, Keith et al. 1986, Lehmann
1992, Moore & Bell 1983). Of these, only Booth et al.
(1993), studying rock doves (Columba livia) and Keith et al.
(1986), studying snowshoe hares
(L. americanus) found
significant negative associations between weight change and
93
parasitism. Other authors have examined changes in metabolic
rate (Booth et al. 1993, Munger & Karasov 1989), organ
pathology (Gellar & Christian 1982, Watkins et al. 1991) and
blood abnormalities (Wiger 1977) with variable results.
None of the factors discussed above addresses the
direct mechanism of host mortality. Among other things,
parasitized animals may succumb to starvation, cannibalism
and predation (Hudson et al. 1992b, Temple 1987, Wiger
1977). In our study, it is unclear which of these factors
was important. In winter deer mice typically weighed 20-25%
less than at other times of the year (pers obs.) indicating
possible nutritional stress. However, some of this weight
loss is probably due to seasonal changes in reproductive
condition (Millar 1989, pers obs). Predators were observed
both in the field (short-tailed weasels, Mustela erminea)
and near enclosures (falcons and hawks). It seems likely
that parasitized deer mice were less able to avoid predation
than unparasitized animals due to poorer general condition
(Wiger 1977, Scott 1988). In addition, adverse weather has
been shown to compound the negative affect of parasites
(Lope et al. 1993).
It is becoming increasingly clear that a wide variety
of parasites (including arthropods, helminths and
protozoans) negatively affect survival of free-living hosts.
In addition, several observational and experimental studies
have documented negative affects of parasites on host
reproduction (Durnin 1993, Lafferty 1993, Moller 1993,
94
Schall 1983). In some cases, the effects appear to be
density dependent (Scott 1988). Thus, there is clear
empirical evidence that parasites in natural populations
have the potential to affect host abundance. Field
experiments at the level of the host population are needed
to determine whether the effects on reproduction and
survival act in a compensatory manner or to regulate host
density.
95
Chapter 5
Variable Levels of Immunity to E. arizonensis
96
Abstract
Acquired immunity to parasites may affect both host and
parasite population dynamics. Although immunity has been
studied experimentally in laboratory-reared hosts, less
attention has focused on free-living animals.
I examined
acquired immunity of free-living deer mice (Peromyscus
maniculatus) to naturally-occurring and experimental
infections of Eimeria arizonensis (Protozoa: Coccidia). In a
mark-recapture study, I found evidence of complete immunity
to natural infections in only 1 of 3 years and evidence of
partial immunity in 2 of 3 years. I subsequently examined
immunity to experimental infections by giving laboratory-
reared, free-living and enclosure populations of deer mice 2
or 3 consecutive E. arizonensis infections. Greater than 90%
(14 of 15) of laboratory-reared animals developed immunity
after only 1 exposure, confirming that E. arizonensis is
immunogenic. However, animals living under natural/semi­
natural conditions developed significantly less immunity in
2 of 3 experiments. These observational and experimental
results suggest that immunocompetence of free-living deer
mice to E. arizonensis may be quite variable. These results
suggest that care should be taken when generalizing from
laboratory to field situations in studies of acquired
immunity.
97
Introduction
The ecology of host-parasite relationships differs from
that of other symbiotic organisms in that hosts may have
varying levels of immunity to their parasites (Quinnell &
Keymer 1990). In vertebrate hosts, acquired immunity may
play a large role in determining parasite and (indirectly)
host population dynamics (Anderson 1991). Two different
types of parasites are recognized: 1) microparasites
(viruses and bacteria) go through multiple reproductive
cycles within the host and usually induce complete immunity
after only one infection (Anderson & May 1979); 2)
macroparasites (helminths and arthropods) shed eggs or
larvae from the host and larvae must infect a new host
before reproduction occurs; acquired immunity is often
partial and the degree of immunity may be dose (density)
dependent (May & Anderson 1979). In addition, acquired
immunity to macroparasites may be lost if enough time passes
between infections. Although protozoans are generally
categorized as microparasites because of their mode of
reproduction, they are often more similar to macroparasites
with respect to acquired immunity (Healing & Nowell 1985,
Quinnell & Keymer 1990).
Acquired immunity to macroparasites and protozoans has
been demonstrated experimentally in many host-parasite
systems in the laboratory (Gregory, Keymer & Clarke 1990,
Higgs & Nowell 1988, Lillehoj 1988, Wakelin 1978, 1987,
98
Wassom et al. 1986) and under semi-natural conditions
(Quinnell 1992). However, it is unclear whether these
results can be generalized to field situations. Laboratory
and semi-natural conditions may differ greatly from field
conditions with respect to temperature, resource
availability and social stress. All of these factors may
reduce the ability of the host to display an immune
response. In addition, genetic variability of host
laboratory stocks may be lower than their free-living
counterparts. Thus, while hosts may be capable of developing
immunity under ideal laboratory conditions, immunocompetence
may be much more variable in the field.
There are several ways to detect acquired immunity in
field populations. Convex age-intensity curves can indicate
the presence of acquired immunity among naturally infected,
free-living hosts (Anderson & May 1985). In a convex age-
intensity curve, parasite intensity or burden is low in
young hosts because of low exposure, increases in young
adults as exposure increases and drops again as animals
acquire immunity. In protozoans, age-prevalence data
(proportion of animals infected) are often used instead of
age-intensity data (Aron & May 1982). A few studies have
found evidence of the presence of acquired immunity using
these methods (Fulford et al. 1992, Gregory, Montgomery &
Montgomery 1992, Woolhouse et al. 1991). However, the same
patterns of infection can also be explained by other
processes such as parasite-induced mortality or behavioral
99
differences among hosts of different ages leading to
differential susceptibility among age classes (Bundy 1988,
Fulford et. al 1992, Pacala & Dobson 1988).
Another way to detect acquired immunity in naturally
infected hosts is to follow individual hosts over time using
mark-recapture techniques. If an animal becomes immune it
should not become reinfected after losing its initial
infection (complete immunity), or subsequent infections
should have a lower intensity (partial immunity). However,
behavioral differences could also lead to observed
differences in parasitism when following individual hosts.
Ideally, both age-intensity/prevalence and mark-recapture
studies should be accompanied by field experiments which
directly test whether free-living animals exhibit immunity.
Eimerians (Apicomplexa: Coccidia) are excellent
parasites to use in observational and experimental field
studies of acquired immunity because large numbers (often >
106)
of oocysts are shed in the feces and infections are of
short duration (< 20 days)
(Long 1982), thus multiple
infections can be followed non-invasively by examination of
feces. In addition, experimental infections of known doses
can be given. Laboratory studies indicate that even small
numbers of eimerians are often enough to induce complete
immunity after as few as four exposures and immunity can be
detected as a drop in fecal oocyst output (Higgs & Nowell
1988, Lillehoj 1988, Rose 1987).
100
I examined acquired immunity to natural and
experimental Eimeria arizonensis infections in deer mice
(Peromyscus maniculatus). I hypothesized that free-living
deer mice become immune to natural infections. If animals
develop complete immunity, they should not become infected
repeatedly (Prediction 1). If animals become partially
immune, there should be a decrease in oocyst output from one
infection to the next (Prediction 2). Observations in a
mark-recapture population in three years suggested that the
proportion of hosts developing complete and partial immunity
was highly variable among years. Several alternative
hypotheses could account for this variability:
1)
animals
may be immunocompromised under some conditions and observed
immunity was due to other factors (see above); 2)
E.
arizonensis may not be immunogenic; 3) animals were immune
but I was unable to detect immunity with the methods I
employed.
I conducted a series of laboratory and field
experiments to differentiate between the following
predictions from the hypotheses: 1)
if variability observed
among naturally infected animals is real, there should be
variability among different experimental conditions; 2) if
E. arizonensis is not immunogenic, none of the
experimentally infected animals should become immune and 3)
if I failed to detect immunity to natural infections,
animals should develop immunity under all experimental
conditions.
101
Methods: Field Observations
A 10 x 12 trapping grid was established in MacDonald
Forest (15 km NW Corvallis, OR) in March 1990. Sherman live-
traps were spaced 10 m apart in 1990 and 15 m apart
thereafter. Traps were set for 2-3 consecutive nights at 2-3
week intervals from 15 April
17 Nov, 1990, 30 March
17
12 July, 1992. All deer mice were
Nov, 1991 and 7 March
either ear-tagged or toe-clipped for individual
identification at first capture. Animals were released each
morning at their point of capture.
Feces were collected from traps, weighed and incubated
in K2Cr207 solution (2.5% w/v) at room temperature for one
week (this procedure facilitates the development of oocysts
into the infective stage and is necessary for identification
to species). Soiled traps were washed before they were
returned to the field.
Fecal samples were macerated and examined by Sheather's
sugar flotation (Ash & Orihel 1987). The number of oocysts
per gram in fecal samples was estimated by either counting
all oocysts present or averaging counts from 10
representative microscope fields and extrapolating to the
entire sample. Thus, counts were estimated as levels 0-6
where 0 = no oocysts detected, 1 = < 10 oocysts/gram, 2 =
10-99 oocysts/gram, 3 = 100-999 oocysts/gram, etc. An
attempt was made to distinguish new infections from
continued infections:
if
an animal which was shedding < 1000
102
oocysts/gram had been shedding oocysts during the previous
trapping session, the second infection was assumed to be a
continuation of the first.
I previously determined that E.
arizonensis oocysts are first shed in deer mouse feces on
day 4 post-inoculation and are shed for 6-8 days, Chapter 2.
Results: Field Observations
Prediction 1, Complete Immunity: animals should not become
infected repeatedly
Only animals which were captured > 5 times (ie, at
least 11 weeks in the population) were used in this analysis
so that they would have adequate opportunities to become
reinfected. Of the 198 deer mice captured over the course of
the study, 42 individual deer mice were caught > 5 times in
3 years and 8 were caught > 5 times in two years. Thus, the
total sample size for this analysis was 50 animal-years.
Figure 5.1 shows that 43-100% of deer mice became infected
repeatedly in all years. However, the modal number of times
animals became infected in 1990 was 1, indicating that
complete immunity may have occurred. In 1991 and 1992, the
modal number of times animals became infected was 3 and 4,
respectively. Only 3 of 29 animals became infected < 2 times
in these two years combined.
103
1990
1991
1992
0
1
3
4
5
6
7
Number of Times Infected
Figure 5.1. Number of times individual, free-living deer
mice were infected with naturally occurring Eimeria
arizonensis each year. Animals were live-trapped, marked and
released. Animals included in this analysis were in the
population at least 11 weeks.
104
Prediction 2, Partial Immunity
All animals which were infected more than once were
used in this analysis. Figure 5.2 shows the median change in
oocyst output from 1 infection to the next (0 = no change,
positive numbers indicate an increase in oocyst output and
negative numbers indicate a decrease). Animals appear to
have developed partial immunity to E. arizonensis after only
one infection in 1990 and after 2 infections in 1991. There
was no evidence of partial immunity in 1992.
Methods: Immunity Experiments
In Chapter 2, we demonstrated that laboratory reared
animals can develop immunity to E. arizonensis. The results
of this experiment are summarized here as a standard to
determine whether the other groups developed immunity (lab­
reared animals will be referred to as the first group). The
second group of animals tested consisted of free-living deer
mice and the third group consisted of animals in large (0.25
ha) field enclosures.
The E. arizonensis oocysts used in these experiments
were obtained from the feces of two deer mice from the
MacDonald Forest population in Spring 1990. The oocysts were
stored at 4°C and fresh oocysts were generated periodically
using laboratory born, Eimeria free deer mice which also
originated from the MacDonald forest population. The oocysts
105
///
Infec 1-2
\\X\ Infec 2-3
AL
A
Infec
3-4
1.0
0.5
0.0
C
a)
C
C
0.5 1
_E
U
0
1 .0
t
a)
75
1990
1991
1992
Figure 5.2. Change in oocyst output with successive
infections. Median change in oocyst output from one
infection to the next in naturally infected, free-living
deer mice. If animals developed partial immunity, there
should be a decrease (negative change) in oocyst output with
each successive infection. If no immunity develops, there
will be no change or an increase in oocyst output (positive
change).
106
used in the experiments were generated 6 days
4 months
before the initial inoculations.
Laboratory Experiment
Laboratory animals originated from MacDonald forest and
had been in captivity 3-5 generations. Table 5.1 shows the
protocol for the laboratory experiment. Deer mice were
lightly anaesthetized with halothane and inoculated with one
of three levels (103,
104,
105)
of E. arizonensis by stomach
intubation. Animals were housed individually in cages with
wire-mesh bottoms. Feces were collected for 20 days post­
inoculation (PI)
in pans suspended under the cages. Animals
were inoculated a second time (given a challenge infection)
on day 21 PI with the same number of oocysts as in the first
inoculation. Feces were again collected for 20 days post-
challenge (PC). Fecal samples were examined by McMaster's
technique (Dunn & Keymer 1986) to determine total number of
oocysts shed over the course of the infection.
Field Experiment
The animals from the mark-recapture population were
used after the observational study had ended. In addition,
to increase sample size, a second mark-recapture grid was
established approx. 1 km from the first. The protocol for
the field experiment is shown in Table 5.1. Animals were
lightly anaesthetized and inoculated by stomach intubation
107
Table 5.1. Protocol for laboratory and field experiments.
Event
Number of oocysts
Days PI*
Treatment
Laboratory (N = 5 in all cases except
Inoculation
Feces collection
Challenge
Feces collection
0
@
Control
[N = 4])
103, 109, 105
20 Days PI
21
103,
104,
105
103,
104,
105
20 Days PC"
Field (sample size in parentheses)
Inoculation
Feces collection
0
2 x 104
(9)
2 x 104
(9)
water
(11)
5 or 6
Challenge
20 or 21
Feces collection
25 or 26
(5-6 PC)
* PI = post-inoculation
** PC = post-challenge
2
x 104
(12)
108
with either 20,000 oocysts (treatment and naive control) or
water (control). Animals were free-living throughout the
experiment. Traps were set overnight on days 5 or 6 PI/PC
(these are the days of peak oocyst output for E.
arizonensis, Chapter 2) and animals were released the
following morning. Feces were collected out of traps,
weighed to the nearest 0.01 g and processed as in the
laboratory experiment. For this experiment and the enclosure
experiment, the total number of oocysts shed per gram of
feces was calculated.
Enclosure Experiment
Free-living deer mice were live-trapped and placed in
4,
50 m x 50 m (2 treatment and 2 control enclosures) field
enclosures in the fall of 1993. There were 100 live-traps
(10 x 10 grid with 5 m spacing) in each enclosure and each
was surrounded by a 1 m high fence above ground and a 0.5 m
fence below ground. Thus, animals were exposed to aerial but
not terrestrial predators and could not disperse out of the
enclosures. The dominant vegetation was alfalfa; deer mice
were not given water or additional food but were provided
with nest boxes (20/enclosure). All treatment animals were
given 50,000 oocysts at each inoculation and all controls
were given water.
The majority of animals were first inoculated in Sept.
1993, given a challenge infection in Oct. 1993 and a second
109
challenge infection in Nov. 1993 at 5 week intervals.
However, a second group of animals was first inoculated in
Oct. 1993 and given a challenge infection in Nov. Thus, the
second group provides a replicate for the first group. Table
5.2 shows the sample sizes and timing of the inoculations
for this experiment. Because sample sizes in each enclosure
were small, data were pooled within treatments. Animals were
trapped and fecal samples were collected as in the field
experiment.
Statistical Analyses
The degree of change in oocyst output for each
treatment animal was calculated as the percent change in
output compared to the mean oocyst output of naive animals
inoculated at the same time and with the same number of
oocysts (eg., oocyst output of an enclosure animal
challenged with 50,000 oocysts in Nov 1993 was compared to
the mean output of naive animals inoculated with 50,000
oocysts for the first time in Nov 1993). This approach was
necessary to have a common basis for comparing the results
of the three experiments. The percent change in oocyst
output was then compared between the laboratory experiment
and the field/enclosure experiments by Mann-Whitney U test.
The proportion of animals that developed immunity (see
below) was compared with Fischer's exact test. Alpha levels
were corrected for the number of comparisons made by the
110
Table 5.2. Sample sizes in enclosure experiments. All
treatment animals were inoculated with 50,000 oocysts, all
control animals were inoculated with water and inoculations
were 5 weeks apart.
3 =
* 1 = initial inoculation, 2 = challenge inoculation,
2nd challenge.
** Because so few naive animals remained in treatment
enclosures in Nov., 8 of 9 naive treatment animals first
inoculated in Nov. were laboratory reared.
Number of Times Inoculated*
Month of 1st
Control
Treatment
Inoculation
1
2
3
1
2
12
23
16
Sept
32
15
Oct
10
6
Nov
9
10
0
3
7
11
111
Bonferroni method (Zar 1984). Alpha oh 0.05 was considered
statistically significant in all cases.
Results: Immunity Experiments
Effectiveness of Inoculations
Animals inoculated for the first time (naive animals)
in all experiments shed large numbers of oocysts in all
cases (Figures 5.3 A,B). Very few sham-inoculated
field/enclosure animals shed oocysts and of those that did,
only one shed at levels close to treatment animals (Table
5.3). Thus, field/enclosure inoculations were successful and
it seems highly unlikely that treatment animals shed oocysts
during the experiment as the result of natural infections.
Laboratory Experiment
All 15 animals showed a large decrease in oocyst output
compared with that of controls. However, 14/15 deer mice
showed a > 90% (Mean = 97.1%) decrease. The 15th animal
showed only a 66 % decrease. Thus, for purposes of
comparison, I considered animals showing a > 90% decrease in
oocyst output from one infection to the next to be immune.
112
30
(1)
0
A
10' oocysts
25
10
tit:2 10
oocysts
oocysts
20
15 ­
10
0
0
Treatment
C
Control
VA
Sept
NN
Oct
ZEI Nov
Cl)
0)
U
0)
LL
O
0
(J)
(f)
L2.)
a
Fld Exp
Encl Exp
Figure 5.3. Experimental infections in naive animals. The
number of Eimeria arizonensis oocysts shed by naive
(previously uninfected) animals at their first experimental
inoculation (+SE). A) results from the laboratory
experiment; treatment deer mice were first inoculated three
weeks prior to the first inoculation of control deer mice.
B) results from field and enclosure experiments. Animals in
the field experiment were given 2 x 104 oocysts; enclosure
animals were given 5 x 104 oocysts. The field and enclosure
experiments were conducted in 1992 and 1993, respectively.
113
Table 5.3. Infections among sham inoculated animals.
Prevalence and intensity of infections among sham inoculated
animals in field and enclosure experiments.
Experiment/
Month
Field Exp
Prevalence:
number of animals
shedding oocysts (total)
4
(12)
Intensity:
oocysts/g feces
<
>
100
106
(3 animals)
(1 animal)
Enclosure Exp
Sept
2
(23)
< 10
Oct
2
(26)
< 10
Nov
1
(18)
< 1000
114
Comparisons Between Laboratory, Field and Enclosure Results
Significantly fewer animals in the field population and
in the first replicate of the enclosure population became
immune than in the lab-reared population (Figure 5.4 A).
However, the difference between the laboratory reared
animals and the enclosure animals was no longer significant
after the second challenge infection was given to enclosure
animals. Thus, these animals developed the same degree of
immunity as their lab-reared counterparts, but only after 2
exposures. In addition, the second group of enclosure
animals (initially inoculated in Oct 1993,
1 month after the
initial inoculation of the first group) all developed
immunity after only one inoculation. The results are similar
if actual decrease in oocyst output is compared (Figure 5.4
B) .
Discussion
In naturally infected, free-living deer mice, there was
evidence of complete immunity in 1 of 3 years and evidence
of partial immunity in 2 of 3 years. In 1 year in which
partial immunity was detected, it was not apparent until
animals had been infected at least twice. Thus, it appears
that immunity to naturally occurring infections was highly
variable between years. My observational results could be
explained by several hypotheses: 1) animals may be
immunocompromised under some conditions; 2) E. arizonensis
115
100 -
A
80
60
40
20
0
100 ­
B
80 -1
-T.
60
40
20
Q)
0
..,c>i,N
.,
(<\<<>/
`'
6c,
..,c;'(t,
or
<<:- \`
,..,Ckce`
'
\c3
c;(
Figure 5.4. Immunity to experimental infections in the
field. Percent of deer mice which became immune to Eimeria
arizonensis (A) and percent decrease (+SE) in oocyst output
(B) after 1 or 2 (enclosure, experiment only) challenge
infections. Experiments were compared using results from the
laboratory as a standard; (*) indicates values significantly
different than laboratory results (P < 0.05).
116
may not be immunogenic; or 3)
I was unable to detect
immunity with the methods I employed. I conducted a series
of laboratory and field experiments to differentiate between
the hypotheses. The experimental results were consistent
with hypothesis 1 but did not support hypotheses 2 and 3;
the laboratory reared animals developed immunity to E.
arizonensis after only one inoculation (not consistent with
H2). Additionally, in 2 of 3 field/enclosure experiments,
animals did not develop the same degree of immunity as in
the laboratory (not consistent with H3).
There are several reasons why free-living animals may
frequently be immunocompromised compared with laboratory-
reared animals. These can be grouped into two categories: 1)
genetic (Wakelin & Blackwell 1988, Wassom et al. 1988) and
2) environmental factors (McCallum 1990). It is possible but
unlikely that my laboratory population had a higher
proportion of genetically "resistant" individuals. The
laboratory colony originated from the same populations as
the animals used in the field experiments, had only been in
captivity 3-5 generations and was maintained as an outbred
colony. It seems more likely that the differences seen
between laboratory and field populations were due to factors
in the host's environment. McCallum (1990) suggests that, in
general, transient factors may play a larger role than
genetics in determining predisposition to parasite
infections in free-living animals.
117
At least one environmental factor would predispose
free-living animals to be more likely to develop immunity:
exposure to natural E. arizonensis infections prior to
experimental inoculation. Despite this, in 2 of 3
experiments, field caught animals exhibited some decrease in
immunocompetence with respect to their laboratory-reared
counterparts. Many environmental factors would predispose
free-living animals to be less likely to develop immunity
than laboratory-reared animals including protein deficiency,
(McMurray 1984, Slater 1988, Slater & Keymer 1986), very low
levels of lipids and some trace elements (Chandra & Amorin
1992, Chandra & Dayton 1982), low night temperature (Noble
1966) and the presence of other concurrent parasitic
infections (Cox 1987, Rose et al. 1994). Stress (e.g. due to
behavioral interactions among conspecifics) can also lead to
impaired immune function especially if it is of short
duration. Chronic stress may actually enhance immune
response if animals can adapt through behavioral or other
means. If animals cannot adapt, chronic stress can also
impair the immune response (reviewed in Griffin 1989).
Although laboratory animals may be under chronic
stress, the environmental conditions were very stable in my
experiments. Animals were fed and given access to water ad
libitum, housed individually and the room temperature was
maintained at 24-27°C. They were also exposed to few other
parasites (colony animals were observed to have fleas and a
nematode). Thus, laboratory animals may have been able to
118
adapt to chronic stress. In contrast, field conditions were
much more variable and animals may not have been able to
adapt to stress. In addition, the field/enclosure
experiments were conducted in the fall and conditions were
harsh. Free-living animals tended to be lighter (11-18 g vs
17.5-24 g in the laboratory), indicating the possibility of
nutritional deficiencies, temperature was highly variable
and population densities were within the normal range for
deer mice (pers obs).
Finally, long-term monitoring of deer
mice indicated that animals were frequently infected with at
least eight other intestinal tract parasites (2 eimerians, 2
isosporans [family Eimeriidae], three pinworms [Nematoda]
and a trematode). These factors may explain their apparent
lowered ability to wage an immune response.
Although laboratory studies of acquired immunity are
quite common, few studies have attempted to determine
whether immunity seen in the laboratory also occurs in the
field. While the results of some field studies of age-
intensity and age-prevalence curves corroborated laboratory
studies (Fulford et al. 1992, Gregory et al. 1992, Woolhouse
et al. 1991), others have not (Behnke & Wakelin 1973,
Gregory 1992). In part, this may be due to the difficulty in
aging field-caught animals (Gregory 1992). A few studies
have documented variability in hematological parameters
associated with acquired immunity in wild rodents
(Dobrowolska et al. 1974, Lochmiller et al. 1994). However,
none of these studies have been experimental. Quinnell
119
(1992) found immunity in an enclosed population of wood mice
(Apodemus sylvaticus) experimentally infected with the
nematode Heliqmosomoides polyqvrus. However, the enclosure
used in this study was small (7 x 6 m), crowded (1 mouse/2
m2), there was no vegetation, predators were excluded,
animals were given food and water ad libitum and they were
given an anthelminthic at the beginning of the experiment.
Thus, while Quinnell (1992) came closer to a natural
situation than a laboratory because animals were exposed to
natural temperature variation, it may not have come close
enough for animals to exhibit natural variation in
immunocompetence. My field experiment represents the first
experimental study of acquired immunity conducted on
completely free-living animals. In addition, the enclosures
used in the enclosure experiment mimicked a natural
situation closely. Although the sample sizes of deer mice in
my field/enclosure experiments were low and both were
conducted in the fall, field observations of naturally-
occurring infections suggest that some degree of
immunocompromise may be common in this system.
Most free-living vertebrates experience more
environmental variability than their captive counterparts.
Although laboratory studies often provide a much needed
starting point for studies of immunity in free-living
animals, the discrepancy between my laboratory and field
results suggests that care should be taken when generalizing
to field situations. The role of acquired immunity to
120
parasites may have ramifications for the population dynamics
of both parasites and hosts. Thus, further study on how
environmental factors interact to mediate immunity in free-
living animals is clearly needed.
121
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