INFLUENCE OF BODY CONDITION AND HOMO- VS. HETEROSUBTYPIC
IMMUNITY ON INFLUENZA A VIRUS INFECTION IN MALLARD DUCKS:
EXPERIMENTAL INFECTION DATA
By
Dustin M. Arsnoe

A THESIS
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
MASTER OF SCIENCE
Fisheries and Wildlife
2010

 

 

ABSTRACT
INFLUENCE OF BODY CONDITION AND HOMO- VS. HETEROSUBTYPIC
IMMUNITY ON INFLUENZA A VIRUS INFECTION IN MALLARD DUCKS:
EXPERIMENTAL INFECTION DATA
By
Dustin M. Arsnoe
Migrating waterfowl, particularly mallards (Anas platyrhynchos), are implicated in the global
spread of influenza A viruses (IAVs). Experimental infection studies are used to assess a bird’s
ability to maintain IAV, but these studies have not represented the natural variation in
physiological condition observed in migrating waterfowl and have only used captive-bred birds.
We assessed how body condition affects susceptibility, viral shedding, and antibody production
in wild-caught and captive-bred juvenile mallards challenged with IAV H5N9. We found wild
mallards fed ad libitum were more susceptible to IAV infection, shed higher virus loads, and
shed virus more frequently compared to birds in poor condition. No difference in viral load was
detected between wild and captive-bred mallards. Antibody production did not vary according to
condition. We conclude captive-bred mallards represent wild mallards during IAV infection
when fed ad libitum, and IAV infection in mallards is largely influenced by body condition.
Waterfowl are frequently exposed to the same or different IAV subtypes during migration.
We examined whether primary H5N9 infection in juvenile mallards provides long-term
immunity towards reinoculation with the same or different IAV subtype. Our findings
demonstrate mallards can mount long-term protective immunological responses against
reinfection with the same H5N9 virus, but were susceptible to reinfection with a different H7N3
subtype 49 days after primary inoculation. Additional IAV immunity studies are needed using
different virus subtypes and different challenge timings.

 

 

 

ACKNOWLEDGEMENTS

I am grateful to the Department of Fisheries and Wildlife at Michigan State University
(MSU) and the American Museum of Natural History for funding this research project. I thank
my committee chair Dr. Jennifer Owen for her never ending encouragement and guidance during
my graduate education. My committee members, Drs. Jean Tsao and Scott Winterstein,
generously provided their time and expertise which undoubtedly improved the quality of my
work.
I thank my colleagues in the Avian Health and Disease Ecology Laboratory, Emily Johnston
and Tiffanie Hamilton, for their assistance with capturing ducks and collecting samples. I am in
debt to Emily Cornelius, Travis Belknap, Isis Kuczaj, Jennifer Sidge, Cole Iaquinto, Sarah
Hamer, and Kaye Ivens for their hard work and dedication as research assistants. This project
would have not been successful without the dependability and skillful help of so many others.
I greatly appreciate the help of several researchers at MSU who donated resources and
expertise towards my laboratory diagnostics. I thank Dr. Roger Maes and Kari Chciuk for
helping me get my hemagglutination inhibition assays running. Pam Campbell generously
donated chicken embryos and provided training for virus propagation techniques. Jeff Landgraf
was most influential in performing RRT-PCR testing.
I owe great thanks to the staff at the MSU Research Containment Facility for their
dependable assistance in caring for my captive ducks. I thank Dr. Hon Ip at the USGS National
Wildlife Health Center for supplying the virus stock for the study. I thank the Michigan
Department of Transportation for allowing me to trap ducks on their mitigation ponds, and the
Michigan DNRE for capturing and donating ducks for my research.

iii

 

 

 

TABLE OF CONTENTS

LIST OF TABLES……………………………………………………………………………… v
LIST OF FIGURES…………………………………………………………………………….. vi
CHAPTER 1
INFLUENCE OF BODY CONDITION ON INFLUENZA A INFECTION IN
MALLARD DUCKS…………………………………………………………………………… 1
Introduction………………………………………………………………………………… 1
Materials and Methods……………………………………………………………………... 4
Animals……………………………………………………………………………... 4
Housing…………………………………………………………………………….. 5
Body condition index……………………………………………………………….. 5
Experimental design………………………………………………………………... 6
Virus………………………………………………………………………………... 7
Matrix gene RTT-PCR……………………………………………………………… 7
Virus titration………………………………………………………………………. 8
IDEXX FlockChek* ELISA…………………………………………………………. 8
Hemagglutination inhibition (HI) assay……………………………………………. 8
Data analysis……………………………………………………………………….. 9
Results………………………………………………………………………………………. 11
Susceptibility to LPAIV (H5N9) infection…………………………………………... 14
Concentration of viral shedding……………………………………………………. 15
Duration of viral shedding………………………………………………………….. 17
Serologic Response…………………………………………………………………. 20
Discussion…………………………………………………………………………………... 22
CHAPTER 2
ROLE OF HOMO- AND HETEROSUBTYPIC IMMUNITY ON REINFECTION
OF MALLARD DUCKS WITH INFLUENZA A VIRUS…………………………………….. 29
Introduction…………………………………………………………………………………. 29
Materials and Methods……………………………………………………………………… 31
Animals……………………………………………………………………………… 31
Experimental design………………………………………………………………… 31
Results………………………………………………………………………………………. 33
Homosubtypic LPAIV (H5N9) reinoculation……………………………………….. 33
Heterosubtypic LPAIV (H7N3) reinoculation……………………………………… 33
Discussion…………………………………………………………………………………... 37
REFERENCES…………………………………………………………………………………. 40

iv

 

 

 

LIST OF TABLES

TABLE 1.1. Body mass (g) for 10 captive-bred mallards (C1 - C10) fed ad libitum from the
time of purchase through LPAIV H5N9 challenge…………………………………………….. 13
TABLE 1.2. Daily mean (± 1 standard error) virus titers (log10 PFU/mL) for mallards inoculated
with LPAIV H5N9. Mean titer includes birds without detectable viral RNA, whereas minimum
and maximum titers are reported for the number of birds (n) with detectable viral RNA on the
given day……………………………………………………………………………………….. 16
TABLE 1.3. Serological status of mallards before and after LPAIV H5N9 challenge
using ELISA and HI tests. Mean (± 1 standard error) test scores include all birds in each
treatment………………………………………………………………………………………... 21
TABLE 2.1. Serological status of mallards before and after homosubtypic LPAIV H5N9 and
heterosubtypic LPAIV H7N3 reinoculation. Mean (± 1 standard error) test scores include all
birds in each treatment………………………………………………………………………….. 35

v

 

 

 

LIST OF FIGURES

FIGURE 1.1. Body condition for wild mallard treatment groups throughout the study. Data
points represent mean condition (± 1 standard error), A = start of diet manipulation, B = LPAIV
H5N9 inoculation……………………………………………………………………………….. 12
FIGURE 1.2. Virus shedding concentration results for captive-bred and wild mallard treatment
groups. Data points represent mean titer (log10 PFU/mL) for each day sampled.……………… 15
FIGURE 1.3. Virus shedding concentration (log10 PFU/mL) results for all mallards with
detectable viral RNA using matrix gene RTT-PCR. A: captive-bred (n = 10), B: wild ad libitum
(n = 10), C: wild lean (n = 10), D: wild poor (n = 6)…………………………………………… 18
FIGURE 1.4. Duration of viral shedding (A) and number of positive samples (B) for captivebred and wild mallard treatment groups. Bars represent means ± 1 standard error, asterisks
identify groups significantly different from one another (Mann-Whitney rank sum, alpha = 0.05).
The number of positive samples was calculated from the 4 sampling days between 1-5 dpi.… 19
FIGURE 1.5. Breast condition score for wild mallard treatment groups at capture vs. LPAIV
H5N9 challenge. Bars represent mean condition score ± 1 standard error (n = 10), asterisks
represent significant within group differences (paired t-test, alpha = 0.05)……………………. 25
FIGURE 2.1. Virus shedding concentration (log10 PFU/mL) for all mallards with detectable
viral RNA after reinoculation with (A) homosubtypic LPAIV H5N9 (n = 1), and (B)
heterosubtypic LPAIV H7N3 (n=10). Birds were challenged 7 weeks after primary LPAIV
H5N9 inoculation……………………………………………………………………………….. 36

vi

 

 

 

CHAPTER 1
INFLUENCE OF BODY CONDITION ON
INFLUENZA A VIRUS INFECTION IN MALLARD DUCKS

INTRODUCTION

Birds associated with aquatic environments including Anseriformes (particularly ducks,
geese, and swans) and Charadriiformes (particularly gulls, terns, and waders) serve as the natural
reservoir for influenza A viruses (IAVs) (Webster et al. 1992). Among these birds, dabbling
ducks within the Anas genus are recognized as the primary reservoir hosts (Olsen et al. 2006).
Prevalence of IAV infection in dabbling ducks peaks during autumn when immunologically
naïve juvenile waterfowl congregate before migrating south (Wallensten et al. 2007). During
migration, many of these birds travel long distances and potentially spread low pathogenic avian
influenza viruses (LPAIVs) among countries or continents. Migration is considered one of the
most physiologically demanding activities in the animal world and animals vary in their ability to
meet the associated energetic challenges. Despite peak IAV prevalence occurring during
migration, studies have not fully evaluated how natural variation in waterfowl condition
influences a bird’s ability to serve as a reservoir host for LPAI.
An understanding of waterfowl host competence during the migratory period is needed to
understand how LPAIV is maintained and transmitted. It has long been assumed that waterfowl
are asymptomatic carriers of LPAI and may transmit the virus during migration (Webster et al.
1992). However, recent examination of migratory behavior in wild Bewick’s swans (Cygnus
columbianus) found that swans infected with LPAIV exhibited delayed migration, reduced
feeding rates, and shorter flight distances compared to uninfected conspecifics (van Gils et al.
1

 

 

 

2007). Furthermore, Latorre-Margalef et al. (2009b) found that migrating mallard ducks (Anas
platyrhynchos) infected with IAVs had significantly lower body mass than did uninfected birds.
These studies concluded that LPAIV infection may incur larger physiological costs to migrating
waterfowl than was previously thought. In response, Flint and Franson (2009) provide an
alternative explanation, suggesting that birds in poorer condition exhibit reduced immune
function and are more susceptible to IAV infection (i.e. “condition-dependent hypothesis”)
(Latorre-Margalef et al. 2009a). If their hypothesis is correct, host condition would predict
susceptibility to infection, and concentration and duration of viral shedding.
Despite the suggested influence of host condition on IAV infection, laboratory experiments
have used birds in normal physiological condition (Sturm-Ramirez et al. 2005, Brown et al.
2006, Keawcharoen et al. 2008, Fereidouni et al. 2009, Jourdain et al. 2010). These studies have
not accurately represented the energetic and immunological condition of migrating ducks. For
example, migrating waterfowl can experience declines in condition due to inclement weather
(Robb et al. 2001) and decreased food availability (Moon et al. 2007). Furthermore, decreases in
condition have been correlated with immunosuppression (Owen and Moore 2008, Buehler et al.
2009). Owen and Moore (2008) found that immune function in migrating thrushes (Family
Turdidae) was positively related to body condition. Common eiders (Somateria mollissima)
experienced reduced immune function during periods of mass loss caused by enhanced
incubation effort (Hanssen et al. 2005). It remains unclear how natural fluctuations in body
condition influence susceptibility and severity of IAV infection.
We evaluated the condition-dependent hypothesis through experimental inoculation of wildcaught juvenile mallards with LPAIV. Mallards were selected as the focal species because they
are the most abundant migratory dabbling duck in North America and Eurasia, and have

2

 

 

 

accounted for more IAV recoveries than any other species of bird (Krauss et al. 2004, Olsen et
al. 2006, Munster et al. 2007). Studies have shown that mallards in normal physiological
condition often remain asymptomatic to IAV infection, but shed high concentrations of the virus
(Sturm-Ramirez et al. 2005, Brown et al. 2006, Keawcharoen et al. 2008). We chose to use wildcaught mallards because previous experimental infection studies have used captive-bred mallards
(Homme and Easterda.Bc 1970, Cooley et al. 1989, Sturm-Ramirez et al. 2005, Brown et al.
2006, Keawcharoen et al. 2008, Fereidouni et al. 2009, Jourdain et al. 2010), which may not be
truly representative of wild mallard host competency. Accordingly, we included a group of
captive-bred mallards for comparison and validation of past research.
In this study, we tested the effect of body condition on susceptibility to infection and viral
shedding patterns in wild-caught juvenile mallards challenged with LPAIV. We hypothesized
food restriction and subsequent reduced body condition will result in (1) increased susceptibility
to infection, (2) increased concentration and duration of viral shedding, and (3) decreased
antibody production. In addition, we compared susceptibility and viral shedding patterns in wildcaught vs. captive-bred juvenile mallards.

3

 

 

 

MATERIALS AND METHODS

Animals:
Wild mallards were trapped in September 2009 to coincide with increased abundance of
staging waterfowl and seasonal peaks in avian influenza prevalence (Halvorson et al. 1985,
Wallensten et al. 2007). Trapping sites (n = 5) were located in shallow marshes surrounded by
cropland within the lower peninsula of Michigan. Birds were captured using portable swim-in
traps (Mauser and Mensik 1992) and rocket nets at sites previously baited with whole kernel
corn.
Mallards were aged in the field as juvenile or adult by examining wing plumage and cloacal
characteristics (Carney 1992). All juvenile birds were immediately transported to a biosafety
level 2 animal containment facility (see Housing, below), weighed to the nearest 1.0 g, and
measured for length of flattened wing chord (nearest 1.0 mm), head (0.1 mm), and
tarsometatarsus (0.1 mm). Upon capture, birds were tested for previous IAV exposure using the
MultiS-Screen ELISA (see Serologic Assays, below). Seronegative birds were isolated from one
another in separate cages to prevent any potential virus transmission within the facility. All birds
were retested with the ELISA at 20 days post-capture to ensure no birds had been recently
infected at time of capture. Previous research indicates 20 days is adequate time for
seroconversion (Brown et al. 2006). Thirty seronegative wild mallards (20 males, 10 females)
remained in the study, all other birds were released.
Once wild mallards were selected, 10 (8 males, 2 females) twelve week-old mallards were
purchased from a closed-flock hatchery (Ridgeway Hatcheries Inc., Ohio, USA). Captive-bred
birds were processed the same as wild mallards and were negative for previous IAV exposure.

4

 

 

 

Bird handling and all experimental procedures were approved by the Institutional Animal Care
and Use Committee of Michigan State University (protocol 03-09-052-00).

Housing:
Mallards were kept in the Michigan State University Research Containment Facility. Birds
were randomly assigned to three identical biosafety level 2 rooms and individually housed in
3

10.6 ft stainless steel rabbit cages at 20°C. Cages were positioned to allow birds to view one
another. Room lighting was adjusted weekly to match the natural photoperiod in Michigan at that
time. Each bird received water and grit ad libitum daily. All birds were acclimated for 30 days
before the start of the study. They were fed ad libitum a commercial maintenance food mash
(21% crude protein, 2.7% crude fat, 4.75% crude fiber; true metabolizable energy = 2.82 kcal/g).

Body condition index:
Capture mass for wild juvenile mallards was adjusted by subtracting the estimated mass of
remaining crop contents (1-25% = -28.2 g, 26-50% = -44.4 g, 51-75% = -68.9 g, 46-100% = 119.5 g), and 3.0 g were added for every hour a bird was held prior to recording its initial mass
(Robb et al. 2001). Body condition was estimated for males and females separately using
residuals from an ordinary least-squares (OLS) regression of adjusted mass vs. an index of body
size (Devries et al. 2008). The body size index was developed by performing a principal
component analysis (PC1) using wing chord, head and tarsus length (PROC PRINCOMP, SAS
institute 2002). Condition scores were calculated individually by dividing a bird’s residual from
the OLS regression by its predicted mass. The condition index was assumed to represent normal
pre-migration staging condition for juvenile mallards in Michigan. Body condition was not

5

 

 

 

estimated for the captive-bred birds. These closed-flock mallards exhibited significant structural
differences (i.e. reduced wing size, large head and tarsus lengths), and sample size (n = 10 birds)
was not large enough to produce a unique and reliable condition model.

Experimental design:
Wild mallards were randomly divided into three treatment groups (n = 10 birds/treatment).
Sex and location of capture were stratified among the groups. After birds acclimated 30 days,
diet treatments were initiated following protocols from a pilot study conducted in spring 2009
(Arsnoe unpublished data). Mean treatment conditions were manipulated through food
availability to relative conditions decided a priori 1) poor treatment = -20% body mass, 2) lean
treatment = -10% body mass, 3) ad libitum treatment = +0-5% body mass. Reduced conditions
were selected to replicate natural (lean) and substantial (poor) decrease in body condition
encountered by migrating waterfowl (Pawlina et al. 1993). Mallards were fed ad libitum to
simulate optimal foraging conditions. Captive-bred mallards (n = 10) were also fed ad libitum.
Body condition was assessed every five days by weighing birds to the nearest 1.0 g. In
addition, we monitored each bird’s body reserves by scoring their keel protuberance and breast
muscle development on a 0-3 point scale (Gregory and Robins 1998). When treatment groups
reached their desired condition levels, all birds were oropharyngeally inoculated with 1.5 mL of
6

10 PFU/mL LPAIV H5N9. Following inoculation birds were maintained on their treatment
diets to keep them at desired conditions. Cloacal and oral swabs were collected the first 3 days
post inoculation (dpi) and every 2 days thereafter until 28 dpi. Swabs from individual birds were
pooled together in 1.5 mL of BHI media with antimicrobial drugs (100X Anti-Anti, 1.0 mL/100
mL BHI media), and transported on dry ice to a -80 °C freezer. Blood serum was collected from

6

 

 

 

the brachial vein on 14, 21, and 28 dpi for serologic testing. At 28 dpi 20 mallards were
euthanized using CO2 asphyxiation, followed by cervical dislocation. The remaining 20
mallards were retained for the study described in Chapter 2 (see below).

Virus:
The LPAI virus used was A/Northern pintail/California/44221-761/2006 (H5N9), obtained
from USGS National Wildlife Health Center, Wisconsin, USA. This strain of IAV was selected
as it has been well characterized and serves as a model waterfowl-derived IAV in our laboratory.
Virus was propagated by inoculating the allantoic cavity of 9-11 day old embryonated chicken
eggs with 200 µL (1:10 dilution in DMEM media) (Woolcock 2008). Allantoic fluid was
harvested after 4 days, centrifuged and stored in 2 mL aliquots at -80 °C. Stock virus was titrated
using MDCK plaque assay as described by Tobita et al. (1975).

Matrix gene RRT-PCR:
Swab samples were thawed at 37 °C and thoroughly homogenized by vortexing. RNA
extractions were performed using the QIAamp viral RNA mini kit (QIAGEN, QIAGEN
Sciences, Maryland, USA) using 140 µl of sample material, according to the manufacturers’
instructions (Jonassen and Handeland 2007). Real-time RT-PCR assays were performed using
protocols targeting the matrix (M) gene (Spackman et al. 2002). Samples were processed at the
Michigan State University Genomics Laboratory on an ABI Prism 7900 Sequence Detection
System machine using the TaqMan One-Step RT-PCR Master Mix (Applied Biosystems, Foster
City, CA, USA). We detected the matrix gene of LPAIV H5N9 at 100nM and 500nM final
concentration, respectively. Two microliters of the final RNA prep were used as template in a 10

7

 

 

 

µl final reaction volume. Cycle threshold (ct) values were standardized by setting the baseline to
a threshold of 0.028 for all runs. All ct-values <40 were considered LPAI virus positive.

Virus titration:
Swab samples were prepared and processed (as described above) with known concentrations
of control viral RNA. We created RNA controls by serially diluting our stock virus of known
1

7

concentration (10 - 10 PFU/mL) in BHI media. RNA controls were used to generate a standard
curve, and sample titers were determined by entering the observed ct-value into the standard
curve equation (Lee and Suarez 2004). Swab samples from all birds were tested through the first
7 dpi. When a bird tested negative for viral RNA on two consecutive sampling events (i.e.
minimum 4 days after last positive sample) it was considered no longer shedding virus.

IDEXX FlockChek* ELISA:
Serum was tested using a commercially available IAV antibody ELISA kit (FlockChek* AI
MultiS-Screen, IDEXX Laboratories Inc., Maine, USA). According to the manufacturers’
instructions, samples with a signal-to-noise ≤ 50% were considered positive. Comparison of the
FlockChek* ELISA with the more recent NP-ELISA revealed both tests are equally reliable in
detecting IAV antibodies, and more specific and sensitive than traditional HI and AGP tests (Wu
et al. 2007).

Hemagglutination inhibition (HI) assay:
To quantify post inoculation serum antibodies, HI assays were performed using standard
protocols (Swayne et al. 1998), using chicken erythrocytes and four HA units of stock virus used
8

 

 

 

for inoculation. Serum samples were treated with 10% erythrocytes solution to remove
nonspecific inhibitors and agglutinins. Samples were processed in duplicate using a 0.5%
suspension of chicken erythrocytes. Antibody titers were expressed as the reciprocal of the
highest serum dilution yielding complete inhibition of hemagglutination. Samples with HI titers
≥ 8 were considered positive.

Data analysis:
The body size index, estimated by PC1, accounted for 60% and 56% of the variance
associated with structural measurements (wing chord, head length, and tarsus length) for females
and males, respectively. Diet treatments were monitored and adjusted using the mean condition
of all birds within a group. Treatment condition scores were compared at the time of capture and
when desired condition levels were met using one-way analysis of variance (ANOVA). Post-hoc
analyses were carried out using two sample t-tests assuming equal variances.
Elevated viral shedding occurred through 5 dpi for most birds. Therefore, analysis of
shedding concentration was performed during the first 5 dpi using repeated-measures ANOVA.
In the analysis, samples that did not exhibit detectable virus were assigned a value of zero, and
birds that did not shed virus during the course of the study were included in the comparisons.
Viral titers were transformed using log (base 10).
Shedding duration was calculated for each bird as the last day post inoculation where viral
RNA was detected. In addition, shedding frequency was calculated for each mallard as the
number of positive samples detected from the 4 sampling days between 1-5 dpi. Mallards that
did not shed detectable virus were excluded from both analyses. Shedding duration and
frequency data were not normally distributed (Shapiro-Wilk, p < 0.05). Therefore, overall group

9

 

 

 

comparisons were done using a nonparametric Kruskal-Wallis ANOVA on ranks, and post-hoc
analyses were conducted using Mann-Whitney rank sum test.
Serologic response was compared among treatment groups on 14, 21, and 28 dpi. For
analysis, negative samples (HI titer < 8) were assigned a value of half the minimum detectable
titer as described by Stephenson et al. (2009). All HI titers were transformed using log (base 2).
Analyses of antibody production were conducted using repeated-measures ANOVA. All
repeated-measures post-hoc analyses were done using paired t-tests. The alpha level was set at
0.05 for all analyses, and derived p values correspond with two-tailed tests. Analyses were
performed using PASW 18.0 (PASW, 2010).

10

 

 

 

RESULTS

A total of 81 (43 males; 38 females) juvenile mallards were captured during September 2009.
Influenza A virus antibodies were detected in 51% of birds (17 males; 24 females) using ELISA.
Of the 40 seronegative mallards available for the study, 10 were released because they had
problems acclimating to captivity. Mean condition scores for the three wild treatment groups
were similar at the time of capture (one-way ANOVA; F2,27 = 1.12, p = 0.34) (Figure 1.1). All
(wild and captive-bred) birds (n = 40) adjusted well to captivity and were eating and drinking
normally at the end of the acclimation period. Food manipulation significantly separated wild
treatment group conditions by day 60 (F2,27 = 18.0, p = < 0.001) (Figure 1.1). Mean condition
score for mallards in the poor treatment (-21%) was significantly reduced compared to birds in
the lean treatment (-13%) (two-sample t-test; t = 2.13, p = 0.046) and ad libitum treatment (2%)
(t = 5.79, p = < 0.001). Condition score for lean birds was lower than for mallards in the ad
libitum treatment (t = 3.77, p = 0.001). Wild mallards in the ad libitum treatment surpassed their
predicted mass during the first five days of diet manipulation, and their mean condition remained
elevated (1.0 - 2.4%) for the remainder of the study. Nine of 10 captive-bred mallards gained
mass during diet manipulation (Table 1.1).

11

 

 

 

 

A

B

 

Percent change in body condition

5
0
-5
-10
-15
ad libitum
lean
poor

-20
-25
0

10

20

30

35

40

45

50

55

60

65

70

75

Days post capture

Figure 1.1. Body condition for wild mallard treatment groups throughout the study. Data points
represent mean condition (± 1 standard error), A = start of diet manipulation, B = LPAIV H5N9
inoculation.

12

Table 1.1. Body mass (g) for 10 captive-bred mallards (C1-C10) fed ad libitum from time of purchase through LPAIV H5N9
challenge.
Bird ID

Acclimation period

Diet manipulation

H5N9 challenge

0*

10

20

30

35

40

45

50

55

60

C1

1139

1249

1264

1219

1216

1186

1221

1211

1204

C2

935

1042

1096

1129

1125

1157

1170

1175

C3

872

853

893

927

929

938

933

C4

1367

1383

1357

1329

1314

1282

C5

1055

1283

1347

1316

1331

C6

796

1042

1133

1171

C7

702

849

932

C8

948

1107

C9

1043

C10

857

1

2

65

70

75

1193

%
4.7

1212

1235

1185

%
-0.7

1167

1148

22.8

1156

1144

1137

-1.0

961

947

938

7.6

935

903

916

-2.3

1276

1245

1219

1217

-11.0

1226

1219

1270

4.4

1345

1333

1356

1333

1331

26.2

1347

1355

1358

2.0

1196

1186

1190

1248

1263

1254

57.5

1290

1277

1304

4.0

946

1001

994

1028

1043

983

1016

44.7

1006

978

1020

0.4

1176

1192

1212

1222

1225

1227

1194

1208

27.4

1236

1213

1226

1.5

1126

1183

1225

1166

1188

1206

1223

1212

1205

15.5

1223

1197

1185

-1.7

950

972

998

994

994

999

1008

993

989

15.4

990

986

1004

1.5

* The numbers indicate days after captive-bred mallards were purchased.
1

Change in mass for each bird at the end of diet manipulation (day 60), expressed as percent change from purchase mass (day 0).

2

Change in mass for each bird 15 days post challenge (day 75), expressed as percent change from inoculation mass (day 60).

13

 

 

Susceptibility to LPAIV (H5N9) infection:
A total of 36 out of 40 (90%) study birds shed detectable virus (≥ 1.0 log10 PFU/mL). All
four birds without detectable viral shedding were wild mallards in the poor condition treatment.
Body condition scores for these birds at the time of LPAIV challenge were -18%, -21%, -24%
and -37%. Thirty-five of 36 (97%) infected birds shed virus by the second day post inoculation.
One captive-bred mallard delayed viral shedding until five days post inoculation. After LPAIV
challenge, no birds exhibited clinical signs of disease.

Concentration of viral shedding:
The mean virus titer (log10 PFU/mL) for all treatment groups peaked at 2 dpi (wild ad libitum
= 3.8; lean = 2.20; poor = 1.33; captive-bred ad libitum = 4.03) and the bulk of shedding
continued through 5 dpi (Figure 1.2). Virus excretion in wild mallards during the first 5 dpi
among treatment groups varied significantly (repeated-measures ANOVA; F2, 27 = 7.24, p =
0.003). In general, higher levels of viral shedding were observed in treatment groups with higher
relative condition scores (i.e. greater food availability) (Table 1.2, Figure 1.2). Wild mallards in
the poor treatment shed less virus than birds fed ad libitum (paired t-test; t = 2.39, p = 0.001) but
not significantly less than lean mallards (t = 1.31, p = 0.065). We were unable to detect a
difference in shedding concentration between ducks in the ad libitum and lean treatments (t =
1.34, p = 0.097). Wild and captive-bred birds fed ad libitum shed similar virus concentrations
(F1, 18 = 0.22, p = 0.64).
Concentration of viral shedding was highly variable within treatment groups during the first
five days of infection. Daily mean titers and standard errors are presented for groups in Table
1.2. Inspection of individual shedding patterns found a total of 14 mallards that excreted virus at
14

 

 

 

 

5

Virus titer, log10 PFU/mL

4

3

2

1

0
0

1

2

3

4

5

6

7

Days after H5N9 inoculation
captive-bred
wild ad libitum

wild lean
wild poor

Figure 1.2. Virus shedding concentration results for captive-bred and wild mallard treatment
groups. Data points represent mean titer (log10 PFU/mL) for each day sampled.

15

 

 

 

Table 1.2. Daily mean (± 1 standard error) virus titers (log10 PFU/mL) for mallards inoculated
with LPAIV H5N9. Mean titer includes birds without detectable viral RNA, whereas minimum
and maximum titers are reported for the number of birds (n) with detectable viral RNA on the
given day.
Treatment group

Days post inoculation
1

2

3

5

7

Captive-bred ad libitum

2.2 ± 0.54

4.0 ± 0.73

3.9 ± 0.87

3.8 ± 0.59

1.2 ± 0.41

min-max (n)

1.6-4.6 (7)

2.8-5.8 (8)

4.5-6.7 (7)

1.1-5.8 (9)

1.7-3.2 (5)

Wild ad libitum

3.3 ± 0.56

3.8 ± 0.52

3.3 ± 0.66

2.1 ± 0.72

1.0 ± 0.45

min-max (n)

1.3-5.4 (9)

1.7-6.2 (10)

1.9-5.7 (8)

3.4-4.9 (5)

1.8-3.3 (4)

Wild lean

2.0 ± 0.42

2.2 ± 0.50

1.9 ± 0.59

2.1 ± 0.72

0.2 ± 0.24

min-max (n)

0.9-4.3 (8)

1.8-5.7 (8)

1.1-6.1 (7)

1.2-6.6 (6)

2.4 (1)

Wild poor

0.4 ± 0.26

1.3 ± 0.69

0.9 ± 0.57

0.3 ± 0.31

-

min-max (n)

1.6-2.3 (2)

2.0-6.8 (4)

1.8-5.6 (3)

3.1 (1)

-

16

 

 

high titer (≥ 5.0 log10 PFU/mL). The majority of these birds were wild and captive-bred mallards
fed ad libitum (wild, n = 5; captive-bred, n = 7), whereas only two birds came from reduced
condition treatments (lean, n = 1; poor, n = 1) (Figure 1.3). However, peak viral titers in the
above mentioned poor bird (6.8 log10 PFU/mL) and lean bird (6.6 log10 PFU/mL) were the
highest observed across all wild mallards (Table 1.2).

Duration of viral shedding:
Duration of infection among all study birds was large (1-20 dpi). Among wild mallards,
mean duration (days) of shedding was largest in groups with higher condition scores (wild ad
libitum = 6.4; lean = 5.4; poor = 3.3) (Figure 1.4). Intermittent shedding beyond 5 dpi was more
common in wild mallards fed ad libitum (n = 5), whereas only two birds in reduced condition
treatments (lean, n = 1; poor, n = 1) shed virus past 5 dpi. Despite these relationships, we were
unable to detect a significant difference in shedding duration among wild mallards (KruskalWallis ANOVA; H = 2.86, 2 d.f., p = 0.24). In addition, mean duration of shedding in captivebred birds (7.8 days) was similar to wild birds fed ad libitum (H = 0.52, 1 d.f., p = 0.47).
The number of positive samples through 5 dpi differed among wild mallard groups (H =
6.20, 2 d.f., p = 0.045) (Figure 1.4). Mean number of positive samples in poor mallards (1.7) was
significantly less than for ad libitum birds (3.2) (Mann-Whitney rank sum; U = 8.0, p = 0.016),
but not less than for lean mallards (2.9) (U = 13.5, p = 0.071). No differences were found
between wild ad libitum and lean treatments (U = 45.0, p = 0.71), or between wild and captivebred birds fed ad libitum (H = 0.007, 1 d.f., p = 0.93).

17

 

Virus titer, log10 PFU/mL

 

 

A

7

B

7

6

6

5

5

4

4

3

3

2

2

1

1

0

0
0

Virus titer, log10 PFU/mL

 

1

2

3

4

5

6

C

7

0

7

1

2

3

6

7

2

1

5

3

2

4

4

3

7

5

4

6

6

5

5

D

7

6

4

1

0

0
0

1

2

3

4

5

6

7

Days after H5N9 inoculation

0

1

2

3

Days after H5N9 inoculation

Figure 1.3. Virus shedding concentration (log10 PFU/mL) results for all mallards with detectable
viral RNA using matrix gene RTT-PCR. A: captive-bred (n = 10), B: wild ad libitum (n = 10),
C: wild lean (n = 10), D: wild poor (n = 6).

18

 

 

A

10
Duration of virus shedding (days)

 

8
6
4
2
0
captive-bred wild ad libitum

wild poor

B

4

Number of positive samples

wild lean

*
3

*

2

1

0
captive-bred wild ad libitum

wild lean

wild poor

Treatment group
Figure 1.4. Duration of viral shedding (A) and number of positive samples (B) for captive-bred
and wild mallard treatment groups. Bars represent means ± 1 standard error, asterisks identify
groups significantly different from one another (Mann-Whitney rank sum, alpha = 0.05). The
number of positive samples was calculated from the 4 sampling days between 1-5 dpi.
19

 

 

Serologic response:
After LPAIV H5N9 challenge, 33 out of 40 (82.5%) study birds tested seropositive using
ELISA, whereas 32 out of 40 (80%) had detectable levels of H5-specific antibodies according to
HI tests. Among wild mallards, mean HI titers peaked at 14 dpi for ad libitum and lean
treatments (3.4, 3.2 log2, respectively), while HI titers in poor mallards were highest at 21 dpi
(3.3 log2) (Table 1.3). Overall antibody production at 14, 21, and 28 dpi was similar across wild
mallard treatments (repeated-measures ANOVA; F2, 23 = 0.29, p = 0.75). Mean HI titer for
captive-bred birds (4.6 log2) peaked at 14 dpi, and did not differ from wild mallards fed ad
libitum (F1, 17 = 2.47, p = 0.13).

20

 

Table 1.3. Serological status of mallards before and after LPAIV H5N9 challenge using ELISA and HI tests. Mean (± 1 standard
error) test scores include all birds in each treatment.
Treatment group

ELISA
B.I.

1

3

H5N9 HI

2

14 *

21

28

B.I.

14

21

28

Captive-bred ad libitum
Seropositive (n)

0.93 ± 0.03
(0)

0.36 ± 0.07
(7)

0.42 ± 0.09
(6)

0.42 ± 0.09
(6)

<3
(0)

4.6 ± 0.70
(7)

4.1 ± 0.55
(7)

3.9 ± 0.53
(7)

Wild ad libitum
Seropositive (n)

0.87 ± 0.05
(0)

0.30 ± 0.06
(9)

0.36 ± 0.07
(6)

0.37 ± 0.07
(6)

<3
(0)

3.4 ± 0.43
(7)

3.2 ± 0.29
(8)

2.8 ± 0.20
(7)

Wild lean
Seropositive (n)

0.86 ± 0.04
(0)

0.21 ± 0.03
(10)

0.33 ± 0.08
(8)

0.36 ± 0.09
(8)

<3
(0)

3.2 ± 0.25
(8)

3.0 ± 0.26
(8)

2.8 ± 0.25
(5)

Wild poor
Seropositive (n)

0.84 ± 0.03
(0)

0.36 ± 0.10
(7)

0.47 ± 0.09
(6)

0.47 ± 0.08
(6)

<3
(0)

3.1 ± 0.38
(6)

3.3 ± 0.58
(5)

3.1 ± 0.31
(6)

* The numbers indicate days after LPAIV H5N9 inoculation.
1

The ELISA scores represent the signal to noise (S/N) ratio where values ≤ 0.50 are considered seropositive.

2

The hemagglutination inhibition values represent the mean titer (log2) of sera samples.

3

Before LPAIV H5N9 inoculation.

21

DISCUSSION

Our study demonstrates body condition significantly influences susceptibility to infection and
viral shedding patterns in wild-caught juvenile mallards challenged with LPAIV H5N9.
However, our findings were opposite of our original predictions based on the conditiondependent hypothesis in which birds in poor condition would experience reduced immune
function and increased susceptibility to infection (Flint and Franson 2009). Here we find birds in
good condition were more susceptible to LPAIV infection, and shed significantly higher virus
titers for a longer duration compared to birds in poor condition. Four mallards in the poor
condition treatment were resistant to infection, whereas all remaining mallards shed detectable
virus. A clear trend was observed among wild birds showing a positive relationship between
body condition and host competence.
Until recently, it has been assumed LPAIV infections remain asymptomatic in wild
waterfowl with little or no effects on life-history parameters. Then van Gils et al. (2007)
determined LPAIV infection decreased migratory performance in Bewick’s swans, and LatorreMargalef et al. (2009b) found mallards infected with IAV were leaner than uninfected
conspecifics. The latter study has generated ongoing debate on whether IAV infection influences
body condition of migrating waterfowl, or vice versa (Flint and Franson 2009, Latorre-Margalef
et al. 2009a). Both sides acknowledge the possibility that birds may become immunosuppressed
during migration due to reduced energy stores, and therefore suggest further studies are needed
to conclusively discriminate between these two hypotheses. To our knowledge, our research
represents the first experimental study that has evaluated the condition-dependent hypothesis
using LPAIV and waterfowl as an experimental model. We have provided evidence that body
22

 

 

condition influences IAV infection in wild juvenile mallards, however, the mechanisms
responsible for our findings remain unclear and contrary to those suggested by the conditiondependent hypothesis (Flint and Franson 2009).
Current research examining host nutrition and susceptibility to infectious disease provides
overwhelming support for our original hypotheses. In general, these studies found malnutrition
increases susceptibility and severity of infection with most microbial agents (Scrimshaw et al.
1968, Louria 2007). In the case of IAV infection, deficiencies in vitamins A and C, selenium,
and protein have increased susceptibility and burden of disease (Stephensen et al. 1993, Beck
2001, Li et al. 2006, Louria 2007, Ritz et al. 2008). These findings were attributed to reduced
immune function caused by limitations in one or several essential nutrients, vitamins, and/or
dietary protein (Pollett et al. 1979, Ritz et al. 2008). If we assume mallards in poor condition
were malnourished, as indicated by significant decrease in breast condition score (Figure 1.5),
then our findings contradict the well established trend described above. Therefore, we propose
the relationship between body condition and LPAIV infection in waterfowl is more complex than
previously thought.
Review of previous research, however, would suggest three additional factors that may
influence the observed relationship: (1) duration of food restriction, (2) depletion of
subcutaneous fat reserves, and (3) changes in intestinal composition due to reduced food intake.
Past studies have shown certain conditions of malnutrition increase hosts resistance to viral
infection (Sprunt 1942, Flanigan and Sprunt 1956). Sprunt and Flanigan (1956) found mice and
chickens fed reduced protein diets exhibited a cyclic pattern of susceptibility relative to those fed
high protein diets. Protein restriction increased susceptibility for the first two weeks, then
decreased susceptibility from three to six weeks. Beyond seven weeks, mice and chickens

23

 

 

 

became more susceptible to viral infection. Animals were less susceptible to infection once they
depleted subcutaneous fat reserves, and initiated catabolism of available protein reserves. Diets
high in protein have been correlated with increased resistance to viral infection (Pollett et al.
1979). Our findings support those of Sprunt and Flanigan (1956), as mallards in poor condition
were less susceptible to infection after four weeks of food restriction. Moreover, birds in the poor
condition treatment had significantly reduced keel scores when challenged with LPAIV (Figure
1.5); thereby suggesting these mallards transitioned to catabolism of protein reserves.
Changes in intestinal composition from reduced food availability may be responsible for
decreased susceptibility and viral titers among mallards in reduced condition. Food deprivation
has been shown to reduce the relative amount of mucin glycoprotein in the intestinal tract.
Smirnov et al. (2004) examined intestinal mucin in chickens fasted for 72 hours, and found acute
food deprivation decreased mucin thickness throughout the intestinal tract. In rats deprived 50%
of their daily intake for five weeks, the concentration of intestinal mucin was significantly
reduced compared to control animals fed normally (Sherman et al. 1985). In waterfowl, LPAI
viruses preferentially bind to sialic acid (SA) receptors which occupy terminal positions on
mucin glycoproteins within the intestinal tract (Webster et al. 1978, Kida et al. 1980). Therefore,
it seems plausible that decreased abundance of mucin may reduce SA expression and inhibit viral
attachment and propagation. Further investigations concerning reduced food availability,
intestinal mucin, and SA expression in waterfowl are needed to evaluate this hypothesized
interaction.

24

 

 

 

 

3.0

Breast condition score (0- 3)

*
2.5

*

2.0

1.5

1.0
ad libitum

lean

poor

Wild treatment group
capture condition

LPAI inoculation condition

Figure 1.5. Breast condition score for wild mallard treatment groups at capture vs. LPAIV H5N9
challenge. Bars represent mean condition score ± 1 standard error (n = 10), asterisks represent
significant within group differences (paired t-test, alpha = 0.05).

25

 

 

We did not detect a significant difference in susceptibility to infection and viral shedding
patterns between wild-caught and captive-bred mallards fed ad libitum. Therefore, we support
the use of captive-bred birds in LPAIV experimental models where birds are fed ad libitum. We
are unable to conclude whether captive-bred birds exhibit similar host competency during
periods of food restriction. Mallards bred and raised in captivity do not encounter prolonged food
shortages, and may respond differently to food deprivation. Additional research is needed to
validate the use of captive-bred mallards in future LPAIV experimental models incorporating
food restriction.
Our serological findings indicate reduced body condition does not affect mallard antibody
production in response to LPAIV H5N9 challenge. These findings do not support our initial
prediction in which resource limited birds experience decreased antibody production
(Hangalapura et al. 2005, Hanssen et al. 2005). It is well understood that maintaining and using
the immune system is energetically costly (Klasing et al. 2004). During periods of limited food
access, Buehler et al. (2009) found migrating shorebirds suppress more costly acute-phase
immune responses (phagocytosis, fever, inflammation) in order to maintain a baseline level of
immune function. Thus, mallards fed restricted diets may have down regulated some aspects of
immune function and retained the ability to produce specific antibodies. Alternatively, mallards
in reduced condition may have adequate resources to enable production of a low-level humoral
response typical of LPAIV infections (Kida et al. 1980, Jourdain et al. 2010).
If the observed influence of body condition on LPAIV infection is at play in nature,
prolonged periods of reduced food availability may inhibit virus transmission. Reduced
transmission of LPAI viruses may decrease the development of highly pathogenic avian
influenza virus (HPAIV). In the past two decades, HPAIV outbreaks have occurred frequently in

26

 

 

 

domestic poultry after introduction of H5 and H7 LPAI viruses (Perdue and Swayne 2005).
These outbreaks resulted in high morbidity, mortality, and severe economic losses (Olsen et al.
2006). Understanding the ecology of LPAIV in waterfowl hosts remains a critical task.
A pattern between host condition and IAV infection has been observed in recent studies
(Hanson et al. 2008, Latorre-Margalef et al. 2009b), and our study provides novel insight
towards explaining these relationships. Hanson et al. (2008) found migrating ruddy turnstones
(Arenaria interpres) with elevated stopover masses exhibited higher IAV prevalence rates. The
researchers attributed their findings to on-site IAV amplification, as lighter birds were presumed
to have recently arrived at the stopover site. However, according to our findings, such
differences could be attributed to decreased IAV susceptibility among birds in poor condition.
On the other hand, Latorre-Margalef et al. (2009b) observed a significant decrease in body mass
in mallards infected with IAV compared to uninfected ducks (mean difference of 20 g).
Although infected birds were lighter, we point out a 20 g reduction in mass would not indicate a
mallard was in poor condition. Mean body mass loss among mallards in our poor treatment
exceeded 200 g. Therefore, the relationship observed by Latorre-Margalef et al. (2009b) does not
refute our findings.
In summary, we have taken a large step in showing how host body condition may play a
significant role in the epidemiology of LPAIV infection in mallards, and presumable other
waterfowl species. We provide evidence that (1) mallards in poor condition are less susceptible
to infection compared to birds in good condition, (2) concentration and duration of viral
shedding are positively correlated with host condition, and (3) body condition does not affect
mallard specific antibody response after LPAIV challenge. The study also suggests that captivebred mallards may replace wild mallards in future experimental models where birds are fed ad

27

 

 

 

libitum. The precise mechanisms of decreased host competence among mallards in reduced
condition remains unknown. If susceptibility follows a cyclic pattern, as indicated by Sprunt and
Flanigan (1956), then birds would first encounter a period of enhanced virus transmission in
response to food deprivation. Additional field and laboratory studies under varying durations of
food restriction are encouraged to clarify this proposed relationship. Furthermore, these studies
should incorporate more comprehensive analyses of immune function and host nutritional
reserves. Such data would help explain how body condition influences waterfowl host
competency during LPAIV infection, and improve future LPAIV transmission models.

28

 

 

 

CHAPTER 2
ROLE OF HOMO- AND HETEROSUBTYPIC IMMUNITY
ON REINFECTION OF MALLARD DUCKS WITH INFLUENZA A VIRUS

INTRODUCTION

In Europe and North America, the prevalence of influenza A virus (IAV) isolation in
waterfowl peaks during autumn migration when juvenile ducks congregate before migrating
south (Hinshaw et al. 1985, Olsen et al. 2006). Low pathogenic avian influenza virus (LPAIV)
prevalence can reach 60% in these birds, and generally includes a large diversity of virus
subtypes (Krauss et al. 2004, Wallensten et al. 2007). As a result, migrating waterfowl are
frequently exposed to the same (homosubtypic) or different (heterosubtypic) IAV subtypes
(Sharp et al. 1997). Recent studies suggest recurring IAV exposures induce transient immunity,
thereby providing an explanation for the low IAV infection prevalence (< 1%) among wintering
waterfowl (Latorre-Margalef et al. 2009b, Jourdain et al. 2010). To better understand
transmission and seasonal maintenance of IAV by migrating waterfowl, we must improve our
understanding of naturally occurring homo- and heterosubtypic immunity.
Previous experimental studies have demonstrated prior LPAIV infection protects against
homosubtypic (Kida et al. 1980) and heterosubtypic (Jourdain et al. 2010) LPAIV reinfections.
Kida et al. (1980) found that after primary LPAIV H7N2 inoculation, ducks resisted
homosubtypic reinfection for 28 days, but were susceptible beyond 46 days post inoculation
(dpi). Jourdain et al. (2010) demonstrated LPAIV H5N7 infection in juvenile mallards (Anas
platyrhynchos) reduced viral RNA excretion during homosubtypic reinfection, and protected
some birds against heterosubtypic LPAIV H5N2 reinfection. These studies demonstrate
29

 

 

 

waterfowl can mount immunological responses against recurring LPAIV exposures; however,
more studies are needed to clarify homo- vs. heterosubtypic interactions and examine duration
between viral challenges.
Understanding the role of homo- and heterosubtypic immunity is essential for several
reasons. Immunity induced by prior IAV infection has been shown to reduce susceptibility and
viral shedding in waterfowl during subsequent infections (Kida et al. 1980, Fereidouni et al.
2009, Jourdain et al. 2010). As a result, maintenance and spread of LPAIVs may be greatly
reduced if herd immunity develops in the population (Latorre-Margalef et al. 2009b, Jourdain et
al. 2010). However, previous LPAIV infection may enhance transmission of highly pathogenic
avian influenza virus (HPAIV) by waterfowl. Recent findings indicate LPAIV infection reduces
disease burden during HPAI reinfection, thereby allowing birds to migrate while shedding
intermediate concentrations of HPAIVs (Fereidouni et al. 2009). In addition, acquired immunity
may reduce the occurrence of simultaneous IAV infections, which play a role in producing novel
virus subtypes through genetic reassortment (Hinshaw et al. 1980, Sharp et al. 1997).
In this study, we tested whether LPAIV H5N9 infection in juvenile mallards 1) provides
long-term homosubtypic immunity to the same LPAIV strain and 2) provides long-term
heterosubtypic immunity to a different LPAIV H7N3 strain. Long-term immunity was defined as
seven weeks between primary and secondary LPAIV inoculation. We hypothesized mallards
would be resistant to reinfection with the homosubtypic H5N9 strain, but susceptible to
heterosubtypic H7N3 reinfection.

30

 

 

 

 

MATERIALS AND METHODS

Animals:
Wild-caught juvenile mallards (n = 20) were retained from the previous study described in
Chapter 1 (hereafter “condition study”). During the condition study, birds were placed on diet
treatments to manipulate their body condition before inoculation with LPAIV H5N9. To
minimize variation in starting condition for the present study, birds with similar body condition
scores were selected. In addition, only birds with detectable levels of H5-specific antibodies
were included in this experiment. Mallards were housed at the Michigan State University
Research Containment Facility using protocols described in Chapter 1 (see Housing).

Experimental design:
Following completion of the condition study (starting at 28 dpi), all birds were maintained
with ad libitum access to water and food. Mallards were randomly divided into two treatment
groups (homosubtypic, n = 10; heterosubtypic, n = 10), each housed in a separate biosafety level
2 room. One day prior to experimental reinoculation (at 48 dpi), cloacal and oral swabs were
collected from all mallards to confirm absence of viral shedding using RRT-PCR as described in
Chapter 1 (see Matrix Gene RRT-PCR). In addition, blood serum was collected to test for IAV
antibodies using ELISA (Chapter 1, see IDEXX FlockChek* ELISA).
At 49 dpi the homosubtypic treatment group was oropharyngeally reinoculated with the same
LPAIV H5N9 virus (A/Northern pintail/California/44221-761/2006) using 1.5 mL of 10

6

PFU/mL. The heterosubtypic treatment group was oropharyngeally reinoculated with a different
6

LPAIV H7N3 virus (A/Northern pintail/Nevada/44252-242/2006) using 1.5 mL of 10 PFU/mL.
31

 

 

Both LPAIV isolates were obtained from USGS National Wildlife Health Center, Wisconsin,
USA. Virus was propagated and titrated as described in Chapter 1 (see Virus). Following
reinoculation, cloacal and oral swabs were collected daily for the first 3 days and then on days 5,
7, 9, 12, and 14 post reinoculation (dpr). Mallards in the homosubtypic treatment were always
sampled before heterosubtypic birds to prevent spillover of the H7N3 LPAIV subtype. Swabs
samples were pooled together for individual birds and tested for the presence of viral RNA.
Blood serum was collected on 7 and 14 dpr for serologic testing. Swab and sera samples were
collected and stored using protocols described in Chapter 1. All mallards were euthanized at 14
dpr using CO2 asphyxiation, followed by cervical dislocation.

32

 

 

 

RESULTS

One day before LPAIV reinoculation (48 dpi), swab samples from all mallards (n = 20)
tested negative for viral RNA using RRT-PCR, indicating birds were not shedding IAV
immediately prior to experimental reinoculation. Serology testing using ELISA found 14 of 20
(70%) mallards remained seropositive 48 days after primary LPAIV H5N9 infection (Table 2.1).
The distribution of seropositive birds was equal among the treatment groups (homosubtypic, n =
7; heterosubtypic, n = 7).

Homosubtypic LPAIV (H5N9) reinoculation:
One of 10 mallards in the homosubtypic treatment shed detectable viral RNA (≥ 1.0 log10
PFU/mL). The positive mallard tested seropositive by ELISA one day prior to reinoculation.
Viral shedding peaked at 2 dpr (5.6 log10 PFU/mL), and viral RNA was detected in all pooled
swabs samples through 9 dpr (Figure 2.1). All birds tested positive for IAV antibodies at 7 and
14 dpr (Table 2.1).

Heterosubtypic LPAIV (H7N3) reinoculation:
All mallards (n = 10) in the heterosubtypic treatment shed detectable viral RNA on the first
date post reinoculation. Mean virus titer peaked at 2 dpr (5.9 log10 PFU/mL) and the majority of
viral shedding occurred through 5 dpr (Figure 2.1). All birds excreted virus concentrations that
exceeded 5.0 log10 PFU/mL, and three birds exhibited viral titers greater than 7.0 log10 PFU/mL.
The duration of infection among mallards ranged from 5-14 dpr, with mean duration being 9.2

33

 

 

 

days. Only one mallard tested positive for viral RNA at 14 dpr. Influenza A virus antibodies
were detected in all mallards at 7 and 14 dpr using ELISA (Table 2.1).

34

 

 

 

 

Table 2.1. Serological status of mallards before and after homosubtypic LPAIV H5N9 and
heterosubtypic LPAIV H7N3 reinoculation using ELISA. Mean (± 1 standard error) test
scores include all birds in each treatment.
1

Treatment group

ELISA

7*

14

Homosubtypic (LPAIV H5N9)
Seropositive (n)

B.I.
0.42 ± 0.07
(7)

0.11 ± 0.02
(10)

0.17 ± 0.04
(10)

Heterosubtypic (LPAIV H7N3)
Seropositive (n)

0.39 ± 0.07
(7)

0.10 ± 0.01
(10)

0.15 ± 0.02
(10)

2

* The numbers indicate days post reinoculation.
1

ELISA scores represent signal to noise ratio (S/N), values ≤ 0.50 are considered seropositive.

2

One day before reinoculation (48 days after primary LPAIV H5N9 challenge).

35

 

 

8

 

A

Virus titer, log10 PFU/mL

7
6
5
4
3
2
1
0
0

2

4

6

8

8

10

12

14

8

10

12

14

B

Virus titer, log10 PFU/mL

7
6
5
4
3
2
1
0
0

2

4

6

Days after reinoculation

Figure 2.1. Virus shedding concentration (log10 PFU/mL) for all mallards with detectable viral
RNA after reinoculation with (A) homosubtypic LPAIV H5N9 (n = 1), and (B) heterosubtypic
LPAIV H7N3 (n=10). Birds were challenged 7 weeks after primary LPAIV H5N9 inoculation.
36

 

 

DISCUSSION

During autumn migration, juvenile mallards congregate with waterfowl from other areas and
are frequently exposed to homo- and heterosubtypic IAVs (Sharp et al. 1997, Krauss et al. 2004,
Wallensten et al. 2007). Our study was designed to test long-term (i.e. 7 weeks after primary
infection) protective immunity throughout the migratory season. We provide evidence that
juvenile mallards can mount an immune response protective against reinfection with the same
LPAI H5N9 virus. Alternatively, we demonstrate mallards were susceptible to infection with a
heterosubtypic LPAI H7N3 virus.
Previous studies successively inoculating juvenile mallards with LPAIV strains have found
birds resistant to homosubtypic reinfection for up to 28 days (Kida et al. 1980, Jourdain et al.
2010). Our findings support these observations, as 9 of 10 mallards resisted reinfection with the
same LPAIV. We did observe one mallard whose positive serological status did not protect it
from homosubtypic reinfection. This observation is in agreement with Kida et al. (1980), who
found Pekin ducks became susceptible to homosubtypic reinfection after 46 days. However, our
findings suggest a greater proportion of wild juvenile mallards resist homosubtypic reinfection
beyond 49 days.
Jourdain et al. (2010) found juvenile mallards are able to mount short-term (i.e. 21 days after
primary infection) immunological responses protective against heterosubtypic LPAIV
reinfections. Our study detected no strong evidence of heterosubtypic immunity, as all mallards
in our heterosubtypic treatment shed detectable viral RNA. We did not have a control group of
birds (not previously exposed to H5N9) to test the response to LPAIV H7N3 alone. However,
the high concentration of virus shedding (≥ 5.0 log10 PFU/mL) observed in all mallards

37

 

 

 

reinfected with heterosubtypic LPAIV H7N3 suggests little or no cross-reactive immunity. We
suggest immunological responses protective against heterosubtypic LPAIV infections may be
short-term (less than 7 weeks).
Current research suggests the transmission dynamics of LPAIVs by migrating waterfowl are
significantly influenced by homo- and heterosubtypic immunity (Kida et al. 1980, Sharp et al.
1997, Jourdain et al. 2010). Our findings contribute to available information in describing the
interaction of more long-term immune responses. In summary, we provide evidence that (1)
juvenile mallards can mount long-term protective immunological responses against reinfection
with the same LPAI H5N9 virus, and (2) are susceptible to heterosubtypic LPAIV H7N3
reinfection 49 days after primary inoculation. Future studies should incorporate new virus
subtypes, different challenge timings, and analysis of immune function to improve our
understanding of homo- and heterosubtypic immunity.

38

 

 

 

REFERENCES

39

 

 

 

REFERENCES

Beck, M. A. 2001. Antioxidants and viral infections: host immune response and viral
pathogenicity. J Am Coll Nutr 20:384S-388S; discussion 396S-397S.
Brown, J. D., D. E. Stallknecht, J. R. Beck, D. L. Suarez, and D. E. Swayne. 2006. Susceptibility
of North American ducks and gulls to H5N1 highly pathogenic avian influenza viruses.
Emerging Infectious Diseases 12:1663-1670.
Buehler, D. M., F. Encinas-Viso, M. Petit, F. Vezina, B. I. Tieleman, and T. Piersma. 2009.
Limited Access to Food and Physiological Trade-Offs in a Long-Distance Migrant
Shorebird. II. Constitutive Immune Function and the Acute-Phase Response.
Physiological and Biochemical Zoology 82:561-571.
Carney, S. M. (1992) U. S. Department of the Interior, U.S. Fish and Wildlife Service,
Washington, D.C. 144pp.
Cooley, A. J., H. Vancampen, M. S. Philpott, B. C. Easterday, and V. S. Hinshaw. 1989.
Pathological Lesions in the Lungs of Ducks Infected with Influenza-a Viruses. Veterinary
Pathology 26:1-5.
Devries, J. H., R. W. Brook, D. W. Howerter, and M. G. Anderson. 2008. Effects of spring body
condition and age on reproduction in Mallards (Anas platyrhynchos). Auk 125:618-628.
Fereidouni, S. R., E. Starick, M. Beer, H. Wilking, D. Kalthoff, C. Grund, R. Hauslaigner, A.
Breithaupt, E. Lange, and T. C. Harder. 2009. Highly Pathogenic Avian Influenza Virus
Infection of Mallards with Homo- and Heterosubtypic Immunity Induced by Low
Pathogenic Avian Influenza Viruses. Plos One 4:-.
Flanigan, C. C., and D. H. Sprunt. 1956. The effect of malnutrition on the susceptibility of the
host to viral infection. J Exp Med 104:687-706.
Flint, P. L., and J. C. Franson. 2009. Does influenza A affect body condition of wild mallard
ducks, or vice versa? Proceedings of the Royal Society B-Biological Sciences 276:23452346.
Gregory, N. G., and J. K. Robins. 1998. A body condition scoring system for layer hens. New
Zealand Journal of Agricultural Research 41:555-559.
Halvorson, D. A., C. J. Kelleher, and D. A. Senne. 1985. Epizootiology of Avian Influenza Effect of Season on Incidence in Sentinel Ducks and Domestic Turkeys in Minnesota.
Applied and Environmental Microbiology 49:914-919.

40

 

 

 

Hangalapura, B. N., M. G. B. Nieuwland, G. D. Reilingh, J. Buyse, H. Van Den Brand, B.
Kemp, and H. K. Parmentier. 2005. Severe feed restriction enhances innate immunity but
suppresses cellular immunity in chicken lines divergently selected for antibody
responses. Poultry Science 84:1520-1529.
Hanson, B. A., M. P. Luttrell, V. H. Goekjian, L. Niles, D. E. Swayne, D. A. Senne, and D. E.
Stallknecht. 2008. Is the occurrence of avian influenza virus in Charadriiformes species
and location dependent? Journal of Wildlife Diseases 44:351-361.
Hanssen, S. A., D. Hasselquist, I. Folstad, and K. E. Erikstad. 2005. Cost of reproduction in a
long-lived bird: incubation effort reduces immune function and future reproduction.
Proceedings of the Royal Society B-Biological Sciences 272:1039-1046.
Hinshaw, V. S., W. J. Bean, R. G. Webster, and G. Sriram. 1980. Genetic reassortment of
influenza A viruses in the intestinal tract of ducks. Virology 102:412-419.
Hinshaw, V. S., J. M. Wood, R. G. Webster, R. Deibel, and B. Turner. 1985. Circulation of
Influenza-Viruses and Paramyxoviruses in Waterfowl Originating from 2 Different Areas
of North-America. Bulletin of the World Health Organization 63:711-719.
Homme, P. J., and Easterda.Bc. 1970. Avian Influenza Virus Infections .4. Response of
Pheasants, Ducks, and Geese to Influenza a/Turkey/Wisconsin/1966 Virus. Avian
Diseases 14:285-&.
Jonassen, C. M., and K. Handeland. 2007. Avian influenza virus screening in wild waterfowl in
Norway, 2005. Avian Diseases 51:425-428.
Jourdain, E., G. Gunnarsson, J. Wahlgren, N. Latorre-Margalef, C. Brojer, S. Sahlin, L.
Svensson, J. Waldenstrom, A. Lundkvist, and B. Olsen. 2010. Influenza Virus in a
Natural Host, the Mallard: Experimental Infection Data. Plos One 5:-.
Keawcharoen, J., D. van Riel, G. van Amerongen, T. Bestebroer, W. E. Beyer, R. van Lavieren,
A. D. M. E. Osterhaus, R. A. M. Fouchier, and T. Kuiken. 2008. Wild ducks as longdistance vectors of highly pathogenic avian influenza virus (H5NI). Emerging Infectious
Diseases 14:600-607.
Kida, H., R. Yanagawa, and Y. Matsuoka. 1980. Duck Influenza Lacking Evidence of Disease
Signs and Immune-Response. Infection and Immunity 30:547-553.
Klasing, K. C., B. D. Humphrey, A. J. Mireles, and E. A. Koutsos. 2004. What are the costs of
immunity? Poultry Science 83:444-444.
Krauss, S., D. Walker, S. P. Pryor, L. Niles, C. H. Li, V. S. Hinshaw, and R. G. Webster. 2004.
Influenza A viruses of migrating wild aquatic birds in North America. Vector-Borne and
Zoonotic Diseases 4:177-189.

41

 

 

 

Latorre-Margalef, N., G. Gunnarsson, V. J. Munster, R. A. M. Fouchier, A. D. M. E. Osterhaus,
J. Elmberg, B. Olsen, A. Wallensten, T. Fransson, L. Brudin, and J. Waldenstrom. 2009a.
Does influenza A affect body condition of wild mallard ducks, or vice versa? A reply to
Flint and Franson. Proceedings of the Royal Society B-Biological Sciences 276:23472349.
Latorre-Margalef, N., G. Gunnarsson, V. J. Munster, R. A. M. Fouchier, A. D. M. E. Osterhaus,
J. Elmberg, B. Olsen, A. Wallensten, P. D. Haemig, T. Fransson, L. Brudin, and J.
Waldenstrom. 2009b. Effects of influenza A virus infection on migrating mallard ducks.
Proceedings of the Royal Society B-Biological Sciences 276:1029-1036.
Lee, C. W., and D. L. Suarez. 2004. Application of real-time RT-PCR for the quantitation and
competitive replication study of H5 and H7 subtype avian influenza virus. Journal of
Virological Methods 119:151-158.
Li, W., N. Maeda, and M. A. Beck. 2006. Vitamin C deficiency increases the lung pathology of
influenza virus-infected gulo-/- mice. J Nutr 136:2611-2616.
Louria, D. B. 2007. Undernutrition can affect the invading microorganism. Clin Infect Dis
45:470-474.
Mauser, D. M., and J. G. Mensik. 1992. A Portable Trap for Ducks. Wildlife Society Bulletin
20:299-302.
Moon, J. A., D. A. Haukos, and L. M. Smith. 2007. Declining body condition of northern pintails
wintering in the Playa Lakes Region. Journal of Wildlife Management 71:218-221.
Munster, V. J., C. Baas, P. Lexmond, J. Waldenstrom, A. Wallensten, T. Fransson, G. F.
Rimmelzwaan, W. E. P. Beyer, M. Schutten, B. Olsen, A. D. M. E. Osterhaus, and R. A.
M. Fouchier. 2007. Spatial, temporal, and species variation in prevalence of influenza A
viruses in wild migratory birds. Plos Pathogens 3:630-638.
Olsen, B., V. J. Munster, A. Wallensten, J. Waldenstrom, A. D. Osterhaus, and R. A. Fouchier.
2006. Global patterns of influenza a virus in wild birds. Science 312:384-388.
Owen, J. C., and F. R. Moore. 2008. Relationship between energetic condition and indicators of
immune function in thrushes during spring migration. Canadian Journal of ZoologyRevue Canadienne De Zoologie 86:638-647.
Pawlina, I. M., D. A. Boag, and F. E. Robinson. 1993. Population-Structure and Changes in
Body-Mass and Composition of Mallards (Anas-Platyrhynchos) Wintering in Edmonton,
Alberta. Canadian Journal of Zoology-Revue Canadienne De Zoologie 71:2275-2281.
Perdue, M. L., and D. E. Swayne. 2005. Public health risk from avian influenza viruses. Avian
Diseases 49:317-327.

42

 

 

 

Pollett, M., J. S. Mackenzie, and K. J. Turner. 1979. The effect of protein-deprivation on the
susceptibility to influenza virus infection: a murine model system. Aust J Exp Biol Med
Sci 57:151-160.
Ritz, B. W., I. Aktan, S. Nogusa, and E. M. Gardner. 2008. Energy restriction impairs natural
killer cell function and increases the severity of influenza infection in young adult male
C57BL/6 mice. J Nutr 138:2269-2275.
Robb, J. R., G. M. Tori, and R. W. Kroll. 2001. Condition indices of live-trapped American
black ducks and mallards. Journal of Wildlife Management 65:755-764.
Scrimshaw, N. S., C. E. Taylor, and J. E. Gordon. 1968. Interactions of nutrition and infection.
Monogr Ser World Health Organ 57:3-329.
Sharp, G. B., Y. Kawaoka, D. J. Jones, W. J. Bean, S. P. Pryor, V. Hinshaw, and R. G. Webster.
1997. Coinfection of wild ducks by influenza A viruses: distribution patterns and
biological significance. Journal of Virology 71:6128-6135.
Sherman, P., J. Forstner, N. Roomi, I. Khatri, and G. Forstner. 1985. Mucin depletion in the
intestine of malnourished rats. Am J Physiol 248:G418-423.
Smirnov, A., D. Sklan, and Z. Uni. 2004. Mucin dynamics in the chick small intestine are altered
by starvation. J Nutr 134:736-742.
Spackman, E., D. A. Senne, T. J. Myers, L. L. Bulaga, L. P. Garber, M. L. Perdue, K. Lohman,
L. T. Daum, and D. L. Suarez. 2002. Development of a real-time reverse transcriptase
PCR assay for type A influenza virus and the avian H5 and H7 hemagglutinin subtypes.
Journal of Clinical Microbiology 40:3256-3260.
Sprunt, D. H. 1942. The Effect of Undernourishment on the Susceptibility of the Rabbit to
Infection with Vaccinia. J Exp Med 75:297-304.
Stephensen, C. B., S. R. Blount, T. R. Schoeb, and J. Y. Park. 1993. Vitamin A deficiency
impairs some aspects of the host response to influenza A virus infection in BALB/c mice.
J Nutr 123:823-833.
Stephenson, I., A. Heath, D. Major, R. W. Newman, K. Hoschler, W. Junzi, J. M. Katz, J. P.
Weir, M. C. Zambon, and J. M. Wood. 2009. Reproducibility of Serologic Assays for
Influenza Virus A (H5N1). Emerging Infectious Diseases 15:1250-1259.
Sturm-Ramirez, K. M., D. J. Hulse-Post, E. A. Govorkova, J. Humberd, P. Seiler, P.
Puthavathana, C. Buranathai, T. D. Nguyen, A. Chaisingh, H. T. Long, T. S. P.
Naipospos, H. Chen, T. M. Ellis, Y. Guan, J. S. M. Peiris, and R. G. Webster. 2005. Are
ducks contributing to the endemicity of highly pathogenic H5N1 influenza virus in Asia?
Journal of Virology 79:11269-11279.

43

 

 

 

Swayne, D., D. Senne, and C. Beard. 1998. Avian influenza 4, 150-155.
Tobita, K., A. Sugiura, C. Enomoto, and M. Furuyama. 1975. Plaque Assay and Primary
Isolation of Influenza a Viruses in an Established Line of Canine Kidney-Cells (Mdck) in
Presence of Trypsin. Medical Microbiology and Immunology 162:9-14.
van Gils, J. A., V. J. Munster, R. Radersma, D. Liefhebber, R. A. M. Fouchier, and M. Klaassen.
2007. Hampered Foraging and Migratory Performance in Swans Infected with LowPathogenic Avian Influenza A Virus. Plos One 2:-.
Wallensten, A., V. J. Munster, N. Latorre-Margalef, M. Brytting, J. Elmberg, R. A. M. Fouchier,
T. Fransson, P. D. Haemig, M. Karlsson, A. Lundkvist, A. D. M. E. Osterhaus, M.
Stervander, J. Waldenstrom, and B. Olsen. 2007. Surveillance of influenza A virus in
migratory waterfowl in northern Europe. Emerging Infectious Diseases 13:404-411.
Webster, R. G., W. J. Bean, O. T. Gorman, T. M. Chambers, and Y. Kawaoka. 1992. Evolution
and Ecology of Influenza-a Viruses. Microbiological Reviews 56:152-179.
Webster, R. G., M. Yakhno, V. S. Hinshaw, W. J. Bean, and K. G. Murti. 1978. Intestinal
Influenza - Replication and Characterization of Influenza-Viruses in Ducks. Virology
84:268-278.
Woolcock, P. R. 2008. Avian influenza virus isolation and propagation in chicken eggs. Methods
Mol Biol 436:35-46.
Wu, R., S. Hu, Y. Xiao, Z. Li, D. Shi, and D. Bi. 2007. Development of indirect enzyme-linked
immunosorbent assay with nucleoprotein as antigen for detection and quantification of
antibodies against avian influenza virus. Veterinary Research Communications 31:631641.

44