Detection of Influenza a Viruses from Environmental Lake and Pond Ice

Home , Antigenic shift, Hemagglutination assay

TITLE “DETECTION OF INFLUENZA A VIRUSES FROM ENVIRONMENTAL LAKE AND

POND ICE”

Zeynep A. Koçer

A Dissertation

Submitted to the Graduate College of Bowling Green

State University in partial fulfillment of

the requirements for the degree of

DOCTOR OF PHILOSOPHY

August 2010

Committee:

Scott O. Rogers, Advisor

W. Robert Midden

Graduate Faculty Representative

John Castello

George Bullerjahn

Paul Morris

ABSTRACT

Scott O. Rogers, Advisor

Environmental ice is an ideal matrix for the long-term protection of organisms due to the limitation of degradative processes. As a result of global climate change, some glaciers and polar ice fields are melting at rapid rates. This process releases viable microorganisms that have been embedded in the ice, sometimes for millions of years. We propose that viral pathogens have adapted to being entrapped in ice, such that they are capable of infecting naïve hosts after melting from the ice. Temporal gene flow, which has been termed genome recycling (Rogers et al., 2004), may allow pathogens to infect large host populations rapidly. Accordingly, we hypothesize that viable influenza A virions are preserved in lake and pond ice. Our main objective was to identify influenza A (H1-H16) from the ice of a few lakes and ponds in Ohio that have high numbers of migratory and local waterfowl visiting the sites. We developed a set of hemagglutinin subtype-specific primers for use in four multiplex RT-PCR reactions. Model studies were developed by seeding environmental lake water samples in vitro with influenza A viruses and subjecting the seeded water to five freeze-thaw cycles at -20oC and -80oC. Results demonstrated that influenza A viruses survive freeze-thaw cycles at both temperatures with a high rate of viability. We then concentrated the meltwater samples from various locations and years by ultracentrifugation and tested for viable influenza A virions using SPF (specifically pathogen free) embryonated chicken egg cultures. Matrix and hemagglutinin genes of the viruses were detected by RT-PCR (reverse transcription-polymerase chain reaction) and sequencing after several passages depending on the starting concentration of the sample. We analyzed the phylogenetic relationships of the viral strains obtained from environmental ice samples with the known strains available in GenBank based on those two different viral genes. These confirmed iii the presence of specific subtypes of influenza A viruses. The increase in positive RT-PCR amplicons indicated that the viruses were replicating in the eggs. Our results demonstrated that viable and infectious influenza A virions are preserved in environmental lake and pond ice.

To my parents,

Necmi & Mücella Koçer;

To my sister, Merve Koçer;

To all musicians who kept me going during all these years; and

To my friends, John Tallman

& Christa L. Bowen, who could not complete their degrees due to unfortunate circumstances

ACKNOWLEDGMENTS

I would like to thank my advisor, Dr. Scott O. Rogers, with great appreciation for his endless support and patience; for trusting me to take this „dark-hole‟ project; for guiding and challenging me through my PhD; and for being a great mentor by encouraging me towards to the light at the end of tunnel. I would also like to thank him for being a best friend and a father to me. There are not enough words to mention my gratitude to you, Scott. Thank you for giving me this great opportunity to work with you. I would like to thank all my committee members Dr. John

Castello, Dr. Paul Morris, Dr. George Bullerjahn, and Dr. Robert Midden for giving me insightful advices, for challenging me and for being very understanding. I would like to acknowledge Dr. Robert Webster and his colleagues for giving me a great opportunity to visit St.

Jude Children‟s Research Hospital facilities and for teaching me how to perform viral culture technique and for sending me the samples I needed; Dr. Yoshihiro Kawaoka and Dr. Ron

Fouchier for providing me samples. I would like to thank them all for giving me great advices, endless help and for making this project possible by sending me the positive viral samples used in this study. This work was partially supported by the National Health Institute. I would like to thank all my lab mates through these years: Seung-Geuk Shin for all his help whenever I needed,

Lorena Harris and Farida Sidiq for being lifetime friends and sisters to me, Ram Veerapaneni,

Tom D‟Elia, Gang Zhang, Yury Shtarkman, Xing Chen, Chia-Jui Tsai, Ameria Vicol, Justin

Zollar, Amal Abu-Almakarem, and the undergraduates Katie Heilman and Clarissa Polen for their great friendship and support during rough days of PhD. Thanks to my dear friends, Mike

Schlais and Tamera Wales, for always being there for me. I would also like to thank all my friends in Turkey for being supportive from across the ocean and the friends in US for making vi me feel like I am home here. Thanks to the Biology Department office staff Lorraine, Linda,

Chris, Steve, Deb, Marsha, Sheila and Chellie for being very friendly and very helpful through my PhD years. Last but not the least, my special thanks to my dad, Necmi, for challenging me and encouraging me to go for my dreams; to my mom, Mücella, for always being there for me and for being a great mom and a best friend and for always helping me in anyway to make my dreams come true. I would love to explain my great appreciation to both my parents for teaching me to love, to be patient, to be understanding and especially to follow your dreams. My special thanks are also to my dearest sister, Merve, for being a great friend and a great listener through all these tough years; and to my other siblings Ilknur, Gonca and Ahmet for their endless support and faith in me.

THANK YOU ALL SO MUCH!! vii

TABLE OF CONTENTS

CHAPTER 1. INTRODUCTION ...... 1

1.1. Genome of influenza A viruses ...... 1

1.2. Genetic events leading to variations, selective pressures and evolution of influenza A ...... 4

1.2.1. Emergence of highly pathogenic avian influenza (HPAI) strains ...... 8

1.3. Host range of avian influenza ...... 9

1.4. Rationale for this dissertation ...... 11

1.5. Objectives ...... 12

1.6. Literature cited ...... 13

CHAPTER 2. PRIMER DEVELOPMENT FOR THE RAPID DETECTION OF HEMAGGLUTININ SUBTYPES OF INFLUENZA A VIRUSES FROM ENVIRONMENTAL SAMPLES ...... 16

1. Introduction ...... 16

2. Methods...... 18

2.1. Primer Design ...... 18

2.2. Sensitivity of the primers ...... 19

2.3. Testing for the cross-reactivity ...... 20

2.4. Development of multiplex RT-PCR conditions ...... 20

3. Results and Discussion ...... 23

3.1. Primer Design ...... 23

3.2. Sensitivity of the primers ...... 29

3.3. Testing for the cross-reactivity ...... 29

3.4. Development of multiplex RT-PCR conditions ...... 30

4. Literature cited ...... 34

CHAPTER 3. EFFECTS OF FREEZING AND THAWING ON THE VIABILITY OF INFLUENZA A VIRUSES ...... 36

1. Abstract ...... 36 viii

2. Introduction ...... 36

3. Materials and Methods ...... 39

3.1. Viral strains ...... 39

3.2. Water sampling and inoculation with the virus mixture ...... 39

3.3. Viral culture ...... 40

3.4. RNA Extraction and RT-PCR amplifications ...... 41

4. Results ...... 42

5. Discussion ...... 46

6. Literature cited ...... 48

CHAPTER 4. DETECTION OF INFLUENZA A VIRUSES FROM ENVIRONMENTAL LAKE AND POND ICE ...... 51

1. Introduction ...... 51

2. Materials and Methods ...... 53

2.1. Ultracentrifugation ...... 53

2.1.1. Viral strains ...... 53

2.1.2. Water sampling and inoculation with the virus ...... 54

2.1.3. Concentrating the viruses from frozen pond water ...... 54

2.1.4. Viral culture ...... 55

2.1.5. RNA Extraction and RT-PCR ...... 56

2.2. Environmental Samples ...... 57

2.2.1. Sampling sites ...... 57

2.2.2. Concentrating the samples ...... 58

2.2.3. Viral culture ...... 58

2.2.4. RNA Extraction, RT-PCR, Cloning and Sequencing ...... 59

3. Results and Discussion ...... 60

3.1. Effects of ultracentrifugation on the viability of influenza A viruses ...... 60

3.2. Assaying the environmental ice and water samples for viable influenza A virions ...... 62 ix

4. Literature cited ...... 71

APPENDIX. SUPPLEMENTARY MATERIAL FOR CHAPTER 2 ...... 74

LIST OF FIGURES

Figure Page

1.1. Influenza A structure with its genome segments ...... 2

1.2. Host range of influenza A ...... 10

2.1. Diagram of the expected sizes of the RT-PCR products for the hemagglutinin genes from each

subtype in four multiplex reactions ...... 22

2.2. Multiplex RT-PCR products amplified using Group 3 and Group 4 primers ...... 33

3.1. The average hemagglutination titers recovered from subsequent freeze-thaw cycles ...... 43

3.2. RT-PCR products amplified based on matrix and hemagglutinin gene of viral nucleic acids extracted

from the allantoic fluids ...... 45

4.1. RT-PCR products amplified by H2 primers from allantoic fluids harvested after the 3rd passage of

the samples ...... 61

4.2. Matrix gene amplification from the first and second passages of allantoic fluids ...... 64

4.3. Maximum parsimony phylogram of a broad selection of matrix gene sequences including the

sequences obtained from different environmental samples ...... 65

4.4. RT-PCR products amplified using Group 1 primers from the viral cultures of concentrated

environmental samples ...... 66

4.5. Neighbor-joining phylogram of the influenza virus hemagglutinin H7 sequence isolated from Ottawa

NWR ...... 68

LIST OF TABLES

Table Page

2.1. Primer sequences designed based on the HA genes of 16 subtypes of influenza A ...... 25

2.2. Detailed information for each primer about the amplified regions of HA gene, the length of the

primer sequences, Tm values, GC content, and the expected PCR product sizes ...... 26

2.3. Analyses of the primers for secondary (2o) structure and primer dimer formation ...... 28

2.4. The sensitivity of the primers...... 31

3.1. Hemagglutinin and matrix primers used to amplify five low pathogenic influenza A strains ...... 42

4.1. Sampling sites, sample type, sampling date, concentration, number of SPF eggs inoculated and

number of passages for viral culture ...... 57

S1. Sequences retrieved from NCBI Influenza Virus Resource for the primer design ...... 74

CHAPTER 1. INTRODUCTION

1.1. Genome of influenza A viruses

Influenza A viruses are enveloped, negative-sense ssRNA viruses belonging to the

Orthomyxoviridae. The genome of the virus is approximately 13 kb long and contains 8 segments of RNA molecules encoding 10 proteins (Fig. 1.1). Viral encoded proteins can be divided into three major groups as the products of nucleocapsid genes (N, NS1/NS2, M1/M2), glycoprotein genes (HA, NA) and polymerase genes (PA, PB1, PB2) which are analogous to the proteins coded by all other negative-sense virus genomes (Strauss & Strauss, 2002). RNA dependent RNA polymerase activity is provided by three gene products. The subunits of viral polymerase are encoded by segment 1 (PB2), segment 2 (PB1) and segment 3 (PA). PB2 involves in the 5‟ cap recognition of host mRNA and initiation of transcription, while PB1 involves in the elongation of nascent viral RNA. It has been assumed that PA is a protease, but its function is unclear (Webster, 1992). Segment 4 encodes hemagglutinin (HA), which is a surface glycoprotein and an integral membrane protein, involved in the recognition of the sialic acid receptor on the host cell and binding. It facilitates the fusion between viral envelope and the host cell membrane. Nucleoprotein (N) is the product of segment 5 and is responsible for encapsidating viral RNA by forming the nucleocapsid. It is also believed that it serves as a part of transcriptase complex by switching the RNA polymerase activity to synthesize viral RNA.

Neuraminidase (NA) is encoded by segment 6 and it is a surface glycoprotein and an integral membrane protein, as well. NA is responsible for cleaving the virus progeny particles from host cell surface during budding. Segment 7 encodes two matrix proteins (M1 and M2). M1 is

Figure 1.1. Influenza A structure with its genome segments: PB2, PB1 and PA - viral polymerases, HA-hemagglutinin, NP-nucleoprotein, NA-neuraminidase, M-Matrix proteins, and

NS-non-structural proteins (adopted from http://www.pasteur.ac.ir/researchDepartment/flu//images/flu_structure.gif)

3 the most abundant protein in the virus particle and it serves to cover the nucleocapsid which will be further covered by the envelope. M2 is also integral membrane protein which serves as ion channel to decrease the interior pH of the cell environment to facilitate viral uncoating after viral entry into the cell. It also inhibits the fusion activity of HA during its transport by equilibrating the pH between the transport vesicle and the cytosol. Segment 8 encodes two nonstructural proteins: NS1 and NS2. In an infected cell, NS1 is abundant in the nucleus, while NS2 is mainly found in the cytoplasm. NS1 is a product of unspliced mRNA and it has several functions. NS1 provides a preference for viral mRNA translation by inhibiting the transport of cellular mRNA and it facilitates the transport of viral mRNA from the nucleus to the cytoplasm after being involved in mRNA splicing as well. It is also known as an anti-interferon protein inhibiting the destruction of viral particles by the host immune system. NS2 is a product of spliced mRNA.

Although its function is unclear, it has been assumed that it facilitates the transport of viral ribonucleoprotein to the cytoplasm (Strauss & Strauss, 2002; Webster et al., 1992). The outer envelope of influenza A virus is acquired from host cell membrane during budding process.

All RNA viruses, including influenza A, replicate their genomes using a replicase (RNA- dependent RNA Polymerase-RdRp) encoded by viral genes, since infected host cells cannot maintain RNA-to-RNA replication efficiently. These viral encoded RNA synthesizing enzymes are also used for the synthesis of mRNA in negative-sense RNA viruses. Therefore, negative sense RNA genomes do not possess the ability to be infectious as purified nucleic acids due to their requirement of viral polymerases (RdRp) for the replication of the genome and transcription of mRNA. Since translation of these genomes is performed by viral proteins, negative-sense ssRNA genomes tend to be larger than positive-sense ssRNA genomes in order to be able to express all required proteins. It is also common to see segmentation of the genome in negative-

4 sense ssRNA viruses, as in influenza A. Although nucleic acid is formed by different segments, they are all packaged into a single virus particle (Cann, 2005).

1.2. Genetic events leading to variations, selective pressures and evolution of influenza A

Due to the segmented pattern of the nucleic acid and the lack of proofreading activity of

RNA polymerase enzyme, the influenza A genome undergoes rapid genetic changes as do all other RNA viruses. They undergo reassortment, recombination, and mutations caused by insertions or deletions. Although these changes occur in all genes encoded by the genome, the major mutations occur in hemagglutinin (HA) and neuraminidase (NA) genes which encode for surface glycoproteins responsible from antigenicity. So far, 16 HA subtypes have been detected based on HA surface antigenicity, while 9 NA subtypes have been identified according to NA surface antigenicity.

During their evolution, the selective pressures imposed on influenza viruses are mainly neutralizing antibodies, chemical antivirals and receptor specificity. As mentioned before, they are subjected to high mutation rates (6.7x10-3 substitutions per nucleotide per year for HA and

3.2x10-3 substitutions per nucleotide per year for NA) (Steinhauer and Skehel, 2002). Influenza

A viruses undergo two main types of genomic changes. These events are antigenic drift and antigenic shift. When an individual is infected with a specific strain of flu, antibodies are produced against the viral antigens. The production of antibodies makes the host organism resistant against the viral infection caused by certain strains for a few years. Therefore, viruses can mutate to eventually escape from immunological recognition by the host organism in an event called antigenic drift. In order to do this, antigenic sites undergo nucleotide substitutions to change the amino acid rather than synonymous mutations which do not change the amino acid

(Earn et al., 2002; Kamps et al., 2006). During adaptation to the new host, newly emerging viral strains with novel antigenicity undergo rapid evolution by being candidates for future pandemics in some cases.

Among all influenza viruses (type A, B, C), influenza A is the only one that undergoes antigenic shift, as well. Antigenic shift gives the viruses an opportunity to jump from one species to another. This type of genetic exchange that facilitates crossing the species barriers occurs in three different ways. When different strains of the virus (generally one from avian strains and one from human strains) infect the same host, they can undergo reassortment by exchanging genes. The segmented nature of the virus genome facilitates the exchange of entire genes between different viral strains. For this type of reassortment, it has been proposed that intermediate host species (such as pigs and chickens) serve as mixing vessels (Webster et al.,

1992; Earn et al., 2002; Kamps et al., 2006). Alternatively, an avian strain can transmit directly to humans without any genetic change, as it has been seen in Asian lineage of highly pathogenic

H5N1 and H9N2. Another way is to transmit the avian strain to an intermediate host and then to humans, again without any genetic change (Kamps et al., 2006). By infecting the intermediate host, receptor specificity can change. During the 20th century, there have been three major antigenic shifts; H2N2 in 1957, H3N2 in 1968 and H1N1 reappearance in 1977. In the 1957 reassortant, HA, NA and PB1 genes were of avian origin, while the other segments were of human origin. In 1968, reassortment occurred when HA and PB1 genes of avian origin became incorporated into a circulating human strain (Steinhauer & Skehel, 2002). Although these virus strains were very aggressive when they first appeared in the human population and caused almost one million mortalities and high morbidity, they have co-existed in human populations since then.

RNA recombination is a rare event among influenza viruses. However, they can undergo non-homologous recombination events within and between gene segments during replication

(Steinhauer & Skehel, 2002). Besides, there are some speculations about the 1918 influenza HA containing an HA2 subunit from a human strain and an HA1 subunit from a swine strain (Gibbs et al., 2001).

The major pandemic in 1918 with the appearance of H1N1 in the human population caused 50-100 million death worldwide (Kamps et al., 2006). When the sequence of the 1918

H1N1 strain was revealed by Taubenberger and his colleagues (1997), it was shown that this pandemic strain was not a reassortant, but it was an entirely avian strain which had adapted to humans gradually. Comparing the mortality rate caused by the 1918 strain and the other pandemics caused by reassortants, it has been proposed that gradual changes in the virus genome by point mutations or antigenic drift are more dangerous to the host species than the reassorted strains (Kamps et al., 2006).

Depending on the migratory flyways of the birds, there are two super families of influenza A viruses: one in Eurasia and one in America. Although these flyways overlap in

Alaska and in Labrador, viruses generally do not mix between the two geographical variations as indicated by the fact that the phylogenetic clades are distinct, and have been for a long period of time (Webster et al., 2007). Although viruses can transmit between these two clades, the frequency is very low. This indicates that influenza A viruses might undergo local adaptation to their hosts.

Considering the fact that influenza A is avirulent in their natural hosts (especially ducks, geese and shorebirds) and they exhibit small amounts of genetic change in their genomes,

7 antigenic drift and antigenic shift in those wild birds, it is suggested that these viruses have been in evolutionary stasis with their natural host reservoir for many centuries (Webster et al., 1992).

They are able to replicate themselves by causing few or mild disease symptoms in wild birds.

This strategy ensures the virus survivability, while giving the viruses an opportunity to infect other organisms by crossing species barriers. Flu has coexisted with the human populations over centuries. Influenza A transmission between species creates temporary lineages, not stable ones.

It is well known that avian influenza uses intermediate host species, such as swine, chicken and quail. These intermediate species serve as mixing vessels between avian and human populations.

As a summary, although they differ from each other, all influenza viruses (type A, B and

C) share a common ancestor that differs from all the other negative-sense ssRNA viruses.

Specifically speaking of influenza A, host-specific strains evolve independently due to the barriers caused by ecological and geographical differences among different host species, immune responses of the host, or lack of infectivity in new host cells. However, viral genes must co- evolve with the host to avoid host immunity and to increase the fitness of progeny virions. While reassortment between co-infecting strains may give rise to the formation of new genotypes in the gene pool, some of these newly emerged genotypes cannot co-exist with the host populations and they are subjected to negative selection. Because neutralizing antibodies produced by host immune system exert high selective pressure on surface glycoproteins (HA and NA), these genes need to evolve faster or be replaced by reassortment. Because of this, reassortant strains may have more selective advantage than the parental viral strains. The other gene segments are under selective pressure caused by roles in efficient virus replication. Part of this is due to the huge number of viral RNAs produced in each infected cell. The probable reason for the variability in the HA and NA genes is that host immune systems kill specific strains, thus removing them from

8 the population. Maintenance of many HA and NA subtypes is necessary to the survival of the virus, while maintenance of low variation in the other genes is necessary to retain functions that ensure virus replication and production. As discussed before, influenza A viruses evolve slowly

(even maintaining evolutionary stasis) in their natural aquatic wild bird reservoir, but they evolve faster in other birds, humans and other mammal populations. As mentioned in Webster et al.

(2007), the development of at least three separate clades in a 3-year period for highly pathogenic

H5N1 indicates the ability of these viruses to evolve rapidly. This is especially true when many versions are able to evade host defenses.

1.2.1. Emergence of highly pathogenic avian influenza (HPAI) strains

When there is an insertion of basic amino acids (especially arginine or lysine) in the cleavage site of the HA protein, some low pathogenic strains become highly pathogenic. This pattern is observed for H5 and H7 subtypes so far. In low pathogenic viruses, the cleavage site of the HA protein contains two basic amino acids and cleavage of the protein is catalyzed by trypsin-like proteases. These proteases are tissue specific and generally found at the surface of respiratory and gastrointestinal epithelia. The insertion of additional amino acid makes the HA protein susceptible to subtilisin-like proteases (eg. furin) which are found in almost every tissue in the body (Kamps et al., 2006). Changing the cleavage site by amino acid insertion is an advantage for highly pathogenic strains to be able to replicate in a systemic manner, since selective pressure is exerted at the HA cleavage site in natural hosts. In its natural host, influenza

A replicates mostly in the intestines after passing through the stomach. Cleaved HA is less stable than uncleaved HA at acidic pHs. Therefore, cleavability of HA by certain types of tissue- specific proteases provides a selective advantage for viral survivability in natural hosts.

1.3. Host range of avian influenza

The natural reservoirs for influenza A viruses are wild aquatic birds for which viral infection is mainly low pathogenic, generally asymptomatic, waterborne and intestinal. It is transmitted via a fecal-oral route through water. However, influenza A viruses can infect many organisms (such as humans, pigs, domestic poultry, cats, sea mammals, horses, etc.) (Fig. 1.2).

Although some subtypes are host specific (such as H1, H2 and H3 for humans, H3N8 for horses and H1 and H3 for pigs) (Webster et al., 2007), all subtypes of influenza A are found in wild birds. Horses are thought be evolutionary dead ends for influenza viruses due to the fact that the interspecies transmission of the virus has not been seen between horses and other animals, although equine influenza A virus acquired its HA (H3) from avian species in the distant past

(Webster et al., 1992).

Although it is not the sole factor, changes in receptor specificity of the virus mainly determines the broad host range. These receptors are tissue and host specific. In avian and equine species, influenza A virus recognizes an α-2-3 sialic acid linkage which is found largely in epithelial tissues of gut and lung, while human strains recognizes an α-2-6 sialic acid linkage which predominates on non-cilial epithelial cells in human airways. In intermediate species, both receptor types are found in higher densities allowing both avian and human strain infections.

However, recent studies have demonstrated that avian like receptors are present in human lungs, as well (Shinya et al., 2006). This indicates that avian viruses can be transmitted to humans directly without an intermediate host. Also, for highly pathogenic H5N1, it has been shown that viruses are shed by a tracheal route in aquatic birds as well indicating that respiratory

10 transmission of viruses is also possible for the aquatic birds, in addition to the standard fecal-oral route (Webster et al., 2007).

Figure 1.2. Influenza A viruses have broad host ranges, including infection of waterfowl, poultry, swine, humans, horses and sea mammals (adopted from http://www.aht.org.uk/images/flu1.gif)

1.4. Rationale for this dissertation

Influenza A viruses are epidemiologically very important pathogens due to their broad host range and their ability to cause pandemics and yearly epidemics. So far, research has been focused on the biotic reservoirs of these viruses. As mentioned before, the RNA genome of the virus is prone to rapid genetic changes. However, some strains have appeared, disappeared and reappeared intermittently, even decades later, almost without any change (Smith et al., 2004;

Shoham, 1993; Shoham, 2005). This fact has raised the question whether these viruses can be preserved in abiotic reservoirs, such as water and ice.

Many researchers have shown high rates of virus survival in lake and river water samples, especially at colder temperatures (4oC) (Hinshaw et al., 1979; Webster et al., 1992;

Markwell and Shortridge, 1982; Stallknecht et al., 1990a; Stallknecht et al., 1990b; Ito et al.,

1995; Brown et al., 2007; Brown et al., 2009). The possibility of environmental ice as abiotic reservoir for these viruses has been proposed by some of these researchers, although little research has been performed.

Environmental ice is an ideal matrix for long-term protection of organisms due to the limitation of degradative processes; light, free water and heat. It has been shown that influenza A

H1 RNA was preserved in Siberian lake ice (Zhang et al., 2006) and virus RNA of various subtypes were recovered from Alaskan lake sediment samples (Lang et al., 2008). It is also well known that virus survivability increases at cold temperatures and below freezing. When water temperatures are warm (22-25oC), 99.9 % of the virus population is inactivated in a week, while it takes several weeks to months if the temperature decreases (increased rate of survivability with decrease temperature) (Parker and Martel, 2002). Also, it has also been shown for influenza A

12 that decreasing temperature makes the lipids in the viral envelope more stable, thereby increasing the chance of virus survival at low temperatures (Polozov et al., 2008). Therefore, we hypothesize that influenza A viruses are able to survive the winter by being entrapped in environmental ice, such that they are capable of infecting naïve hosts after melting from the ice.

This project is the first study on the detection of viable influenza A viruses from environmental ice samples. The data obtained from this project will shed light onto the importance of abiotic environmental sources, such as ice, as alternative mixing vessels for these viruses and will provide additional information about the ecology and evolution of influenza A viruses to understand the potential origin of new pathogenic strains.

1.5. Objectives

In the present research, the viability of influenza A viruses in environmental lake and

pond ice has been assessed. Specific objectives that were addressed are as follows:

 to optimize a rapid, sensitive and accurate molecular technique by the

development and use of hemagglutinin subtype-specific primers in multiplex RT-

PCR reactions,

 to assess the effect of freezing and thawing on the viability of influenza A viruses,

 to analyze various environmental water and ice samples collected at various times

of the year, and from various locations to determine the concentrations of

infectious viral particles,

 to assess the importance of influenza A surveillance to understand the ecology

and evolution of these viruses.

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Brown, J.D., G. Goekjiana, R. Poulsona, S. Valeikab, and D.E. Stallknecht. 2009. Avian

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Zhang, G., D. Shoham, D. Gilichinsky, S. Davydov, J.D. Castello, and S.O. Rogers.

2006. Evidence of influenza A virus RNA in Siberian lake ice. Journal of Virology, 80:

12229-12235.

CHAPTER 2. PRIMER DEVELOPMENT FOR THE RAPID DETECTION OF

HEMAGGLUTININ SUBTYPES OF INFLUENZA A VIRUSES FROM

ENVIRONMENTAL SAMPLES

1. Introduction

Influenza A viruses circulate constantly in animals and humans annually. Approximately

35,000 people in the US die from the effects of this virus each year. Occasionally, highly pathogenic strains cause mortality in birds, and some of these can be transmitted to humans, often with high rates of morbidity and mortality. Being able to identify these viruses accurately and rapidly is crucial to being able to applying preventative and therapeutic measures to control the spread and severity of this disease. To identify avian influenza viruses during an outbreak, a basic approach would be the observation of clinical signs followed by serological diagnosis.

However, it is not an easy task to follow the clinical signs from infected birds, since the majority of the time symptoms are not detectable without another diagnostic test for confirmation.

Serological diagnosis of highly pathogenic avian influenza is difficult due to the fact that, in most cases, antibody development is not observed by the time birds die. Besides, even if the antibodies are detectable before the death of an infected bird, this detection takes considerable time during which infection spreads among the population and may move to distant location. To date, several serological and molecular diagnostic tests have been developed for these viruses.

Still today, one of the most valuable diagnostic tests is the isolation of virus in embryonated chicken eggs or in tissue culture using Madin-Darby canine kidney (MDCK) cells. Although it is very helpful and required confirmation of virus infectivity, the virus culturing methods are time consuming. Besides, in addition to the requirement of having a ready supply of cell lines or eggs,

17 the work might require the use of higher biosecurity levels when attempting to culture or amplify potentially highly pathogenic influenza (Suarez et al., 2007). Embryonated egg or cell culture studies are also followed by hemagglutinin inhibition and/or hemagglutination assays to confirm the virus subtype and virus titer, respectively. Although hemagglutination assay can give rapid results about the virus titer, virus titers must be quite high for detection (Rimmelzwaan et al.,

1998). On the other hand, hemagglutinin inhibition assay requires the use of antisera raised against each subtype. While it is a widely used method for subtyping the avian influenza, it is labor intensive, higher levels of biosecurity are necessary, and it is specific for particular subtypes of the virus. In general, although serological methods can detect the virus accurately, the time required to perform these tests and the instant availability of biosecurity conditions are major drawbacks during outbreaks.

Molecular diagnostic tools target the viral nucleic acids. There have been several published reports of methods developed to identify specific strains of influenza A, as well as a few reports that describe methods to identify broad ranges of influenza A viruses. Although very sensitive primers were designed for the detection of matrix genes (Fouchier et al., 2000; Di Trani et al.,

2006), these amplifications do not give any information about the subtype of influenza A viruses.

Hoffman et al. (2001) developed the universal primer set for the detection of 8 segments of the virus genome. While the universal primer set amplifies the each segment to their full lengths, the amplified products, especially hemagglutinin and neraminidase, need to be sequenced to determine the subtype of the virus.

Based on the objective of rapid detection of the hemagglutinin subtype of these viruses by

RT-PCR method, several hemagglutinin-specific primers have been also designed by different research groups. The primers developed specific for 15 HA-subtypes were sensitive for

18 subtyping, but require separate RT-PCR amplifications (Lee et al., 2001), which is labor intensive. In another study, a single set of primers were developed based on the conserved region in the HA2 part of hemagglutinin gene for the detection of various subtypes of influenza A in a single-step reaction (Phipps et al. 2004). However, this amplification attempt requires the sequencing of the PCR products to determine the exact subtype due to the fact that all amplicons are of similar size. In addition, some variations were detected within that conserved regions, especially in H5 sequences, possibly reducing the sensitivity of this primer set. Due to the high virulence of H5 and H7 in poultry and increased number of outbreaks, single step multiplex RT-

PCR was optimized for the rapid detection of these specific subtypes. However, these multiplex reactions were either limited to detect H5 and N1 subtypes in one step reactions (Wei et al.,

2006) or limited for the simultaneous amplification of H5 and H7 subtypes (Thontiravong et al.,

2007).

In this study, we have developed a set of primers that can be used in four multiplex RT-PCR amplification reactions to produce amplicons of different sizes for all 16 subtypes. These primers can be used as an initial diagnostic test to determine the hemagglutinin subtype of avian influenza, either directly from environmental samples or from extracted viral nucleic acids from virus cultures. The sensitivity and specificity of the primers allows the detection of viral nucleic acids even in low titers. The method is accurate, simple, sensitive, and rapid.

2. Methods

2.1. Primer Design

Twenty primer sets were developed to detect 12 hemagglutinin (HA) subtypes. For the other four subtypes, H1, H2, H5 and H6, previously designed primers were used (H1 primers-

Zhang et al., 2006; H2, H5 and H6 primers - G. Zhang, unpublished). For each subtype, complete hemagglutinin gene sequences were retrieved from NCBI Influenza Virus Resource

(Table S1). Sequences were aligned with ClustalW2

(http://www.ebi.ac.uk/Tools/clustalw2/index.html) to determine the conserved nucleotide sequences. Primers were designed based on the conserved regions of the HA genes. Primer sequences were selected based on their GC content, dissimilarities at their 5‟ and 3‟ ends to avoid non-specific priming and their Tm (melting/annealing temperatures) values. Tm and GC content of these sequences were determined using an online oligo calculator

(http://www.cnr.berkeley.edu/~zimmer/oligoTMcalc.html). Each primer individually and each primer set (forward and reverse) were analyzed for secondary structure and primer dimer formation to avoid mispriming using another online DNA calculator (http://www.sigma- genosys.com/calc/DNACalc.asp). More than one primer set was designed to detect certain subtypes, such as H3, H4, H7 and H11 due to the sequence variations in the hemagglutinin genes among different strains of these subtypes.

2.2. Sensitivity of the primers

Positive viral RNA nucleic acids were obtained from Dr. Yoshihiro Kawaoka, School of

Veterinary Medicine, University of Wisconsin, Madison, WI; and from Dr. Robert G. Webster,

Division of Virology, Department of Infectious Diseases, St. Jude Children‟s Research Hospital,

Memphis, TN. The sensitivity of the primers was determined by RT-PCR amplification of the part of the HA gene from positive viral nucleic acids for all 16 HA subtypes in 10-fold dilutions up to 10-10.

RT-PCR amplifications were performed using GeneAmp EZ rTth RNA PCR Kit

(Applied Biosystems, Branchburg, NJ). The composition of each 25 µl reaction was as follows:

1X EZ Buffer, 300 µM of each dNTPs, 5 units/50 µl of rTth DNA Polymerase, 2.5 mM of 25 mM Mn(OAc)2, 0.45 µM of each primer and 1 µl of each viral RNA extract. Reverse transcription was performed at 60oC for 45 min followed by PCR amplification of the cDNA at

95 oC for 4 min, and 40 cycles of 94 oC for 1 min; 53, 56, 58 or 60 oC for 2 min; and 72 oC for 2 min. Final extension was performed at 72 oC for 10 min and cooled to 4 oC. PCR products were subjected to electrophoresis on 1% agarose gels in TBE (89 mM Tris-base, 89 mM borate, 2 mM

EDTA, pH 8.0), containing 0.5 µg/ml ethidium bromide. After electrophoresis, gels were visualized using UV irradiation, and digitally photographed.

2.3. Testing for the cross-reactivity

Following the sensitivity tests, each primer set was tested with the other 15 viral RNAs to avoid cross-amplification among the HA genes from different subtypes. In order to do so, each viral nucleic acid was subjected to RT-PCR amplifications using all 23 other primer sets individually. RT-PCR amplifications were performed as previously described. PCR products were subjected to electrophoresis on 1% agarose gels in TBE (89 mM Tris-base, 89 mM borate,

2 mM EDTA, pH 8.0), containing 0.5 µg/ml ethidium bromide. Gels were visualized using UV irradiation, and digitally photographed following electrophoresis.

2.4. Development of multiplex RT-PCR conditions

Depending on their annealing temperatures (53, 56, 58 and 60 oC), the twenty-four primer sets were categorized and optimized in four major groups to be used in multiplex RT-PCR reactions. The diagram of the expected amplicons of each primer set for each group of multiplex

RT-PCR is given in Fig. 2.1. Each group of primers was tested in RT-PCR reactions without template to confirm that they do not self-prime each other. Positive viral nucleic acids were subjected to RT-PCR amplifications using a combination of all primer sets in each group to confirm that multiplex amplifications of different strains can be detected in single reactions.

RT-PCR amplifications were performed as previously described. PCR products were subjected to electrophoresis on 1% agarose gels in TBE, containing 0.5 µg/ml ethidium bromide.

Following electrophoresis, gels were visualized using UV irradiation, and digitally photographed.

Figure 2.1. Diagram of the expected sizes of the RT-PCR products for the hemagglutinin genes from each subtype in four multiplex reactions. Annealing temperatures are given for each group. Group 1 primers: H3-1: 413 bp, H3-3: 597 bp, H4-2: 450 bp, H7-2: 491 bp, H11-1: 447 bp, H11-2: 416 bp and

H16: 616 bp; Group 2 primers: H3-4: 491 bp, H4-1:578 bp, H7-3: 633 bp, H7-4: 643 bp, H15: 746 bp;

Group 3 primers: H7-1: 719 bp, H8: 645 bp, H9: 569 bp, H10: 643 bp, H12: 501 bp, H13: 357 bp, H14:

500 bp; Group 4 primers: H1: 677 bp, H2: 509 bp, H3-2: 479 bp, H5: 513 bp, and H6: 375 bp. Molecular weight marker of 100 bp ladder is shown on the left side of each diagram.

3. Results and Discussion

3.1.Primer Design

Although there are other widely used methods to determine the subtypes of influenza A viruses, a few obstacles limit their utility for certain applications. These methods are more labor intensive; less sensitive, or they require the use of antisera raised for each specific strain. In the case of highly pathogenic strains, production of antisera is limited to a small number of authorized laboratories. Also the majority of these methods require the presence of high viral titers. In this study, we attempted to develop specific and sensitive primer pairs for each hemagglutinin subtype of influenza A viruses. The aim was immediate detection of each HA subtype from environmental water and ice samples, in which the virus titers are expectedly lower than in biological samples obtained from birds in the field.

In order to develop efficient primers for the rapid detection of influenza A hemagglutinin

(HA) subtypes from environmental water and ice samples, complete HA gene sequences of all subtypes of influenza A viruses were retrieved from the NCBI Influenza Virus Resource. The

HA gene sequences of each subtype were aligned using ClustalW2

(http://www.ebi.ac.uk/Tools/clustalw2/index.html) in order to locate conserved nucleotide sequences in the hemagglutinin genes of different strains of the same subtype so that primer pairs could be designed. Due to sequence variations in the HA gene, more than one primer pair were developed for some subtypes (e.g. H3, H4, H7 and H11). Each primer sequence to be used in this study is given in Table 2.1. Detailed information about their Tm (melting/annealing temperatures) values,

GC contents, amplified regions and the expected amplicon sizes are given in Table 2.2. Based on this information, each primer was tested with the viral nucleic acids of known subtypes, which will be discussed in the next section.

Each primer and primer set was also examined to avoid any secondary structure formations and/or primer dimer formation by an online DNA calculator tool at Sigma Aldrich website (http://www.sigma-genosys.com/calc/DNACalc.asp). When a problem was observed with the possible primer/primer pair, an alternate site was selected and examined for secondary structure, until a suitable site had been located. The ultimate details of this analysis are shown in

Table 2.3.

Table 2.1. Primer sequences designed based on the hemagglutinin genes of 16 subtypes of influenza A viruses (H1 primers- Zhang et al., 2006; H2, H5 and H6 primers - G. Zhang, unpublished). All others were designed for this project.

Forward (5’-3’) Reverse (5’-3’)

H1 ATGCSAACAACTCAACCGACAC GGGTTCYAGCARRGTCCAGTARTA

H2 TATGCAGCAGACAAAGAATCCAC CYATCATGATTGCCAGTGACA

H3-1 GCAGGTTTCATAGAAAATGG TCACATTTGTGGTATATTTTGAA

H3-2 GCAGCAGACCTTAAAAGCACTCA GATATGGCAAAGGAAATCCACAG H3-3 GGTCAGAGGYCAATCAGG CATAGRTCTATTTTGGTGTCTTC

H3-4 GATGACCAAATTGAGGTGACCAA CATTATTTGGCATAGTCACATTCA

H4-1 ATAGCTGGRTTCATAGAGAATGG AGCAAAAAGCATGATATGGAGAA

H4-2 ACAAATGAGAAATACCACCA CCCACAAAATAAAGGCTAAAA

H5 TACGCTGCAGACAAAGAATCCACTC ATGATTGCCAGTGCTAGGGAACTC

H6 GGACATACAATGCTGAACTKYTGGTT GCATTGAACCATTTGARCACATCC

H7-1 TTCAATGGGGCATTCATAGC GTGTTGTTCCTTATGCTCTCCAT

H7-2 GTAGAAGTTGTCAATGCAAC CCATATAGTTTGGTCTGYTC

H7-3 AGATCAGGATCTTCATTCTATGC ACCTTCCCATCCATTTTCAAT

H7-4 TAATTATTGAGAGGCGAGAAGGA CTATTTATGTTCTGAAAGGGCAA

H8 TGGTTGACCAAAAAGAAACCAGA TGCCATTCCTGTTCCCTCTGA

H9 GCTGGATTCATAGARGGAGGTTGG GATGAGGCGACAGTCGAATAAAT

H10 ATAGAAAATGGATGGGAAGGRATGGT TACAGATTGTGCATCGCATGTT

H11-1 GATATGCTTTTGAGATTGTCTC CTATTTCACTGAATTCGTGTTG

H11-2 TATGTAGCAGTGGGTTCAGA CCAGCTATTGCACCAAACAG

GGCAATAGACAATATGCAGAACAAACT CCGAAAATGAAACCCCCAATAATC H12 H13 ATGCTTGCAGACGAAATAGATGA ACTGCTTGCAATGCARCTGTA

H14 TACAAGGTGGCAACAGGGAGAGT CCTTCAGCATTTTGGTGCCTGAA

H15 ACTTTTACCTTCAATGGTGCATT ATGCTCTCCATACATTGATCGTC

H16 TGGCYAAATGCAACACAAAATG TCCATRCATGAGTCATTACATTT

Table 2.2. Detailed information for each primer about the amplified regions of HA gene, the length of the primer sequences, Tm values, GC content, and the expected PCR product sizes.

Forward Reverse

Conserved GC Conserved Length GC Exp. PCR Length T T region on m region on m (bp) (0C) (0C) HA gene (%) HA gene (bp) (%) Product

H1 79-100 22 68.24 50 733-756 24 68 54 677 bp

H2 1150-1172 23 65 43 1640-1660 21 66.97 48 509 bp

H3-1 1083-1102 20 58.61 40 1476-1498 23 58.38 26 416 bp

H3-2 1167-1189 23 66.5 48 1623-1645 23 65.23 43 479 bp

H3-3 711-728 18 63.37 61 1285-1307 23 58.17 39 597 bp

H3-4 168-190 23 66.8 43 568-591 24 62.3 33 491 bp

H4-1 1063-1085 23 62.04 43 1618-1640 23 63.46 35 578 bp

H4-2 1223-1242 20 56.84 35 1652-1672 21 59.5 33 450 bp

H5 1144-1168 25 69.18 48 1633-1656 24 68.18 50 513 bp

H6 1310-1335 26 66.89 42 1661-1684 24 70.17 46 375 bp

H7-1 766-785 20 64.54 45 1462-1484 23 63.03 43 719 bp

H7-2 145-164 20 54.63 40 616-635 20 56.21 45 491 bp

H7-3 466-488 23 60.09 39 1078-1098 21 63.07 38 633 bp

H7-4 281-303 23 62.69 39 901-923 23 60.8 35 643 bp

H8 512-534 23 66.35 39 1136-1156 21 69.03 52 645 bp

H9 1033-1056 24 69.44 54 1579-1601 23 65.05 43 569 bp

H10 1053-1078 26 69.43 42 1665-1686 22 65.21 41 634 bp

H11-1 832-853 22 56.95 36 1257-1278 22 58.73 36 447 bp

H11-2 669-688 20 58.26 45 1062-1081 20 63.17 50 413 bp

H12 1174-1200 27 68.55 41 1651-1674 24 68.35 42 501 bp

H13 1305-1327 23 62.60 39 1641-1661 21 65.84 48 357 bp

H14 619-641 23 67.81 52 1096-1118 23 70.87 48 500 bp

H15 778-800 23 62.30 35 1501-1523 23 63.94 43 746 bp H16 896-917 22 69.45 41 1489-1511 23 62.93 35 616 bp

Table 2.3. Analyses of the primers for secondary (2o) structure and primer dimer formation. All these analyses were performed at http://www.sigma-genosys.com/calc/DNACalc.asp

Forward Reverse Forward and Reverse

2o Primer Primer Primer 2o structure 2o structure structure dimer dimer dimer

H1 None No None No Moderate No

H2 None No Very weak No Weak No

H3-1 Weak No Weak No Weak No

H3-2 None No Weak No Moderate No

H3-3 None No None No Very weak No

H3-4 Weak No None No Moderate No

H4-1 Very weak No Very weak No Weak No

H4-2 None No None No None No

H5 None No Very weak No Moderate No

H6 Very weak No None No Moderate No

H7-1 Weak No Very weak No Moderate No

H7-2 Moderate No Weak No Moderate No

H7-3 Weak No None No Moderate No

H7-4 None No None No Weak No

H8 Moderate No None No Moderate No

H9 None No Weak No Weak No

H10 None No Weak No Moderate No

H11-1 Moderate No Weak No Moderate No

H11-2 None No Very weak No Moderate No

H12 Weak No None No Weak No

H13 None No Weak No Strong No

H14 None No Moderate No Strong No

H15 Weak No Very weak No Moderate No

H16 None No Very weak No Weak No

3.2. Sensitivity of the primers

The sensitivity of the primers was tested with the positive viral nucleic acids. Each viral

RNA was diluted with RNase-free water in 10-fold dilutions up to 10-10. Each dilution was subjected to RT-PCR amplification with specific primer sets to determine the sensitivity.

According to the sensitivity results of these primers, the majority of the primer pairs were able to amplify dilutions of 10-4 or higher of the stock viral RNAs (Table 2.4). In fact, a few of them were sensitive down to a few viral particles (e.g. H4-1, H9, H12, H13). The sensitivity of a few primers was only at 10-2 dilutions of the stock RNA. This might have been caused by lower than expected original concentration of stock viral nucleic acids or degradation of RNA during transport or processing in the laboratory. However, overall, the majority of the primers were 100 fold or more sensitive than the hemagglutination assay which can detect down to approximately

106 viral particles.

3.3. Testing for the cross-reactivity

To avoid cross amplification of the viral RNAs of different subtypes, each primer set was tested with the RNAs of the other 15 subtypes. No amplification was observed in any of the RT-

PCR attempts using other viral nucleic acids as template rather than the specified subtype.

Therefore, no cross-amplification occurred with any combination of primers and viral subtypes.

3.4. Development of multiplex RT-PCR conditions

Due to the large set of primer pairs used in this study, multiplex RT-PCR conditions were optimized to reduce preparation time and to increase the possibility of recovering viral RNAs of influenza A from low titer environmental samples. As explained in the previous section, primers were grouped into four major groups based on their annealing temperatures at 53, 56, 58 and 60 oC (Fig. 2.1). Prior to testing with the viral nucleic acids, these four groups of primers were subjected to RT-PCR without using a template to see if there is any self-priming to each other.

According to these amplifications, no amplicon was observed due to self priming of the primers used together in a certain group.

In order to confirm that each expected amplicon of different subtypes can be detected in one- step multiplex reactions, each group of primers was tested using known viral nucleic acids in

RT-PCR amplifications with a combination of all primer sets in each group. RT-PCR products amplified by all four groups in multiplex reactions exhibited amplicons of the predicted sizes.

The amplicons obtained by Group 3 and 4 primers are shown in Fig. 2.2. Since some amplicons are almost the same size, the resolution of these products was challenging using 1% agarose gels.

Due to this obstacle, the number of the bands observed on the gels was less than the number of the bands expected for these groups as shown in Fig. 2.1.

Table 2.4. The sensitivity of the primers determined by the 10-fold dilutions of positive viral nucleic acids.

RNA Highest RNA primers Subtype concentration sensitivity samples (ng/µl) detected

H1 Wisconsin H1N1 A/PR/8/34 265 10-6

H2 Wisconsin H2N2 A/Japan/305/57 64 10-4

H3-1 St. Jude H3N8 A/duck/Hokkaido/33/1980 N/A 10-2

H3-2 Wisconsin H3N8 A/equine/Miami/63 87 10-2

H3-3 Wisconsin H3N8 A/equine/Miami/63 87 10-4

H3-4 St. Jude H3N1 A/blue-winged teal/ALB/452/1983 N/A 10-4

H4-1 Wisconsin H4N6 A/duck/Czech/56 136 10-10

H4-2 St. Jude H4N1 A/mallard/ALB/47/1998 N/A 10-2

H5 Wisconsin H5N3 A/tern/South Africa/61 93 10-4

H6 Wisconsin H6N2 A/turkey/MA/65 377 10-6

H7-1 St. Jude H7N1 A/mallard/Alberta/34/2001 N/A 10-2

H7-2 St. Jude H7N7 A/duck/Heinersdorf/S495/6/86 N/A N/D

H7-3 Wisconsin H7N7 A/equine/Prague/56 78 10-2

H7-4 St. Jude H7N3 A/mallard/Netherlands/12/2000 N/A 10-4

H8 Wisconsin H8N4 A/turkey/Ontario/6118/68 145 10-4

H9 Wisconsin H9N2 A/turkey/WI/66 157 10-10

H10 Wisconsin H10N8 A/chicken/N/Germany/49 616 10-4

H11-1 Wisconsin H11N9 A/duck/Memphis/546/74 100 10-2

H11-2 St. Jude H11N9 A/mallard/Alberta/245/2003 N/A 10-4

H12 Wisconsin H12N5 A/duck/Alberta/60/76 65 10-9

H13 Wisconsin H13N6 A/gull/MD/707/77 59 10-9

H14 Wisconsin H14N5 A/mallard/Gurjev/263/82 88 10-6

H15 St. Jude H15N8 A/duck/Australia/341/83 N/A 100

H16 St. Jude H16N3 A/shorebird/DE/72/06 N/A N/D

*N/A- not applicable; N/D- not detected

Development of the hemagglutinin subtype-specific primers and the optimization of these into multiplex RT-PCR reactions decreased the workload; while it increased the possibility of detecting viruses at low titers in the samples. That is, only four reactions are needed rather than twenty-four. Also, since the primer pair to sample ratio is higher than in single primer pair reactions, there is a greater chance of detecting rare molecules. This is very helpful when dealing with the environmental ice samples in which the expected viral titers are lower than the biological specimens collected from birds directly.

Figure 2.2. Multiplex RT-PCR products amplified using Group 3 and Group 4 primers (Group 3 primers: H7-1: 719 bp, H8: 645 bp, H9: 569 bp, H10: 643 bp, H12: 501 bp, H13: 357 bp, H14:

500 bp; Group 4 primers: H1: 677 bp, H2: 509 bp, H3-2: 479 bp, H5: 513 bp, and H6: 375 bp).

We have demonstrated a method that provides high sensitivity and specificity. It is especially useful for amplification of influenza A RNA in dilute samples, as well as in samples where multiple subtypes are present. Sensitivities to a few particles are possible. While hemagglutinin subtype can be inferred from the amplicon sizes and primers used, sequencing is needed to confirm the exact identity of the HA gene.

4. Literature cited

Di Trani, L., B. Bedini, I. Donatelli, L. Campitelli, B. Chiappini, M.A. De Marco, M.

Delogu, C. Buonavoglia, and G. Vaccari. 2006. A sensitive one-step real-time PCR for

detection of avian influenza viruses using a MGB probe and an internal positive control.

BMC Infect. Dis. 6: 87

Fouchier, R.A.M., T.M. Bestebroer, S. Herfst, L. Van Der Kemp, G.F.

Rimmelzwaan, and A.D.M.E. Osterhaus. 2000. Detection of influenza A viruses from

different species by PCR amplification of conserved sequences in the matrix gene. J. of

Clinical Microbiology, 38 (11): 4096-4101.

Hoffmann, E., J. Stech, Y. Guan, R.G. Webster, and D.R. Perez. 2001. Univrsal

primer set for the full-length amplification of all influenza A viruses. Archives of

Virology, 146: 2275-2289.

Lee, M.-S., P.-C. Chang, J.-H. Shien, M.-C. Cheng, and H.K. Shieh. 2001.

Identification and subtyping of avian influenza viruses by reverse transcription-PCR.

Journal of Virological Methods, 97: 13-22.

Phipps, L.P., S.C. Essen and I.H. Brown. 2004. Genetic subtyping of influenza A

viruses using RT-PCR with a single set of primers based on conserved sequences within

the HA2 coding region. Journal of Virological Methods, 122: 119-122.

Rimmelzwaan, G.F., M. Baars, E.C.J. Claas, and A.D.M.E. Osterhaus. 1998.

Comparison of RNA hybridization, hemagglutination assay, titration of infectious virus

35 and immunofluorescence as methods for monitoring influenza virus replication in vitro.

Journal of Virological Methods, 74: 57-66.

Suarez, D.L., A. Das, and E. Ellis. 2007. Review of rapid molecular diagnostic tools for avian influenza virus. Avian Diseases, 51: 201-208.

Thontiravong, A., S. Payungporn, J. Keawcharoen, S. Chutinimitkul, S.

Wattanodorn, S. Damrongwatanapokin, A. Chaisingh, A. Theamboonlers, Y.

Poovorawan, and K. Oraveerakul. 2007. The single-step multiplex reverse transcription-polymerase chain reaction assay for detecting H5 and H7 avian influenza A viruses. Tohoku J. Exp. Med., 211: 75-79.

Wei, H.-L., G.-R. Bai, A.S. Mweene, Y.-C. Zhou, Y.-L. Cong, J. Pu, S. Wang, H.

Kida, J.-H. Liu. 2006. Rapid detection of avian influenza virus A and subtype H5N1 by single step multiplex reverse transcription-polymerase chain reaction. Virus Genes, 32:

261-267.

Zhang, G., D. Shoham, D. Gilichinsky, S. Davydov, J.D. Castello, and S.O. Rogers.

2006. Evidence of influenza A virus RNA in Siberian lake ice. Journal of Virology, 80:

12229-12235.

CHAPTER 3. EFFECTS OF FREEZING AND THAWING ON THE VIABILITY OF

INFLUENZA A VIRUSES

1. Abstract

Freezing and thawing present challenges to the retention of viability in viruses such as influenza

A. The ability to survive these processes has distinct advantages for these viruses in the environment. Viability has been assumed to decrease with each freeze-thaw cycle. Also, freezing at -80°C and storing samples at these temperatures are believed to preserve viability more than freezing and storing at -20°C. To test these assumptions, five low pathogenic strains of influenza

A viruses were seeded into pond water samples and subjected to five freeze-thaw cycles either at

-20oC or at -80oC, and were tested for viability after each freeze-thaw cycle by inoculating into fertilized chicken eggs. Hemagglutination assays using chicken red blood cells and RT-PCR

(reverse transcription-polymerase chain reaction) indicated steady rates of viability through all five freeze-thaw cycles at -20oC and -80oC. These results suggest that freezing and thawing have little or no effect on the viability of influenza A viruses at either temperature. This indicates that influenza A viruses are capable of survival after entrapment in environmental ice, thus rendering them potentially infectious after melting from pond, lake or other environmental ice.

2. Introduction

Microbes face many challenges due to a variety of environmental conditions, including exposure to solar (ultraviolet) irradiation, desiccation, and temperature extremes. When microbes are exposed to freezing and thawing, the phase changes and crystal formation of the water molecules can cause disruption of many classes of biomolecules, including those that form

37 integrated membranes. Under environmental conditions, microbes often are exposed to multiple freeze-thaw cycles. Their survival may be threatened, especially during the transition phase from liquid water to ice and back to liquid. Microbes that can survive multiple freeze-thaw cycles have a selective advantage over those that perish under these conditions. Influenza A viruses are surrounded by a portion of the host membrane as they exit the host cells. When they are excreted from the hosts, the membrane-surrounded virions often are exposed to multiple freeze-thaw cycles, especially when they are excreted into bodies of water in temperate, subarctic and arctic regions. The viral envelope must remain intact in order for the viruses to maintain infectivity.

Influenza A viruses are enveloped, negative sense ssRNA viruses belonging to the

Orthomyxoviridae. The genome of the virus is approximately 13 kb long and contains 8 segments of RNA encoding 10 proteins. Due to the lack of proofreading activity of RNA polymerase enzyme and to the segmented pattern of the viral RNA genome, the influenza A genome undergoes a rapid genetic change, as do all RNA viruses. Although these changes occur in all genes encoded by the genome, the major changes occur in hemagglutinin (HA) and neuraminidase (NA) genes that encode for surface glycoproteins responsible for antigenicity and host cell recognition. So far, 16 subtypes have been detected with respect to the differences in

HA surface antigen, while 9 subtypes have been identified according to NA surface antigen.

The natural reservoirs of influenza A viruses are wild aquatic birds for which viral infection is mainly of low pathogenicity, generally asymptomatic, waterborne and intestinal. It is primarily transmitted via a fecal-oral route through water among waterfowl. The waterborne transmission of influenza A viruses has already been shown by many researchers (Hinshaw et al.,

1979; Webster et al., 1992; Markwell and Shortridge, 1982; Stallknecht et al., 1990a; Stallknecht et al., 1990b; Ito et al., 1995; Brown et al., 2007). The possibility of environmental ice serving as

38 an abiotic reservoir for these viruses has been proposed by some researchers, although little research has been performed. Environmental ice is an ideal matrix for long-term protection of organisms due to the limitation of degradative processes; light, free water and heat (Rogers et al.

2004, Castello and Rogers (ed.), 2005; Smith et al. 2004). It has been shown that influenza A H1

RNA was preserved in Siberian lake ice (Zhang et al., 2006) and virus RNA of various subtypes was recovered from frozen Alaskan lake sediment samples (Lang et al., 2008). It is also well known that the survivability of many types of viruses increases at cold temperatures and below freezing. When water temperatures are warm (22-25oC), 99.9 % of the virus population is inactivated within a week, while it takes several weeks to months if the temperature decreases, and this trend continues to well below freezing for many types of viruses (Parker and Martel,

2002, Kamps et al, 2006). It has also been shown for influenza A that decreasing temperature stabilizes the lipids in the viral envelope thereby increasing the chance of virus survival at low temperatures (Polozov et al., 2008).

Although environmental ice might serve as a preservation reservoir for influenza A viruses, the freeze-thaw cycles occurring in the lake surface ice due to temperature fluctuations during winter and early spring could have adverse effects on the survival of these viruses.

Additionally, it has generally been assumed that influenza A viruses survive better when frozen at -80°C (a temperature restricted primarily to extreme polar zones) than at -20°C (a common surface temperature at the surface of higher latitude lakes in the winter). In this study, the effects on the viability of influenza A virions through multiple freeze-thaw events were examined at both temperatures. We hypothesize that the influenza A viruses are able to survive through several freeze-thaw cycles in natural environmental conditions throughout the winter and early spring. Natural pond water was seeded with five strains of influenza A virions, and then was

39 exposed to five freeze-thaw cycles at -20°C and -80°C, followed by viral culture, hemagglutination and RT-PCR (reverse transcription PCR) assays.

3. Materials and Methods

3.1. Viral strains

Low-pathogenic influenza A strains were obtained from Dr. Robert G. Webster, Division of Virology, Department of Infectious Diseases, St. Jude Children‟s Research Hospital,

Memphis, TN. The strains used to determine the effects of the freeze-thaw process on the viability of the viruses are as follows: H2N2 (A/Blue-winged teal/Alberta/604/78), H3N2

(A/Mallard/Alberta/85/76), H5N2 (A/Mallard/Alberta/7/76), H6N2 (A/Mallard/Alberta/98/85), and H7N1 (A/Mallard/Alberta (Eden)/28/92).

3.2. Water sampling and inoculation with the virus mixture

A pond water sample was collected on May 15, 2008 from the Rec Center pond (41.38o

N, 83.63o W) at Bowling Green State University, Bowling Green, Ohio campus. A mixture of five subtypes of low pathogenic influenza A strains was prepared by adding 0.1 ml of each virus to a final volume of 0.5 ml. The pond water sample was seeded with this virus mixture at a 1/10 dilution. One half of the seeded pond water was subjected to five freeze-thaw cycles at -80oC and the other half was subjected to five freeze-thaw cycles at -20oC. Thawing was carried out at 4°C in each case. Viral culture was performed before and after each freeze-thaw cycle. The freeze- thaw experiment was replicated 3 times at -80oC and 2 times at -20oC to confirm the results.

3.3. Viral culture

Fresh embryonated specific pathogen free (SPF) chicken eggs (0-day old) (Charles River

Laboratories International, Inc., Wilmington, MA) were incubated at 37oC with 72% humidity for 10 days, tilting the eggs every hour using Humidaire incubator/hatcher, Model 21 (Humidaire

Incubator Company, New Madison, OH). Embryo development was observed each day by candling. Non-developing embryos were discarded. In order to eliminate the risk of bacterial contamination, an antibiotic mixture was prepared at a final concentration of 5 mg gentamicin sulfate, 40,000 units streptomycin sulfate, 20,000 units polymixin B and 200,000 units penicillin

G (potassium salt) in 1 ml concentrated stock solution. From this antibiotic stock solution, 0.5 ml was added to 50 ml 1X sterile PBS to be used during the injection procedure. Ten-day old embryonated SPF chicken eggs were injected with 0.1 ml of the seeded water sample together with 0.1 ml of the antibiotic/PBS solution per egg through the allantoic cavity. In total, 15 eggs were inoculated per freeze-thaw cycle for -80oC and 9 eggs were inoculated for -20oC.

o Inoculated eggs were incubated at 35 C (without CO2) for 3 days. Embryo death was monitored for each day of incubation by candling. Since virus growth takes approximately 48 hrs, embryos that were dead at the end of the first day of incubation were discarded. At the end of the incubation period, the eggs were chilled overnight at 4oC before harvesting.

Allantoic fluids were collected from eggs for further analysis. Blood agar plates (10 g/l meat extract, 10 g/l peptone, 5 g/l NaCl, 15 g/l agar, and 5% defibrinated sheep blood; pH 7.3) were inoculated immediately after harvesting with allantoic fluids and incubated at 35oC for 3 days to test for bacterial contamination in the samples. An aliquot of 100 µl from each allantoic sample was transferred into 96-well microtiter plates for hemagglutination assays. From these,

50 µl of the sample was transferred to next row of wells to make 2-fold dilutions of these

41 samples in 1X sterile PBS. A small amount (50 µl) of previously prepared 0.5 % chicken red blood cells (CRBC) (Charles River Laboratories International, Inc., Wilmington, MA) was added to each well and incubated at room temperature for 30 min. Hemagglutination titers were determined at the end of the incubation period based on the agglutination of CRBCs in the presence of viruses.

3.4. RNA Extraction and RT-PCR amplifications

Viral RNAs were extracted from allantoic fluids of each chicken egg separately using a

QIAamp Viral RNA Kit (QIAGEN, Valencia, CA) according to the manufacturer‟s instructions.

Extracted viral nucleic acids were analyzed by RT-PCR based on amplification of part of the hemagglutinin (HA) and matrix (M) genes. The HA and M primers used for the amplification of low pathogenic influenza A strains are given in Table 3.1. Based on their annealing temperatures

(56oC and 60oC), HA primers were used in two multiplex reactions, while the amplifications using M-primers were performed at 56oC separately. The positive controls used in the hemagglutinin amplifications are the positive strains used in the experiment, while sterile distilled water was used as a negative control. RT-PCR amplifications were performed using a

GeneAmp EZ rTth RNA PCR Kit (Applied Biosystems, Branchburg, NJ). The composition of each 25 µl reaction was as follows: 1X EZ Buffer, 300 µM of each dNTPs, 2.5 units of rTth

DNA Polymerase, 2.5 mM Mn(OAc)2, 0.45 µM of each primer and 1 µl of each viral RNA extract. Reverse transcription was performed at 60oC for 45 min followed by PCR amplification of the cDNA at 95oC for 4 min and 40 cycles of 94oC for 1 min, 56 or 60oC for 2 min, 72oC for 2 min. Final extension was performed at 72oC for 10 min and cooled to 4oC. RT-PCR products were subjected to electrophoresis on 1% agarose gels in TBE (89 mM Tris-base, 89 mM borate,

2 mM EDTA, pH 8.0), containing 0.5 µg/ml ethidium bromide. After electrophoresis, gels were visualized using UV irradiation, and digitally photographed.

Table 3.1. Hemagglutinin and matrix primers used to amplify five low pathogenic influenza A strainsa

Product Annealing Primer Forward (5’-3’) Reverse (5’-3’) size temperature (bp)

H2 TATGCAGCAGACAAAGAATCCAC CYATCATGATTGCCAGTGACA 60oC 509

H3-4 GATGACCAAATTGAGGTGACCAA CATTATTTGGCATAGTCACATTCA 56oC 491

H5 TACGCTGCAGACAAAGAATCCACTC ATGATTGCCAGTGCTAGGGAACTC 60oC 513

H6 GGACATACAATGCTGAACTKYTGGTT GCATTGAACCATTTGARCACATCC 60oC 375

H7-4 TAATTATTGAGAGGCGAGAAGGA CTATTTATGTTCTGAAAGGGCAA 56oC 643

M* CTTCTAACCGAGGTCGAAACGTA GGATTGGTCTTGTCTTTAGCCA 56oC 147

a H2-H5-H6 primers were designed by G. Zhang (unpublished), H3-4 and H7-4 primers were designed by

Z. Koçer. *Previously designed matrix primers were used to amplify the matrix gene (DiTrani et al., 2006).

4. Results

According to the hemagglutination assay results shown in Fig. 3.1, no substantial differences in virus viability were observed among the freeze-thaw cycles. The mean hemagglutination titers were almost the same for the samples inoculated prior to freezing as for the samples after five freeze-thaw cycles. In fact, virus titers were slightly higher at the end of fifth cycle than initial values. The difference between freezing at -20°C and -80°C and thawing

43 was negligible in almost every case. Although it seems that there was a significant difference in the virus titer after freeze-thaw cycle 3 for -80°C, this resulted from a very high titer from a single egg in that sample group. This might have resulted from differences in the original chicken flock (genotype), differences in individual chicken eggs in terms of susceptibility to influenza A, or simply non-homogeneity in the injected samples. Virus titers returned to mean levels in freeze-thaw cycle 4, indicating an experimental anomaly in cycle 3. We consider this to be related to experimental variability rather than an effect from freezing and thawing.

6000

5000

4000

3000 -80°C 2000 -20°C

mean HA HA titermean (in 0.05 ml) 1000

0 BF FT-1 FT-2 FT-3 FT-4 FT-5 freeze-thaw cycles

Figure 3.1. The average hemagglutination titers recovered from subsequent freeze-thaw cycles.

BF is before freezing; FT-1, FT-2, FT-3, FT-4, and FT-5 are freeze-thaw cycles 1, 2, 3, 4, and 5, respectively. The lines designate the standard error values.

RT-PCR amplifications of the matrix and hemagglutinin genes were performed on the extracted viral RNA genome. As shown in Fig. 3.2A, the amplicons for the matrix gene obtained from each sample group were consistent at both temperatures. The viral RNA extracts were further tested by the amplification of hemagglutinin genes. As mentioned previously, HA amplicons were produced from two separate multiplex RT-PCR reactions, because of differences in the annealing temperatures of the primers. The strains H2N2, H5N2 and H6N2 were amplified in one reaction, while H3N2 and H7N1 strains were amplified in a separate reaction. According to these amplification results, H2N2, H3N2, H5N2 and H6N2 were successfully recovered in the freeze-thaw cycles, and almost all amplicons were of the same intensity for each sample group at both temperatures (Fig. 3.2B and 3.2C). No amplification was observed for H7N1 (Fig. 3.2B) suggesting that this certain strain did not result in a high infection rates in the chicken embryos.

It should be noted that this was also the case prior to freezing.

Figure 3.2. RT-PCR products amplified from viral nucleic acids extracted from the allantoic fluids. (A)

Viral RNA amplified with primers for the matrix gene (expected size of 147 bp). (B) Viral RNA amplified with HA primers for H3 and H7 (expected sizes were 491 and 643, respectively; H7 amplification was not detected). (C) Viral RNA amplified with HA primers for H2, H5, and H6

(expected sizes were 509, 513 and 375 bp, respectively). In each case, upper panel shows the amplifications from -80oC sample group, while the lower panel shows -20oC sample group. BF is before freezing; FT-1, FT-2, FT-3, FT-4, and FT-5 are freeze-thaw cycles 1, 2, 3, 4, and 5, respectively; “m” indicates the molecular weight standards; (+) and (-) designate positive and negative controls.

5. Discussion

Influenza A viruses are epidemiologically important pathogens due to their broad host range and their ability to cause pandemics and yearly epidemics. It is important to understand the effects of the variety of environmental parameters to fully understand the annual outbreaks and emergence of new strains. Many researchers have shown high rates of virus survival in lake and river water samples under varied environmental conditions (e.g. low temperatures, low salinity and slightly basic pH) (Hinshaw et al., 1979; Webster et al., 1992; Markwell and Shortridge,

1982; Stallknecht et al., 1990a; Stallknecht et al., 1990b; Ito et al., 1995; Brown et al., 2007;

Brown et al., 2009). Viral nucleic acids have been detected in the lake ice and lake sediment samples from Siberia and Alaska (Zhang et al., 2006; Lang et al., 2008). Here, we have demonstrated that influenza A virions are stable through multiple cycles of freezing and thawing under environmentally relevant conditions (i.e., freezing at -20°C).

Due to temperature fluctuations, temperate, subarctic, and arctic bodies of water usually go through several freeze-thaw cycles during the winter and early spring months. This process might affect the survivability of viruses unless they are resistant to the potentially damaging effects of freezing and thawing. If viruses such as influenza A can survive these processes, then they may be capable of infecting local and migrating animals that come into contact with the ice and melt water. We tested the effects of freezing and thawing on the survival of influenza A virions in order to understand the virus response to conditions common in natural settings. Five subtypes of low pathogenic influenza A strains were used to perform model freeze-thaw experiments at two different temperatures. Pond water was used instead of distilled water,

47 because this better mimics environmental conditions (eg. in terms of pH, salinity, and osmolarity).

A temperature of -80°C is rare on Earth, while -20°C is common, especially in higher latitudes and at high elevations. Thus, while the assumption has been that -80°C preserves this virus better than -20°C, this was unsupported by the results of this study. The amplicons for both genes obtained from each sample group supported the hemagglutination assay results by giving almost the same intensity of amplifications throughout subsequent cycles of freeze-thaw at both temperatures (Fig. 3.2A, 3.2B and 3.2C). Therefore, we conclude that viability of influenza A viruses is not affected by freeze-thaw events, at least up to five cycles at both temperatures.

Virus levels, as indicated by hemagglutination, and genotype, as indicated by RT-PCR, remain stable through all five cycles at -20°C and at -80°C.

Our results are confirmatory of the report that the membrane surrounding influenza A virions becomes more stable, and almost crystalline as the temperature is decreased (Polozov et al.

2008). The results indicate that several freezing and thawing events throughout the winter and early spring due to temperature fluctuations should not affect influenza A survival. This supports the possibility of environmental ice acting as an abiotic reservoir for viable influenza A virions that pose a potential risk for infection of local and migratory waterfowl visiting that area, along with other susceptible hosts (including humans). As a result, the infection risk hidden in the environmental ice and water may facilitate infection, as well as the emergence of new strains, and possibly new subtypes, following recombination and reassortment with other strains carried by the visiting animals. Therefore, environmental ice may act as a reservoir of viable influenza A virions. The meltwater can then become a source for mixed strains that are capable of infection of susceptible hosts.

6. Literature cited

Brown, J.D., D.E. Swayne, R.J. Cooper, R.E. Burns, and D.E. Stallknecht. 2007.

Persistence of H5 and H7 avian influenza viruses in water. Avian Diseases, 51: 285-289.

Brown, J.D., G. Goekjiana, R. Poulsona, S. Valeikab, and D.E. Stallknecht. 2009. Avian

influenza virus in water: Infectivity is dependent on pH, salinity and temperature. Vet.

Microbiol. 136:20-26.

Castello, J.D., and S.O. Rogers (ed). 2005. Life in Ancient Ice. Princeton Univ. Press,

Princeton, NJ.

Di Trani, L., B. Bedini, I. Donatelli, L. Campitelli, B. Chiappini, M.A. De Marco, M.

Delogu, C. Buonavoglia, and G. Vaccari. 2006. A sensitive one-step real-time PCR for

detection of avian influenza viruses using a MGB probe and an internal positive control.

BMC Infectious Disease, 6: 87

Hinshaw, V.S., R.G. Webster, and B. Turner. 1979. Water-borne transmission of influenza

A viruses? Intervirology, 11: 66-68.

Ito, T., K. Okazaki, Y. Kawaoka, A. Takada, R.G. Webster, and H. Kida. 1995.

Perpetuation of influenza A viruses in Alaskan waterfowl reservoirs. Archives of Virology,

140: 1163-1172.

Kamps, B.S., C. Hoffmann, and W. Preiser. 2006. Influenza Report 2006. Flying

Publisher.

Lang, A. S., A. Kelly, and J. Runstandler. 2008. Prevalence and diversity of avian influenza viruses in environmental reservoirs. Journal of General Virology, 89: 509-519.

Markwell, D.D., and K.F. Shortridge. 1982. Possible waterborne transmission and maintenance of influenza viruses in domestic ducks. Applied and Environmental

Microbiology, 43(1): 110-116.

Parker, L.V., and C.J. Martel. 2002. Long-term survival of enteric microorganisms in frozen wastewater. US Army Corps of Engineers, Engineer Research and Development

Center. ERDC/CRREL TR-02-16.

Polozov, I.V., L. Bezrukov, K. Gawrisch, and J. Zimmerberg. 2008. Progressive ordering with decreasing temperature of the phospholipids of influenza virus. Nature Chemical

Biology, 4: 248-255.

Rogers, S.O., W.T. Starmer, and J.D. Castello. 2004. Recycling of pathogenic microbes through survival in ice. Medical Hypotheses, 63(5): 773-777.

Smith, A.W., D.E. Skilling, J.D. Castello, and S.O. Rogers. 2004. Ice as a reservoir for pathogenic animal viruses. Medical Hypotheses, 63: 560-566.

Stallknecht, D.E., M.T. Kearney, S.M. Shane, and P.J. Zwank. 1990a. Effects of pH, temperature and salinity on persistence of avian influenza viruses. Avian Diseases, 34(2):

412-418.

Stallknecht, D.E., S.M. Shane, M.T. Kearney, and P.J. Zwank. 1990b. Persistence of avian influenza viruses in water. Avian Diseases, 34: 406-411.

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(1): 152-179.

Zhang, G., D. Shoham, D. Gilichinsky, S. Davydov, J.D. Castello, and S.O. Rogers.

2006. Evidence of influenza A virus RNA in Siberian lake ice. Journal of Virology, 80:

12229-12235.

CHAPTER 4. DETECTION OF INFLUENZA A VIRUSES FROM ENVIRONMENTAL

LAKE AND POND ICE

1. Introduction

Influenza A viruses are epidemiologically very important pathogens due to their broad host range, their ability to infect many species, and their ability to cause pandemics and yearly epidemics by avoidance of host immune systems through continuous changes in their genomes.

As mentioned in previous chapters, they are enveloped, negative sense ssRNA viruses belonging to the Orthomyxoviridae. The genome of the virus is approximately 13 kb long and contains 8 segments of RNA encoding 10 proteins. The segmented nature of the viral genome and lack of a proofreading function in the RNA polymerase allow the majority of rapid genetic changes, such as antigenic drift via point mutations, or antigenic shift via reassortment and recombination between gene segments. Avian influenza is subtyped based on two surface antigens, hemagglutinin and neuraminidase (Strauss & Strauss, 2002; Webster et al., 1992). The majority of genetic changes are observed in these two genes due to the fact that viruses can escape from host immunological recognition and this allows them to enter into the host cells and maintain the viral fitness (Earn et al., 2002; Kamps et al., 2006). The envelope they acquire from the host cell during budding gives them another protection layer while they are outside the host cell.

Due to tissue and host specificities of the receptors, influenza A has the ability to infect a broad range of host species, including waterfowl, migratory birds, shorebirds, poultry, swine, humans, sea and land mammals, and horses. However, wild aquatic birds are known as the natural reservoir for these viruses (Webster et al., 1992). Therefore, influenza A viruses can cross species barriers in some circumstances by modifying their surface antigens and changing the

52 receptor specificity. The presence of two different types of receptors, α-2-3 sialic acid and α-2-6 sialic acid, in some species (e.g. swine, quail, and chicken) facilitates the interspecies transmission of these viruses (Webster et al., 1992; Earn et al., 2002; Kamps et al., 2006). These intermediate species are known as biotic mixing vessels for the virus between human and avian strains; and biotic reservoirs help maintain viral diversity.

Due to the importance of avian influenza for causing human disease, and the fact that it causes more than 35,000 deaths annual in the US alone, research have been mainly focused on the biotic reservoirs. Although the RNA genome of the virus undergoes rapid genetic changes due to the reasons mentioned above, appearance and disappearance of some strains without any change over long periods of time (Smith et al., 2004; Shoham, 1993; Shoham, 2005) the possibility of abiotic reservoirs, such as water and ice, have been proposed.

Avian influenza causes low pathogenicity and usually asymptomatic infections in wild aquatic birds, and it is primarily transmitted via a fecal-oral route through water (Webster et al.,

1992). Transmission of the viruses through the water has been well documented (Hinshaw et al.,

1979; Webster et al., 1992; Markwell and Shortridge, 1982; Stallknecht et al., 1990a; Stallknecht et al., 1990b; Ito et al., 1995; Brown et al., 2007; Brown et al., 2009). However, little research has been performed to test the hypothesis that environmental ice serves as an abiotic reservoir for avian influenza. In fact, due to the limitation of degradative processes (such as light, free water and heat) environmental ice is an ideal matrix for long-term protection of organisms. It has already been shown that influenza A H1 RNA was preserved in Siberian lake ice (Zhang et al.,

2006) and several subtypes of virus RNA were recovered from Alaskan lake sediment samples

(Lang et al., 2008). It is also well known that virus survivability increases at cold temperatures and below freezing. Inactivation of 99.9 % of the virus population takes a week at warmer water

53 temperatures (22-25oC), while inactivation of the same amount of virus population takes several weeks to months if the temperature decreases (Parker and Martel, 2002). In addition, a decrease in temperature increases the stability of the lipids in the viral envelope allowing survival of the viruses at low temperatures (Polozov et al., 2008). Therefore, we hypothesize that influenza A viruses are able to survive the winter by being entrapped in environmental ice, such that they are capable of infecting naïve hosts after melting from the ice.

In this study, we sampled water and ice samples from lake and ponds during various seasons, and assayed these environmental samples to determine whether influenza A viruses maintain their viability in the ice. We inoculated environmental water and ice samples into embryonated chicken eggs, and monitored viral multiplication using hemagglutination assays and reverse transcription-PCR amplifications of the viral nucleic acid extracted from the allantoic fluids. This study is the first research focused on the detection of viable influenza A viruses from environmental ice samples.

2. Materials and Methods

2.1. Ultracentrifugation

2.1.1. Viral strains

Low-pathogenic influenza A strains were obtained from Dr. Robert G. Webster, Division of Virology, Department of Infectious Diseases, St. Jude Children‟s Research Hospital,

Memphis, TN. The strains used to determine the effect of ultracentrifugation on the viability of the viruses were as follows: H2N2 (A/Blue-winged teal/Alberta/604/78), H3N2

(A/Mallard/Alberta/85/76), H6N2 (A/Mallard/Alberta/98/85). The initial hemagglutination titers of these viruses were 2-10, 2-8 and 2-7, respectively.

2.1.2. Water sampling and inoculation with the virus

Seven bottles of pond water sample were collected from Ottawa National Wildlife

Refuge, OH on November 18, 2008 (41.63o N, 83.22o W). Water samples were inoculated with the virus(es) as follow: Bottle 1-no virus added in vitro; Bottle 2- H2N2 at 1 to 300 dilution;

Bottle 3- H2N2 at 1 to 150 dilution; Bottle 4- H2N2 at 1 to 60 dilution; Bottle 5- H3N2 and

H6N2 at a final dilution of 1 to 300; Bottle 6- H3N2 and H6N2 at a final dilution of 1 to 100;

Bottle 7- H3N2 and H6N2 at a final dilution of 1 to 50. All pond water samples were stored at -

20oC for 2 days to obtain evenly frozen samples. After 2 days, frozen pond water samples were transferred to 4oC subsequently for thawing under stable cold conditions.

2.1.3. Concentrating the viruses from frozen pond water

After thawing, frozen pond water samples were filtered through 1.2 µm glass fiber filters, then 0.45 µm and 0.22 µm Durapore membranes to eliminate the contamination risk and to obtain only viral size particles in the filtrates. Filters were changed for every 100 ml of water sample to avoid clogging of the filters. Each filtrate was centrifuged at 113,000 xg for 5 hrs at

4oC using sterile polyallomer centrifuge tubes. Ultracentrifugation was performed using 60Ti rotor - model 2601 at L8-70M ultracentrifuge (Beckman Coulter Inc, Brea, CA). Each pellet was resuspended in sterile 1X PBS and stored at -20oC until viral culture was performed. Samples were concentrated to 100 times of their initial concentrations.

2.1.4. Viral culture

Fresh embryonated specific pathogen free (SPF) chicken eggs (0-day old) (Charles River

Laboratories International, Inc., Wilmington, MA) were incubated at 37oC with 72% humidity for 10 days, tilting the eggs every hour using Humidaire incubator/hatcher, Model 21 (Humidaire

Incubator Company, New Madison, OH). Embryo development was monitored daily by candling and non-developing embryos were discarded. In order to eliminate the risk of bacterial contamination, an antibiotic mixture was prepared at a final concentration of 5 mg gentamicin sulfate, 40,000 units streptomycin sulfate, 20,000 units polymixin B and 200,000 units penicillin

G (potassium salt) in 1 ml concentrated stock solution. From this antibiotic stock solution, 0.5 ml was added to 50 ml 1X sterile PBS to be used during the injection procedure. Ten-day old embryonated SPF chicken eggs were injected with 0.1 ml of the seeded water sample together with 0.1 ml of the antibiotic/PBS solution per egg through the allantoic cavity. Five chicken eggs were used for each water sample. Allantoic fluids collected at the end of incubation were transferred to another set of eggs up to a 3rd passage through their allantoic cavities. Between each passage, allantoic samples were stored at -80oC. Inoculated eggs were incubated at 35oC

(without CO2) for 3 days. Embryo death was monitored for each day of incubation by candling.

Embryos that were dead at the end of the first day of incubation were discarded based on the fact that virus growth takes approximately 48 hrs. At the end of incubation period, SPF chicken eggs were chilled overnight at 4oC.

56 sample was transferred into 96-well microtiter plates for hemagglutination assays. From these,

50 µl of the sample was transferred to next row of wells to make 2-fold dilutions of these samples in 1X sterile PBS. A small amount (50 µl) of previously prepared 0.5 % chicken red blood cells (CRBC) (Charles River Laboratories International, Inc., Wilmington, MA) was added to each well and incubated at room temperature for 30 min. Hemagglutination titers were determined at the end of the incubation period based on the agglutination of CRBCs in the presence of viruses.

2.1.5. RNA Extraction and RT-PCR

Viral RNAs were extracted from allantoic fluids of each chicken egg separately using a

QIAamp Viral RNA Kit (QIAGEN, Valencia, CA) according to the manufacturer‟s instructions.

Extracted viral nucleic acids were analyzed by RT-PCR based on amplification of part of the hemagglutinin (HA) gene. The specific HA primers H2, H3-4 and H6 were used for the amplification of low pathogenic influenza A strains H2N2, H3N2 and H6N2 (please refer to

Table 2.1 in Chapter 2 for the primer sequences). Based on their annealing temperatures, H2 and

H6 primers were used in one multiplex reaction at 60 oC, while the amplifications of H3N2 using

H3-4 primers were performed at 56 oC, separately. RT-PCR amplifications were performed using a GeneAmp EZ rTth RNA PCR Kit (Applied Biosystems, Branchburg, NJ). The composition of each 25 µl reaction was as follows: 1X EZ Buffer, 300 µM of each dNTPs, 2.5 units of rTth

72oC for 2 min. Final extension was performed at 72oC for 10 min and cooled to 4oC. RT-PCR products were subjected to electrophoresis on 1% agarose gels in TBE (89 mM Tris-base, 89

57 mM borate, 2 mM EDTA, pH 8.0), containing 0.5 µg/ml ethidium bromide. After electrophoresis, gels were visualized using UV irradiation, and digitally photographed.

2.2. Environmental Samples

2.2.1. Sampling sites

Currently, six different environmental ice and water samples have been analyzed. These samples were obtained from the Ottawa National Wildlife Refuge (NWR), the BGSU Golf

Course pond and the Rec Center pond. The sample details are given in Table 4.1. Environmental water and ice samples were stored at -80oC until they were processed.

Table 4.1. Sampling sites, sample type, sampling date, concentration, number of SPF eggs inoculated and number of passages for viral culture (OttNWR: Ottawa National Wildlife Refuge;

MS 4 and MS 5 are ponds within OttNWR).

Latitude/ Number Original Sampling Concentrated Number of eggs Location longitude of source date by inoculated/passage passages BGSU 41.38° N (Golf 83.62° W Water Jan, 2008 29X 10-10-10-9-9 5 course) BGSU 41.38° N (Rec 83.63° W Water Jan, 2008 30X 10-10-10-9-9 5 Center) OttNWR- 41.63° N Water Feb, 2008 27X 10-10-10-9-7 5 MS 4 & 5 83.22° W OttNWR- 41.63° N Ice with Feb, 2008 27X 10-10-10-9-9 5 MS 4 & 5 83.22° W feces OttNWR- 41.63° N Water Nov,2008 100X 5-8-9-11-12-12 6 MS 5 83.22° W OttNWR- 41.63° N March, Ice 250X 29-29 2 MS 5 83.22° W 2009

2.2.2. Concentrating the samples

Environmental frozen water and ice samples were held at 4oC for thawing under stable cold conditions. The melting procedure was performed under sterile conditions. Each meltwater sample was divided into sterile polyallomer centrifuge tubes and concentrated by ultracentrifugation at 113,000 xg at 4oC for 5-10 hours/run. Ultracentrifugation was performed using 60Ti rotor - model 2601 at L8-70M ultracentrifuge (Beckman Coulter Inc, Brea, CA).

Depending on the initial volume of the sample, several runs of ultracentrifugation were performed until the entire sample was concentrated. Final concentrations of each sample are given in Table 4.1. At the end, pellets were resuspended in sterile 1X PBS and stored at -20oC until they were injected into chicken eggs.

2.2.3. Viral culture

Viral culture from each sample set was performed as outlined previously. Ten-day old

SPF embryonated chicken eggs were inoculated with the concentrated samples along with antibiotic/PBS mix through their allantoic cavities. Ten to thirty chicken eggs were used for each water sample. Allantoic fluids collected at the end of incubation were transferred into another set of eggs. The number of inoculated eggs and the number of subsequent passages for viral culture for each sample set is shown in Table 4.1. Between each passage, allantoic samples were stored

o o at -80 C. Inoculated eggs were incubated at 35 C (without CO2) for 3 days. At the end of the incubation period, SPF chicken eggs were chilled overnight at 4oC. Hemagglutination assays and blood agar tests for bacterial contamination were performed from allantoic fluids as described above.

2.2.4. RNA Extraction, RT-PCR, Cloning and Sequencing

Viral RNAs were extracted from allantoic fluids of each chicken egg separately using a

QIAamp Viral RNA Kit (QIAGEN) according to the manufacturer‟s instructions. Extracted viral nucleic acids were analyzed by RT-PCR assay based on amplification of part of the hemagglutinin (HA) and matrix (M) genes. For the hemagglutinin amplification, previously developed subtype-specific primers were used in four multiplex reactions at different annealing temperatures as explained in Chapter 2. Hemagglutinin subtype-specific primers are given in

Table 2.1 (Chapter 2). For the amplification of matrix gene, a primer set developed by Di Trani et al. (2006) was used at an annealing temperature of 56 oC. The composition of RT-PCR reactions and the thermocycler conditions were described previously. PCR products were subjected to electrophoresis on 1% agarose gels with TBE and ethidium bromide (0.5 µg/ml).

After electrophoresis, gels were visualized using UV irradiation, and digitally photographed.

A TOPO TA Cloning Kit (Invitrogen Corporation, Carlsbad, CA) was used for the cloning of positive amplicons. The amplified products were ligated into a PCR2.1 TOPO vector according to manufacturer‟s instructions. For each ligation reaction, 4 µl of PCR product was used together with 1 µl of salt solution (1.2 M NaCl and 0.06 M MgCl2) and 1 µl of vector (10 ng/µl). Plasmids were transformed into host bacteria (One Shot ® TOP10 Competent E. coli cells) and grown on selective media. The clones were further analyzed by colony PCR to determine the positive clones. PCR amplifications were performed using a native Taq DNA

Polymerase (Fermentas Inc., Glen Burnie, MD). The composition of each 50 µl reaction was as

o follows: 1X Taq buffer with (NH4)2SO4 [750 mM Tris-HCl (pH 8.8 at 25 C), 200 mM

(NH4)2SO4, 0.1 % Tween 20], 200 µM of each dNTPs, 1.25 units of Taq DNA Polymerase, 1.5 mM MgCl2, 0.5 µM of M13 forward and reverse primers. Each clone was resuspended in this 50

µl reaction. PCR amplifications were carried out at 95 oC for 4 min; and 30 cycles of 94oC for 1 min, 45 oC for 2 min, and 72 oC for 2 min. Final extension was performed at 72 oC for 10 min and cooled to 4oC. PCR products were subjected to electrophoresis on 1% agarose gels in TBE

(89 mM Tris-base, 89 mM borate, 2 mM EDTA, pH 8.0), containing 0.5 µg/ml ethidium bromide. After electrophoresis, gels were visualized using UV irradiation, and digitally photographed.

Positive PCR products were purified using QIAquick PCR Purification Kit (QIAGEN,

Valencia, CA) according to manufacturer‟s instructions. Purified PCR products then were sent to

Geneway Research (Hayward, CA) for sequencing. Phylogenetic analyses were performed with

PAUP (Phylogenetic Analysis Using Parsimony; Swofford, 2001) using Maximum Parsimony and Neighbor Joining.

3. Results and Discussion

3.1. Effects of ultracentrifugation on the viability of influenza A viruses

Considering the possibility of low virus titers embedded in ice, we decided to concentrate the environmental samples. Ultracentrifugation is one method for this purpose. For this part of the project, we tested the effects of ultracentrifugation conditions on the viability of viruses.

Additionally, influenza A viruses are known to attach to solid particles in the water. Therefore, we also tested how filtering and ultracentrifugation would affect the viability of the virus in the pellet after these two processes were applied. For this purpose, 7 bottles of pond water samples were prepared in vitro using different subtypes of influenza A viruses at different concentrations.

We used 3 different subtypes (H2N2 individually; H3N2 and H6N2 in combination); since different subtypes might react differently in chicken egg cultures. Additionally, we analyzed

61 several different dilutions of these viruses in the pond water samples to determine the lowest detectable concentration of the viruses in environmental samples.

Results demonstrated that ultracentrifugation is an effective method for concentrating the viruses without causing adverse effects on the viability based on growth in chicken eggs and hemagglutination assays. At the end of the first passages of the samples through allantoic cavities of SPF chicken eggs, hemagglutination tests were negative for the lower concentrations, although we were able to detect the viruses by the amplification of HA genes using RT-PCR for a few samples. The detection rates for HA gene amplification increased by the end of 3rd passage of the samples through subsequent viral cultures in embryonated eggs (Fig. 4.1).

Figure 4.1. RT-PCR products amplified by H2 primers from allantoic fluids harvested after the

3rd passage of the samples. Expected amplicon size is 509 bp. As molecular marker, 100 bp ladder was used to monitor the amplicon size. Positive control was the nucleic acid extracted from known H2N2 used in the experiment.

Although the initial concentrations of viruses at different dilutions were not hugely different, hemagglutination and RT-PCR assays revealed a rapid loss of viral detection following dilution. The abnormally low detection rates at lower dilutions could be mainly related to two factors: the ability of the chicken embryo‟s immune system to inhibit viral replication at low concentrations; or decreasing the expected virus particles in the sample by filtering. In other words, some of viruses might have been eliminated on the filters if they were attached to debris in the pond water samples).

3.2. Assaying the environmental ice and water samples for viable influenza A virions

Six environmental water and ice samples collected from 3 different locations at different times were assayed for the presence of influenza A viruses (Table 4.1). Samples were concentrated by ultracentrifugation, without filtering, so that no viruses were lost by the filtering system, as mentioned in the previous section. The final concentrations of the samples varied depending on their initial volumes (Table 4.1). For instance, the sample concentrated by 250 times had an initial volume of 1500 ml, while the samples concentrated by ~30 times had initial volumes of approximately 80-150 ml. The number of subsequent viral cultures in embryonated chicken eggs for each sample is indicated in Table 4.1.

Although influenza A viruses could not be detected by hemagglutination assays following viral cultures, we were able to detect the viruses by the amplification of two different genes, the matrix and hemagglutinin (HA) genes. For HA gene amplification, the 24 influenza A subtype-specific primers developed previously (see Chapter 2) were used. The presence of viral nucleic acid was also confirmed by the amplification of the matrix gene, which is relatively more conserved among influenza A subtypes. The detection levels of the amplification products varied

63 depending on the initial concentration of injected sample into chicken embryos. The more concentrated the sample, the earlier the gene products were amplified when transferred through chicken eggs. For instance, for the sample concentrated by 250 times, the HA gene products were visible after the first viral culture passage (but not for the matrix gene amplicons). Once these samples were transferred to another set of chicken eggs for the second passage, we observed an increase in the amplification of matrix gene products (Fig. 4.2). This indicates that the number of virus particles increased due to replication in the host cell, which supports the viability of the virions in the original sample. If no replication occurred, the virions would have been diluted many times as it was passed into the next set of eggs. Therefore, amplification frequency would decrease with each passage. However, amplification frequencies increased for many samples, while no decrease was observed.

Figure 4.2. Matrix gene amplification from the first and second passages of allantoic fluids (red and yellow arrows show the positive and negative samples after cloning, respectively). Expected amplicon size is 147 bp. As molecular marker, 100 bp ladders were used to monitor the amplicon size. Red arrows indicate the samples from which positive clones were obtained; while no clone was obtained from the samples indicated by yellow arrows. Positive control was H1N1-VR1520 strain from ATCC, and negative control was distilled water.

The clone sequences obtained using matrix gene products were aligned with the original matrix gene sequences of each subtype retrieved from the NCBI influenza database. According to the phylogenetic relationships, the clones obtained from this study were closely related to other influenza A subtypes based on the matrix gene sequence. The clones of different sample sets (Fig. 4.3) grouped together, while they all came closest either to H1N1 and H5N2. This may indicate the most prevalent subtypes in the ponds at the time of sampling, although there is no strong correlation between the matrix genes and the hemagglutinin subtypes of influenza A.

Figure 4.3. Maximum parsimony phylogram of a broad selection of matrix gene sequences including the sequences obtained from different environmental samples (red: Ottawa NWR,

2/08-ice, 3rd passage; blue: Ottawa NWR, 11/08-water, 4th passage; green: Ottawa NWR, 03/09- ice, 2nd passage).

We also analyzed the hemagglutinin sequences cloned from the environmental samples.

So far, several sequences indicate subtypes of hemagglutinin genes, including H3, H4, H7 and

H11 (Fig. 4.4).

Figure 4.4. RT-PCR products amplified using Group 1 primers from the viral cultures of concentrated environmental samples. Group 1 primers and the expected amplicons are as follows: H3-1: 413 bp, H3-3: 597 bp, H4-2: 450 bp, H7-2: 491 bp, H11-1: 447 bp, H11-2: 416 bp and H16: 616 bp.

All of these sequences were aligned separately with the previously published sequences retrieved from the influenza database and the phylogenetic relationships were analyzed.

Although some sequences are shorter than expected, we were still able to group them after manual alignment. For instance, one of the H7 clones was aligned with the known HA-gene sequences of each subtype. Interestingly, among all these subtypes, this specific clone was grouped with H7 subtypes (Fig. 4.5), even though it was shorter than expected.

Figure 4.5. Neighbor-joining phylogram of the influenza virus hemagglutinin H7 sequence isolated from Ottawa NWR (ice collected in March, 2009).

The hemagglutination assay requires high concentration of virus for the successful detection of the virus presence in the sample (Rimmelzwaan et al., 1998). However, polymerase chain reaction amplification is a very sensitive technique for the detection of many organisms in environmental samples at low concentrations. As mentioned in Chapter 2, the HA primers used to amplify the hemagglutinin genes by RT-PCR are sensitive approximately to 104 virus particles, while a few primer sets were sensitive down to a few virus particles. On the other hand, the lowest detection level for hemagglutination assay is 106 virus particles/ml.

Influenza A viruses are epidemiologically very important pathogens due to their broad host range and their ability to cause pandemics and yearly epidemics. To date, research has been focused on the biotic reservoirs of these viruses. As mentioned before, the RNA genome of the virus is prone to rapid genetic changes. However, some strains have appeared, disappeared and reappeared intermittently, even decades later, almost without any change (Rogers et al., 2004;

Smith et al., 2004; Shoham, 1993; Shoham, 2005). This fact has raised the question whether these viruses can be preserved in abiotic reservoirs, such as water and ice. Many researchers have shown high rates of virus survival in lake and river water samples, especially at colder temperatures (4oC) (Hinshaw et al., 1979; Webster et al., 1992; Markwell and Shortridge, 1982;

Stallknecht et al., 1990a; Stallknecht et al., 1990b; Ito et al., 1995; Brown et al., 2007; Brown et al., 2009). Also, viral nucleic acid was detected in the lake ice and lake sediment samples from

Siberia and Alaska (Zhang et al., 2006; Lang et al., 2008). All of these findings are consistent with the possibility of environmental ice acting as an abiotic reservoir for influenza A viruses.

Our findings demonstrate that environmental ice and water samples serve as abiotic reservoirs for viable influenza A virions that pose the potential risk of infection to local and migratory waterfowl visiting that area, along with other susceptible hosts (including humans).

Once the ice thaws, the viable virions embedded in ice may mix with other subtypes during host infection. The mixed genotypes may yield novel gene combinations. As a result, this infection risk hidden in the environmental ice and water will facilitate the emergence of new strains, and possibly new subtypes.

This project is the first study on the detection of viable influenza A viruses from environmental ice samples. The data obtained from this project will shed light onto the importance of abiotic environmental sources, such as ice, as alternative mixing vessels for influenza A viruses and will provide additional information about the ecology and evolution of avian influenza to understand the potential origin of new pathogenic strains.

4. Literature cited

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Brown, J.D., G. Goekjiana, R. Poulsona, S. Valeikab, and D.E. Stallknecht. 2009.

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Vet. Microbiol. 136: 20-26.

Earn, D.J.D., J. Dushoff, S.A. Lewin. 2002. Ecology and evolution of flu. Trends in

Ecology and Evolution, 17(7): 334-340.

Hinshaw, V.S., R.G. Webster, B. Turner. 1979. Water-borne transmission of influenza

A viruses? Intervirology, 11: 66-68.

Ito, T., K. Okazaki, Y. Kawaoka, A. Takada, R.G. Webster and H. Kida. 1995.

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Virology, 140: 1163-1172.

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influenza viruses in environmental reservoirs. Journal of General Virology, 89: 509-519.

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Center. ERDC/CRREL TR-02-16.

Polozov, I.V., L. Bezrukov, K. Gawrisch and J. Zimmerberg. 2008. Progressive ordering with decreasing temperature of the phospholipids of influenza virus. Nature

Chemical Biology, 4: 248-255.

Rimmelzwaan, G.F., M. Baars, E.C.J. Claas, and A.D.M.E. Osterhaus. 1998.

Comparison of RNA hybridization, hemagglutination assay, titration of infectious virus and immunofluorescence as methods for monitoring influenza virus replication in vitro.

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Rogers, S.O., W.T. Starmer and J.D. Castello. 2004. Recycling of pathogenic microbes through survival in ice. Medical Hypotheses, 63(5): 773-777.

Shoham, D. 1993. Biotic-abiotic mechanisms for long-term preservation and reemergence of influenza type A virus genes. Prog. Medical Virology. 40: 178-192.

Shoham, D. 2005. Viral pathogens of humans likely to be preserved in natural ice, p.

208-226. In J.D. Castello and S.O. Rogers (ed.), Life in Ancient Ice. Princeton Univ.

Press, Princeton, NJ.

Smith, A.W., D.E. Skilling, J.D. Castello, and S.O. Rogers. 2004. Ice as a reservoir for pathogenic animal viruses. Medical Hypotheses, 63: 560-566.

Stallknecht, D.E., M.T. Kearney, S.M. Shane, and P.J. Zwank. 1990a. Effects of pH, temperature and salinity on persistence of avian influenza viruses. Avian Diseases, 34(2):

412-418.

Stallknecht, D.E., S.M. Shane, M.T. Kearney, and P.J. Zwank. 1990b. Persistence of avian influenza viruses in water. Avian Diseases, 34: 406-411.

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2006. Evidence of influenza A virus RNA in Siberian lake ice. Journal of Virology, 80:

12229-12235.

APPENDIX. SUPPLEMENTARY MATERIAL FOR CHAPTER 2

Table S1. Sequences retrieved from NCBI Influenza Virus Resource for the primer design.

Gene length Primer Accession number Subtype (bp)

H3-1 AB292402 A/duck/Hong Kong/836/1980 (H3N1) 1731

AB292410 A/duck/Hong Kong/22A/1976 (H3N3) 1731

AB277754 A/duck/Hokkaido/5/1977 (H3N2) 1731

AB292660 A/duck/Hong Kong/22B/1976 (H3N6) 1731

AY779254 A/turkey/Minnesota/764-2/03 (H3N2) 1714

M16739 A/duck/33/1980 (H3) 1653

M16737 A/duck/5/1977(H3) 1653

D00931 A/duck/Hong Kong/64/76 (H3) 1653

D00932 A/duck/Hong Kong/231/77 (H3) 1653

DQ150425 A/swine/MI/PU243/04 (H3N1) 1765

DQ923506 A/swine/Korea/PZ72-1/2006 (H3N1) 1701

CY000257 A/New York/52/2004 (H3N2) 1762

H3-2 AB289341 A/swan/Shimane/227/01 (H3N9) 1731

AB292668 A/duck/Ukraine/1963 (H3N8) 1731

CY006011 A/Pigeon/Nanchang/9-058/2000 (H3N3) 1764

CY006012 A/bantam/Nanchang/9-366/2000 (H3N3) 1764

CY006013 A/Chicken/Nanchang/7-010/2000 (H3N6) 1764

CY006035 A/duck/NZL/38/1984 (H3N8) 1765

H3-3 DQ146419 A/canine/Iowa/13628/2005 (H3N8) 1698

M24723 A/equine/Fontainebleau/1976 (H3N8) 1762

M24721 A/equine/Algiers/1972 (H3N8) 1762

CY011048 A/blue-winged teal/Ohio/908/2002 (H3N2) 1731

CY017709 A/mallard/Ohio/156/1990 (H3N6) 1731

CY021277 A/mallard/Maryland/631/2005 (H3N2) 1729

H3-4 M16741 A/duck/21/1982 (H3) 1653

CY005943 A/mallard duck/ALB/26/1976 (H3N1) 1765

CY005940 A/blue-winged teal/ALB/452/1983 (H3N1) 1765

CY005935 A/pintail duck/ALB/86/1976 (H3N2) 1765

CY014548 A/mallard duck/Alberta/331/1985 (H3N6) 1765

CY005936 A/mallard duck/ALB/712/1978 (H3N3) 1765

CY005942 A/mallard/ALB/394/1988 (H3N3) 1765

CY004657 A/mallard duck/ALB/438/1985 (H3N4) 1765

CY004692 A/mallard/ALB/118/1995 (H3N5) 1765

CY013263 A/pintail/Alaska/53/2005 (H3N6) 1732

CY005941 A/mallard duck/ALB/635/1983 (H3N5) 1765

H4-1 AB288842 A/duck/Hokkaido/1058/01 (H4N5) 1704

AB289333 A/duck/Mongolia/583/02 (H4N7) 1704

AB289331 A/duck/Hong Kong/438/1977 (H4N8) 1704

AF290436 A/Duck/Czechoslovakia/56 (H4N6) 1714

CY006030 A/duck/Hong Kong/24/1976 (H4N2) 1738

CY005672 A/grey teal/Australia/2/1979 (H4N4) 1738

CY006017 A/Duck/Nanchang/4-165/2000 (H4N6) 1738

H4-1 CY006027 A/duck/Hong Kong/365/1978 (H4N6) 1738

CY005679 A/gray teal/AUS/3/1979 (H4N6) 1738

CY014630 A/red-necked stint/Australia/4189/1980 (H4N8) 1739

M25285 A/budgerigar/Hokkaido/1/1977 (H4N6) 1738

M25286 A/duck/New Zealand/31/1976 (H4N6) 1738

M25283 A/duck/Czeckoslovakia/1956 (H4N6) 1738

DQ327834 A/red-necked Stint/Australia/1/2004 (H4N8) 1710

H4-2 CY004925 A/mallard/ALB/47/1998 (H4N1) 1738

CY004939 A/mallard/Alberta/30/2001 (H4N8) 1738

CY005964 A/blue-winged teal/ALB/136/1990 (H4N3) 1738

CY014857 A/mallard duck/New York/180/1986 (H4N9) 1738

CY005953 A/mallard duck/ALB/354/1978 (H4N2) 1738

CY015459 A/green-winged teal/Ohio/344/1986 (H4N2) 1704

CY005948 A/mallard/Alberta/300/1977 (H4N3) 1738

CY005957 A/mallard duck/ALB/581/1983 (H4N4) 1738

CY005963 A/blue-winged teal/ALB/103/1990 (H4N5) 1738

CY014562 A/mallard/Alberta/254/2003 (H4N6) 1738

CY005962 A/ruddy turnstone/DE/512/1988 (H4N6) 1738

M25288 A/chicken/Alabama/1/1975 (H4N8) 1737

M25291 A/seal/Massachussetts/133/1982 (H4N5) 1737

DQ236166 A/blue-winged teal/Barbados/21/04 (H4N3) 1707

H7-1 AF071776 A/chicken/New York/1995 (H7) 1629

CY014896 A/chicken/Pennsylvania/143586/2002 (H7N2) 1707

H7-1 CY005983 A/mallard/Alberta/34/2001 (H7N1) 1731

CY005975 A/widgeon/ALB/284/1977 (H7N3) 1731

CY005973 A/mallard duck/ALB/224/1977 (H7N5) 1731

CY005928 A/ruddy turnstone/NJ/65/1985 (H7N3) 1731

CY014786 A/turkey/Minnesota/1/1988 (H7N9) 1731

CY015027 A/chicken/Chile/184240-4322/2002 (H7N3) 1760

DQ525411 A/cinnamon teal/Bolivia/4537/2001 (H7N3) 1712

H7-2 AF202227 A/chicken/Victoria/1/92 (H7N3) 1737

AY943924 A/chicken/NSW/1/97 (H7N4) 1738

M17736 A/starling/Victoria/1/1985 (H7N7) 1692

M24457 A/FPV/Rostock/1934 (H7N1) 1742

CY014992 A/fowl/Dobson/1927 (H7N7) 1741

L37794 A/FPV/Weybridge (H7N7) 1741

Z47199 A/chicken/Victoria/75 (H7N7) 1638

H7-3 AY724257 A/chicken/Hebei/1/2002 (H7N2) 1732

AF202235 A/turkey/Israel/Ramon/79 (H7N2) 1732

AF202248 A/gull/Italy/692-2/93 (H7N2) 1732

AF202234 A/ostrich/Zimbabwe/222/96 (H7N1) 1731

AF202253 A/ostrich/South Africa/M320/96 (H7N7) 1732

AF028021 A/turkey/Ireland/PV74/1995 (H7N7) 1732

AF202228 A/common iora /Singapore/F89/95 (H7N1) 1732

CY015033 A/chicken/Pakistan/34669/1995 (H7N3) 1741

CY006029 A/dk/Hong Kong/293/1978 (H7N2) 1732

L43914 A/goose/Leipzig/187/7/1979 (H7N7) 1743

H7-4 AY586408 A/turkey/Italy/214845/02 (H7N3) 1689

AY338459 A/Netherlands/219/03 (H7N7) 1737

AY338460 A/mallard/Netherlands/12/00 (H7N3) 1732

AY999985 A/Mallard/Sweden/100/02 (H7N7) 1684

AY999981 A/Mallard/Sweden/91/02 (H7N9) 1684

CY014718 A/mallard/Netherlands/12/2000 (H7N3) 1726

H8 AB289343 A/turkey/Ontario/6118/1968 (H8N4) 1710

AF310989 A/Red Kont/Delaware/254/94 (H8N4) 1229

AF310987 A/Pintail Duck/Alberta/114/79 (H8N4) 1237

AF310988 A/Mallard Duck/Alberta/357/84 (H8N4) 1226

CY005972 A/mallard/ALB/194/1992 (H8N4) 1744

CY017749 A/mallard/Alaska/708/2005 (H8N4) 1705

CY015173 A/duck/Alaska/702/1991 (H8N2) 1744

CY014583 A/mallard duck/Alberta/7/1987 (H8N4) 1744

CY005971 A/pintail duck/Alberta/114/1979 (H8N4) 1744

CY014659 A/turkey/Ontario/6118/1968 (H8N4) 1744

CY005970 A/mallard/Alberta/283/1977 (H8N4) 1744

D90304 A/turkey/Ontario/6118/1968 (H8N4) 1698

EF061122 A/duck/Yangzhou/02/2005 (H8N4) 1744

H9 AB125928 A/duck/Hokkaido/49/98 (H9N2) 1707

AF461524 A/Chicken/Shanghai/4-1/01 (H9N2) 1683

AF523389 A/Duck/Shantou/1588/00 (H9N1) 1626

AF523390 A/Duck/Shantou/2030/00 (H9N1) 1626

AF203008 A/ck/Korea/ms96/96 (H9) 1683

H9 AJ291392 A/Chicken/Pakistan/2/99 (H9N2) 1701

AY633276 A/mallard/Alberta/321/88 (H9N2) 1716

AY633116 A/mallard/Alberta/743/83 (H9N1) 1721

AY949989 A/chicken/Shandong/2/02 (H9N?) 1704

AY206673 A/duck/Hong Kong/448/78 (H9N2) 1683

AY206671 A/duck/Hong Kong/147/77 (H9N6) 1683

AY862599 A/chicken/Korea/S4/03 (H9N2) 1683

AY862606 A/chicken/Korea/S18/03 (H9N2) 1690

CY005985 A/ruddy turnstone/DE/2576/1987 (H9N5) 1742

CY005986 A/laughing gull/DE/2718/1987 (H9N5) 1742

CY005987 A/ruddy turnstone/DE/2731/1987 (H9N1) 1742

CY005929 A/knot/DE/2552/1987 (H9N5) 1742

CY005934 A/ruddy turnstone/DE/773/1988 (H9N6) 1742

CY005988 A/ruddy turnstone/DE/510/1988 (H9N6) 1742

CY005984 A/mallard duck/ALB/743/1983 (H9N1) 1742

CY005919 A/mallard duck/ALB/396/1983 (H9N1) 1742

CY004642 A/mallard duck/ALB/506/1983 (H9N1) 1742

CY014663 A/turkey/Wisconsin/1/1966 (H9N2) 1742

CY006021 A/Wild Duck/Nanchang/2-0480/2000 (H9N2) 1742

CY005639 A/duck/HK/147/1977 (H9N6) 1742

CY005746 A/duck/NZL/76/1984 (H9N1) 1742

CY004420 A/laughing gull/DE/5/2003 (H9N1) 1736

CY005992 A/shorebird/DE/261/2003 (H9N5) 1742

DQ064369 A/chicken/Henan/43/02 (H9N2) 1742

DQ885991 A/chicken/Korea/164/04 (H9N8) 1640

EF063514 A/chicken/Dubai/339/2001 (H9N2) 1722

H10 AB271117 A/duck/Hong Kong/938/80 (H10N1) 1694

AB292412 A/duck/Hong Kong/786/1979 (H10N3) 1694

AB289339 A/swan/Shimane/1331/1981 (H10N6) 1694

AB274041 A/duck/Hokkaido/18/00 (H10N4) 1694

CY005995 A/mallard duck/ALB/40/1984 (H10N6) 1728

CY005996 A/pintail duck/ALB/584/1984 (H10N6) 1728

CY014739 A/mallard duck/Minnesota/19/1979 (H10N7) 1728

CY005930 A/shorebird/DE/10/2004 (H10N7) 1728

CY006001 A/shorebird/DE/122/2004 (H10N7) 1728

CY005922 A/mallard/Alberta/209/2003 (H10N7) 1728

CY005999 A/pintail/Alberta/202/2000 (H10N7) 1728

CY006000 A/mallard/Alberta/208/2000 (H10N7) 1728

CY005998 A/mallard/ALB/196/1996 (H10N7) 1728

CY005997 A/mallard/ALB/5/1995 (H10N1) 1728

CY005982 A/pintail/Alberta/129/1993 (H10N7) 1728

CY017781 A/mallard/Ohio/99/1989 (H10N7) 1694

CY005921 A/pintail duck/ALB/303/1977 (H10N7) 1728

CY005993 A/mallard duck/ALB/302/1977 (H10N7) 1728

CY005994 A/blue-winged teal/ALB/778/1978 (H10N3) 1728

CY014671 A/chicken/Germany/n/1949 (H10N7) 1728

CY014644 A/quail/Italy/1117/1965 (H10N8) 1728

CY014619 A/duck/Hong Kong/562/1979 (H10N9) 1728

M21646 A/mink/Sweden/1984 (H10N4) 1727

M21647 A/chicken/Germany/N/1949 (H10N7) 1727

H11-1 AB288845 A/duck/England/1/1956 (H11N6) 1726

AB277756 A/swan/Shimane/48/1997 (H11N2) 1726

AY684895 A/mallard/Netherlands/7/99 (H11N2) 1760

CY014719 A/shoveler/Netherlands/19/1999 (H11N9) 1760

CY014679 A/duck/England/1956 (H11N6) 1760

D90306 A/duck/England/1/1956 (H11N6) 1698

DQ080993 A/duck/Yangzhou/906/2002 (H11N2) 1698

DQ327835 A/sharp-tailed sandpiper/Australia/6/2004(H11N9) 1710

DQ482667 A/mallard/Xuyi/8/2004 (H11N?) 1698

H11-2 CY006004 A/pintail/Alberta/84/2000 (H11N9) 1760

CY017845 A/green-winged teal/Ohio/81/1999 (H11N2) 1727

CY005924 A/mallard/Alberta/245/2003 (H11N9) 1760

CY006003 A/mallard/ALB/124/1991 (H11N2) 1760

CY005923 A/mallard duck/ALB/294/1977 (H11N9) 1760

CY014687 A/duck/Memphis/546/1974 (H11N9) 1760

CY017075 A/bufflehead/Ohio/246/1986 (H11N2) 1727

CY018015 A/mallard/Ohio/102/1986 (H11N3) 1727

CY017765 A/black duck/Ohio/194/1986 (H11N1) 1726

CY006002 A/mallard duck/ALB/797/1983 (H11N3) 1760

CY014593 A/redhead duck/Alberta/357/1983 (H11N9) 1760

CY014595 A/ruddy turnstone/Delaware/2762/1987 (H11N2) 1757

CY014806 A/mallard duck/Tennessee/11457/1985 (H11N9) 1760

CY006005 A/shorebird/DE/236/2003 (H11N9) 1760

H12 AB288843 A/duck/Hokkaido/66/01 (H12N5) 1703

AB288334 A/duck/Alberta/60/1976 (H12N5) 1703

CY016419 A/mallard/Ohio/409/1988 (H12N5) 1698

CY017853 A/mallard/Ohio/407/1987 (H12N5) 1699

CY006006 A/mallard duck/Alberta/342/1983 (H12N1) 1737

CY006008 A/ruddy turnstone/DE/97/2000 (H12N5) 1737

CY014598 A/laughing gull/Delaware/94/2000 (H12N4) 1737

CY005925 A/mallard/ALB/52/1997 (H12N5) 1737

CY006007 A/green-winged teal/ALB/199/1991 (H12N5) 1737

CY012840 A/pintail/Alaska/20/2005 (H12N5) 1703

CY017733 A/pintail/Alaska/102/2005 (H12N5) 1701

CY005920 A/pintail/Alberta/49/2003 (H12N5) 1737

CY014636 A/red-necked stint/Australia/5745/1981 (H12N9) 1737

D90307 A/duck/Alberta/60/1976 (H12N5) 1695

DQ787811 A/duck/Primorie/3691/02 (H12N2) 1663

H13 AB284988 A/duck/Siberia/272/1998 (H13N6) 1732

AB285094 A/duck/Siberia/272PF/1998 (H13N6) 1746

AY684886 A/black-headed gull/Netherlands/1/00 (H13N8) 1740

AY684887 A/black-headed gull/Sweden/1/99 (H13N6) 1765

CY005914 A/herring gull/DE/475/1986 (H13N2) 1767

CY005932 A/herring gull/NJ/782/1986 (H13N2) 1767

CY005979 A/laughing gull/DE/2838/1987 (H13N2) 1768

CY014603 A/herring gull/Delaware/660/1988 (H13N6) 1773

CY014720 A/gull/Minnesota/945/1980 (H13N6) 1766

H13 CY005931 A/shorebird/DE/68/2004 (H13N9) 1767

CY014694 A/gull/Maryland/704/1977 (H13N6) 1768

D90308 A/gull/Maryland/704/1977 (H13N6) 1701

M26089 A/black-headed gull/Astrakhan/227/84 (H13N6) 1765

M26090 A/ring-billed gull/Maryland/704/1977 (H13N6) 1768

M26091 A/pilot whale/Maine/328 HN/1984 (H13N2) 1768

H14 AB289335 A/mallard/Astrakhan/263/1982 (H14N5) 1714

CY014604 A/mallard duck/Astrakhan/263/1982 (H14N5) 1748

M35996 A/Mallard/Gurjev/244/82 (H14) 1716

M35997 A/Mallard/Gurjev/263/82 (H14) 1749

H15 A/wedge-tailed shearwater/Western CY006010 1763 Australia/2576/1979 (H15N9)

A/wedge-tailed shearwater/Western CY006034 1763 Australia/2327/1983 (H15N9)

A/sooty tern/Western CY006033 1763 Australia/2190/1983(H15N9)

A/Australian shelduck/Western CY006032 1763 Australia/1756/1983 (H15N2)

CY006009 A/duck/AUS/341/1983 (H15N8) 1762

L43916 A/duck/Australia/341/83 (H15N8) 1762

L43917 A/shearwater/West Australia/2576/79 (H15N9) 1762

H16 AY684889 A/black-headed gull/Sweden/3/99 (H16N3) 1695

AY684890 A/black-headed gull/Sweden/4/99 (H16N3) 1753

AY684888 A/black-headed gull/Sweden/2/99 (H16N3) 1753

H16 AY684891 A/black-headed gull/Sweden/5/99 (H16N3) 1760

AX411576 Sequence 11 from Patent WO0224734 (H16) 1753

AX411574 Sequence 9 from Patent WO0224734 (H16) 1753

AX411575 Sequence 10 from Patent WO0224734 (H16) 1753

AX411577 Sequence 12 from Patent WO0224734 (H16) 1757

CY005933 A/herring gull/DE/712/1988 (H16N3) 1758

CY014599 A/shorebird/New Jersey/840/1986 (H16N3) 1758

A/black-legged kittywake/Alaska/295/1975 CY015160 1758 (H16N3)