The Pennsylvania State University
The Graduate School
Department of Veterinary and Biomedical Sciences
A COMPREHENSIVE STUDY OF THE HEALTH OF FARM-RAISED WHITE-
TAILED DEER (ODOCOILEUS VIRGINIANUS) WITH EMPHASIS ON
RESPIRATORY TRACT INFECTION BY FUSOBACTERIUM SPP.
A Dissertation in
Pathobiology
by
Jason W. Brooks
© 2010 Jason W. Brooks
Submitted in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
August 2010
ii
The dissertation of Jason W. Brooks was reviewed and approved* by the following:
Bhushan Jayarao Professor of Veterinary Science Dissertation Advisor Chair of Committee
Arthur Hattel Senior Research Associate
Avery August Professor of Immunology
Gary San Julian Professor of Wildlife Resources
Sanjeev Narayanan Assistant Professor College of Veterinary Medicine, Kansas State University Special Member
Vivek Kapur Professor and Head of the Department of Veterinary and Biomedical Sciences
*Signatures are on file in the Graduate School.
iii
ABSTRACT
White-tailed deer farming is an established and growing industry in Pennsylvania.
Managers of deer farming operations often struggle with animal health problems, the
most common of which is pneumonia associated with Fusobacterium sp. infection.
Fusobacterium is a genus of anaerobic, gram negative, rod-shaped bacteria that have
been associated with many infectious disease processes in humans and animals. It is
important to the deer industry, as well as the cattle and sheep industries, to more clearly
understand fusobacterial disease pathogenesis and determine effective treatment and
prevention strategies. The objectives of the following series of studies were: 1) to
determine the current management practices and animal health problems on white-tailed
deer farms in Pennsylvania, 2) to phenotypically and genotypically characterize a set of
Fusobacterium isolates from white-tailed deer and evaluate their association with disease
in the host, and 3) to determine the pathogenicity of Fusobacterium varium to the respiratory tract of mice and to evaluate this system as a model for pulmonary fusobacterial infection of deer.
In order to determine the current management practices used by white-tailed deer farms and identify animal health problems in these herds, a self-administered questionnaire was mailed to managers of 235 farms in Pennsylvania that raise white- tailed deer. Herds ranged in size from 1 to 350 deer. Land holdings ranged from 0.07 to
607 hectares (0.17 to 1,500 acres). Stocking density ranged from 0.1 to 118.6 deer/hectare (0.04 to 48 deer/acre). Most (84%) respondents raised deer for breeding or hunting stock; 13% raised deer exclusively as pets or for hobby purposes, and purpose iv
varied by herd size. Multiple associations were identified between management or
disease factors and herd size. The use of vaccines, veterinary and diagnostic services,
pasture, and artificial insemination increased as herd size increased. The most common
disease conditions in herds of all sizes were respiratory tract disease, diarrhea, parasitism,
and sudden death. The prevalence of respiratory tract disease increased as herd size
increased. Results suggested that many aspects of herd management for white-tailed deer
farms in Pennsylvania were associated with herd size, but that regardless of herd size,
many preventive medicine practices were improperly used or underused in many herds.
A total of 28 clinical strains of Fusobacterium spp. were isolated at necropsy over a two-year period from the respiratory tract of cervids, including white-tailed deer, elk,
and reindeer. Isolates were identified as F. varium (21/28, 75%), F. necrophorum subsp.
funduliforme (5/28, 17.9%), and F. necrophorum subsp. necrophorum (2/28, 7.1%).
Using PCR-based detection of virulence genes, all F. varium isolates were negative for
the promoter region of the leukotoxin operon of F. necrophorum and the hemagglutinin-
related protein gene of F. necrophorum. In the necropsy population no significant
differences in gross or microscopic lesions were detected across Fusobacterium species,
suggesting similar potential for virulence, however, toxicity to bovine polymorphonuclear leukocytes was not observed in any F. varium strains, perhaps
indicating that a virulence factor other than leukotoxin is involved in the pathogenesis of
F. varium infection. F. varium was less susceptible to many antimicrobials than were the
F. necrophorum subspecies. These data suggest that F. varium may be a significant
pathogen in deer and may require different treatment and prevention methods than F.
necrophorum. v
C57BL/6 mice and BALB/c mice were inoculated intranasally with various
strains and varying dosages of F. varium to evaluate the pathogenicity of F. varium to the respiratory tract of mice and to determine the utility of this system as model of fusobacterial pneumonia. Prior to inoculation, mice were pre-treated with either 0.2 mg
dexamethasone intraperitoneally once daily for four days, 10 µg of lipopolysaccharide
(LPS) of E. coli 055:B5 intranasally one time, or no pre-treatment. Following
inoculation mice were observed for morbidity and mortality for fourteen days. No mice
infected with F. varium showed clinical illness or died. No mice infected with F. varium
developed gross or microscopic lesions. The bacterium was recovered from the blood of
one mouse, but was not recovered from blood or lung of any other mice. Anti-
fusobacterial IgM or IgG were not produced in serum by 14 days in response to infection.
Pre-existing antibodies detected in pooled serum bound similarly with proteins of two F.
necrophorum subspecies and three F. varium strains, but did not bind to proteins of
similar size from other common bacterial pathogens including Escherichia coli,
Pasteurella pneumotropica, Arcanobacterium pyogenes, and Clostridium perfringens.
No serum antibodies were detected in BALB/c mice. These results suggest that F.
varium is not highly pathogenic to the respiratory tract of mice, and does not result in a
humoral immune response following intranasal inoculation.
In conclusion, the results of this series of studies suggest that fusobacterial
species, specifically F. varium, are not highly pathogenic to the respiratory tract and are
unable to establish lung infection alone without some severe predisposing condition.
Management changes on deer farms should be instituted in an effort to minimize stress,
overcrowding, nutritional imbalance, and environmental contamination, while vi
maximizing immunity through colostrum and vaccination, and developing effective preventive health and biosecurity programs with routine veterinary consultation. vii
TABLE OF CONTENTS
LIST OF FIGURES……………………………………………………………….. x
LIST OF TABLES………………………………………………………………… xii
ACKNOWLEDGEMENTS………………………………………………………. xiv
Chapter 1. INTRODUCTION…………………………………………………….. 1
1.1 Statement of the Problem……………………………………………… 2
1.2 Initial Observations……………………………………………………. 2
1.3 Retrospective Study of Deer Mortality………………………………... 3
1.4 Evaluation of Viral Etiologies with emphasis on BVDV and BRSV… 5
1.5 Objectives……………………………………………………………... 7
1.6 References…………………………………………………………….. 8
Chapter 2. REVIEW OF LITERATURE…………………………………………. 9
2.1 Deer Farming Industry………………………………………………… 10
2.2 Animal Health in Deer Populations…………………………………… 11
2.3 Anaerobic Bacteria……………………………………………………. 13
2.3.1 The Fusobacteria…………………………………………….. 15
2.3.2 Fusobacterium necrophorum………………………………... 16
2.3.3 Fusobacterium varium………………………………………. 21
2.4 Lung Infection and Immunity…………………………………………. 22
2.5 Animal Models of Bacterial Pulmonary Infection…………………….. 25
2.6 Animal Models of Anaerobic Bacterial Infection…………………….. 27
2.7 References…………………………………………………………….. 31 viii
Chapter 3. MANAGEMENT PRACTICES USED BY WHITE-TAILED DEER FARMS
IN PENNSYLVANIA AND HERD HEALTH PROBLEMS……………. 53
3.1 Acknowledgements…………………………………………………… 54
3.2 Abstract……………………………………………………………….. 54
3.3 Introduction…………………………………………………………… 54
3.4 Materials and Methods………………………………………………… 54
3.5 Results…………………………………………………………………. 55
3.6 Discussion…………………………………………………………….. 58
3.7 References…………………………………………………………….. 60
Chapter 4. PHENOTYPIC AND GENOTYPIC CHARACTERIZATION OF
FUSOBACTERIUM ISOLATES FROM THE RESPIRATORY TRACT OF
DEER………………………………………………………………………. 61
4.1 Abstract……………………………………………………………….. 62
4.2 Introduction……………………………………………………………. 62
4.3 Materials and Methods………………………………………………… 64
4.4 Results…………………………………………………………………. 69
4.5 Discussion…………………………………………………………….. 74
4.6 Acknowledgements…………………………………………………… 76
4.7 References…………………………………………………………….. 77
Chapter 5. EFFECTS OF FUSOBACTERIUM VARIUM ON THE RESPIRATORY
TRACT OF MICE FOLLOWING INTRANASAL INOCULATION…………… 88
5.1 Abstract……………………………………………………………….. 89
5.2 Introduction…………………………………………………………… 90 ix
5.3 Materials and Methods………………………………………………… 91
5.4 Results…………………………………………………………………. 96
5.5 Discussion…………………………………………………………….. 99
5.6 Acknowledgements…………………………………………………… 103
5.7 References……………………………………………………………... 103
Chapter 6. SUMMARY AND CONCLUSIONS………………………………….. 117
Appendix A. SURVEY QUESTIONNAIRE……………………………………… 122
Appendix B. SUPPLEMENTAL PHENOTYPIC DATA………………………… 126
Appendix C. MOUSE INFECTIVITY TRIALS RAW DATA…………………… 132 x
LIST OF FIGURES
Figure 2.1. F. necrophorum subsp. necrophorum growth on blood agar...... 46
Figure 2.2. F. necrophorum subsp. funduliforme growth on blood agar.………….. 47
Figure 2.3. F. varium growth on blood agar……………………………………….. 48
Figure 2.4. Fusobacterium spp. growth in PRAS-BHI broth……………………… 49
Figure 2.5. F. necrophorum subsp. necrophorum cellular morphology (Gram
Stain).…………………………………………………………………... 50
Figure 2.6. F. necrophorum subsp. funduliforme cellular morphology (Gram
Stain)…………………………………………………………………… 51
Figure 2.7. F. varium cellular morphology (Gram Stain)…………………………. 52
Figure 4.1. Phylogenetic analysis of the 16S rRNA gene sequence of
Fusobacterium spp…...... 82
Figure 5.1. Liver – C57BL/6 mouse (H&E)……………………………………….. 111
Figure 5.2. Liver – C57BL/6 mouse (Brown & Brenn), (Warthin-Starry)………… 112
Figure 5.3. Lung – C57BL/6 mouse (H&E) ………………………………………. 113
Figure 5.4. Western blot of pre-infection mouse serum…………………………… 114
Figure 5.5. Western blot of post-infection mouse serum……………………………115
Figure 5.6. Western blot of pre-infection mouse serum from C57BL/6 with protein from
Fusobacterium and other bacteria……….……………………………... 116
Figure B.1. Growth curves of Fusobacterium spp. in brain-heart infusion broth…. 126 xi
Figure B.2. Summarized leukotoxin production by species as determined by toxicity to
bovine PMNs following exposure to culture supernatant as measured by
propidium iodide (PI) staining on flow cytometry…………………….. 127
xii
LIST OF TABLES
Table 3.1. Descriptive characteristics of white-tailed deer farms in Pennsylvania... 55
Table 3.2. Use of preventive medicine practices on white-tailed deer farms in
Pennsylvania…………………………………………………………….. 56
Table 3.3. Use of veterinary and diagnostic services on white-tailed deer farms in
Pennsylvania…………………………………………………………….. 57
Table 3.4. Feeding practices on white-tailed deer farms in Pennsylvania…………. 57
Table 3.5. Disease conditions affecting herd health reported by owners and managers of
white-tailed deer farms in Pennsylvania………………………………... 58
Table 4.1. Fusobacterium strains characterized in this study……………………… 83
Table 4.2. Phenotypic and Genotypic Identification of Fusobacterium isolates…... 84
Table 4.3. Antimicrobial Susceptibility of Fusobacterium spp……………………. 85
Table 4.4. Antibiogram of Fusobacterium spp…………………………………….. 86
Table 4.5. Profiles of virulence gene PCR products………………………………. 87
Table 5.1. Study design of trial 1………………………………………………….. 108
Table 5.2. Study design of trial 2………………………………………………….. 109
Table 5.3. Study design of trial 3…………………………………………………... 110
Table B.1. Results for Fusobacterium isolates using Remel Rapid ANA II and
Biomerieux RapID 32A anaerobic bacterial identification kits………... 128
Table B.2. Results of χ2 analysis of association between lesion production or
co-pathogen and fusobacterial infection….……………………………. 129 xiii
Table B.3. Results of contingency analysis of association between pneumonia and
respiratory fusobacterial infection in all deer on which anaerobic cultures were
performed………………………………………………………………. 130
Table B.4. Results of contingency analysis of association between pneumonia and
respiratory infection with Arcanobacterium pyogenes in all deer on which
anaerobic cultures were performed….…………………………………. 131
Table C.1. Bacteriology results of trial 1…………………………………………... 132
Table C.2. Necropsy results of trial 1……………………………………………… 133
Table C.3. Histology results of trial 1……………………………………………… 133
Table C.4. Bacteriology results of trial 2…………………………………………... 134
Table C.5. Necropsy results of trial 2……………………………………………… 135
Table C.6. Histology results of trial 2……………………………………………… 135
Table C.7. Morbidity and mortality results of trial 3……………………………… 136
Table C.8. Bacteriology results of trial 3………………………………………….. 137
Table C.9. Necropsy results of trial 3……………………………………………… 138
Table C.10. Histology results of trial 3……………………………………………. 138
xiv
ACKNOWLEDGEMENTS
There are many people to whom I am indebted for their constant support during my graduate studies. First and foremost, I wish to express my deepest gratitude to my dissertation advisor, Dr. Bhushan Jayarao, for his unwavering support of my graduate research and professional development during the last six years. His perpetual confidence in my abilities was at times the only remaining spark to fuel my efforts. Likewise, I would like to sincerely thank all of the additional members of my thesis committee, including Dr. Avery August, Dr. Gary San Julian, Dr. Art Hattel, and Dr. Sanjeev Narayanan. Guided by their wisdom and insight, my graduate studies have allowed me to develop professionally and to conduct meaningful research with practical results. Dr. Art Hattel, in particular, has been a true mentor to me and successfully guided me into the fascinating world of veterinary pathology. The countless hours that he spent teaching me on the multi-headed microscope have doubtlessly molded me into the pathologist that I have become. I have had the very good fortune to become acquainted with a very generous and talented group of professionals at the College of Veterinary Medicine at Kansas State University. Dr. Sanjeev Narayanan, Dr. T.G. Nagaraja, and Mr. Amit Kumar have all helped me tremendously in my studies of Fusobacterium. I can only hope to emulate their kindness and generosity throughout my career and personal life. Dr. Subhashinie Kariyawasam and Mrs. Valerie Lintner of the Section of Bacteriology, and Dr. Suzanne Myers and Ms. Rhiannon Schneider of the Section of Molecular Biology at the Animal Diagnostic Laboratory at Penn State University provided critical assistance and training and generously provided essential resources. Mrs. Lola Hubler and Mr. Bill Weaver provided a tremendous amount of technical support at nearly every stage of my research, from necropsy assistance to photography to anesthesia and more. Without their assistance this work would not have been possible. Dr. Vivek Kapur, Professor and Head of the Department of Veterinary and Biomedical Sciences has been very supportive of my efforts to work as a member of his faculty while pursuing this degree. I wish to thank him for his encouragement and for his critical support. I sincerely thank the Missouri Whitetail Breeders and Hunting Ranch Association for their financial support, without which I would not have been able to complete this research. Finally, above all I must thank my faith and my family, including my mother and father, my brother and sister in law, and my wife and children. Mom and Dad, you shaped me from my earliest days into the person who I am today. Without your constant unconditional love and support I would have failed many years ago. I am truly grateful to have received such a gift of familial love and guidance. It is with the utmost humility that I give the most sincere thanks to my wife, Judy, who has, without hesitation, whole-heartedly supported me throughout many long years of education from veterinary medical school to pathology residency to graduate school, all the while tolerating my incessant volunteer activities. Through it all, she has been my guidepost, my rock, and my foundation. She and our three children, Miles, Clara, and Elena, have been my true inspiration to continue. They have all made tremendous sacrifices for my sake in an effort to help me succeed…I only hope that I never disappoint them. I love you all and I thank you for carrying me through to the end. 1
Chapter 1
INTRODUCTION
2
1.1 Statement of the Problem
White-tailed deer farming has become an established and growing nationwide
industry in which Pennsylvania has long been a leader. Managers of deer farming
operations often struggle with animal health problems and one of the most common is
pneumonia which is often associated with Fusobacterium sp. infection. Another
common form of Fusobacterium sp. infection in deer is a condition known as
necrobacillosis, which results in purulent necrotic lesions that may involve most of the
body. The infection often spreads to multiple organs resulting in death, as antibiotic
therapy is difficult. The underlying cause of this deer respiratory disease complex and
necrobacillosis is not completely understood, and Fusobacterium sp. may represent either
a primary pathogen or a secondary invader. With the future of the last remaining
commercial fusobacterial vaccines in question, it is important to the deer industry, as well as the cattle and sheep industries, to more clearly understand fusobacterial disease pathogenesis and determine effective treatment and prevention strategies.
1.2 Initial Observations
During the summer and fall of 2003 the author began to observe that a large proportion of deer submitted for post mortem examination to the Animal Diagnostic
Laboratory at Penn State University were diagnosed with respiratory tract infection. The inflammation was consistently severe and extensive, with nearly diffuse pulmonary consolidation and multifocal necrotic cavitations replaced by copious volumes of purulent exudate and extensive adherent fibrin throughout the chest bilaterally.
Interestingly, in the majority of such cases the owners reported sudden unexpected death 3
with little or no observed clinical disease in affected animals. The most commonly isolated bacterial pathogens were Fusobacterium spp. and Arcanobacterium pyogenes.
Despite repeated efforts to identify viral agents by virus isolation, tissue immunofluorescence assay (IFA), immunohistochemistry (IHC), or polymerase chain reaction (PCR), viral organisms were almost never identified, and parasitic and fungal pathogens were ruled out. These initial observations prompted the author to further investigate this respiratory disease complex of farm-raised white-tailed deer.
1.3 Retrospective Study of Deer Mortality
As the initial step of this investigation of the respiratory disease complex, the author was a co-investigator of a study of mortality in farm-raised white-tailed deer.1 In this study, the postmortem records of 160 white-tailed deer that were submitted for necropsy from 59 distinct Pennsylvania deer farms over a 3.5-year period were reviewed to determine the primary cause of death. The most common causes of death were bronchopneumonia (39/160; 24.4%), enterocolitis (30/160; 18.8%), malnutrition (13/160;
8.1%), and trauma (11/160; 6.9%). Other causes of mortality during the study period included gastrointestinal parasitism, cellulitis with septicemia, degenerative myopathy, ruminal acidosis, and nephritis. Arcanobacterium pyogenes, Fusobacterium necrophorum, Escherichia coli, and Mannheimia haemolytica were the most commonly isolated bacteria from the pneumonic lungs. The majority (52.2%) of all death loss in deer of known ages occurred in animals 1 year of age or less, with 46.2% of fatal bronchopneumonia cases occurring during this interval. An even greater proportion of bronchopneumonia cases (64.1%) occurred in deer 2 years of age or less. 4
In this dataset, F. necrophorum accounted for 27.8% of the cases of
bronchopneumonia in which anaerobic bacterial cultures were attempted, however
anaerobic cultures were performed on lung tissue in only 23 of the 36 deer with
bronchopneumonia. Of these 23 deer, 10 (43.5%) were positive for F. necrophorum.
Thus, the prevalence of F. necrophorum in pneumonic lungs may have been
underrepresented in this study.
Results of this study indicated that the first year of life for farm-raised deer is
especially critical, for the majority of total death loss for all age groups occurred during this age interval. Contributing factors suggested by the authors may include poor colostrum management leading to reduced passive immunity, poor maternal care, inadequate milk production by the doe, high deer population density, inappropriate postweaning nutrition, inadequate shelter, dirty or excessively soiled fawning areas, and lack of adequate vaccination programs. By determining the major causes of age-specific mortality, a variety of preventative measures can be recommended to producers including implementation of vaccination programs, ensuring that newborns receive adequate volumes of high-quality colostrum, and properly balancing rations. Avoidance of overcrowding, periodic movement of feeders to uncontaminated areas, periodic herd health checks throughout the day, especially during the fawning season, paddock rotation, and provision of clean and adequate shelters should be instituted. Management techniques that may reduce stress and risk of trauma may include allowing animals to become accustomed to handling areas, fence lines, and gates prior to handling; proper removal of antlers of aggressive animals and members of the bachelor herd; and protection of chemically immobilized deer from herdmates. 5
1.4 Evaluation of Viral Etiologies with emphasis on BVDV and BRSV
Having determined that bronchopneumonia was the leading cause of death in the necropsy population of farmed deer, the author began investigating the etiologic agents responsible for the infection. Since it is well known in cattle and other animal populations that bacterial lung infections often follow primary viral infections, the author and his colleagues began testing necropsy cases for viruses known to cause pneumonia in other livestock species, including bovine viral diarrhea virus (BVDV), bovine respiratory syncytial virus (BRSV), bovine herpesvirus type 1 (BHV1), and parainfluenzavirus type
3 (PI3). With the exception of two cases that will be subsequently discussed, all cases were negative for known viruses for which analyses were performed.
In 2004 the author identified the presence of BVDV in a white-tailed deer shortly following the unusual diagnosis by one of his colleagues of BVDV infection in two captive elk. As a result, in 2005 the author and his co-investigators conducted a study using the carcasses of 61 farm-raised white-tailed deer originating from Pennsylvania that were submitted for post mortem examination.2 Complete gross and histologic examinations were performed. Single-tube real-time reverse transcription polymerase chain reaction (real-time RT-PCR) for the detection of BVDV type 1 (BVDV-1) and type
2 (BVDV-2) and IHC staining of ear-notch skin to identify BVDV antigen was performed on each animal. Virus isolation was performed on tissue samples from 25 of
61 animals. All tissues samples tested negative for both BVDV-1 and BVDV-2 by real- time RT-PCR, virus isolation, and IHC. Gross or histopathologic lesions suggestive of 6
BVDV infection were not detected. Results of this study suggested that BVD is not a common cause of mortality in farm-raised white-tailed deer in Pennsylvania.
The absence of BVDV in all tissues in this study suggested that fatal acute and persistent BVDV infections are uncommon in farm-raised white-tailed deer in
Pennsylvania. The samples were collected from deer that died of unknown causes, frequently after a period of illness and it was hypothesized that some cases may have been the result of acute or persistent BVDV infections. With a sample size of 61 animals out of an estimated population of 25,000, the authors concluded with 95% confidence that the prevalence of BVDV in the necropsy population of captive white-tailed deer was not greater than 1.0%. Assuming that the presence of BVDV is positively associated with the risk of mortality in captive deer, infected animals would more likely be detected in this necropsy population than in animals randomly selected from the captive population. These findings suggested that, although cervids are susceptible to infection with BVDV as demonstrated in previous studies, BVDV is not a common cause of mortality in Pennsylvania farm-raised white-tailed deer, and that the prevalence of infection in the general population is less than 1%.
Having found no supportive evidence of any viral etiology, the author began to observe microscopic lung changes in several deer with pneumonia which resembled those of cattle with BRSV infection. Thus, the author considered that BRSV, a virus which commonly causes pneumonia in cattle, should be further investigated in deer. In 2008, an undergraduate student investigator and the author evaluated tissues from 53 wild and captive white-tailed deer for the presence of BRSV using PCR and IHC.3 All samples were negative by PCR, and 2/49 (4.1%) tested positive for BRSV by IHC. One positive 7
case was a wild deer with no gross or microscopic evidence of pneumonia and unrelated
traumatic cause of death, and the other positive case was a captive deer with pneumonia,
but lacking the characteristic microscopic changes of BRSV infection. There was no
significant association between the presence of BRSV and pneumonia, and it was
determined that the prevalence of BRSV infection in the Pennsylvania deer population is
<8% using IHC results or <0.55% using PCR results. Thus the significance of this
finding is undetermined and may represent a transient infection.
As a result of these preliminary analyses, no strong evidence of a viral cause for respiratory infection in deer was identified. Therefore, the author decided to pursue the hypothesis that the cause of the respiratory disease complex was primarily bacterial.
1.5 Objectives
These initial observations and preliminary studies resulted in a series of additional studies of various aspects of respiratory disease and fusobacterial infection of deer. The objectives of the following series of studies were:
1) To determine the current management practices and animal health problems on white-tailed deer farms in Pennsylvania.
2) To phenotypically and genotypically characterize a set of Fusobacterium isolates from white-tailed deer and evaluate their association with disease in the host.
3) To determine the pathogenicity of Fusobacterium varium to the respiratory tract of mice and to evaluate this system as a model for pulmonary fusobacterial infection of deer.
8
1.6 References
1. Hattel AL, Shaw DP, Love BC, et al. A retrospective study of mortality in
Pennsylvania captive white-tailed deer (Odocoileus virginianus): 20000--2003. J Vet
Diagn Invest 2004;16:515-521.
2. Brooks JW, Key DW, Hattel AL, et al. Failure to detect bovine viral diarrhea virus in necropsied farm-raised white-tailed deer (Odocoileus virginianus) in
Pennsylvania. J Vet Diagn Invest 2007;19:298-300.
3. Nau M. Identifying the presence of bovine respiratory syncytial virus in
Pennsylvania white-tailed deer. Undergraduate Honors Thesis, Department of Veterinary and Biomedical Sciences/Schreyer Honors College. University Park, PA 16801: The
Pennsylvania State University, May 2009.
9
Chapter 2
REVIEW OF LITERATURE
10
2.1 Deer Farming Industry
White-tailed deer (Odocoileus virginianus) are a species of wild ruminant
commonly found in much of North America. They belong to the order Artiodactyla,
suborder Ruminantia, and family Cervidae, within which there are 17 genera comprising
40 species of deer world-wide.1 There are 30 recognized subspecies of white-tailed deer, the largest of which, Odocoileus virginianus borealis or the northern woodland white- tail, is found in Pennsylvania and across the northeastern United States and southeastern
Canada.1 White-tailed deer have traditionally been prized by such varied groups as
outdoorsmen, naturalists, sportsmen, gourmands, craftsmen, chefs, and conservationists
for their many attributes including large antlers, healthful savory venison, and attractive
sturdy hides, in addition to their natural wild majesty and gamesmanship. Consequently,
within the past two decades, the farming of white-tailed deer has become an established
and growing industry in North America in which Pennsylvania is a leader.2,3 In the most
recent survey conducted by the Pennsylvania Game Commission in 2002, the agency that
regulated white-tailed deer farms in the state at that time, there were 25,600 captive
cervids on 743 farms in Pennsylvania.4 In a 2006 industry report, Pennsylvania was
ranked 2nd in the United States in numbers of commercial deer and elk farms, 3rd in number of deer and elk sold, and 5th in total deer and elk kept; deer and elk farm total
sales were growing at a rate of 12% per year and were expected to reach $50 million by
2010.3 By the 2007 Census of Agriculture, the number of deer farms in Pennsylvania had increased to 810, equaling a 54% increase in farm numbers and a 33% increase in deer numbers since the previous Census of Agriculture in 2002.5 Products sold by deer farms
include animal sales such as breeding stock or hunting stock, materials such as antlers, 11
venison, hides, urine, or crafts made from deer tissues, and services such as hunting, deer observation, lodging, and dining.3
2.2 Animal Health in Deer Populations
As intensively managed livestock operations, white-tailed deer farms are subject to a large number of animal health risks including infectious diseases, nutritional imbalances, exposure to toxic substances, and behavioral or management errors. Several studies have been published in attempts to characterize the disease conditions and management practices on white-tailed deer farms 2,6-8, however most studies of deer health have examined either the longer standing red deer and elk farming industry popular in New Zealand, Canada, and the United Kingdom 9-12 or have examined only wild populations of white-tailed deer.13-16 Studies of farm-raised white-tailed deer from the United States and Canada have resulted in similar findings that identify pneumonia, gastrointestinal disease/diarrhea, trauma, and parasitism as the most common causes of death in these farmed populations.2,6,7 Among the most commonly isolated pathogenic organisms are Fusobacterium necrophorum and Arcanobacterium pyogenes which are commonly recovered from abscesses or lung lesions.2,7,17 Interestingly, there is a striking absence of any mention of viral pathogens in the bulk of the literature pertaining to farm- raised white-tailed deer. This author has observed a small number of cases each of rabies virus infection, bovine viral diarrhea virus (BVDV) infection, and bovine respiratory syncytial virus (BRSV) infection in deer and elk from Pennsylvania, but other suggestions of viral disease are largely absent with the exception of obvious outbreaks of 12
such viral diseases as epizootic hemorrhagic disease (EHDV), or the usually insignificant
skin lesions caused by deer papillomavirus.18-21
Disease conditions among farmed red deer and elk vary somewhat according to geographic location, for many of these herds are located in New Zealand, the United
Kingdom, or Canada, but, similar to white-tailed deer, consistently among the most prevalent were trauma, pneumonia, gastrointestinal disease/diarrhea, and parasitism.9-12
Diseases and mortality in wild populations of white-tailed deer have also been
extensively studied and differ significantly from those of farm-raised deer. Leading
causes of death among wild deer have been reported as hunting, predation by wild and
domestic canids, and motor vehicle-related trauma.13-16,22,23 Contagious diseases,
including bacterial, viral, or parasitic infections, occur in smaller numbers of wild deer.24
Similar to what is observed in farmed deer, viral infection does not appear to be a significant cause of clinical disease with the exception of such viruses as rabies virus,
EHDV, bluetongue virus (BTV), eastern equine encephalitis virus (EEEV), malignant catarrhal fever virus (MCFV), vesicular stomatitis virus (VSV), or rarely occurring viral foreign animal diseases such as foot and mouth disease (FMD), rinderpest, and peste de petite ruminants, all of which tend to cause infrequent but significant outbreaks with severe morbidity and/or mortality.19,25-29 Serologic studies of wild white-tailed deer populations have reported varied results, but percentages of animals with positive serum
titers for antibodies against BVDV, bovine herpesvirus type 1 (BHV1), or
parainfluenzavirus type 3 (PI3) were generally very low with the exception of an isolated
wild herd on a densely populated island.30-32 Among viral organisms, serum antibody
production to BVDV has been the most extensively studied. Although seropositivity in 13
white-tailed deer is typically low, rates of seropositivity in other cervid species have been
quite variable ranging from zero in some populations to 69% in caribou in Canada.33,34
Although clinical disease attributed to BVDV infection has rarely been reported, some cases of clinical illness and death have been described.18,35 Thus, the clinical significance of most viral infections in white-tailed deer, with the exception of those severe infections previously described, remains to be determined. Regardless, the common causes of mortality in wild deer, primarily various forms of trauma, are very different from the common causes of mortality in farm-raised deer.
2.3 Anaerobic Bacteria
Anaerobic bacteria are organisms that do not require molecular oxygen for growth and reproduction. Although oxygen is not required by any anaerobic species, various degrees of tolerance to oxygen exist among groups of bacteria. Those with the greatest tolerance for oxygen are the facultative anaerobes, capable of growing equally well in the presence or absence of oxygen. At the opposite extreme, those with the lowest tolerance to oxygen are the obligate anaerobes which can grow only in an anaerobic environment and will be killed by exposure to air for an amount of time that varies between species, but is typically between several minutes to several hours.36 Their intolerance to oxygen
likely stems in part from their lack of such enzymes as superoxide dismutase and
catalase, however some species do produce these enzymes and, thus, the mechanisms are
oxygen toxicity are not completely understood.36,37 Anaerobic microenvironments in healthy animals and humans are typically found in biofilms of the oronasal cavity, gastrointestinal tract, and urogenital tract where facultative anaerobes of the normal flora 14
scavenge the majority of available oxygen.36,38,39 Biofilms are microbial communities
that attach to surfaces and produce a supportive matrix of polysaccharides, nucleic acids,
and extracellular polymeric substances typically resulting in a polymicrobial population
containing both aerobes and anaerobes.40 On the mucosal surfaces of the body anaerobic
bacteria typically outnumber aerobes, thus, most anaerobic infections arise in proximity
to mucosal surfaces or extend into the deeper tissues or bloodstream from a portal of
entry within a mucosal surface.36,41 Anaerobic infections may be caused by either endogenous anaerobes that are part of the normal body flora or by exogenous anaerobes
that are typically found only outside of the body, although infection by endogenous
organisms is more common. Many anaerobic infections are polymicrobial, involving
multiple organisms that are typically mixtures of obligate anaerobes and facultative
anaerobes36,39 and symbiotic relationships have been demonstrated between various
bacterial species that can favor the development of the disease process and have
significance for their response to medical therapy.42-46
Perhaps some of the first anaerobic bacteria were discovered during the late 19th century when Loeffler identified filamentous gram-negative rods in necrotic tissues from calves with calf diphtheria in 1884.47 By injecting the organism into mice, he was able to produce necrotic purulent lesions from which he could recover bacteria similar in appearance. He was able to culture the organism in serum broth, but was unable to cultivate it on agar. In turn, Bang, Schmorl, Hallé, Veillon and Züber, and Courmont and
Cade also identified similar organisms from various human and animal tissues in the late
1800s.47 Despite the prior discovery of several clinical diseases associated with anaerobic bacterial infection in animals and humans, physicians largely dismissed 15
anaerobes as harmless commensal organisms until the mid to late 20th century.48 Prior to that time, most clinical bacteriology did not attempt to cultivate obligate anaerobic bacteria. During the 1970s, it was discovered that abscesses in mice could be produced by pure infection with single or combined species of obligate anaerobic bacteria.49,50
Soon after, the significance of anaerobic infections was perhaps most fully realized by the discovery that intra-abdominal sepsis in a rat model produced a two-phase disease process. The first phase of the disease was predominated by facultative bacteria such as
E. coli and other enterococci, while the second phase was predominated by obligate anaerobes such as Bacteroides fragilis and Fusobacterium sp.48,51,52 The two phases of
the disease process were clinically distinct and responded very differently to antimicrobial therapy. The awareness of the clinical significance of anaerobic infections, coupled with revolutionary developments in microbiologic techniques by Hungate and
Holdman and Moore, led to the dawn of the era of anaerobic bacteriology.53
2.3.1 The Fusobacteria
Historically there has been much controversy and confusion regarding the
taxonomy of anaerobic organisms, in part due to the limited methodologies for
cultivating and evaluating anaerobic bacteria. Today, owing largely to the advent of
molecular techniques for analyzing the similarity of conserved gene sequences of
bacterial isolates, the classification of the fusobacteria has been much improved, resulting
in a more concise genus.54 The genus Fusobacterium has long been classified as one of
several genera under the family Bacteroidaceae which included all gram-negative, non-
spore-forming, rod-shaped, anaerobic bacteria,53,55 however more recently the genus has 16
been reclassified under a new scheme that considers Fusobacteria as a unique phylum within which are found the class Fusobacteria, order Fusobacteriales, and family
Fusobacteriaceae which includes the genus Fusobacterium and ten other genera almost none of which are medically significant.56,57 Members of the genus Fusobacterium have classically been sub-defined as those gram-negative, non-spore-forming, rod-shaped, anaerobic bacteria that either had fusiform ends or produced large amounts of butyric acid.54 By using 16S rRNA gene sequences or 16S-23S rDNA internal transcribed spacer regions (ITS) sequencing, the genus has been more clearly defined and now includes 13 species.54,57 F. mortiferum, F. naviforme, F. nucleatum, F. periodonticum, F. ulcerans, and F. varium are primarily isolated from human clinical samples, while F. necrogenes,
F. equinum, F. simiae are primarily from animal infections. F. russi along with F. nucleatum are commonly found in cat abscesses, canine and feline flora, and dog and cat bite wounds of humans. F. gonidiaformans is occasionally isolated from normal and abnormal urogenital and intestinal tract samples.58 F. perfoetans has been isolated only from feces.54 F. equinum is a newly recognized species isolated from the normal and abnormal oral cavity of horses and is phenotypically and biochemically similar to F. necrophorum.59,60 F. necrophorum is a major human and animal pathogen as will be subsequently described.
2.3.2 Fusobacterium necrophorum
F. necrophorum is a pleomorphic fusobacterial species that is a normal inhabitant of the gastrointestinal tract, oropharyngeal cavity, and urogenital tract of animals and humans.61 As a result of the long-standing confusion surrounding the 17
nomenclature of the fusobacteria and other anaerobes, F. necrophorum in particular has
had a rather lengthy and tortuous succession of names perhaps numbering as many as
52.47 At various times the organism has been known as Actinomyces necrophorus,
Actinomyces pseudonecrophorus, Bacillus filiformis, Bacillus funduliformis, Bacillus fundibuliformis, Bacillus necrophorus, Bacillus necrosus, Bacillus pyogenes anaerobius,
Bacillus symbiophiles, Bacterium funduliformis, Bacterium fundibuliformis, Bacterium
necrophorum, Bacterium necrophorus, Bacteroides funduliformis, Bacteroides fundibuliformis, Bacteroides necrophorus, Bang’s necrosis bacillus, Corynebacterium necrophorum, Fusiformis hemolyticus, Fusiformis necrophorus, Necrobacterium
funduliforme, Necrobacterium necrophorus, Proactinomyces necrophorus,
Pseudobacterium funduliformis, Schmorl’s bacillus, Sphaerophorus funduliforme,
Sphaerophorus funduliformis, Sphaerophorus necrophorus, Sphaerophorus
pseudonecrophorus, Streptothrix cuniculi, Streptothrix necrophora, Streptothrix
necrophorus, and Streptothrix necupthora among others.47,53,62 Since the era of anaerobic
bacteriology of the mid to late 20th century, F. necrophorum was traditionally sub-
classified into the biotypes A, B, C, and AB. By more recent analysis via 16S rRNA
gene sequencing, these biotype designations have been replaced by new species and
subspecies assignments. F. necrophorum biotype A is now known as F. necrophorum
subspecies necrophorum, whereas biotype B is now known as F. necrophorum
subspecies funduliforme. F. necrophorum biotype C has been reclassified as F. varium
while the taxonomic status of biotype AB remains unresolved.58
F. necrophorum is well established as a causative agent of many necrotizing infections in animals and humans including liver abscesses in cattle, foot rot in domestic 18
ruminants, stomatitis and laryngitis in calves, dermatitis in horses, and necrotizing
infections of the head and neck in multiple species including pigs, marsupials, and many
species of deer.53,54,63,64 While the organism has long been known to plague domestic
cattle and sheep, it has only more recently become recognized as a significant pathogen
in cervids resulting in a severe infection known as necrobacillosis. This infection is a
syndrome characterized by purulent necrotic lesions most commonly affecting the mouth,
pharynx, head, neck, lung, liver, or feet and is one of the most important diseases of
farm-raised deer.2,17,64-68 In humans the organism is known to cause cancrum oris (noma) and a group of severe septicemic infections including necrobacillosis, post-anginal sepsis,
and Lemierre’s syndrome which are variously described clinical diagnoses within the
spectrum of disease caused by infection with F. necrophorum that results in some
combination of fusobacterial septicemia, thrombophlebitis especially of the internal
jugular vein, and metastatic infection in the lungs or other distant location.47,69 It has been recently ascertained that F. n. subsp. necrophorum is the major pathogenic subspecies found in animal infections, whereas F. n. subspecies funduliforme is the major subspecies found in human infections.47
F. necrophorum is believed to possess several virulence factors that may play various roles in the pathogenesis of infection. These include leukotoxin, lipopolysaccharide (LPS), hemolysin, hemagglutinin, dermonecrotic toxin, capsule, adhesions or pili, and several additional enzymes such as proteases and deoxyribonucleases. While each of these may contribute to entry and establishment of infection, leukotoxin is considered to be the major virulence factor involved in animal infections based on correlation between toxin production and abscess formation and 19
correlation between anti-leukotoxin antibodies and protection in challenge studies.58,61
Leukotoxin is a water soluble secreted protein exotoxin with a high molecular weight of approximately 336,000 Da.70 Leukotoxin activity in culture supernatant tends to peak
during the late log and early stationary growth phases, followed by a rapid decline.71 The leukotoxin protein most severely affects bovine polymorphonuclear cells (PMNs), causing activation and apoptosis at low concentrations or necrosis at higher concentrations.72 Leukotoxin production by F. n. subsp. necrophorum is typically many
times greater than leukotoxin production by F. n. subsp. funduliforme which may account
for the increased virulence of the former in animal infections.71,73 The role of leukotoxin,
however, in naturally occurring non-bovine animal infections or human infections
remains undetermined as one study reported that strains recovered from these infections
less commonly possessed a leukotoxin gene that was able to be detected by PCR.74
The gene encoding leukotoxin is located within the lktBAC operon which consists
of three genes, lktB, lktA, and lktC. These three genes correspond respectively to an outer
membrane protein with putative transporter function, the leukotoxin protein, and a
protein of unknown function. The leukotoxin gene consists of 9,726 bp encoding a protein of 3,241 amino acids that does not have any significant sequence similarity to any known leukotoxin of other bacteria.45,58,75 The lktBAC operon promoter region is distinct
for each of the two subspecies of F. necrophorum, differing in sequence, length, and
strength allowing for differentiation of the two subspecies by PCR based detection of the
leukotoxin gene promoter.73,76
Isolation and cultivation of F. necrophorum requires strict anaerobic conditions.
The organism usually grows well on a variety of media including blood agar, Brucella 20
agar, brain-heart infusion broth, and chopped meat medium. Colony morphology at 48 –
72 hours is notably different between the two subspecies (Figure 2.1). On blood agar F. n. subsp. necrophorum forms colonies that are circular with erose edges, umbonate,
approximately 3 – 5 mm in diameter, and grey, while F. n. subsp. funduliforme forms
colonies that are circular with entire edges, convex, approximately 1 mm in diameter,
sticky and yellow-grey. In PRAS-BHI broth at 12 – 24 hours F. n. subsp. necrophorum
forms turbid growth throughout the medium, while F. n. subsp. funduliforme forms a sediment at the bottom of the tube (Figure 2.2).55,60,77 Cellular morphology of F. n.
subsp. necrophorum is characterized by pleomorphic gram-negative bacilli
approximately 2 – 100 µm in length with round to tapered ends and a tendency to form
long filaments, while that of F. n. subsp. funduliforme is described as short curved bacilli
approximately 1 – 10 µm in length (Figure 2.3).36,77 Ultrastructural detail by electron
microscopy confirms these differences in cellular morphology and also demonstrates the
presence of extracellular material and interlinking cell surface projections on F. n. subsp.
funduliforme that may be associated with its tendency to form sediment in broth culture.78
Metabolic and biochemical characteristics of F. necrophorum that may be useful in clinical identification include the production of butyrate, the conversion of lactate and threonine to propionate, a positive indole reaction, and inability to grow in 20% bile.41
An additional aid in differentiation of the two subtypes is that F. n. subsp. necrophorum is both lipase positive and alkaline phosphatase positive while F. n. subsp. funduliforme is negative for alkaline phosphatase and has a variable lipase reaction.59,77
21
2.3.3 Fusobacterium varium
F. varium is an obligate anaerobe that is known to be an integral component of the
microflora of the gastrointestinal tract and may also be found in the oral cavity. It has an
antagonistic effect on such enteric pathogens as Shigella and Salmonella, and produces
large quantities of butyrate which has a protective anti-inflammatory effect on the colonic
mucosa.79 Although infection caused by F. varium is rare, it may become an
opportunistic pathogen and has been associated in humans with ulcerative colitis,
decubitus ulcers, intra-abdominal infections, and less frequently with colon cancer, intra-
ocular infections, and conjunctivitis.79-81 Animal infections with F. varium are similarly
rare. In one study the organism represented 11 of 1,097 (1%) Fusobacterium isolates
from clinical specimens taken mostly from abscesses and respiratory tract samples of a
group of animals including dogs, cats, horses, cattle, sheep, goats, and a variety of exotic
species.82 F. varium has been recovered from a lamb with necrotizing hepatitis and from multiple tissues of farm-raised white-tailed deer with necrobacillosis.17,83 Despite a small
number of clinical cases of F. varium associated disease, the organism has been
demonstrated to be non-pathogenic when injected intraperitoneally or intraportally into
mice.84 Little is known of any virulence factors of F. varium, although, in contrast to the
findings of other studies that suggest a protective effect of butyrate, one recent study has
identified butyric acid as the major toxin in the culture supernatant, implicating that it
may be a major factor in the pathogenesis of some F. varium related disease. This effect,
however, may be influenced by butyrate concentration or defective butyrate metabolism
by host cells.85 22
Isolation and cultivation of F. varium requires strict anaerobic conditions. Similar
to other fusobacteria, the organism usually grows well on a variety of media including
blood agar, Brucella agar, brain-heart infusion broth, and chopped meat medium. Growth
on blood agar at 48 – 72 hours results in colonies that are circular with entire to erose
margins, low convex, 1 – 2 mm in diameter, and translucent with grey centers (Figure
2.1). In PRAS-BHI broth at 12 – 24 hours F. varium forms turbid growth throughout the
medium similar to F. n. subsp. necrophorum (Figure 2.2). Cellular morphology is
described as gram-negative pleomorphic short bacilli of approximately 1 – 2 µm length
with a tendency to form pairs or short filaments (Figure 2.3).36 Metabolic and
biochemical characteristics of F. varium that may be useful in clinical identification
include the production of butyrate, the conversion of threonine to propionate, the
fermentation of glucose, fructose, and mannose, and the ability to grow in 20% bile.41
2.4 Lung Infection and Immunity
Throughout much of history, pneumonia has been one of the most significant causes of illness and death in humans and animals. In humans, pneumonia is the leading cause of death due to infectious disease in the industrialized world and the 3rd leading overall cause of death worldwide.86 In domestic livestock, pneumonia is one of the most significant causes of death and economic loss in herds of cattle, farm-raised deer, and
other species.2,87 The essential function of the lung requires that it be exposed to large
quantities of air inhaled from the external environment along with abundant particulate
matter, potentially containing pathogenic microorganisms or contamination from the flora of the upper respiratory tract. Yet the lung must clear any inhaled foreign materials and 23
organisms and maintain a sterile environment in order for respiration and gas exchange to
occur.88 In order to achieve this, the lung is well protected from infection by a
multifaceted system of innate and adaptive defense mechanisms that must be tightly
regulated to prevent infection while minimizing damage to the pulmonary tissues.
Among the first defenders of the respiratory tract are the mucociliary elevator of
the major airways, the surfactant layer of the alveoli, complement, bronchial and lung epithelium, alveolar macrophages, and PMNs.87,89 The mucociliary elevator is composed
of ciliated pseudostratified or columnar epithelial cells of the trachea and bronchi coated
by a layer of mucus secreted by goblet cells within the epithelium that is capable of
carrying debris upward out of the respiratory tract. Surfactant proteins A and D are
produced by alveolar type II pneumocytes and play a role in reducing surface tension and
maintaining the patency of the alveolar lumen. Alveolar macrophages reside within the
surfactant layer along the alveolar epithelium and, as the first line of phagocytic cells,
effectively phagocytize low numbers of invading microorganisms; when larger numbers of microbes are present, PMNs predominate as the major phagocytic cell in the lung.89,90
Alveolar macrophages and epithelial cells are able to recognize pathogens by interaction between their transmembrane and intracellular pattern recognition receptors (PRRs) and a number of conserved pathogen-associated molecular patterns (PAMPs) found on microbial surfaces.86 Activation of PRRs by PAMPs signals the release of chemokines,
cytokines, prostaglandins, leukotrienes, and antimicrobial proteins such as defensins,
cathelicidins, lysozyme, and lactoferrin which play roles in killing or inhibiting microbes,
cell signaling, and chemotaxis of phagocytic cells.89,90 A major family of PRRs are the
Toll-like receptors (TLRs) which differentially recognize conserved molecular structures 24
of bacterial and viral organisms such as lipoteichoic acid and peptidoglycan (TLR-1,2,6),
lipopolysaccharide (LPS) (TLR-4), double-stranded RNA (TLR-3), and flagellin (TLR-
5).86,91 Within the interstitium is an additional group of cells that can participate if an
organism invades the epithelium. These cells include interstitial macrophages, natural
killer cells, invariant natural killer cells, and mast cells which play variable roles in
microbial killing, cell lysis, and production of cytokines such as interferon-γ (IFN- γ),
interleukin (IL) -1, IL-4, IL-6, IL-18, and tumor necrosis factor-α (TNFα).89-91
The adaptive immune response is slower than the innate response, but it is more
specific and its effects are longer lasting with memory to protect against future
exposure.92 The major links between the innate and adaptive immune responses are
provided by invariant natural killer T cells (iNKT) and subepithelial dendritic cells (DC).
These cells are among the first cells to recognize organisms in the lower respiratory tract
and engage in antigen processing following PAMP recognition by PRR interaction. They
then, respectively, produce cytokines such as IFN- γ and IL-4 and traffic to draining lymph nodes to present antigen to T lymphocytes which traffic to the lung through the bloodstream where they will mature into CD4+ or CD8+ T cells.89,93,94 There CD4+ T cells generate a polarized type 1 or type 2 response by the profile of cytokines they secrete. Type 1 responses are typically characterized by production of IFN- γ, IL-2,
TNFα, and granulocyte/macrophage colony-stimulating factor (GM-CSF) resulting in
activation of a cell mediated response whereas type 2 responses are typically
characterized by production of IL-4, IL-5, IL-9, IL-10, and IL-13 and a subsequent
humoral response.94,95 Likewise B lymphocytes in draining lymph nodes become
activated, and as plasma cells in the lymph node or having migrated to the lung via the 25
bloodstream, produce immunoglobulin IgA predominately in the upper respiratory tract,
and IgM, and IgG predominately in the lower respiratory tract.89,95
Throughout the immune response it is critical that the pro-inflammatory and anti-
inflammatory effects be modulated to maximize control of infection and minimize host
tissue damage.96 There are multiple regulatory mechanisms in both the innate and
adaptive immune responses. The innate response is modulated in part by the IL-1
receptor-associated kinase (IRAK)-like molecule which inhibits signaling from PRRs and
cytokine receptors and down-regulates the TLR signaling cascade by blocking nuclear factor κB (NF-κB).89,91 The adaptive response is regulated through regulatory T lymphocytes and cytokines such as IL-10 and transforming growth factor-β (TGF-β).89
2.5 Animal Models of Bacterial Pulmonary Infection
As previously described, the upper and lower respiratory tract possesses multiple defense mechanisms to protect against microbial infection, however pneumonia continues to be a severe and common problem in humans and domestic animals. Failure of a single one of these defense mechanisms should not lead to systemic failure resulting in lung infection, but rather it is believed that multiple defense mechanisms must be negatively affected simultaneously. Furthermore, the number and virulence of the organism entering the lung will influence the effectiveness of the host response and resulting disease.88 Highly pathogenic organisms may alone be capable of inducing pneumonia following introduction to the lower respiratory tract, while less virulent opportunistic
pathogens require that several lung defense mechanisms become impaired in order to
establish infection.88,96 Lung defenses may fail for a number of reasons, but among the 26
most common known reasons are: failure of the mucociliary elevator as a result of
primary viral or bacterial infection or inhaled pollutants or very cold air that inhibits ciliated epithelial cells; inhibition of alveolar macrophages or neutrophils by viral infection or bacterial leukotoxins; and reduction of antimicrobial protein expression in response to hypercortisolemia resulting from physiologic stress.87,88
The most common cause of pneumonia in adult humans is bacterial infection,
most frequently by Streptococcus pneumoniae, Haemophilus influenzae, and
Mycoplasma pneumoniae.86 Although aspiration pneumonia and lung abscesses are most
frequently caused by infection with anaerobic bacteria,36 most cases of lung infection are associated with aerobes, thus most animal models of lung infection have been developed with aerobic bacteria. Perhaps the predominant animal model of bacterial lung infection is Streptococcus pneumoniae infection in the mouse. This highly virulent pathogen is
capable in many experimental systems of producing severe or fatal lung infections by
intranasal or aerosol inoculation alone, but the severity of the infection is enhanced by
pre-infection with influenza virus or by the depletion of neutrophils.97-99 Adherence of
Streptococcus pneumoniae to tracheal epithelium was enhanced and bacterial clearance
from the lung was impaired following exposure of tracheal epithelium to hydrochloric
acid as may occur with aspiration of gastric content.100,101 In a murine model of lung
infection with Pseudomonas aeruginosa, impairment of pulmonary defense mechanism
including suppression of both the inducible isoform of nitric oxide synthase (iNOS) and
TNF-α production resulted in fatal pneumonia following dexamethasone treatment.102
Despite the focus on innate immune response in pulmonary infection, in mouse models of lung infection by Bordetella parapertussis and Bordetella pertussis it has been 27
demonstrated that both innate and adaptive immunity, including both the cellular and
humoral arms, are required for clearance of the organism from the lung emphasizing the
importance of the memory response and vaccination in protection against pulmonary
pathogens.103
In addition to bacterial infection of the lung, acute lung injury (ALI) and the more
severe acute respiratory distress syndrome (ARDS) represent other often non-infectious
conditions in which there is lung injury, disruption of respiratory epithelium, and
pulmonary inflammation which may lead to sepsis and multiple organ dysfunction
syndrome and death.104 Lipopolysaccharide, a surface antigen from the wall of gram-
negative bacteria is known as a strong pro-inflammatory compound and has been used
intranasally or intratracheally to induce the neutrophil infiltration and lung injury of
ALI.105,106 Similarly, inhaled irritants such as nickel sulfate, polytetrafluoroethylene, and ozone have also produced varying degrees of ALI in susceptible mouse strains.107 These mechanisms of lung damage may also represent significant obstacles for pulmonary defense mechanisms and may be associated with the pathogenesis of some lung infections.
2.6 Animal Models of Anaerobic Bacterial Infection
The obligate anaerobe most commonly isolated from human clinical samples is
Bacteroides fragilis, which is associated with a variety of infectious processes including bacteremia, endocarditis, urogenital/perineal infections, intra-abdominal infections, pneumonia, and abscess formation in many tissues.36,108 Consequently, B. fragilis
infection is one of the most common animal models of anaerobic infection. Since, 28
however, it is well known that most anaerobic infections are polymicrobial, it naturally
follows that animals models including infection with other commonly encountered
anaerobes such as Fusobacterium have also been developed. Most anaerobic animal
models have used mice, rats, or rabbits to model such disease processes as intra-
abdominal sepsis, intra-abdominal abscess formation, lung abscess formation, antibiotic-
associated colitis, inflammatory bowel disease, and microbial synergy in polymicrobial
infections.46,48,109 Additive or synergistic interactions between and various facultative
anaerobic and obligate anaerobic bacteria have been demonstrated including A. pyogenes,
E. coli, B. fragilis, Fusobacterium and others.42,44,46,109 Interestingly, one study reports
that while clinical samples containing Bacteroides typically contained multiple species of
Bacteroides, the vast majority of samples containing Fusobacterium contained only a
single fusobacterial species.82 A model of anaerobic lung abscessation has been developed in the New Zealand white rabbit using intratracheal inoculation of endogenous oral anaerobes including Peptococcus morbillorum, F. nucleatum, Eubacterium lentum,
and B. fragilis, however no mouse or rat models of anaerobic pneumonia following nasal
or tracheal exposure have been developed.109
Numerous animal models of infection with F. necrophorum have been developed
using cattle, mice such as BALB/c, ICR (CD-1), or Swiss Webster, or rabbits. The
majority of these studies have inoculated cattle by intraportal injection110 or mice by
intraperitoneal or intravenous routes to produce septicemia and intrahepatic or
intraabdominal abscesses53,61,77,111-118 or by subcutaneous route to produce localized or
multisystemic soft tissue infection.42,53,77,115-117,119 While some of these investigators
have also attempted intraportal, intrahepatic, or footpad inoculation of mice, no attempt at 29
intranasal or intratracheal inoculation with Fusobacterium has been reported. One study
evaluated the effects of intrathoracic administration of F. necrophorum in mice which resulted in a 3-4 day course of illness and death with necrotizing lung abscesses near the injection site.115 In most animal models, F. n. subsp. necrophorum has been
demonstrated to be more virulent than F. n. subsp. funduliforme, resulting in more severe lesions and mortality.74 Very few animal studies have been conducted using F. varium,
but the organism has long been considered non-pathogenic and not of great medical importance except for rarely occurring opportunistic infections. One study confirmed the non-virulent nature of F. varium infection by intraperitoneal and intraportal injection in mice, which failed to produce disease in any mice following intraportal infection and only produced liver abscessation in one of 15 mice following intraperitoneal infection.84
One study demonstrated that F. varium was capable of inducing subcutaneous abscesses in mice following subcutaneous inoculation.42 More recently a model of ulcerative colitis
has been proposed by intrarectal inoculation of F. varium in mice which suggests that
butyric acid may be an important virulence factor of this organism.85
The immune response to fusobacterial infection has been the focus of some
studies attempting to develop effective vaccine strategies for reducing the incidence and
severity of hepatic abscessation or footrot in feedlot cattle. It has been demonstrated in mice that severe F. necrophorum infection that was cleared by antimicrobial therapy provided no protective immunity to future F. necrophorum infection,119 but that
intraperitoneal immunization with killed whole cell F. necrophorum bacterin was
somewhat protective against challenge especially when enhanced by an aluminum
hydroxide adjuvant.111 Following a report that anti-leukotoxin immunity reduced liver 30
abscesses and footrot in cattle,120 most researchers have focused vaccination efforts on inducing immunity to leukotoxin. Since then, numerous other studies have demonstrated
the serum neutralizing antibody production and protective effects against liver abscesses
and footrot in cattle following administration of F. necrophorum toxoid vaccine.121-124
Protective effects and anti-leukotoxin antibody response have also been demonstrated in mice following immunization with various leukotoxin preparations including F. necrophorum culture supernatant, affinity purified leukotoxin, and numerous truncated recombinant leukotoxin proteins.61 One study, however, suggests that vaccination of non-bovine animal species against leukotoxin may be less protective against fusobacterial disease than in cattle.74 F. necrophorum bacterin vaccines have also reduced the prevalence of liver abscesses and footrot in cattle, although no direct comparative studies between toxoid vaccines and bacterin vaccines have been conducted.125 Similarly, F.
nucleatum bacterins have been protective against oral F. nucleatum infection and have
elicited serum neutralizing antibodies.126 Although much has been learned about the
immune response to fusobacterial infection, clearly much remains to be determined.
31
2.7 References
1. Rue LL, III. Varieties and Distribution. The Deer of North America. Danbury,
CT: Outdoor Life Books, 1989;18-38.
2. Hattel AL, Shaw DP, Love BC, et al. A retrospective study of mortality in
Pennsylvania captive white-tailed deer (Odocoileus virginianus): 2000-2003. J
Vet Diagn Invest 2004;16:515-521.
3. PDFA. The economic impact of Pennsylvania's deer farms: Pennsylvania Deer
Farmers Association, Nov 2006.
4. Rosenberry C, Boyd R, Houghton G. 2002 white-tailed deer and elk propagator
inventory: Pennsylvania Game Commission, June 2003.
5. United States Department of Agriculture NASS. 2007 Census of Agriculture, Dec
2009.
6. Bruning-Fann CS, Shank KL, Kaneene JB. Descriptive epidemiology of captive
cervid herds in Michigan, USA. Vet Res 1997;28:295-302.
7. Haigh J, Berezowski J, Woodbury MR. A cross-sectional study of the causes of
morbidity and mortality in farmed white-tailed deer. Can Vet J 2005;46:507-512.
8. Haigh J, Berezowski J, Woodbury MR. A cross-sectional study of reproductive
indices and fawn mortality in farmed white-tailed deer. Can Vet J 2005;46:413-
416.
9. Audige L, Wilson PR, Morris RS. Disease and mortality on red deer farms in
New Zealand. Vet Rec 2001;148:334-340.
10. Fletcher TJ. Management problems and disease in farmed deer. Vet Rec
1982;111:219-223. 32
11. Pople NC, Allen AL, Woodbury MR. A retrospective study of neonatal mortality
in farmed elk. Can Vet J 2001;42:925-928.
12. Woodbury MR, Berezowski J, Haigh J. A retrospective study of the causes of
morbidity and mortality in farmed elk (Cervus elaphus). Can Vet J 2005;46:1108-
1113, 1120-1101.
13. DelGiudice GD, Riggs MR, Joly P, et al. Winter severity, survival, and cause-
specific mortality of female white-tailed deer in north-central Minnesota. Journal
of Wildlife Management 2002;66:698-717.
14. Nixon CM, Hansen LP, Brewer PA, et al. Survival of white-tailed deer in
intensively farmed areas of Illinois. Canadian Journal of Zoology 2001;79:581-
588.
15. Van Deelen TR, Campa H, III, Haufler JB, et al. Mortality patterns of white-tailed
deer in Michigan's Upper Peninsula. Journal of Wildlife Management
1997;61:903-910.
16. Whitlaw HA, Ballard WB, Sabine DL, et al. Survival and cause-specific mortality
rates of adult white-tailed deer in New Brunswick. Journal of Wildlife
Management 1998;62:1335-1341.
17. Chirino-Trejo M, Woodbury MR, Huang F. Antibiotic sensitivity and biochemical
characterization of Fusobacterium spp. and Arcanobacterium pyogenes isolated
from farmed white-tailed deer (Odocoileus virginianus) with necrobacillosis. J
Zoo Wildl Med 2003;34:262-268. 33
18. Brooks JW, Key DW, Hattel AL, et al. Failure to detect bovine viral diarrhea
virus in necropsied farm-raised white-tailed deer (Odocoileus virginianus) in
Pennsylvania. J Vet Diagn Invest 2007;19:298-300.
19. Howerth EW, Stallknecht DE, Kirkland PD. Bluetongue, Epizootic Hemorrhagic
Disease, and Other Orbivirus-Related Diseases In: Williams ES,Barker IK, eds.
Infectious Diseases of Wild Mammals. Ames, IA: Iowas State Press, 2001;77-97.
20. Nau M. Identifying the presence of bovine respiratory syncytial virus in
Pennsylvania white-tailed deer. Undergraduate Honors Thesis, Department of
Veterinary and Biomedical Sciences/Schreyer Honors College. University Park,
PA 16801: The Pennsylvania State University, May 2009.
21. Sundberg JP, Ranst MV, Jenson AB. Papillomavirus Infections In: Williams
ES,Barker IK, eds. Infectious Diseases of Wild Mammals. Ames, IA: Iowa State
Press, 2001;223-231.
22. Nelson ME, Mech LD. Mortality of white-tailed deer in northeastern Minnesota.
Journal of Wildlife Management 1986;50:691-698.
23. Nelson TA, Woolf A. Mortality of white-tailed deer fawns in southern Illinois.
Journal of Wildlife Management 1987;51:326-329.
24. Reed DE, Shave H, Bergeland ME, et al. Necropsy and laboratory findings in
free-living deer in South Dakota. J Am Vet Med Assoc 1976;169:975-979.
25. Heuschele WP, Reid HW. Malignant Catarrhal Fever In: Williams ES,Barker IK,
eds. Infectious Diseases of Wild Mammals. Ames, IA: Iowa State Press,
2001;157-164. 34
26. Rossiter P. Morbilliviral Diseases In: Williams ES,Barker IK, eds. Infectious
Diseases of Wild Mammals. Ames, IA: Iowa State Press, 2001;37-76.
27. Schmitt SM, Cooley TM, Fitzgerald SD, et al. An outbreak of Eastern equine
encephalitis virus in free-ranging white-tailed deer in Michigan. J Wildl Dis
2007;43:635-644.
28. Thompson GR, Bengis RG, Brown CC. Picornavirus Infections In: Williams
ES,Barker IK, eds. Infectious Diseases of Wild Mammals. Ames, IA: Iowa State
Press, 2001;119-130.
29. Yuill TM, Seymour C. Arbovirus Infections In: Williams ES,Barker IK, eds.
Infectious Diseases of Wild Mammals. Ames, IA: Iowa State Press, 2001.
30. Davidson WR, Crow CB. Parasites, diseases, and health status of sympatric
populations of sika deer and white-tailed deer in Maryland and Virginia. J Wildl
Dis 1983;19:345-348.
31. Kahrs R, Atkinson G, Baker JA, et al. Serological Studies on the Incidence of
Bovine Virus Diarrhea, Infectious Bovine Rhinotracheitis, Bovine Myxovirus
Parainfluenza-3, and Leptospira Pomona in New York State. Cornell Vet
1964;54:360-369.
32. Sadi L, Joyal R, St-Georges M, et al. Serologic survey of white-tailed deer on
Anticosti Island, Quebec for bovine herpesvirus 1, bovine viral diarrhea, and
parainfluenza 3. J Wildl Dis 1991;27:569-577.
33. Elazhary MA, Frechette JL, Silim A, et al. Serological evidence of some bovine
viruses in the caribou (Rangifer tarandus caribou) in Quebec. J Wildl Dis
1981;17:609-612. 35
34. Van Campen H, Williams ES, Edwards J, et al. Experimental infection of deer
with bovine viral diarrhea virus. J Wildl Dis 1997;33:567-573.
35. Chase CCL, Braun LJ, Leslie-Steen P, et al. Evidence of bovine viral diarrhea
virus persistent infection in two white-tailed deer in southeastern South Dakota.
American Association of Bovine Practitioners Annual Conference 2004;169.
36. Englekirk PG, Duben-Englekirk J, Dowell VR. Principles and Practice of
Clinical Anaerobic Bacteriology. Belmont, CA: Star Publishing Co., 1992.
37. Quinn PJ, Markey BK, Carter ME, et al. Veterinary Microbiology and Microbial
Disease. Ames, IA: Blackwell, 2004.
38. Carter GR, Chengappa MM, Roberts AW. Essentials of Veterinary Microbiology.
5th ed. Philadelphia, PA: William and Wilkins, 1995.
39. Donlan RM, Costerton JW. Biofilms: survival mechanisms of clinically relevant
microorganisms. Clin Microbiol Rev 2002;15:167-193.
40. Macassey E, Dawes P. Biofilms and their role in otorhinolaryngological disease. J
Laryngol Otol 2008;122:1273-1278.
41. Jousimies-Somer H, Summanen P, Citron DM, et al. Wadsworth-KTL Anaerobic
Bacteriology Manual. 6th ed. Belmont, CA, U.S.A.: Star Pub. Co., 2002.
42. Brook I, Walker RI. The relationship between Fusobacterium species and other
flora in mixed infection. J Med Microbiol 1986;21:93-100.
43. Periasamy S, Kolenbrander PE. Aggregatibacter actinomycetemcomitans builds
mutualistic biofilm communities in saliva with Fusobacterium nucleatum and
Veillonella sp. Infect Immun 2009. 36
44. Smith GR, Till D, Wallace LM, et al. Enhancement of the infectivity of
Fusobacterium necrophorum by other bacteria. Epidemiol Infect 1989;102:447-
458.
45. Tadepalli S, Narayanan SK, Stewart GC, et al. Fusobacterium necrophorum: a
ruminal bacterium that invades liver to cause abscesses in cattle. Anaerobe
2009;15:36-43.
46. Onderdonk AB, Bartlett JG, Louie T, et al. Microbial synergy in experimental
intra-abdominal abscess. Infect Immun 1976;13:22-26.
47. Riordan T. Human infection with Fusobacterium necrophorum (Necrobacillosis),
with a focus on Lemierre's syndrome. Clin Microbiol Rev 2007;20:622-659.
48. Onderdonk AB. Animal models simulating anaerobic infections. Anaerobe
2005;11:189-195.
49. Hill GB, Osterhout S, Pratt PC. Liver abscess production by non-spore-forming
anaerobic bacteria in a mouse model. Infect Immun 1974;9:599-603.
50. Wilkins TD, Walker CB, Nitzan D, et al. Experimental infections with anaerobic
bacteria in mice. J Infect Dis 1977;135 Suppl:S13-17.
51. Onderdonk AB, Weinstein WM, Sullivan NM, et al. Experimental intra-
abdominal abscesses in rats: quantitative bacteriology of infected animals. Infect
Immun 1974;10:1256-1259.
52. Weinstein WM, Onderdonk AB, Bartlett JG, et al. Experimental intra-abdominal
abscesses in rats: development of an experimental model. Infect Immun
1974;10:1250-1255. 37
53. Langworth BF. Fusobacterium necrophorum: its characteristics and role as an
animal pathogen. Bacteriol Rev 1977;41:373-390.
54. Citron DM. Update on the taxonomy and clinical aspects of the genus
Fusobacterium. Clin Infect Dis 2002;35:S22-27.
55. Moore WEC, Holdeman LV, Kelley RW. Fusobacterium In: Krieg NR,Hold J,
G., eds. Bergey's Manual of Systemic Bacteriology. Baltimore, MD: Williams and
Wilkins, 1984;631-637.
56. National Library of Medicine NIH. NCBI taxonomy browser.
http://ncbi.nlm.nih.gov/Taxonomy/Browser, April 9, 2010.
57. Songer JG, Post KW. Veterinary Microbiology, Bacterial and Fungal Agents of
Animal Disease. St. Louis, MO: Elsevier Saunders, 2005.
58. Nagaraja TG, Narayanan SK, Stewart GC, et al. Fusobacterium necrophorum
infections in animals: pathogenesis and pathogenic mechanisms. Anaerobe
2005;11:239-246.
59. Dorsch M, Lovet DN, Bailey GD. Fusobacterium equinum sp. nov., from the oral
cavity of horses. Int J Syst Evol Microbiol 2001;51:1959-1963.
60. Tadepalli S, Stewart GC, Nagaraja TG, et al. Fusobacterium equinum possesses a
leukotoxin gene and exhibits leukotoxin activity. Vet Microbiol 2008;127:89-96.
61. Narayanan SK, Chengappa MM, Stewart GC, et al. Immunogenicity and
protective effects of truncated recombinant leukotoxin proteins of Fusobacterium
necrophorum in mice. Vet Microbiol 2003;93:335-347.
62. Brazier JS. Human infections with Fusobacterium necrophorum. Anaerobe
2006;12:165-172. 38
63. Ramos-Vara JA, Duran O, Render JA, et al. Necrotising stomatitis associated
with Fusobacterium necrophorum in three sows. Vet Rec 1998;143:282-283.
64. Leighton FA. Fusobacterium necrophorum infection In: Williams ES,Barker IK,
eds. Infectious Diseases of Wild Mammals. Ames, Iowa: Iowa State Press,
2001;493-496.
65. Edwards JF, Davis DS, Roffe TJ, et al. Fusobacteriosis in captive wild-caught
pronghorns (Antilocapra americana). Vet Pathol 2001;38:549-552.
66. Ramos-Vara JA, Ganjam I, Fales W. Pulmonary necrobacillosis in a white-tailed
deer. Vet Rec 2003;152:401-403.
67. Roeder BL, Chengappa MM, Lechtenberg KF, et al. Fusobacterium necrophorum
and Actinomyces pyogenes associated facial and mandibular abscesses in blue
duiker. J Wildl Dis 1989;25:370-377.
68. Wobeser G, Runge W, Noble D. Necrobacillosis in deer and pronghorn antelope
in Saskatchewan. Can Vet J 1975;16:3-9.
69. Enwonwu CO, Falkler WA, Jr., Idigbe EO, et al. Pathogenesis of cancrum oris
(noma): confounding interactions of malnutrition with infection. Am J Trop Med
Hyg 1999;60:223-232.
70. Tan ZL, Nagaraja TG, Chengappa MM, et al. Biological and biochemical
characterization of Fusobacterium necrophorum leukotoxin. Am J Vet Res
1994;55:515-521.
71. Tan ZL, Nagaraja TG, Chengappa MM. Factors affecting the leukotoxin activity
of Fusobacterium necrophorum. Vet Microbiol 1992;32:15-28. 39
72. Narayanan S, Stewart GC, Chengappa MM, et al. Fusobacterium necrophorum
leukotoxin induces activation and apoptosis of bovine leukocytes. Infect Immun
2002;70:4609-4620.
73. Tadepalli S, Stewart GC, Nagaraja TG, et al. Leukotoxin operon and differential
expressions of the leukotoxin gene in bovine Fusobacterium necrophorum
subspecies. Anaerobe 2008;14:13-18.
74. Ludlam HA, Milner NJ, Brazier JS, et al. lktA-encoded leukotoxin is not a
universal virulence factor in invasive Fusobacterium necrophorum infections in
animals and man. J Med Microbiol 2009;58:529-530.
75. Narayanan SK, Nagaraja TG, Chengappa MM, et al. Cloning, sequencing, and
expression of the leukotoxin gene from Fusobacterium necrophorum. Infect
Immun 2001;69:5447-5455.
76. Zhang F, Nagaraja TG, George D, et al. The two major subspecies of
Fusobacterium necrophorum have distinct leukotoxin operon promoter regions.
Vet Microbiol 2006;112:73-78.
77. Tan ZL, Nagaraja TG, Chengappa MM. Fusobacterium necrophorum infections:
virulence factors, pathogenic mechanism and control measures. Vet Res Commun
1996;20:113-140.
78. Garcia MM, Becker SA, Brooks BW, et al. Ultrastructure and molecular
characterization of Fusobacterium necrophorum biovars. Can J Vet Res
1992;56:318-325. 40
79. Potrykus J, White RL, Bearne SL. Proteomic investigation of amino acid
catabolism in the indigenous gut anaerobe Fusobacterium varium. Proteomics
2008;8:2691-2703.
80. Legaria MC, Lumelsky G, Rodriguez V, et al. Clindamycin-resistant
Fusobacterium varium bacteremia and decubitus ulcer infection. J Clin Microbiol
2005;43:4293-4295.
81. Minami M, Ando T, Okamoto A, et al. Seroprevalence of Fusobacterium varium
in ulcerative colitis patients in Japan. FEMS Immunol Med Microbiol 2009;56:67-
72.
82. Jang SS, Hirsh DC. Characterization, distribution, and microbiological
associations of Fusobacterium spp. in clinical specimens of animal origin. J Clin
Microbiol 1994;32:384-387.
83. Foster AP, Otter A, Naylor R, et al. Hepatitis in a six-month-old lamb with
Fusobacterium varium infection. Vet Rec 2009;164:98.
84. Hiraiwa K, Shinjo T. Pathogenicity of Fusobacterium necrophorum biovar C in
mice by intraperitoneal and intraportal injections. Nippon Juigaku Zasshi
1989;51:1111-1114.
85. Ohkusa T, Okayasu I, Ogihara T, et al. Induction of experimental ulcerative
colitis by Fusobacterium varium isolated from colonic mucosa of patients with
ulcerative colitis. Gut 2003;52:79-83.
86. Hippenstiel S, Opitz B, Schmeck B, et al. Lung epithelium as a sentinel and
effector system in pneumonia--molecular mechanisms of pathogen recognition
and signal transduction. Respir Res 2006;7:97. 41
87. Al-Haddawi M, Mitchell GB, Clark ME, et al. Impairment of innate immune
responses of airway epithelium by infection with bovine viral diarrhea virus. Vet
Immunol Immunopathol 2007;116:153-162.
88. Caswell JL, al-Haddawi M. If lung defenses are this sophisticated, why is
bronchopneumonia so pervasive? ACVP Annual Conference 2006;166-171.
89. Lipscomb MF, Hutt J, Lovchik J, et al. The pathogenesis of acute pulmonary viral
and bacterial infections: investigations in animal models. Annu Rev Pathol
2010;5:223-252.
90. van der Poll T, Opal SM. Pathogenesis, treatment, and prevention of
pneumococcal pneumonia. Lancet 2009;374:1543-1556.
91. Calbo E, Garau J. Of mice and men: innate immunity in pneumococcal
pneumonia. Int J Antimicrob Agents 2010;35:107-113.
92. Tsai KS, Grayson MH. Pulmonary defense mechanisms against pneumonia and
sepsis. Curr Opin Pulm Med 2008;14:260-265.
93. Boyton R. The role of natural killer T cells in lung inflammation. J Pathol
2008;214:276-282.
94. Curtis JL. Cell-mediated adaptive immune defense of the lungs. Proc Am Thorac
Soc 2005;2:412-416.
95. Goldsby RA, Kindt TJ, Osborne BA, et al. Immunology. 5th ed. New York, NY:
W.H. Freeman and Co., 2003.
96. Si-Tahar M, Touqui L, Chignard M. Innate immunity and inflammation--two
facets of the same anti-infectious reaction. Clin Exp Immunol 2009;156:194-198. 42
97. Bergeron Y, Ouellet N, Deslauriers AM, et al. Cytokine kinetics and other host
factors in response to pneumococcal pulmonary infection in mice. Infect Immun
1998;66:912-922.
98. Nuermberger E, Helke K, Bishai WR. Low-dose aerosol model of pneumococcal
pneumonia in the mouse: utility for evaluation of antimicrobial efficacy. Int J
Antimicrob Agents 2005;26:497-503.
99. Smith MW, Schmidt JE, Rehg JE, et al. Induction of pro- and anti-inflammatory
molecules in a mouse model of pneumococcal pneumonia after influenza. Comp
Med 2007;57:82-89.
100. Ishizuka S, Yamaya M, Suzuki T, et al. Acid exposure stimulates the adherence of
Streptococcus pneumoniae to cultured human airway epithelial cells: effects on
platelet-activating factor receptor expression. Am J Respir Cell Mol Biol
2001;24:459-468.
101. Johanson WG, Jr., Stephen JJ, Pierce AK. Bacterial growth in vivo. An important
determinant of the pulmonary clearance of Diplococcus pneumoniae in rats. J
Clin Invest 1974;53:1320-1325.
102. Satoh S, Oishi K, Iwagaki A, et al. Dexamethasone impairs pulmonary defence
against Pseudomonas aeruginosa through suppressing iNOS gene expression and
peroxynitrite production in mice. Clin Exp Immunol 2001;126:266-273.
103. Wolfe DN, Kirimanjeswara GS, Harvill ET. Clearance of Bordetella
parapertussis from the lower respiratory tract requires humoral and cellular
immunity. Infect Immun 2005;73:6508-6513. 43
104. Zhang X, Song K, Xiong H, et al. Protective effect of florfenicol on acute lung
injury induced by lipopolysaccharide in mice. Int Immunopharmacol 2009.
105. Starcher B, Williams I. A method for intratracheal instillation of endotoxin into
the lungs of mice. Lab Anim 1989;23:234-240.
106. Szarka RJ, Wang N, Gordon L, et al. A murine model of pulmonary damage
induced by lipopolysaccharide via intranasal instillation. J Immunol Methods
1997;202:49-57.
107. Wesselkamper SC, Prows DR, Biswas P, et al. Genetic susceptibility to irritant-
induced acute lung injury in mice. Am J Physiol Lung Cell Mol Physiol
2000;279:L575-582.
108. Onderdonk AB, Markham RB, Zaleznik DF, et al. Evidence for T cell-dependent
immunity to Bacteroides fragilis in an intraabdominal abscess model. J Clin
Invest 1982;69:9-16.
109. Kannangara DW, Thadepalli H, Bach VT, et al. Animal model for anaerobic lung
abscess. Infect Immun 1981;31:592-597.
110. Lechtenberg KF, Nagaraja TG. Hepatic ultrasonography and blood changes in
cattle with experimentally induced hepatic abscesses. Am J Vet Res 1991;52:803-
809.
111. Abe PM, Holland JW, Stauffer LR. Immunization of mice against Fusobacterium
necrophorum infection by parenteral or oral administration of vaccine. Am J Vet
Res 1978;39:115-118. 44
112. Abe PM, Lennard ES, Holland JW. Fusobacterium necrophorum infection in
mice as a model for the study of liver abscess formation and induction of
immunity. Infect Immun 1976;13:1473-1478.
113. Abe PM, Majeski JA, Lennard ES. Pathological changes produced by
Fusobacterium necrophorum in experimental infection of mice. J Comp Pathol
1976;86:365-369.
114. Conlon PJ, Hepper KP, Teresa GW. Evaluation of experimentally induced
Fusobacterium necrophorum infections in mice. Infect Immun 1977;15:510-517.
115. Maestrone G, Sadek S, Kubacki E, et al. Sphaerophorus necrophorus: laboratory
model for the evaluation of chemotherapeutic agents in mice. Cornell Vet
1975;65:187-204.
116. Smith GR, Thornton EA. Pathogenicity of Fusobacterium necrophorum strains
from man and animals. Epidemiol Infect 1993;110:499-506.
117. Smith GR, Thornton EA. Classification of human and animal strains of
Fusobacterium necrophorum by their pathogenic effects in mice. J Med
Microbiol 1997;46:879-882.
118. Berg JN, Scanlan CM. Studies of Fusobacterium necrophorum from bovine
hepatic abscesses: biotypes, quantitation, virulence, and antibiotic susceptibility.
Am J Vet Res 1982;43:1580-1586.
119. Smith GR, Wallace LM, Till D. Necrobacillosis and immunity in mice. Epidemiol
Infect 1989;103:211-215. 45
120. Garcia MM, Dorward WJ, Alexander DC, et al. Results of a preliminary trial with
Sphaerophorus necrophorus toxoids to control liver abscesses in feedlot cattle.
Can J Comp Med 1974;38:222-226.
121. Clark BL, Emery DL, Stewart DJ, et al. Studies into immunisation of cattle
against interdigital necrobacillosis. Aust Vet J 1986;63:107-110.
122. Saginala S, Nagaraja TG, Lechtenberg KF, et al. Effect of Fusobacterium
necrophorum leukotoxoid vaccine on susceptibility to experimentally induced
liver abscesses in cattle. J Anim Sci 1997;75:1160-1166.
123. Saginala S, Nagaraja TG, Tan ZL, et al. The serum neutralizing antibody response
in cattle to Fusobacterium necrophorum leukotoxoid and possible protection
against experimentally induced hepatic abscesses. Vet Res Commun 1996;20:493-
504.
124. Saginala S, Nagaraja TG, Tan ZL, et al. Serum neutralizing antibody response and
protection against experimentally induced liver abscesses in steers vaccinated
with Fusobacterium necrophorum. Am J Vet Res 1996;57:483-488.
125. Checkley SL, Janzen ED, Campbell JR, et al. Efficacy of vaccination against
Fusobacterium necrophorum infection for control of liver abscesses and footrot in
feedlot cattle in western Canada. Can Vet J 2005;46:1002-1007.
126. Liu PF, Haake SK, Gallo RL, et al. A novel vaccine targeting Fusobacterium
nucleatum against abscesses and halitosis. Vaccine 2009;27:1589-1595.
46
Figure 2.1
Fusobacterium necrophorum subsp. necrophorum growth on blood agar. 47
Figure 2.2
Fusobacterium necrophorum subsp. funduliforme growth on blood agar.
48
Figure 2.3
Fusobacterium varium growth on blood agar.
49
Figure 2.4
Fusobacterium spp. growth in PRAS-BHI broth: Left vial F. n. necrophorum, center vial F. n. funduliforme, right vial F. varium.
50
Figure 2.5
F. n. necrophorum cellular morphology (Gram Stain).
51
Figure 2.6
F. n. funduliforme cellular morphology (Gram Stain).
52
Figure 2.7
F. varium cellular morphology (Gram Stain).
53
Chapter 3
MANAGEMENT PRACTICES USED BY WHITE-TAILED DEER FARMS IN
PENNSYLVANIA AND HERD HEALTH PROBLEMS
Jason W. Brooks and Bhushan M. Jayarao
The Animal Diagnostic Laboratory (Brooks) and the Department of Veterinary and
Biomedical Sciences (Jayarao), College of Agricultural Sciences, The Pennsylvania State
University, University Park, PA 16802.
Published in JAVMA, Vol 232, No. 1, January 2008, 98-104.
54 55 56 57 58 59 60 61
Chapter 4
PHENOTYPIC AND GENOTYPIC CHARACTERIZATION OF
FUSOBACTERIUM ISOLATES FROM THE RESPIRATORY TRACT OF DEER
Jason W. Brooks a,*, Bhushan M. Jayarao a, Amit Kumar b, Sanjeev Narayanan b,
Suzanne Myers a, T.G. Nagaraja b
a Department of Veterinary and Biomedical Sciences, College of Agricultural Sciences,
The Pennsylvania State University, University Park, PA 16802
b Department of Diagnostic Medicine/Pathobiology, College of Veterinary Medicine,
Kansas State University, Manhattan, KS, 66506
Submitted for Publication to Veterinary Microbiology, February 2010.
62
4.1 Abstract
A total of 28 clinical strains of Fusobacterium spp. were isolated at necropsy over a two-year period from the respiratory tract of cervids, including white-tailed deer, elk, and reindeer. Isolates were identified as F. varium (21/28, 75%), F. necrophorum subsp. funduliforme (5/28, 17.9%), and F. necrophorum subsp. necrophorum (2/28, 7.1%).
Using PCR-based detection of virulence genes, all F. varium isolates were negative for the promoter region of the leukotoxin operon of F. necrophorum and the hemagglutinin- related protein gene of F. necrophorum. In the necropsy population no significant differences in gross or microscopic lesions were detected across Fusobacterium species, suggesting similar potential for virulence, however toxicity to bovine polymorphonuclear leukocytes was not observed in any F. varium strains, perhaps indicating that a virulence factor other than leukotoxin is involved in the pathogenesis of F. varium infection. F. varium was less susceptible to many antimicrobials than were the F. necrophorum subspecies. These data suggest that F. varium may be a significant pathogen in deer and may require different treatment and prevention methods than F. necrophorum.
Keywords: Fusobacterium necrophorum, Fusobacterium varium, necrobacillosis, antimicrobial susceptibility, leukotoxin, pneumonia, white-tailed deer, Odocoileus virginianus
4.2 Introduction
Fusobacterium spp. are gram negative, non-spore-forming, anaerobic, rod-shaped bacteria of the Bacteroidaceae family. F. necrophorum, which is currently recognized as 63
two subspecies, F. n. subsp. necrophorum and F. n. subsp. funduliforme, is one of the
most common anaerobic bacteria isolated from abscesses, respiratory tract infections, and
other necrotizing infections in domestic livestock, wild mammals, and humans.1-4 While its role in hepatic abscessation and foot rot in cattle and small ruminants is well recognized, its role in severe respiratory tract infections in deer is less completely characterized.5,6 Reports of infection with F. varium are relatively rare in the literature,
and the organism has been associated with ulcerative colitis, decubitus ulcers, and
respiratory tract infections in humans and with suppurative or ulcerative lesions of the
gastrointestinal tract or oral cavity in animals.7-10 In addition, various Fusobacterium spp. have been isolated from many wild ruminants affected by necrobacillosis, a syndrome that is characterized by purulent necrotic lesions most commonly affecting the mouth, pharynx, lung, liver, or feet and is one of the most important diseases of farm-
raised deer.5,8,11-15 Some of the most frequently isolated pathogens in these cases are F.
necrophorum and F. varium with common co-infection by Arcanobacterium pyogenes.5,8
The infection often spreads to multiple organs resulting in death, as antibiotic therapy is often not successful.11
F. necrophorum possesses several virulence factors, however, leukotoxin is
considered to be the major virulence factor involved in animal infections.3,4 The leukotoxin operon consists of three genes, lktB, lktA, and lktC corresponding respectively to an outer membrane protein with putative transporter function, the leukotoxin protein, and a protein of unknown function.3,4,16 The lkt operon promoter region is distinct for
each of the two subspecies of F. necrophorum and allows for differentiation of the two
subspecies.17,18 The virulence factors of F. varium have not been well characterized. 64
The objective of the current study was to phenotypically and genotypically
characterize clinical isolates of Fusobacterium spp. from deer and to evaluate their association with respiratory infections.
4.3 Materials and Methods
4.3.1 Collection and identification of isolates
A total of 28 clinical isolates of Fusobacterium spp. were used in this study. The isolates included two (7.1%) F. n. subsp. necrophorum, five (17.9%) F. n. subsp.
funduliforme, and 21 (75.0%) F. varium. Additionally, four ATCC reference strains were
included in the study (Table 1). All 28 clinical strains were isolated at necropsy from the
upper or lower respiratory tract of cervids including white-tailed deer (23/28, 82.1%), elk
(3/28, 10.7%), and reindeer (2/28, 7.1%) at the Animal Diagnostic Laboratory at The
Pennsylvania State University between October 2006 and October 2008. Gross lesions, histologic lesions, and complete bacteriological culture results were retrieved for each clinical case. The cervids originated from 18 independent farms across the state of
Pennsylvania.
Tissue samples were cultured on pre-reduced Brucella agar plates supplemented with 5% sheep blood, hemin, and vitamin K (Remel, Lenexa, KS, USA), incubated at
37˚C in an anaerobic chamber (Shel Lab, Cornelius, OR, USA). Suspect colonies of anaerobic, catalase-negative, gram negative rods were subcultured onto pre-reduced
Brucella agar plates, pre-reduced blood agar plates and pre-reduced, anaerobically- sterilized, brain-heart infusion (PRAS-BHI) broth. Aerobic tolerance was determined by culturing isolates on blood agar plates (Remel) at 37˚C incubated aerobically. Three-way 65
antibiotic disk diffusion susceptibility test and nitrate test were done on Brucella agar plates with vancomycin, kanamycin, colistin, and nitrate disks (Anaerobe Systems,
Morgan Hill, CA, USA). Nitrate negative isolates showing resistance to vancomycin and susceptibilities to kanamycin and colistin were considered to be Fusobacterium sp.19 The indole test was performed by placing one drop of indole reagent (p- dimethylaminocinnamaldehyde, Anaerobe Systems) onto one of the antibiotic disks. A pre-reduced McClung-Toabe egg yolk agar plate (Remel) was inoculated with the isolate and incubated anaerobically at 37˚C for 48-72 h to determine lipase activity. Bile tolerance was determined by plating colonies onto pre-reduced Bacteroides fragilis isolation (BBE) agar plates (Remel) containing 20% bile and incubating anaerobically at
37˚C for 48-72 h. Overnight cultures in PRAS-BHI broth were observed for growth sedimentation. All isolates were analyzed with Rapid ANAII (Remel) and RapID 32A
(Biomerieux, Marcy l’Etoile, France) kits according to the manufacturers’ instructions.
4.3.2 DNA extraction and 16S rDNA sequencing
Chromosomal DNA was extracted from all 28 Fusobacterium isolates. Briefly, cultures were grown anaerobically on pre-reduced blood agar plates at 37˚C for 48-72 h.
The DNA was isolated using the DNeasy Blood and Tissue Kit (Qiagen, Hilden,
Germany) according to the manufacturer’s instructions. The extracts were stored at -
20°C. Universal primers (p515FPL – fwd GCG GAT CCT CTA GAC TGC AGT GCC
AGC AGC CGC GGT AA and p13B – rev CGG GAT CCC AGG CCC GGG AAC GTA
TTC AC) were used to amplify a region of the 16S rRNA gene.20 Amplified products
were electrophoresed in 1.5% agarose gels and purified using QIAquick Gel Extraction 66
Kit (Qiagen). The purified PCR product was submitted to a commercial laboratory
(Davis Sequencing, Davis, CA, USA) for sequence analysis; all isolates were sequenced at least two times. Consensus sequences of >99% agreement were calculated using
Vector NTI Advance 11.0 Align X (Invitrogen, Carlsbad, CA, USA). Sequence alignment and phylogenetic analysis were conducted using Mega version 4 by Clustal W and the neighbor-joining method.21
4.3.3 Antimicrobial susceptibility determination
The broth micro-dilution method of antimicrobial susceptibility was performed on all 28 clinical strains and the four ATCC reference strains of Fusobacterium spp. by the
Sensititre System (Trek Diagnostic Systems, Cleveland, OH, USA) according to the manufacturer’s instructions, which conforms to the CLSI recommendations.22 Colonies were selected from 48 h cultures on blood agar plates, inoculated into Sensititre Mueller-
Hinton broth tubes (Trek), and transferred into Sensititre supplemented Brucella broth tubes (Trek) adjusted to 1 × 106 CFU/ml. Aliqouts (100 µl) were transferred to wells of a
96-well BOPO6F antimicrobial susceptibility plate (Trek). Plates were covered by perforated adhesive seals (Trek), incubated anaerobically at 37°C for 48 h, and the minimal concentration of antimicrobial agents that inhibited bacterial growth was determined by visual examination. Breakpoints and interpretations were determined by
Trek Diagnostic Systems software. When no breakpoint was provided, interpretations and breakpoints followed CLSI guidelines (danofloxacin and tulathromycin) or product technical insert (tylosin).23
67
4.3.4 Preparation of culture supernatants
Fusobacterium strains were grown in PRAS-BHI at 37°C to an absorbance of
0.7-0.8 at 600 nm, placed on ice, and pelleted by centrifugation at 10,000 × g for 10 min
at 4°C. Supernatants were filter sterilized with 0.22 µm membrane filters (GE Osmonics,
Minnetonka, MN, USA) and stored at -80°C.
4.3.5 Preparation of white blood cells
Bovine peripheral polymorphonuclear leukocytes (PMNs) were prepared as
previously described.24 Briefly, blood was collected from healthy cattle by venipuncture
into heparinized tubes (BD, Franklin Lakes, NJ, USA). Blood was transported on ice and
centrifuged at 1,500 × g for 10 min at 4°C. The plasma, buffy coat, and top 1/3 of RBC
layer were discarded. Remaining erythrocytes were lysed with RBC lysis buffer
(BioLegend, San Diego, CA, USA) at room temperature for 10-15 min. Purified
leukocytes were washed twice and resuspended in 0.01 M PBS. Viable cell
concentration was determined with a hemacytometer by the trypan blue dye exclusion
method.
4.3.6 Cell viability assay
Leukotoxicity was determined by incubation of bovine PMNs with sterile culture
supernatant and propidium iodide (PI) as previously described.2 Approximately 2 × 106
PMNs were placed into sterile 5 ml polystyrene culture tubes (VWR, West Chester, PA,
USA) and incubated with an equal volume of filter sterilized culture supernatant. Cells were incubated for 45 min at 37°C and 5% CO2, washed twice, and resuspended in PBS. 68
Five µl of 1 mg/ml PI was added to each tube and incubated for 5-10 min in the dark.
Samples were analyzed on a Cytomics FC500 flow cytometer (Beckman Coulter,
Fullerton, CA, USA). The proportion of cells stained with PI was determined by the detection of red fluorescence of cells in the PMN gate.
4.3.7 Lkt promoter and hemagglutinin gene PCR
Chromosomal DNA was extracted from all isolates as before. Primers specific
for the promoter region upstream of lktB gene (lktp and fund) and the hemagglutinin- related protein gene (haem), previously identified in F. necrophorum were used.18,25 The lkt promoter sequence differs between the two F. necrophorum subspecies, so two different primer sets were used. The lktp primer pair detects the lkt promoter in both subspecies, whereas the fund primer pair detects the lkt promoter of only the funduliforme subspecies. Primers were as follows: lktp forward 5’-TCT CCC GGG CTC GAG GAA
ATC TTT AAA GCA C-3’, lktp reverse 5’-TCT CCC GGG CAT AAT TTC TCC CAA
TTT TAT T-3’, fund forward 5’-CTC AAT TTT TGT TGG AAG CGA G-3’, fund reverse 5’-CAT TAT CAA AAT AAC ATA TTT CTC AC-3’, haem forward 5’-CAT
TGG GTT GGA TAA CGA CTC CTA C-3’, and haem reverse 5’-CAA TTC TTT GTC
TAA GAT GGA AGC GG-3’. Amplification was performed in an iCycler thermal cycler
(Bio-Rad Laboratories, Hercules, CA, USA) using a modification of the conditions used by Zhang et al. (2006): initial denaturation at 94°C for 3 min, followed by 35 cycles of denaturation at 94°C for 1 min, annealing at 45°C for 30 sec, and 62.4°C (lktp) or 55.1°C
(haem) or 50.8°C (fund) for 30 sec, extension at 72°C for 1 min, and the final extension at 72°C for 4 min. The amplified products were electrophoresed in 1.5% agarose gel 69
containing ethidium bromide. Products to be sequenced were purified using QIAquick
Gel Extraction Kit and submitted with the respective forward primer to a commercial laboratory (Davis Sequencing, Davis, CA, USA) for sequence analysis.
4.3.8 Statistical analysis
Flow cytometry data were evaluated by one-way ANOVA with comparisons for each pair using Student’s t test. In order to evaluate the association between lesions observed and Fusobacterium type isolated, a t test was used to detect differences in single proportions and a χ2 test or Fisher’s exact test was used to determine dependence for
bacterial type against lesion type and co-pathogen type. A Cochran-Mantel-Haenszel test
of association was applied to each contingency table in order to detect associations
between Fusobacterium type and lesion type while stratified by co-pathogen type. Odds
ratios were calculated for the association between respiratory pathogen and the presence
of pneumonia. Standard statistical software (JMP, Cary, NC, USA) was used. For all
analyses, a value of P < 0.05 was considered significant.
4.4 Results
4.4.1 Phenotypic characterization of isolates
Colony morphology varied among Fusobacterium types. At 48 – 72 h on
blood agar, F. n. subsp. necrophorum formed colonies that were circular with erose
edges, umbonate, approximately 3 – 5 mm in diameter, and grey, while F. n. subsp.
funduliforme formed colonies that were circular with entire edges, convex, approximately
1 mm in diameter, and yellow-grey. F. varium formed colonies that were circular with 70
entire to erose margins, low convex, 1 – 2 mm in diameter, and translucent with grey centers. Colonies of F. nucleatum were circular with entire edges, raised, 1 – 2 mm in diameter, and dull grey.
Cellular morphology was also distinct for each Fusobacterium type. While all types evaluated were gram negative bacilli, their length and tendency to form filaments or to aggregate differed. Bacterial length was approximately 3 – 4 µm for F. varium, 2 – 4
µm for F. n. subsp. necrophorum, 1 – 2.5 µm for F. n. subsp. funduliforme, and 3 – 7 µm with tapered ends for F. nucleatum. F. n. subsp. necrophorum was pleomorphic and formed many long filaments while F. n. subsp. funduliforme formed many shorter filaments and tended to aggregate heavily forming dense clusters. F. varium and F. nucleatum tended to form short filaments only infrequently. Growth and biochemical characteristics differed across fusobacterial types as summarized in Table 2 and
Appendix B.
Bacterial identification with Rapid ANAII or RapID 32A alone was not reliable for species-level identification of many Fusobacterium strains evaluated (Table 2).
Although F. n. subsp. necrophorum was consistently correctly identified at the species level with both test kits, neither kit was able to distinguish between F. n. subsp. funduliforme and F. nucleatum. Additionally, F. varium was sometimes misidentified as
F. nucleatum with Rapid ANAII.
4.4.2 16S rDNA sequencing
The 16S rDNA sequences of the nine F. necrophorum isolates clustered together, however, the two subspecies were not distinguishable by 16S rDNA sequences alone 71
(Figure 1). All 21 F. varium isolates clustered apart from the F. necrophorum and F. nucleatum groups, and from an F. equinum group included for reference. All previously reported sequences obtained from GenBank clustered with study strains of the same subtype, however some published F. varium sequences did not cluster with the larger F. varium group.
4.4.3 Antimicrobial susceptibility
Results of broth micro-dilution antimicrobial susceptibility test are summarized in
Table 3. While F. n. subsp. necrophorum and F. n. subsp. funduliforme showed similar patterns of susceptibility, the susceptibility of F. varium was markedly different for many antimicrobials. All three fusobacterial types were susceptible to ampicillin, florfenicol, and trimethoprim/sulfamethoxazole. F. varium was much less susceptible than F. necrophorum to clindamycin, tilmicosin, tulathromycin, and tylosin and moderately less susceptible to chlortetracycline, oxytetracycline, penicillin, and tiamulin. Antimicrobials to which isolates demonstrated intermediate or variable susceptibilities included ceftiofur, danofloxacin, spectinomycin, and sulfadimethoxine. All isolates of these three types were resistant to enrofloxacin, gentamycin, and neomycin. The single F. nucleatum isolate was resistant only to gentamycin and neomycin. Antibiograms of antimicrobial resistance profiles are summarized in Table 4.
4.4.4 Leukotoxin activity by flow cytometry
Toxicity of culture supernatants (secreted leukotoxin) to bovine PMNs, as determined by PI staining, was highly dependent upon bacterial type (Appendix B). The 72
uptake of PI by PMNs was highest in cells incubated with supernatant of F. n. subsp.
necrophorum (mean 67.0%, SE 7.3), intermediate with F. n. subsp. funduliforme (mean
27.6%, SE 5.0), and very low for F. varium (mean 2.2%, SE 2.7) and F. nucleatum (mean
2.2%, SE 12.7), which approximated the negative control (mean 1.2%, SE 9.0). Means
for F. n. subsp. necrophorum and F. n. subsp. funduliforme differed significantly from
each other (P < 0.0001) and from the mean of F. varium (P < 0.0001). The means of F. varium, F. nucleatum, and negative control were not significantly different (P > 0.05).
4.4.5 Virulence gene PCR analysis
PCR analysis consistently identified isolates of F. n. subsp. necrophorum and F. n. subsp. funduliforme, and was able to distinguish F. varium strains from F. necrophorum or F. nucleatum (Table 5). Using the lktp primers for the promoter region upstream of lktB, all F. n. subsp. necrophorum strains amplified the expected product of approximately 571 bp while all F. n. subsp. funduliforme strains amplified a slightly smaller product of approximately 449 bp as expected. All F. varium strains and F. nucleatum were negative for the lkt promoter. With the fund primers specific for the lkt promoter region of F. n. subsp. funduliforme, four of six (67%) F. n. subsp. funduliforme strains amplified the expected 337 bp product, while two of six (33%) failed to amplify the product. All F. n. subsp. necrophorum strains, all F. varium strains, and F. nucleatum were negative for the lkt promoter region of F. n. subsp. funduliforme. Using the haem primers for the hemagglutinin-related protein gene, all F. n. subsp. necrophorum strains amplified the expected 311 bp product, while all F. n. subsp. funduliforme, F. varium, and F. nucleatum isolates were negative. Each of the three primer sets resulted in a non- 73
specific product in many of the F. varium isolates. Similarly, the haem primers produced a non-specific product in several F. n. subsp. funduliforme isolates. These products were sequenced and determined to be the result of non-specific amplification of genomic
DNA.
4.4.6 Association of bacterial type with post mortem lesions
Numerous gross and microscopic lesions were detected in cases from which
Fusobacterium was isolated. Among the 26 cases for which pathology data were available, lesions included pneumonia (16, 61.5%), rumenitis (6, 23.0%), oropharyngitis
(5, 19.2%), nephritis (5, 19.2%), hepatitis (4, 15.4%), carditis (3, 11.5%), abomasitis (2,
7.7%), enteritis (2, 7.7%), meningitis/encephalitis (2, 7.7%), peritonitis (1, 3.9%), keratoconjunctivitis (1, 3.9%), myositis (1, 3.9%), splenitis (1, 3.9%), and osteomyelitis
(1, 3.9%). Co-pathogens in 27 cases included E. coli (18, 66.7%), Arcanobacterium pyogenes (9, 33.3%), Bacteroides fragilis (7, 25.9%), Bacteroides spp. (5, 18.5%),
Streptococcus spp. (5, 18.5%), Pasteurella spp. (4, 14.8%), Enterococcus spp. (3,
11.1%), Clostridium perfringens (3, 11.1%), Staphylococcus spp. (2, 7.4%),
Actinobacillus spp. (2, 7.4%), Proteus spp. (2, 7.4%), Peptostreptococcus spp. (1, 3.7%),
Aeromonas spp. (1, 3.7%), Bacillus spp. (1, 3.7%), Pseudomonas spp. (1, 3.7%),
Aspergillus spp. (1, 3.7%), Serratia spp. (1, 3.7%), and Acinetobacter spp. (1, 3.7%).
Although the prevalence of pneumonia was significantly higher (P = 0.001) in
Fusobacterium sp. positive cases (61.5%) than in the general necropsy population of farm-raised white-tailed deer (24.4%) as previously reported by Hattel et al. (2004), no significant association was detected between the occurrence of lesion and Fusobacterium 74
type, between co-pathogen and Fusobacterium type, or between lesion and co-pathogen
(Appendix B). The odds ratio for the association between respiratory Fusobacterium infection and the presence of pneumonia in all deer in which lung was anaerobically
cultured was not remarkable (1.04, P = 1.0), however the odds ratio for Arcanobacterium
pyogenes infection and pneumonia was high (6.3, P = 0.001) also while stratified by co-
infection with Fusobacterium sp. (P = 0.001) (Appendix B).
4.5 Discussion
Respiratory tract infection is one of the primary causes of mortality in farm-raised
white-tailed deer.5,6 The cause of such infections remains undetermined, but one of the
most commonly isolated pathogens is Fusobacterium. It is important to the deer farming
industry and the animal health community to more fully characterize the etiology and
pathogenesis of this disease in order to develop appropriate treatment and prevention
measures. This study shows that F. varium comprises a large proportion of the clinical
isolates from cases of fusobacterial respiratory tract infections in deer. Standard
phenotypic methods were sufficient for identification of the clinical Fusobacterium types
used in this study, however such methods are labor intensive and time consuming and
may be insufficient for speciating or subtyping some Fusobacterium spp. isolates.
Commercial bacterial identification kits such as Rapid ANAII and RapID 32A were
useful as part of a diagnostic scheme, but were inadequate as the sole means of
identification of many Fusobacterium spp. to the species or subspecies level. Genotypic
analysis by PCR or ribosomal gene sequencing may be preferable for diagnosis or
confirmation. 75
PCR amplification of the lkt promoter and the hemagglutinin gene differentiated
F. necrophorum subtypes from other Fusobacterium spp. Variations in bands produced
by F. n. subsp. funduliforme with fund primers may represent polymorphisms in the sequences of this virulence gene. No evidence of leukotoxin production by any strain of
F. varium was detected by virulence gene PCR analysis or by the leukocyte viability
assay. Despite the lack of leukotoxin production by F. varium isolates, no significant difference was detected between F. varium associated lesion scores and F. necrophorum associated lesion scores. It is possible that the pathogenicity of F. varium is associated not with leukotoxin production but with some other virulence factor. A recent study suggests that the presence of the leukotoxin gene is much less prevalent in non-bovine, invasive animal and human strains and, thus, should not be considered the primary virulence factor in all hosts.26 Additionally, butyric acid has been identified as a potential
virulence factor in F. varium based on cytotoxicity to Vero cells and production of
colonic lesions in mice.27
Co-infection with Arcanobacterium pyogenes was associated with an increased
likelihood of pneumonia among cases that were positive for Fusobacterium spp. perhaps
indicating some synergistic effect between Fusobacterium spp. and A. pyogenes. This
association was not observed for any other bacteria in this study. Previous reports have
demonstrated interaction between Fusobacterium and various aerobic and anaerobic
bacteria including A. pyogenes, E. coli, Bacteroides fragilis, and others.28-30 The significance of such co-infections must be further investigated in order to better understand the pathogenesis of the infection and to identify risk factors and effective therapeutic strategies. 76
Few studies on antimicrobial susceptibilities of Fusobacterium spp., particularly
F. varium, have been reported. Results of this study suggest that the susceptibility to
antimicrobials is markedly different among the Fusobacterium species evaluated, and are
consistent with previous studies.8,9,31-33 It has been reported that most Fusobacterium
spp. are sensitive to β-lactams and clindamycin, but that F. varium tends to be more
resistant.9 Our data support the increased resistance of F. varium to clindamycin as well as chlortetracycline, oxytetracycline, tiamulin, tilmicosin, tulathromycin, and tylosin with respect to F. necrophorum. This finding is of clinical relevance, because all treatment and prevention efforts have been directed toward F. necrophorum infections, selecting antibiotics and formulating vaccines accordingly, as most fusobacterial infections in animals have been attributed to F. necrophorum. However, in the dataset used in this study, the majority (75.0%) of clinical isolates were F. varium, which suggests that most cases of necrobacillosis may be caused by a different species of Fusobacterium with a different antibiotic resistance profile and, potentially, a different antigenic profile.
Therefore, in order to properly diagnose and provide critical information for treatment and preventive measures, it is essential that clinical Fusobacterium sp. isolates are identified to the species or subspecies level. Additional studies are required to determine virulence factors of F. varium and the pathogenicity and host immune response to F. varium infection in order to develop effective treatment and prevention strategies.
4.6 Acknowledgements
The authors thank the Missouri Whitetail Breeders and Hunting Ranch
Association; Dr. Vivek Kapur, Dr. Arthur Hattel, Dr. Jenny Fisher, Dr. Subhashinie 77
Kariyawasam, Valerie Linter, and Rhiannon Schneider of The Pennsylvania State
University Department of Veterinary and Biomedical Sciences; and Greg Peterson and
Elena Gart of Kansas State University College of Veterinary Medicine for their
assistance.
4.7 References
1. Citron DM. Update on the taxonomy and clinical aspects of the genus
Fusobacterium. Clin Infect Dis 2002;35:S22-27.
2. Narayanan S, Stewart GC, Chengappa MM, et al. Fusobacterium necrophorum
leukotoxin induces activation and apoptosis of bovine leukocytes. Infect Immun
2002;70:4609-4620.
3. Nagaraja TG, Narayanan SK, Stewart GC, et al. Fusobacterium necrophorum
infections in animals: pathogenesis and pathogenic mechanisms. Anaerobe
2005;11:239-246.
4. Tadepalli S, Narayanan SK, Stewart GC, et al. Fusobacterium necrophorum: a
ruminal bacterium that invades liver to cause abscesses in cattle. Anaerobe
2009;15:36-43.
5. Hattel AL, Shaw DP, Love BC, et al. A retrospective study of mortality in
Pennsylvania captive white-tailed deer (Odocoileus virginianus): 20000--2003. J
Vet Diagn Invest 2004;16:515-521.
6. Brooks JW, Jayarao BM. Management practices used by white-tailed deer farms
in Pennsylvania and herd health problems. J Am Vet Med Assoc 2008;232:98-104. 78
7. Jang SS, Hirsh DC. Characterization, distribution, and microbiological
associations of Fusobacterium spp. in clinical specimens of animal origin. J Clin
Microbiol 1994;32:384-387.
8. Chirino-Trejo M, Woodbury MR, Huang F. Antibiotic sensitivity and biochemical
characterization of Fusobacterium spp. and Arcanobacterium pyogenes isolated
from farmed white-tailed deer (Odocoileus virginianus) with necrobacillosis. J
Zoo Wildl Med 2003;34:262-268.
9. Legaria MC, Lumelsky G, Rodriguez V, et al. Clindamycin-resistant
Fusobacterium varium bacteremia and decubitus ulcer infection. J Clin Microbiol
2005;43:4293-4295.
10. Minami M, Ando T, Okamoto A, et al. Seroprevalence of Fusobacterium varium
in ulcerative colitis patients in Japan. FEMS Immunol Med Microbiol 2009;56:67-
72.
11. Leighton FA. Fusobacterium necrophorum infection In: Williams ES,Barker IK,
eds. Infectious Diseases of Wild Mammals. Ames, Iowa: Iowa State Press,
2001;493-496.
12. Edwards JF, Davis DS, Roffe TJ, et al. Fusobacteriosis in captive wild-caught
pronghorns (Antilocapra americana). Vet Pathol 2001;38:549-552.
13. Ramos-Vara JA, Ganjam I, Fales W. Pulmonary necrobacillosis in a white-tailed
deer. Vet Rec 2003;152:401-403.
14. Roeder BL, Chengappa MM, Lechtenberg KF, et al. Fusobacterium necrophorum
and Actinomyces pyogenes associated facial and mandibular abscesses in blue
duiker. J Wildl Dis 1989;25:370-377. 79
15. Wobeser G, Runge W, Noble D. Necrobacillosis in deer and pronghorn antelope
in Saskatchewan. Can Vet J 1975;16:3-9.
16. Narayanan SK, Nagaraja TG, Chengappa MM, et al. Cloning, sequencing, and
expression of the leukotoxin gene from Fusobacterium necrophorum. Infect
Immun 2001;69:5447-5455.
17. Tadepalli S, Stewart GC, Nagaraja TG, et al. Human Fusobacterium
necrophorum strains have a leukotoxin gene and exhibit leukotoxic activity. J
Med Microbiol 2008;57:225-231.
18. Zhang F, Nagaraja TG, George D, et al. The two major subspecies of
Fusobacterium necrophorum have distinct leukotoxin operon promoter regions.
Vet Microbiol 2006;112:73-78.
19. Jousimies-Somer H, Summanen P, Citron DM, et al. Wadsworth-KTL Anaerobic
Bacteriology Manual. 6th ed. Belmont, CA, U.S.A.: Star Pub. Co., 2002.
20. Relman DA, Schmidt TM, MacDermott RP, et al. Identification of the uncultured
bacillus of Whipple's disease. N Engl J Med 1992;327:293-301.
21. Tamura K, Dudley J, Nei M, et al. MEGA4: Molecular Evolutionary Genetics
Analysis (MEGA) software version 4.0. Molecular Biology and Evolution
2007;24:1596-1599.
22. CLSI. Methods for Antimicrobial Susceptibility Testing of Anaerobic Bacteria;
Approved Standard - Seventh Edition. CLSI document M11-A7. Wayne, PA:
Clinical and Laboratory Standards Institute, 2007.
23. CLSI. Performance Standards for Antimicrobial Disk and Dilution Susceptibility
Tests for Bacteria Isolated From Animals; Approved Standard - Third Edition. 80
CLSI document M31-A3. Wayne, PA: Clinical and Laboratory Standards Institute,
2008.
24. Tan ZL, Nagaraja TG, Chengappa MM. Factors affecting the leukotoxin activity
of Fusobacterium necrophorum. Vet Microbiol 1992;32:15-28.
25. Aliyu SH, Marriott RK, Curran MD, et al. Real-time PCR investigation into the
importance of Fusobacterium necrophorum as a cause of acute pharyngitis in
general practice. J Med Microbiol 2004;53:1029-1035.
26. Ludlam HA, Milner NJ, Brazier JS, et al. lktA-encoded leukotoxin is not a
universal virulence factor in invasive Fusobacterium necrophorum infections in
animals and man. J Med Microbiol 2009;58:529-530.
27. Ohkusa T, Okayasu I, Ogihara T, et al. Induction of experimental ulcerative
colitis by Fusobacterium varium isolated from colonic mucosa of patients with
ulcerative colitis. Gut 2003;52:79-83.
28. Brook I, Walker RI. The relationship between Fusobacterium species and other
flora in mixed infection. J Med Microbiol 1986;21:93-100.
29. Kannangara DW, Thadepalli H, Bach VT, et al. Animal model for anaerobic lung
abscess. Infect Immun 1981;31:592-597.
30. Smith GR, Till D, Wallace LM, et al. Enhancement of the infectivity of
Fusobacterium necrophorum by other bacteria. Epidemiol Infect 1989;102:447-
458.
31. Courcol RJ, Lee KW, Downes J, et al. In-vitro susceptibilities of Bacteroides
gracilis, Fusobacterium mortiferum and F. varium to 17 antimicrobial agents. J
Antimicrob Chemother 1990;26:157-158. 81
32. Piriz Duran S, Valle Manzano J, Cuenca Valera R, et al. Susceptibilities of
Bacteroides and Fusobacterium spp. from foot rot in goats to 10 beta-lactam
antibiotics. Antimicrob Agents Chemother 1990;34:657-659.
33. Simon PC. Susceptibility of Fusobacterium necrophorum to antimicrobials. Part
I: as determined by the disc method. Can J Comp Med 1977;41:166-168.
82
F.necrophorum funduliforme (EF447425) F.necrophorum funduliforme (AM905356) F.necrophorum funduliforme ATCC 51357 F.necrophorum funduliforme P18B F.necrophorum necrophorum P38A
79 F.necrophorum funduliforme P21V F.necrophorum necrophorum ATCC 25286 F.necrophorum funduliforme P30B F.necrophorum funduliforme P20B 51 F.necrophorum funduliforme P27B F.necrophorum necrophorum P26A 75 F.necrophorum necrophorum (AJ867039) F.necrophorum necrophorum (X55411) F.equinum (AJ295750)
76 F.equinum (EF447429) 55 F.equinum (EF447428)
95 F.nucleatum nucleatum ATCC 25586 96 F.nucleatum nucleatum (AJ133496) F.nucleatum polymorphum (AF287812) F.varium (X55413) F.varium P39V F.varium P14V F.varium P3V F.varium P7V F.varium P24V F.varium P12V 98 F.varium P41V F.varium P17V F.varium P34V F.varium P22V F.varium (M58686) F.varium P11V 64 F.varium P35V F.varium P36V F.varium P8V F.varium P9V F.varium ATCC 8501 F.varium P29V F.varium P1V F.varium P23V F.varium P16V F.varium P37V F.varium P10V F.varium (AJ867035) 100 F.varium (AJ867036)
Figure 4.1. Phylogenetic analysis of the 16S rRNA gene sequence of Fusobacterium spp. 83
Table 4.1 Fusobacterium strains characterized in this study Species Strain Host species Source F. n. necrophorum P26A White‐tailed deer Lung P38A White‐tailed deer Oral abscess ATCC 25286 Bovine Liver abscess F. n. funduliforme P18B White‐tailed deer Lung P20B White‐tailed deer Lung P21B Elk Lung P27B Elk Lung P30B White‐tailed deer Oral abscess ATCC 51357 Bovine Liver abscess F. varium P1V Elk (wild) Pharynx P3V White‐tailed deer Lung P7V White‐tailed deer Pharynx P8V Reindeer Lung P9V White‐tailed deer Pharynx P10V White‐tailed deer Pharynx P11V White‐tailed deer Lung P12V White‐tailed deer Pharynx P14V White‐tailed deer (wild) Lung P16V White‐tailed deer Lung P17V Reindeer Lung P22V White‐tailed deer Lung P23V White‐tailed deer Lung P24V White‐tailed deer Lung P29V White‐tailed deer Lung P34V White‐tailed deer Lung P35V White‐tailed deer Lung P36V White‐tailed deer Lung P37V White‐tailed deer Oral abscess P39V White‐tailed deer Lung P41V White‐tailed deer Lung ATCC 8501 NA NA F. nucleatum nucleatum ATCC 25586 NA Cervico‐facial lesion
Table 4.2 Phenotypic and Genotypic Identification of Fusobacterium spp. isolates Rapid ANA II Result (quality of ID %) RapID 32A Result (quality of ID %) 16S rRNA Result Supplemental Analysis Final ID # (%) Sediment Lipase Bile Indole Phosphatase of isolates F. necrophorum (99.8) F. necrophorum (70.7)/C. tetani (29) F. necrophorum ‐ + ‐ + + F.n. necrophorum 3 (9.4) F. necrophorum (60)/F. nucleatum (40) F. nucleatum (62)/ C. tetani (20.7) F. necrophorum + + ‐ + ‐ F.n. funduliforme 6 (18.8) F. varium (>99.9) C. histolyticum (72)/F. varium (11.3) F. varium ‐ ‐ + ‐ ‐ F. varium 9 (28.1) F. nucleatum (>99.9) F. varium (92.5) F. varium ‐ ‐ + + ‐ F. varium 6 (18.8) F. varium (98.9) F. varium (99.9) F. varium ‐ ‐ + + ‐ F. varium 3 (9.4) F. varium (>99.9) F. varium (99.9) F. varium ‐ ‐ + + ‐ F. varium 3 (9.4) F. nucleatum (99.4) F. varium (92.5) F. varium ‐ ‐ + + ‐ F. varium 1 (3.1) F. necrophorum (60)/F. nucleatum (40) F. nucleatum (62)/ C. tetani (20.7) F. nucleatum ‐ ‐ ‐ + ‐ F. nucleatum 1 (3.1)
84 Table 4.3
Antimicrobial Susceptibility of Fusobacterium spp. F.n. necrophorum F.n. funduliforme F. varium F. nucleatum MIC Range # (%) MIC Range # (%) MIC Range # (%) MIC Range # (%) Antimicrobial (µg/ml) Resistant (µg/ml) Resistant (µg/ml) Resistant (µg/ml) Resistant Ampicillin ≤0.25 0 (0) ≤0.25 0 (0) 1 – 2 0 (0) ≤0.25 0 (0) Ceftiofur ≤0.25 – 8 1 (33.3) ≤0.25 0 (0) 0.5 – 8 9 (40.9) ≤0.25 0 (0) Chlortetracycline ≤0.5 0 (0) ≤0.5 – 4 0 (0) ≤0.5 – 8 0 (0) ≤0.5 0 (0) Clindamycin ≤0.25 0 (0) ≤0.25 0 (0) 1 – 16 17 (77.3) ≤0.25 0 (0) Danofloxacin 0.5 – 1 0 (0) 0.25 – 1 0 (0) 1 0 (0) 0.5 0 (0) Enrofloxacin 1 – 2 2 (66.7) 2 6 (100) 2 22 (100) 0.5 0 (0) Florfenicol ≤0.25 0 (0) ≤0.25 0 (0) ≤0.25 0 (0) ≤0.25 0 (0) Gentamicin 4 – 16 2 (66.7) 8 – 16 5 (83.3) 16 22 (100) 16 1 (100) Neomycin 16 – 32 3 (100) 32 6 (100) 32 22 (100) 32 1 (100) Oxytetracycline ≤0.5 – 2 0 (0) ≤0.5 – 8 0 (0) ≤0.5 – 8 0 (0) ≤0.25 0 (0) Penicillin ≤0.12 0 (0) ≤0.12 0 (0) 0.25 – 1 0 (0) ≤0.12 0 (0) Spectinomycin ≤8 – 32 0 (0) 32 0 (0) 32 – 64 20 (90.9) ≤0.8 0 (0) Sulfadimethoxine ≤256 – >256 1 (33.3) ≤256 – >256 1 (16.7) ≤256 – >256 2 (9.1) ≤256 0 (0) Tiamulin ≤0.5 0 (0) ≤0.5 0 (0) 32 0 (0) ≤0.5 0 (0) Tilmicosin ≤4 0 (0) ≤4 0 (0) 16 – 64 17 (77.3) ≤4 0 (0) Trimethoprim/Sulfa ≤2/38 0 (0) ≤2/38 0 (0) ≤2/38 0 (0) ≤2/38 0 (0) Tulathromycin ≤1 0 (0) ≤1 0 (0) 16 – 64 17 (77.3) ≤1 0 (0) Tylosin ≤0.5 – 1 0 (0) ≤0.5 – 2 0 (0) 32 22 (100) ≤0.5 0 (0)
85 86
Table 4.4 Antibiogram of Fusobacterium spp. Organism Antibiograma # (%) of isolates F. n. necrophorum CeEGN 1 (3.1) GNSu 1 (3.1) EN 1 (3.1) F. n. funduliforme EGNSu 1 (3.1) EGN 4 (12.5) EN 1 (3.1) F. varium CeClEGNSpTiTuTy 3 (9.4) CeClEGNSpTiTy 3 (9.4) CeClEGNSpTuTy 1 (3.1) CeEGNSpTuTy 1 (3.1) CeEGNTiTuTy 1 (3.1) ClEGNSpSuTiTuTy 1 (3.1) ClEGNSpTiTuTy 6 (18.8) ClEGNSpTiTy 1 (3.1) ClEGNSpTuTy 2 (6.3) EGNSpSuTiTuTy 1 (3.1) EGNSpTy 1 (3.1) EGNTiTuTy 1 (3.1) F. nucleatum GN 1 (3.1) a Indicates resistance to Ce (ceftiofur), Cl (clindamycin), E (enrofloxacin), G (gentamicin), N (neomycin), Sp (spectinomycin), Su (sulfadimethoxine), Ti (tilmicosin), Tu (tulathromycin), Ty (tylosin) 87 Table 4.5 Profiles of virulence gene PCR products (bp) Organism Lktpa Fundb Haemc Number of isolates (%) F. n. necrophorum 571 ‐ 311 3/3 (100.0) F. n. funduliforme 449 337 ‐ 4/6 (66.7) 449 ‐ ‐ 2/6 (33.3) F. varium ‐ ‐ ‐ 22/22 (100.0)
F. nucleatum ‐ ‐ ‐ 1/1 (100.0) a F. necrophorum leukotoxin promoter region (5’lktpXmXh‐3’lktpXm) b Leukotoxin promoter region of F.n. funduliforme (Fund5p‐Fund3p) c Hemagglutinin‐related protein gene (Haem forward‐Haem reverse)
88
Chapter 5
EFFECTS OF FUSOBACTERIUM VARIUM ON THE RESPIRATORY TRACT
OF MICE FOLLOWING INTRANASAL INOCULATION
89
5.1 Abstract
It has been reported that Fusobacterium varium is the predominant fusobacterial species isolated from respiratory tract lesions of white-tailed deer. The objective of this study was to evaluate the pathogenicity of F. varium to the respiratory tract of mice and to determine the utility of this system as model of fusobacterial pneumonia. C57BL/6 mice and BALB/c mice were inoculated intranasally with various strains and varying dosages of F. varium. Prior to inoculation, mice were pre-treated with either 0.2 mg dexamethasone intraperitoneally once daily for four days, 10 µg of lipopolysaccharide
(LPS) of E. coli O55:B5 intranasally one time, or no pre-treatment. Following inoculation mice were observed for morbidity and mortality for fourteen days. No mice infected with F. varium showed clinical illness or died. No mice infected with F. varium developed gross or microscopic lesions. The bacterium was recovered from the blood of one C57BL/6 mouse, but was not recovered from blood or lung of any other mice. Anti- fusobacterial IgM or IgG were not produced in serum by 14 days in response to infection.
Pre-existing antibodies detected in pooled serum bound with proteins of two F. necrophorum subspecies and three F. varium strains, but did not bind to proteins of similar size from other common bacterial pathogens including Escherichia coli,
Pasteurella pneumotropica, Arcanobacterium pyogenes, and Clostridium perfringens.
No serum antibodies were detected in any BALB/c mice at any time. These results suggest that F. varium is not highly pathogenic to the respiratory tract of mice, and does not result in a humoral immune response following intranasal inoculation.
90
5.2 Introduction
Fusobacterium is a genus of anaerobic, gram negative, rod-shaped bacteria that
have been associated with many infectious disease processes in humans and animals.
Although there are approximately 13 species within the genus, Fusobacterium necrophorum has classically received the most attention as a serious pathogen as a result of the severe necrotizing infections it causes in humans and ruminants.1 F. necrophorum
has recently been subdivided into two subspecies, F. necrophorum subsp. necrophorum
and F. necrophorum subsp. funduliforme, the former of which has been demonstrated in
animal infections to be more highly virulent due to increased leukotoxin production.2,3
Diseases attributed to F. necrophorum infection include liver abscesses and foot rot in cattle and sheep, stomatitis and laryngitis in calves, pigs, and marsupials, dermatitis in horses, and necrobacillosis of wild ruminants.4-6 The latter condition is characterized by
multifocal necrotizing lesions and abscesses in multiple tissues of the body. Specifically,
in white-tailed deer (Odocoileus virginianus) that are examined through the necropsy
service at the Animal Diagnostic Laboratory at The Pennsylvania State University, the
disease tends to most severely affect the lungs and pharynx of adult deer or, somewhat
less frequently, the head and neck of juvenile deer. In contrast, infection with F. varium
is reported rarely, and is most often associated with ulcerative colitis or decubitus ulcers
in humans and with lesions of the gastrointestinal tract or oral cavity in animals.7-10
In a recent study by the author, a set of fusobacterial isolates from deer were characterized, and F. varium was found to be the predominant species.11 Although both
F. n. subsp. necrophorum and F. n. subsp. funduliforme were also identified in several 91
cases, no significant differences in lesion severity were detected between any
fusobacterial species. Despite the similarities in clinical disease in all of the cases,
evidence of leukotoxin production was not identified in any F. varium isolates, leading the author to speculate that an unidentified virulence factor may be involved in the pathogenesis of F. varium infection. This hypothesis is supported by the findings of
Ludlam et al., (2009) which suggest that leukotoxin should not be considered the major virulence factor in non-bovine fusobacterial infections.12
The objective of this study was to evaluate the pathogenicity of F. varium in mice
following intranasal inoculation and to develop a murine model of respiratory tract
infection with F. varium that mimics the natural presumed route of infection and pathology of deer. Results of this study were expected to direct future studies in which the investigators would prepare various types of vaccines and challenge vaccinated mice with various species of Fusobacterium to determine the efficacy of such vaccines in preventing lesions caused by Fusobacterium infection and to determine if any cross protection exists. Furthermore, these results would expedite and target specific future studies on captive cervids and domestic ruminants and allow the investigators to better advise producers regarding the use of fusobacterial vaccines, specifically with regard to respiratory tract infection.
5.3 Materials and Methods
5.3.1 Bacterial strains
All bacterial strains used in these experiments were obtained from the author’s collection of Fusobacterium spp. obtained through the necropsy service at the Animal 92
Diagnostic Laboratory at The Pennsylvania State University. All isolates were recovered
from the lungs of white-tailed deer between October 2006 and October 2008. Three F.
varium strains (P11V, P16V, P23V) and one strain of F. n. subsp. necrophorum (P26A)
were selected based upon lesions in the respective source clinical cases and leukotoxin
production as determined by propidium iodide (PI) staining of polymorphonuclear cells
(PMNs) by flow cytometry.11 These strains were grown overnight at 37°C in Hungate
tubes (Bellco Glass, Vineland, NJ, USA) containing pre-reduced anaerobically sterilized
brain-heart infusion (PRAS-BHI) broth (BD, Franklin Lakes, NJ, USA) to an absorbance
of 0.8 at 600 nm and serially diluted in PRAS-BHI under anaerobic conditions to produce
inoculation mixtures containing between 103 and 109 colony forming units (CFU) as
determined by spread plating. Inoculation mixtures were placed in anaerobic 10 ml glass serum vials (Wheaton, Millville, NJ, USA) capped with butyl stoppers, crimped with aluminum seals, and transported on ice.
5.3.2 Mice
Eight week old female C57BL/6 mice (19 to 20 g) and female BALB/c mice (19 to 21 g) were purchased from a commercial vendor (Jackson Laboratory, Bar Harbor,
ME, USA). Mice were housed in microisolator cages containing two mice per cage and
provided food and water ad libitum. The animal room temperature was 21.1°C (70°F) and the relative humidity was 50% with a 12:12 hour light:dark cycle. All animals were housed for an acclimation period of 1 to 2 weeks prior to experimental use. All procedures were conducted in compliance with IACUC protocols.
5.3.3 Animal experiments
93
Trial 1
Nine groups of four C57BL/6 mice were inoculated intranasally with either 25 μl
or 50 μl at the tip of the nares (half in each nostril) of F. varium at a dose of 103, 105, or
107 CFU of one of three strains of F. varium following light anesthesia with isoflurane
(Table 5.1). Mice were held upright for approximately 30 seconds to allow for inhalation
of the inoculum.
Trial 2
Four groups of four C57BL/6 mice were inoculated intranasally with either 25 μl
or 50 μl at the tip of the nares (half in each nostril) of inoculum at a dose of 107 or 109
CFU of one strain of F. varium (P11V) or F. n. subsp. necrophorum (P26A) as previously described (Table 5.2).
Trial 3
Twelve groups of two C57BL/6 mice and eight groups of two BALB/c mice were inoculated intranasally with 50 μl at the tip of the nares (half in each nostril) of F. varium at a dose of 109 CFU of one of three strains of F. varium as previously described (Table
5.3). Prior to inoculation, mice were placed into one of three pre-treatment groups. To
simulate exposure to elevated serum cortisol associated with physiologic stress, mice in
the dexamethasone group were administered 0.1 ml of 2 mg/ml dexamethasone (0.2 mg)
(Vedco, Saint Joseph, MO, USA) intraperitoneally once daily for four days beginning on
day -3 and ending on day 0 immediately prior to infection. To simulate pulmonary injury
for a primary pathogen, mice in the LPS group were given 60 µl of 167 µg/ml (10 µg) of
E. coli O55:B5 lipopolysaccharide (LPS) (Sigma-Aldrich, Saint Louis, MO, USA) in
phosphate buffered saline (PBS) intranasally following light sedation by isoflurane one 94
time 24 hours prior to infection. Following intranasal administration of LPS, mice were
held upright for approx 30 seconds. Mice in the no pre-treatment group were not pre-
treated in any way prior to inoculation. Additionally, four mice that received no pre-
treatment, two BALB/c and two C57BL/6, were inoculated with F. varium and
euthanized approximately one hour post infection to assess the delivery of the inoculum
to the lungs.
5.3.4 Clinical signs
Following inoculation, mice were observed at least once daily for morbidity and
mortality for a fourteen day period. Mice that became clinically ill and reached surrogate
endpoints, including labored respiration, loss of ability to ambulate, or dehydration, were euthanized by CO2 inhalation. All surviving mice were euthanized by CO2 inhalation at
14 days post infection.
5.3.5 Gross necropsy and histopathology
Following death or euthanasia complete necropsies were performed on all mice
with most emphasis placed on respiratory tract lesions. Representative sections of lung,
heart, liver, kidney, and spleen from all mice were placed in 10% neutral buffered
formalin.
5.3.6 Bacterial quantitation
All remaining lung tissue from each mouse that was not used for histopathology
was weighed and placed in 1 ml PRAS-BHI in sterile stomacher bags (Seward, Worthing,
West Sussex, UK). Samples were homogenized manually with a syringe barrel for 30
seconds under anaerobic conditions. Serial dilutions of lung elution were made using
PRAS-BHI broth, and 100 µl of each dilution was plated onto blood agar (Remel, 95
Lenexa, KS, USA) and Fusobacterium selective agar (FSA) (Anaerobe Systems, Morgan
Hill, CA, USA) plates. Heart blood was collected in a heparinized syringe created by drawing approx 0.3 ml sodium heparin (APP Pharmaceuticals, Schaumburg, IL, USA) into a tuberculin syringe and then emptying the syringe.13 A 0.1 ml aliquot of heart blood
was diluted in 0.9 ml PRAS-BHI broth in Hungate tubes and placed on ice for transport,
and 100 µl of this dilution was plated onto blood agar. All agar plates were incubated
anaerobically at 37°C for 48 hrs.
5.3.7 Western blot assay of serum antibodies
Immediately prior to infection and immediately following euthanasia, blood was
collected by mandibular vein technique and by post mortem cardiac puncture respectively
from all mice for serum anti - F. varium antibody analysis by western blotting. Blood
was centrifuged at 1,500 × g for 10 minutes, serum was separated and pooled by
treatment group, and frozen at -80°C. PRAS-BHI broth in Hungate tubes containing
overnight growth of Fusobacterium was centrifuged at 10,000 × g for 10 minutes. The
pellet was resuspended in a small amount of PBS and Laemmli sample buffer (Bio-Rad,
Hercules, CA, USA) containing β-mercaptoethanol and warmed on a heat block at 95°C
for 10 minutes. Colonies of E. coli, Pasteurella pneumotropica, or Arcanobacterium
pyogenes were collected from blood agar plates, placed directly into PBS and Laemmli
sample buffer, and treated as previously described. Resulting bacterial protein lysate was
loaded onto a 12% tris-HCl polyacrylamide gel (Bio-Rad) and electrophoresed at 160 V
for approximately one hour. Proteins were transferred overnight onto 0.45 µm
nitrocellulose membrane (Bio-Rad) at 22 V in a 4°C environment. The membrane was
incubated for 3 hours at room temperature with blocking solution containing 5% non-fat 96
dry milk in tris buffered saline (TBS) on an orbital shaker. The membrane was washed
with 0.05% TBS-Tween (TBST) and incubated with 1:50 mouse serum in a multiscreen
apparatus (Bio-Rad) for 1 hour at room temperature on an orbital shaker. The membrane
was washed with TBST and incubated with alkaline phosphatase (AP) conjugated goat
anti-mouse IgG or goat anti-mouse IgM (Santa Cruz Biotechnology, Santa Cruz, CA,
USA) each diluted 1:1000 for 1 hour at room temperature on an orbital shaker. The
membrane was washed with TBST, color developed with BCIP/NBT (Bio-Rad), and
development stopped with 5% acetic acid.
5.4 Results
5.4.1 Clinical effect of intranasal challenge
No mice infected with F. varium became clinically affected at any time in any of
the three trials. As summarized in Appendix C, no morbidity or mortality were observed
in F. varium infected mice, and a single mouse infected with F. n. subsp. necrophorum
became moribund at 4 days post infection, requiring euthanasia.
5.4.2 Gross and microscopic lesions
No mice infected with F. varium developed any gross or microscopic lesions that were attributed to fusobacterial infection. The one mouse that was euthanized in trial 2
following infection with F. n. subsp. necrophorum developed severe multifocal hepatic
necrosis that was grossly visible at necropsy. Histologically the lesion was severe
multifocal necrotizing hepatitis characterized by extensive multifocal random
hepatocellular necrosis with infiltration by neutrophils (Figure 5.1). Within the affected foci there were numerous gram negative filamentous bacteria consistent with F. n. subsp. 97
necrophorum detected by the Brown and Brenn method and the Warthin-Starry method
(Figure 5.2).
In the LPS pre-treatment group, 7 of 7 (100%) BALB/c mice that were euthanized at 14 days post infection displayed moderate splenomegaly on gross exam.
Microscopically, the splenomegaly resulted from lymphoid hyperplasia, consistent with an LPS-associated response. 1 of 8 (12.5%) C57BL/6 mice in the LPS pre-treatment group histologically displayed mild splenic lymphoid necrosis. Four uninfected mice, one BALB/c and one C57BL/6 from each of the two pre-treatment groups, were euthanized on day 3 post infection to monitor the effects of pre-treatment. Although no gross lesions were detected and no Fusobacterium was recovered, both mice (2 of 2,
100%) in the LPS group displayed microscopic changes consistent with LPS-associated injury including mild interstitial pneumonia characterized by multifocal mild to moderate thickening of the alveolar septa, moderate infiltration of alveolar septa by macrophages and rare neutrophils, marked type II pneumocyte hypertrophy, and mild vascular endothelial hypertrophy (Figure 5.3). These changes were not detected in any LPS- treated mice at 14 days post infection. In the two mice that were euthanized at day 3 from the dexamethasone pre-treatment groups, no significant gross or microscopic changes were detected and no Fusobacterium was recovered.
Finally, two background lesions were detected in mice of multiple treatment and control groups in all experiments. Minimal focal subacute to chronic hepatitis was detected in 8% to 69% of C57BL/6 mice in each of the three trials. Lesions were composed of a focal accumulation of lymphocytes and macrophages, occasionally with few neutrophils, within the hepatic parenchyma suggesting a chronic nature. The lesion 98
was not detected in any BALB/c mice. Additionally, moderate splenic hemosiderosis
was detected in 4 of 36 (11%) C57BL/6 mice pre-treated with IP dexamethsone. The
lesion was not detected in any C57BL/6 mice that were not treated with dexamethasone
or in any BALB/c mice.
5.4.3 Recovery of bacteria
F. varium (1.9 × 102 CFU/ml) was recovered from the blood of one mouse in trial
1, but no F. varium was recovered from any other tissues from any other mouse in any
trial except the positive controls of trial 3 (Appendix C). F. n. subsp. necrophorum too numerous to count was recovered from the blood, lung, and liver of the affected mouse from trial 2.
5.4.4 Western blot analysis of serum antibodies
Pre-infection pooled sera from day 0 from 8 of 12 (66.7%) C57BL/6 groups of trial 3 contained both IgM and IgG that bound to a fusobacterial protein approximately 45 kDa in size (Figure 5.4). Post-infection pooled sera from day 14 from each of the
C57BL/6 groups of trial 3 were nearly identical to pre-infection sera with similar bands present in 10 of 12 (83.3%) groups (Figure 5.5). These serum antibodies bound to a protein of similar size in protein lysates from F. n. subsp. necrophorum, F. n. subsp. funduliforme, and three different strains of F. varium. Serum antibodies also bound to an
Escherichia coli protein of approximately 50 kDa, but did not bind to any protein in lysates of Pasteurella pneumotropica, Arcanobacterium pyogenes, or Clostridium perfringens (Figure 5.6). No immunoglobulin that demonstrated affinity to fusobacterial protein lysate was detected in any pre-infection or post-infection serum from BALB/c mice. 99
5.5 Discussion
Immune defense mechanisms of the respiratory tract are numerous and complex
and involve both innate and adaptive defense mechanisms that must be tightly regulated
to prevent infection while minimizing damage to the respiratory tissues. Failure of a
single one of these defense mechanisms does not result in lung infection, but rather it is
believed that multiple defense mechanisms must be simultaneously affected.14 Because of this, establishment of infection in the lung by a bacterial pathogen is difficult. For this reason, many models of pulmonary bacterial infection have relied on methods of suppressing the host immune function, damaging the physical barriers of the respiratory immune system, or enhancing bacterial growth and survival. Systemic administration of dexamethasone, a synthetic glucocorticoid that simulates the effect of cortisol, has long been used for this purpose. Although the effects of glucocorticoid administration are numerous, the desired effect in the pulmonary infection model is non-specific immunosuppression which may allow for inoculated bacteria to evade depressed host immune mechanisms and establish infection.15 Creating damage to the physical epithelial
barrier of the lung is another common approach to inducing pulmonary bacterial
infection, often by creating a primary lung infection with a pneumotropic virus such as
influenza.16 It has been demonstrated that intranasal LPS is capable of causing similar
epithelial necrosis and inflammation as occur with some primary viral infections, thus it
may be possible to produce conditions amenable to bacterial infection following LPS
administration.17,18 100
Numerous studies have demonstrated the pathogenic effects of F. necrophorum in
cattle, mice, and rabbits.19-27 Very few studies, however, have been done on F. varium,
as this organism has long been considered nonpathogenic. Furthermore, aside from one
study that produced lung abscesses in mice following intrathoracic injection28, no
previous infectivity studies have been performed using respiratory tract infection with
Fusobacterium. Fortunately, numerous other models of pulmonary bacterial infection have been established in mice including Streptococcus pneumoniae, Pseudomonas aeruginosa, Bordetella pertussis, Bordetella parapertussis, Actinobacillus suis, and
Acinetobacter baumannii among others. Many of these models rely on immunosuppression or primary viral infection to establish conditions that will support bacterial infection in the lung or hasten to the disease process.15,16,29-31 The methods used
in this set of experiments were based on existing models of pulmonary bacterial infection
in mice. In the first two trials it was hypothesized that the bacterium was capable of
establishing lung infection without the need of an immunosuppressant or primary
infectious agent. When this approach failed to produce infection, immunosuppression
was induced by dexamethasone and pulmonary epithelial damage was produced by LPS.
Infection of the lung by F. varium was not able to be established in this study by
intranasal inoculation even by pre-treatment with dexamethasone or LPS. When F.
varium was replaced by a highly leukotoxigenic strain of F. n. subsp. necrophorum in trial 2, still no lung infection was produced, but rather hepatic infection developed in one animal. Lack of infection in all cases was demonstrated by failure to produce any of the following evidence: clinical signs, mortality, gross lesions, microscopic lesions, or the recovery of the bacterium from the lung, although gross and histologic lesions of LPS 101
exposure were present at day 3 post-infection.17,18,32 This suggests that fusobacterial
species, specifically F. varium, are not highly pathogenic and are unable to establish
infection alone without some severe predisposing condition.
In the mice of trial 3, even chemically induced immunosuppression or
pneumopathic LPS pre-treatments did not allow for bacterial infection to develop,
however the extent of immunosuppression resulting from the treatment regimen was not
determined. It has been reported that children with oral fusobacterial infections had
multiple nutrient deficiencies and widespread viral infections compared to healthy
counterparts.33 Perhaps in cases of lung infection in deer, a more severe insult to the respiratory immune defenses is produced by either a primary pathogen, such as an
undetected virus, or by dietary complications such as ruminal acidosis or nutrient deficiency. Alternatively, the mouse may not be the ideal species in which to develop the model. This species was selected not only for its ease of use, technical familiarity, and low cost, but also because numerous models of both fusobacterial infection and respiratory bacterial infection have been developed in the mouse. Future studies should consider the use of an alternative animal model such as the rabbit or natural host species such as domestic or wild ruminants. Furthermore, the route of exposure in natural infections of deer may not in fact be respiratory. It is possible that lung infection is the end result of hematogenous or systemic infection that originates from the gastrointestinal tract as in domestic ruminants, or by some other means.
Previous studies in mice and cattle using systemic infection with F. necrophorum
have demonstrated anti-fusobacterial serum antibody production.20-23,34 In this study, anti-fusobacterial IgM or IgG were not produced in serum by 14 days in response to 102
infection with F. varium. Pre-existing antibodies were detected in most pooled serum
samples; these antibodies bound identically with proteins of two F. necrophorum
subspecies and multiple F. varium isolates, but did not produce a 45 kDa band against protein lysates from other common bacterial pathogens including Escherichia coli,
Pasteurella pneumotropica, Arcanobacterium pyogenes, and Clostridium perfringens.
This suggests that these represent naturally occurring anti-fusobacterial antibodies, likely from the indigenous microflora of the oropharyngeal cavity, gastrointestinal tract, or urogenital tract. Interestingly, no such antibodies were detected in any BALB/c mice at any time, however no differences in response to infection were detected. Thus, the significance of these pre-existing antibodies is uncertain, but no protective effect was observed in this trial. Although antibodies against Fusobacterium were not produced in any mice in response to inoculation, many C57BL/6 mice may possess pre-existing anti- fusobacterial antibodies. That the BALB/c mice in this study did not possess such antibodies may suggest some difference in immune response or prior exposure to bacterial pathogens between mouse strains.
In conclusion, this series of experiments failed to produce pulmonary infection with F. varium or F. necrophorum in mice, and infected mice failed to produce serum antibodies in response to the exposure. Pulmonary infection was not able to be established even with pre-treatment with dexamethsone or LPS, suggesting that F. varium is not highly pathogenic in mice. Future studies should consider the use of alternative routes of infection, alternative animal species such as rabbits, cattle, or deer, or genetically modified mouse strains.
103
5.6 Acknowledgements
The author thanks the Missouri Whitetail Breeders and Hunting Ranch
Association; Dr. Vivek Kapur, Dr. Bhushan Jayarao, Dr. Subhashinie Kariyawasam, Lola
Hubler, and Bill Weaver of The Pennsylvania State University Department of Veterinary and Biomedical Sciences; and Dr. Sanjeev Narayanan and Amit Kumar of Kansas State
University College of Veterinary Medicine for their assistance.
5.7 References
1. Nagaraja TG, Narayanan SK, Stewart GC, et al. Fusobacterium necrophorum
infections in animals: pathogenesis and pathogenic mechanisms. Anaerobe
2005;11:239-246.
2. Tan ZL, Nagaraja TG, Chengappa MM. Factors affecting the leukotoxin activity
of Fusobacterium necrophorum. Vet Microbiol 1992;32:15-28.
3. Tadepalli S, Stewart GC, Nagaraja TG, et al. Leukotoxin operon and differential
expressions of the leukotoxin gene in bovine Fusobacterium necrophorum
subspecies. Anaerobe 2008;14:13-18.
4. Citron DM. Update on the taxonomy and clinical aspects of the genus
Fusobacterium. Clin Infect Dis 2002;35:S22-27.
5. Hattel AL, Shaw DP, Love BC, et al. A retrospective study of mortality in
Pennsylvania captive white-tailed deer (Odocoileus virginianus): 20000--2003. J
Vet Diagn Invest 2004;16:515-521. 104
6. Leighton FA. Fusobacterium necrophorum infection In: Williams ES,Barker IK,
eds. Infectious Diseases of Wild Mammals. Ames, Iowa: Iowa State Press,
2001;493-496.
7. Chirino-Trejo M, Woodbury MR, Huang F. Antibiotic sensitivity and biochemical
characterization of Fusobacterium spp. and Arcanobacterium pyogenes isolated
from farmed white-tailed deer (Odocoileus virginianus) with necrobacillosis. J
Zoo Wildl Med 2003;34:262-268.
8. Jang SS, Hirsh DC. Characterization, distribution, and microbiological
associations of Fusobacterium spp. in clinical specimens of animal origin. J Clin
Microbiol 1994;32:384-387.
9. Legaria MC, Lumelsky G, Rodriguez V, et al. Clindamycin-resistant
Fusobacterium varium bacteremia and decubitus ulcer infection. J Clin Microbiol
2005;43:4293-4295.
10. Minami M, Ando T, Okamoto A, et al. Seroprevalence of Fusobacterium varium
in ulcerative colitis patients in Japan. FEMS Immunol Med Microbiol 2009;56:67-
72.
11. Brooks JW, Jayarao BM, Kumar A, et al. Phenotypic and genotypic
characterization of Fusobacterium isolates from the respiratory tract of deer. Vet
Microbiol Submitted for Publication, February 2010.
12. Ludlam HA, Milner NJ, Brazier JS, et al. lktA-encoded leukotoxin is not a
universal virulence factor in invasive Fusobacterium necrophorum infections in
animals and man. J Med Microbiol 2009;58:529-530. 105
13. Chantler J, Cox DJ. Self-prepared heparinized syringes for measuring ionized
magnesium in critical care patients. Br J Anaesth 1999;83:810-812.
14. Caswell JL, al-Haddawi M. If lung defenses are this sophisticated, why is
bronchopneumonia so pervasive? ACVP Annual Conference 2006;166-171.
15. Satoh S, Oishi K, Iwagaki A, et al. Dexamethasone impairs pulmonary defence
against Pseudomonas aeruginosa through suppressing iNOS gene expression and
peroxynitrite production in mice. Clin Exp Immunol 2001;126:266-273.
16. Smith MW, Schmidt JE, Rehg JE, et al. Induction of pro- and anti-inflammatory
molecules in a mouse model of pneumococcal pneumonia after influenza. Comp
Med 2007;57:82-89.
17. Szarka RJ, Wang N, Gordon L, et al. A murine model of pulmonary damage
induced by lipopolysaccharide via intranasal instillation. J Immunol Methods
1997;202:49-57.
18. Zhang X, Song K, Xiong H, et al. Protective effect of florfenicol on acute lung
injury induced by lipopolysaccharide in mice. Int Immunopharmacol 2009.
19. Lechtenberg KF, Nagaraja TG. Hepatic ultrasonography and blood changes in
cattle with experimentally induced hepatic abscesses. Am J Vet Res 1991;52:803-
809.
20. Saginala S, Nagaraja TG, Lechtenberg KF, et al. Effect of Fusobacterium
necrophorum leukotoxoid vaccine on susceptibility to experimentally induced
liver abscesses in cattle. J Anim Sci 1997;75:1160-1166.
21. Saginala S, Nagaraja TG, Tan ZL, et al. The serum neutralizing antibody response
in cattle to Fusobacterium necrophorum leukotoxoid and possible protection 106
against experimentally induced hepatic abscesses. Vet Res Commun 1996;20:493-
504.
22. Saginala S, Nagaraja TG, Tan ZL, et al. Serum neutralizing antibody response and
protection against experimentally induced liver abscesses in steers vaccinated
with Fusobacterium necrophorum. Am J Vet Res 1996;57:483-488.
23. Narayanan SK, Chengappa MM, Stewart GC, et al. Immunogenicity and
protective effects of truncated recombinant leukotoxin proteins of Fusobacterium
necrophorum in mice. Vet Microbiol 2003;93:335-347.
24. Smith GR, Thornton EA. Pathogenicity of Fusobacterium necrophorum strains
from man and animals. Epidemiol Infect 1993;110:499-506.
25. Smith GR, Thornton EA. Classification of human and animal strains of
Fusobacterium necrophorum by their pathogenic effects in mice. J Med
Microbiol 1997;46:879-882.
26. Smith GR, Till D, Wallace LM, et al. Enhancement of the infectivity of
Fusobacterium necrophorum by other bacteria. Epidemiol Infect 1989;102:447-
458.
27. Smith GR, Wallace LM, Till D. Necrobacillosis and immunity in mice. Epidemiol
Infect 1989;103:211-215.
28. Maestrone G, Sadek S, Kubacki E, et al. Sphaerophorus necrophorus: laboratory
model for the evaluation of chemotherapeutic agents in mice. Cornell Vet
1975;65:187-204. 107
29. Joly-Guillou ML, Wolff M, Pocidalo JJ, et al. Use of a new mouse model of
Acinetobacter baumannii pneumonia to evaluate the postantibiotic effect of
imipenem. Antimicrob Agents Chemother 1997;41:345-351.
30. Nuermberger E, Helke K, Bishai WR. Low-dose aerosol model of pneumococcal
pneumonia in the mouse: utility for evaluation of antimicrobial efficacy. Int J
Antimicrob Agents 2005;26:497-503.
31. Ojha S, Hayes MA, Turner PV, et al. An experimental model of Actinobacillus
suis infection in mice. Comp Med 2007;57:340-348.
32. Sagara-Ishijima N, Furuhama K. Toxic characteristics of the synthetic lipid A
derivative DT-5461 in rats and monkeys. Toxicol Sci 1999;49:324-331.
33. Enwonwu CO, Falkler WA, Jr., Idigbe EO, et al. Pathogenesis of cancrum oris
(noma): confounding interactions of malnutrition with infection. Am J Trop Med
Hyg 1999;60:223-232.
34. Clark BL, Emery DL, Stewart DJ, et al. Studies into immunisation of cattle
against interdigital necrobacillosis. Aust Vet J 1986;63:107-110.
108
Trial 1 Design – # of mice CFU Inoc Vol Bacterial Strain (µl) 103 105 107 25 2 2 2 F varium P11V 50 2 2 2 25 2 2 2 F varium P16V 50 2 2 2 25 2 2 2 F varium P23V 50 2 2 2 12 12 12 Total mice = 36
Table 5.1 - Study design of trial 1.
109
Trial 2 Design – # of mice CFU Inoc Vol Bacterial Strain (µl) 107 109 25 2 2 F varium P11V 50 2 2 F necrophorum 25 2 2 P26A 50 2 2 8 8 Total mice = 16
Table 5.2 - Study design of trial 2.
110
Trial 3 Design – # of mice Pre‐Treatment IN Inoc Dex LPS None C57BL/6 F varium P11V 2 2 4 F varium P16V 2 2 2 F varium P23V 2 2 2 PRAS‐BHI 2 2 2 BALB/c F varium P11V 2 2 2 F varium P16V 2 2 F varium P23V 2 2 PRAS‐BHI 2 2 16 16 12 Total mice = 44
Table 5.3 - Study design of trial 3.
111
Figure 5.1
Liver – C57BL/6 mouse (H&E): Top panel shows focal necrosis rimmed by inflammatory cells and cellular debris. Bottom panel shows normal liver.
112
Figure 5.2
Liver – C57BL/6 mouse: Top panel shows gram negative bacilli in necrotic focus (Brown & Brenn). Bottom panel shows filamentous morphology of bacilli in necrotic focus (Warthin-Starry). 113
Figure 5.3
Lung – C57BL/6 mouse (H&E): Top panel shows diffuse interstitial pneumonia with alveolar septal thickening and type II pneumocyte hypertrophy. Bottom panel shows normal lung. 114
250 150 100 75 50
37
25
20
15
10 kD M 1 2 3 4 5 6 7 8 9 10 11 12
250 150 100 75
50
37
25
20
15
10
Figure 5.4
Western blot of pre-infection mouse serum. IgM (top) and IgG (bottom) serum antibodies were detected in C57BL6 mice following infection with F. varium. Lanes are as follows: M, protein marker; 1-4, no pre-treatment (P11V, P16V, P23V, PRAS-BHI); 5-8, dexamethasone (P11V, P16V, P23V, PRAS-BHI); 9-12, LPS (P11V, P16V, P23V, PRAS-BHI). 115
250 150 100 75 50
37
25
20
15
10 kD M 1 2 3 4 5 6 7 8 9 10 11 12 13 14
250 150 100 75 50
37
25
20
15
10
Figure 5.5
Western blot of post-infection mouse serum. No changes in IgM or IgG serum antibodies were detected in mice following infection with F. varium. Lanes are as follows: M, protein marker; 1, rabbit anti-fusobacterial antiserum; 2, pooled pre-infection serum; 3-6, no pre-treatment (P11V, P16V, P23V, PRAS-BHI); 7-10, dexamethasone (P11V, P16V, P23V, PRAS-BHI); 11-14, LPS (P11V, P16V, P23V, PRAS-BHI). 116
kD M 1 2 3 4 56 7 8 9 150 100 75
50
37
25
20
15
10
Figure 5.6
Western blot of pre-infection mouse serum from C57BL/6. Serum antibodies bound similarly to protein from Fusobacterium spp. but not to protein from other bacteria. Lanes are as follows: M, protein marker; 1, F. necrophorum subsp. necrophorum; 2, F. necrophorum subsp. funduliforme; 3, F. varium P11V; 4, F. varium P16V; 5, F. varium P23V; 6, Escherichia coli; 7, Pasteurella pneumotropica; 8, Arcanobacterium pyogenes, 9, Clostridium perfringens. 117
Chapter 6
SUMMARY AND CONCLUSIONS
118
The results of this series of studies are preliminary and need to be interpreted as an
instrument to help direct additional future investigation of the subject. Nonetheless, some
practical conclusions can be drawn that may have immediate implications both in the field and in the laboratory.
Results of the producer survey suggest that preventive medicine practices are improperly
used or underused in many herds. The frequency of vaccination and the number of vaccines
used increases as herd size increases, however this effect also parallels an increase in reporting of
most disease conditions, questioning the practical efficacy of many vaccination programs.
Vitamin E and selenium supplements are regularly administered on less than half of the farms
despite that fact that vitamin E or selenium deficiency is a common cause of death in deer.
Although most respondents routinely administer anthelmintics, the efficacy of parasite control
programs on many of the farms is questionable due to the high frequency of anthelmintic
administration, random use of multiple products, predominant use of ivermectin, or too
infrequent use of anthelmintics. Most farms are deficient in their recognition of biosecurity
issues, and many respondents take no specific biosecurity measures when purchasing animals, a
serious risk for introduction of infectious diseases. Thus, it is essential that veterinary
practitioners and deer farmers collaborate to design and implement effective animal health and
biosecurity programs.
The disease conditions identified by the producer survey correlate well with those of the
retrospective mortality study by Hattel et al (2004) and several other studies, highlighting
respiratory tract disease as the most frequent disease of farmed deer. The underlying cause of
respiratory tract disease in deer is not entirely understood, although one of the most commonly
isolated pathogens is Fusobacterium. Through the characterization of a set of 28 clinical 119
Fusobacterium isolates from the respiratory tract of white-tailed deer, the author has determined that F. varium, not F. necrophorum, comprises the majority (75.0%) of the clinical isolates from cases of fusobacterial respiratory tract infections in deer. Although no leukotoxin production by
F. varium was detected by PCR or by the leukocyte viability assay, no difference was detected between lesion scores caused by F. varium and F. necrophorum suggesting that infection with either species may result in production of similar lesions. Perhaps of greater immediate significance in the field is that the antimicrobial susceptibility is quite different between the two species. In this study all fusobacteria were susceptible to ampicillin, florfenicol, and trimethoprim/sulfamethoxazole, however F. varium was much less susceptible than F. necrophorum to clindamycin, tilmicosin, tulathromycin, and tylosin and moderately less susceptible to chlortetracycline, oxytetracycline, penicillin, and tiamulin. This is of great clinical relevance, as all existing treatment and prevention protocols have been developed against F. necrophorum infection. In order to correctly diagnose and treat fusobacterial infection, it is recommended that clinical isolates be identified to the species or subspecies level and that an antimicrobial susceptibility panel be performed.
Experimental infection of the lung by F. varium was not able to be established by intranasal inoculation of mice in this series of studies. In subsequent experiments, infection was not able to be established even following immunosuppressive pre-treatment with dexamethasone or pro-inflammatory pre-treatment with bacterial LPS. When F. varium was replaced by F. necrophorum subsp. necrophorum in one trial, still no lung infection was produced, but liver infection developed in one animal. The inability to produce lung infection in any mice suggests that fusobacterial species, specifically F. varium, are not highly pathogenic to the respiratory tract and are unable to establish lung infection alone without some severe predisposing condition. 120
Such conditions in deer may arise following infection by a primary pathogen, such as an
undetected virus or bacterium, by dietary complications such as ruminal acidosis, by a deficiency
of an essential nutrient such as copper or selenium, or in response to chronic physiologic stress.
Additional studies of fusobacterial disease are clearly required. Future studies should
consider the use of alternative routes of infection, the effects of co-infection with mixed bacterial
species, and the use of alternative animal species such as rabbits or even naturally affected
ruminant species such as cattle and deer. Epidemiologic studies of deer populations should be
conducted to better determine the risk factors and predisposing conditions that may lead to
fusobacterial pneumonia. The effects of infection and vaccination upon the immune response
should be more thoroughly studied in order to determine the efficacy of currently available
commercial vaccines and to assess the need for developing more effective vaccines to prevent
fusobacterial disease.
In conclusion, while F. varium and F. necrophorum can clearly cause severe soft tissue
infection in multiple species of animals and humans, the pathogenesis of such infections remains
undetermined. Based upon the results of this comprehensive study, although much remains to be
understood about fusobacterial infection, the following recommendations can be made for deer farmers and veterinarians while additional research continues. Management changes should be
made as described in an effort to minimize stress, overcrowding, ruminal acidosis, and
environmental contamination, while maximizing immunity through colostrum and vaccination,
and developing effective preventive health and biosecurity programs with routine veterinary
consultation. Cases of fusobacterial infection should be tested by a veterinary diagnostic
laboratory and the organism identified to the species level. Antibiotics effective for the
infectious strain should be administered, but empirical use of ampicillin, florfenicol, or 121
trimethoprim/sulfamethoxazole may be the most appropriate selection in the absence of an antimicrobial susceptibility profile. Although no direct evaluation of vaccine efficacy was made in these studies, the lack of leukotoxin production by F. varium and the cross-reactivity of murine serum antibodies to various Fusobacterium types suggest that bacterin vaccines may be protective against fusobacterial infection, therefore the use of anti-fusobacterial bacterin vaccines is recommended at this time. 122
Appendix A
SELF-ADMINISTERED QUESTIONNAIRE
Pennsylvania Deer Farming Survey
Please respond to each of the following questions and return this form in the envelope provided. Your completion and return of this questionnaire implies consent for Penn State University to use this information for the purposes described in the attached letter of consent.
Select only one answer for each question unless otherwise indicated.
1. What species of deer do you raise? (check all that apply) White-tailed deer ____ Red deer ____ Elk ____ Reindeer ____ Other ______
2. a. How many total deer do you have at your facility? ______b. Number of adult females (over 1 year of age) ______c. Number of adult males (over 1 year of age) ______d. Number of juveniles (less than 1 year of age) ______
3. For what purpose(s) do you raise deer? (check all that apply) Breeding stock ____ Hunting stock ____ Venison ____ Hard antlers ____ Velvet antlers ____ Other ______
4. How many acres do you have dedicated to deer farming? ______
5. a. Do you intend to add more animals in the next five years? Yes/No b. Do you intend to add more acreage in the next five years? Yes/No c. Do you intend to reduce the number of animals in the next five years? Yes/No d. Do you intend to reduce the number of acres in the next five years? Yes/No
6. How frequently do you vaccinate your animals? Less than once per year ____ Once per year ____ Twice per year ____ More than twice per year ____ Never ____
123
7. What vaccine(s) do you use on your animals? (check all that apply) Clostridium (ex: Covexin 8, Ultrabac 7) ____ Leptospirosis (ex: Lepto 5, Leptoferm 5) ____ Fusobacterium (ex: Volar, Fusogard) ____ Other ______None ____
8. Do you give selenium or vitamin E injections (ex: Bo-Se, Vital E)? Yes/No a. If yes, please fill in the following table:
To what animals do you When do you give How do you give the What product do you give injections? injections? medicine? use?
ex: juveniles ex: spring and fall ex: injection ex: Bo-Se
9. Do you use dewormers on your animals? Yes/No a. If yes, please fill in the following table:
How do you give the What product do you When do you deworm? medicine? use?
ex: spring and fall ex: injection ex: Ivomec
10. Which of the following is/are used in the breeding program on your farm? (check all that apply) Natural breeding by male(s) in pen with females ____ Artificial insemination using purchased semen ____ Artificial insemination using semen collected on your farm ____ Sale of semen collected on your farm ____ Embryo transfer ____ Other ______
11. Do you have a veterinarian regularly visit your farm? Yes/No a. If yes, how often? Less than once per year ____ Once per year ____ Twice per year ____ More than twice per year ____ 124
b. If yes, what service(s) do you regularly request of your veterinarian? (check all that apply) Treatment of sick animals ____ Nutritional advice ____ Disease prevention (vaccination or deworming advice) ____ Reproduction ____ Antler removal ____ Health forms for transport of animals ____ Necropsy of dead animals ____ Sedation of animals for handling ____ Other ______
12. Are there additional veterinary services that are not currently offered that you would you like to receive? Yes/No a. If yes, what services? ______
13. Have you ever had an animal that has died examined after death (necropsied) either by your veterinarian or at an animal diagnostic laboratory? Yes/No a. If yes, how often do you request a necropsy after an animal dies? Always ____ Most of the time ____ Rarely ____ b. If yes, where do you prefer to have your animal examined (necropsied)? On your farm by your regular veterinarian ____ At the Animal Diagnostic Lab at Penn State ____ At the Pennsylvania Veterinary Lab in Harrisburg ____ At the School of Veterinary Medicine at New Bolton Center ____ Other ______
14. Do you purchase animals from other farms and bring them to your farm? Yes/No a. If yes, please fill in the following table:
What health What type of animals Where do you buy When do you buy precautions do you do you buy? them? them? take?
ex: adult bucks ex: New York ex: every september ex: blood testing for__
15. Do your deer have access to pasture? Yes/No a. If yes, is the pasture: (check all that apply) Grass pasture ____ Alfalfa or clover pasture ____ Mixed pasture ____ please describe ______125
16. Do you feed hay? Yes/No a. If yes, is the hay: (check all that apply) Grass hay ____ Alfalfa or clover hay ____ Mixed hay ____ please describe ______
17. Do you feed grain? Yes/No a. If yes, is the grain: (check all that apply) Grain mixed on your farm____ Grain mixed at feed mill ____ Commercial deer pellet ____
18. Do you provide minerals? Yes/No a. If yes, are the minerals: (check all that apply) Mixed with the feed on your farm ____ Mixed with the feed at feed mill ____ Offered free choice as loose salt ____ Offered free choice as lick block ____
19. Which disease problem(s) would you consider to be the most common on your farm? (check all that apply and indicate which one is the biggest problem) Pneumonia ____ Diarrhea ____ Parasites ____ Weight loss ____ Foot rot ____ Sudden death ____ Abortion ____ Other ______
20. In what aspects of deer management and disease prevention would you most like to see research performed? (check all that apply) Nutrition ____ Infectious diseases ____ Parasite control ____ Reproduction ____ Antler growth ____ Economics of deer farming ____ Other ______
By completing and returning this questionnaire, you certify that you have read and understood all of the terms of the letter of consent and that you grant permission to Penn State University to use this information for the purposes described in the letter of consent.
Please return questionnaires to: Dr. Jason Brooks Animal Diagnostic Laboratory The Pennsylvania State University University Park, PA 16802-1110
126
Appendix B
SUPPLEMENTAL PHENOTYPIC DATA
Figure B.1 – Growth curves of Fusobacterium spp. in brain-heart infusion broth.
127
Figure B.2 – Summarized leukotoxin production by species as determined by toxicity to bovine PMNs following exposure to culture supernatant as measured by propidium iodide (PI) staining on flow cytometry.
128
Identification with Rapid ANA and RapID 32A Species/type Rapid Rapid ANA Result RapID 32A RapID 32A Result ANAII Code (quality of ID %) Code (quality of ID %) F. n. necrophorum 26 000104 F. necrophorum (99.8) 0000600000 F. necrophorum (70.7)/C. tetani (29) 38 000104 F. necrophorum (99.8) 0000600000 F. necrophorum (70.7)/C. tetani (29) ATCC25286 000104 F. necrophorum (99.8) 0000600000 F. necrophorum (70.7)/C. tetani (29) F. n. funduliforme 18 000004 F. necrophorum (60)/F. nucleatum (40) 0000200000 F. nucleatum (62)/ C. tetani (20.7) 20 000004 F. necrophorum (60)/F. nucleatum (40) 0000200000 F. nucleatum (62)/ C. tetani (20.7) 21 000004 F. necrophorum (60)/F. nucleatum (40) 0000200000 F. nucleatum (62)/ C. tetani (20.7) 27 000004 F. necrophorum (60)/F. nucleatum (40) 0000200000 F. nucleatum (62)/ C. tetani (20.7) 30 000004 F. necrophorum (60)/F. nucleatum (40) 0000200000 F. nucleatum (62)/ C. tetani (20.7) ATCC51357 000004 F. necrophorum (60)/F. nucleatum (40) 0000200000 F. nucleatum (62)/ C. tetani (20.7) F. varium 1 000007 F. varium (98.9) 0002204000 F. varium (99.9) 3 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 7 000006 F. nucleatum (>99.9) 0000204000 F. varium (92.5) 8 000057 F. varium (>99.9) 0002204000 F. varium (99.9) 9 000007 F. varium (98.9) 0002204000 F. varium (99.9) 10 000006 F. nucleatum (>99.9) 0000204000 F. varium (92.5) 11 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 12 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 14 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 16 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 17 000006 F. nucleatum (>99.9) 0000204000 F. varium (92.5) 22 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 23 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 24 000006 F. nucleatum (>99.9) 0000204000 F. varium (92.5) 29 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) 34 000047 F. varium (>99.9) 0002204000 F. varium (99.9) 35 000007 F. varium (98.9) 0002204000 F. varium (99.9) 36 000046 F. nucleatum (99.4) 0000204000 F. varium (92.5) 37 000006 F. nucleatum (>99.9) 0000204000 F. varium (92.5) 39 000047 F. varium (>99.9) 0002204000 F. varium (99.9) 41 000043 F. varium (>99.9) 0000004000 C. histolyticum (72)/F. varium (11.3) ATCC8501 000006 F. nucleatum (>99.9) 0000204000 F. varium (92.5) F. nucleatum ATCC25586 000004 F. necrophorum (60)/F. nucleatum (40) 0000200000 F. nucleatum (62)/ C. tetani (20.7)
Table B.1 – Results for Fusobacterium isolates using Remel Rapid ANA II and Biomerieux RapID 32A anaerobic bacterial identification kits.
129
Gross and microscopic lesions and co-pathogens associated with Fusobacterium infection Fnn Fnf Var Lesion (n = 2) P (n = 5) P (n = 20) P Total Pneumonia 1 1.000 4 0.617 11 0.352 16 Rumenitis 1 0.231 1 1.000 4 0.596 6 Oropharyngitis 0 1.000 1 1.000 4 1.000 5 Nephritis 0 1.000 2 0.236 3 0.558 5 Hepatitis 0 1.000 1 1.000 3 1.000 4 Carditis 0 1.000 0 1.000 3 1.000 3
Fnn Fnf Var Co-pathogen (n = 2) P (n = 5) P (n = 20) P Total E. coli 1 1.000 3 0.628 14 1.000 18 A. pyogenes 1 0.346 2 1.000 6 0.628 9 B. fragilis 0 1.000 2 0.588 5 1.000 7 Bacteroides 0 1.000 1 1.000 4 1.000 5 Streptococcus 0 1.000 1 1.000 4 1.000 5 Pasteurella 0 1.000 0 0.555 4 0.543 4
Table B.2 – Results of χ2 analysis of association between lesion production or co- pathogen and fusobacterial infection. No significant associations were detected.
Table B.3– performed. respiratory fusobacterial infection ina ContingencyAnalysis of Resp FusoBy Pneumonia Exact Test 's r he Fis Test 2-Tail Right Left erson s Pear Likelihood Ratio Odds Ratio Tests Contingency Table Odds Ratio Pneumonia 1.042453 1 0 % Row % Col % Total Count 104 N 0 Results ofcontingencyanalysis 68.00 24.29 16.35 67.09 75.71 50.96 67.31 e u o Fus p Res Lower 95% Lower 1.0000 0.5691 0.6236 17 53 70 DF Prob ChiSquare 0.398157 1 1 0.00359335 32.00 23.53 32.91 76.47 25.00 32.69 7.69 -LogLike 0.007 0.007 26 34 8 Alternative Hypothesis Prob(Resp Fuso=1) is different across Pneumonia across Prob(Respdifferent Fuso=1)is Pneumonia=1 for Prob(Respgreater Fuso=1)is than0 Pneumonia=0 for Prob(Respgreater Fuso=1)is than1 Upper 95% 75.96 24.04 Prob>ChiSq 2.729347 104 79 25 Sure(U) e RSquar 0.9325 0.9324 0.0001 ll deeronwhichanaerobiccultureswere associationbetweenpneumoniaand
130 131
Contingency Analysis of Resp A pyo By Pneumonia Contingency Table Res p A py o Count 0 1 Total % Col % Row % 0 21 4 25 20.19 3.85 24.04 36.84 8.51 84.00 16.00 1 36 43 79
Pneumonia 34.62 41.35 75.96 63.16 91.49 45.57 54.43 57 47 104 54.81 45.19 Tests
N DF -LogLike RSquare (U) 104 1 6.1659534 0.0861
Test ChiSquare Prob>ChiSq Likelihood Ratio 12.332 0.0004* Pearson 11.323 0.0008*
Fisher's Exact Test Prob Alternative Hypothesis Left 0.9999 Prob(Resp A pyo=1) is greater for Pneumonia=0 than 1 Right 0.0006* Prob(Resp A pyo=1) is greater for Pneumonia=1 than 0 2-Tail 0.0010* Prob(Resp A pyo=1) is different across Pneumonia
Odds Ratio Odds Ratio Lower 95% Upper 95% 6.270833 1.971142 19.94953 Contingency Analysis of Resp A pyo By Pneumonia Cochran-Mantel-Haenszel Tests Stratified by Resp Fuso CMH Test ChiSquare DF Prob>Chisq Correlation of Scores 11.1175 1 0.0009* Row Score by Col Categories 11.1175 1 0.0009* Col Score by Row Categories 11.1175 1 0.0009* General Assoc. of Categories 11.1175 1 0.0009* Frequency Counts
Res p Fuso=0 Pneumonia Re s p A pyo 0 1 0 14 24 1 3 29 Res p Fuso=1 Pneumonia Re s p A pyo 0 1 0 7 12 1 1 14
Table B.4 – Results of contingency analysis of association between pneumonia and respiratory infection with Arcanobacterium pyogenes in all deer on which anaerobic cultures were performed. 132
Appendix C
BACTERIOLOGY, GROSS, AND MICROSCOPIC PATHOLOGY RESULTS
FROM TRIALS 1, 2, AND 3
Trial 1 – Bacteriology (day 14) Lung Colonies per plate Approximate Dosage (CFU) Bacterial Inoc Vol Strain (µl) 10Exp3 10Exp5 10Exp7 25 0 0 0 P11V 50 0 0 0 25 ‐ ‐ 0 P23V 50 ‐ ‐ 0 25 ‐ ‐ 0 P16V 50 ‐ ‐ 0
Blood Colonies per plate Approximate Dosage (CFU) Bacterial Inoc Vol Strain (µl) 10Exp3 10Exp5 10Exp7 25 0 19 0 P11V Direct 50 0 ‐ 0 25 0 0 0 P11V BHI 50 0 0 0
Table C.1 – Bacteriology results of trial 1.
133
Trial 1 – Gross Pathology Gross Lesions (All mice euthanized on day 14)
Bacterial Strain Inoc Vol (µl) 103 105 107 25 0 0 0 P11V 50 0 0 0 25 0 0 0 P23V 50 0 0 0 25 0 0 0 P16V 50 0 0 0
Table C.2 – Necropsy results of trial 1.
Trial 1 – Histopathology (day 14)
Bacterial Inoc Vol Strain (µl) 103 105 107 25 ‐ ‐ 1 of 2 mild hepatitis P11V 50 ‐ ‐ 0 25 ‐ ‐ 0 P23V 50 ‐ ‐ 0 25 ‐ ‐ 1 of 2 mild hepatitis P16V 50 ‐ ‐ 1 of 2 mild hepatitis
Table C.3 – Histology results of trial 1.
134
Trial 2 – Bacteriology (day 14 unless indicated by *) Lung Colonies per plate Approximate Dosage (CFU) Bacterial Strain Inoc Vol (µl) 107 109 25 0 0 P11V 50 0 0 25 0 TNTC* P26A 50 0 0
Blood Colonies per plate Approximate Dosage (CFU) Bacterial Strain Inoc Vol (µl) 107 109 P11V Blood 25 0 0 Agar 50 0 0 P26A Blood 25 0 0 Agar 50 0 0 25 0 0 P11V FSA 50 0 0 25 0 0 P26A FSA 50 0 0 25 0 0 P11V BHI 50 0 0 25 0 TNTC* P26A BHI 50 0 0 * Euthanized day 4
Table C.4– Bacteriology results of trial 2.
135
Trial 2 – Gross Pathology Gross Lesions (euthanized on day 14 unless indicated by *)
Bacterial Strain Inoc Vol (µl) 107 109 25 0 0 P11V 50 0 0 25 0 1 of 2 hepatic necrosis* P26A 50 0 0 * Euthanized day 4
Table C.5 – Necropsy results of trial 2.
Trial 2 – Histopathology (day 14 unless indicated by *)
Bacterial Inoc Vol Strain (µl) 107 109 25 2 of 2 mild hepatitis 2 of 2 mild hepatitis P11V 50 1 of 2 mild hepatitis 1 of 2 mild hepatitis 1 of 2 mild hepatitis; 1 of 2 severe 25 2 of 2 mild hepatitis necrotizing hepatitis* P26A 50 2 of 2 mild hepatitis 0 * Euthanized day 4
Table C.6 – Histology results of trial 2.
136
Trial 3 ‐ Morbidity and Mortality Clinical Signs
Mouse Strain Bacterial Strain Dexamethasone Dexamethasone LPS LPS None None P11V 0 0 0 0 0 0 P23V 0 0 0 0 C57BL/6 P16V 0 0 0 0 BHI 0 0 0 0 P11V 0 0 0 0 0 0 P23V 0 0 0 0 BALB/c P16V 0 0 0 0 BHI 0 0 0 0
Death
Mouse Strain Bacterial Strain Dexamethasone Dexamethasone LPS LPS None None P11V Euth d14 Euth d14 Euth d14 Euth d14 Euth d0 Euth d0 P23V Euth d14 Euth d14 Euth d14 Euth d14 C57BL/6 P16V Euth d14 Euth d14 Euth d14 Euth d14 BHI Euth d3 Euth d14 Euth d3 Euth d14 P11V Euth d14 Euth d14 Euth d14 Euth d14 Euth d0 Euth d0 P23V Euth d14 Euth d14 Euth d14 Euth d14 BALB/c P16V Euth d14 Euth d14 Euth d14 Euth d14 BHI Euth d3 Euth d14 Euth d3 Euth d14
Table C.7 – Morbidity and mortality results of trial 3.
137
Trial 3 – Bacteriology Lung Fusobacterium CFU/g lung
Mouse Bacterial Strain Strain Dexamethasone Dexamethasone LPS LPS None None P11V 0 0 0 0 4.3 X 10exp5 2.1 X 10exp4 P23V 0 0 0 0 C57BL/6 P16V 0 0 0 0 BHI 0 0 0 0 P11V 0 0 0 0 1.1 X 10exp6 9.1 X 10exp5 P23V 0 0 0 0 BALB/c P16V 0 0 0 0 BHI 0 0 0 0
Blood Fusobacterium CFU/ml blood
Mouse Bacterial Strain Strain Dexamethasone Dexamethasone LPS LPS None None P11V 0 ‐ 0 ‐ 0 ‐ P23V 0 ‐ 0 ‐ C57BL/6 P16V 0 ‐ 0 ‐ BHI ‐ ‐ ‐ ‐ P11V 0 ‐ 0 ‐ 0 ‐ P23V 0 ‐ 0 ‐ BALB/c P16V 0 ‐ 0 ‐ BHI ‐ ‐ ‐ ‐
Table C.8 – Bacteriology results of trial 3.
138
Trial 3 – Gross Pathology Gross Lesions
Mouse Strain Bacterial Strain Dexameth Dexameth LPS LPS None None P11V 0 0 0 0 0 0 P23V 0 0 0 0 C57BL/6 P16V 0 0 0 0 BHI 0 0 0 0 P11V 0 0 Splenomegaly Splenomegaly 0 0 P23V 0 0 Splenomegaly Splenomegaly BALB/c P16V 0 0 Splenomegaly Splenomegaly BHI 0 0 0 Splenomegaly
Table C.9 – Necropsy results of trial 3.
Trial 3 – Histopathology
Mouse Bacterial Strain Strain Dexamethasone Dexamethasone LPS LPS None None P11V Spl hemosiderosis 0 Mild hepatitis 0 Mild hepatitis 0 P23V Mild hepatitis 0 Spl necrosis 0 C57BL/6 Mild hepatitis, Spl P16V hemosiderosis Mild hepatitis 0 0 BHI Spl hemosiderosis Spl hemosiderosis Int Pneumo 0 P11V 0 0 0 0 0 0 P23V 0 0 0 0 BALB/c P16V 0 0 0 0 BHI 0 0 Int Pneumo 0
Table C.10 – Histology results of trial 3.
VITA Jason W. Brooks Education: • Ph.D. (Pathobiology) – The Pennsylvania State University – 2010 • Residency (Veterinary Pathology) – The Pennsylvania State University – 2003 through 2007 • V.M.D. – University of Pennsylvania, School of Veterinary Medicine – 2000 • B.S. (Biology) – Juniata College – 1996
Selected Publications and Presentations: • Brooks JW, Jayarao BM, Kumar A, Narayanan S, Myers S, Nagaraja TG. Phenotypic and genotypic characterization of Fusobacterium isolates from the respiratory tract of deer. Veterinary Microbiology. Submitted February 2010. • Shock BC, Brooks JW, Pokorny E, Mattive T, Yabsley MJ. Clinical challenge (Taenia crassiceps infection in a ring-tailed lemur). Journal of Zoo and Wildlife Medicine, 41(2): 379–381, 2010. • Johnson AJ, Pessier AP, Wellehan JFX, Childress A , Norton TM, Stedman NL, Bloom DC, Belzer W, Titus VR, Wagner R, Brooks JW, Spratt J, Jacobson ER. Ranavirus infection of free-ranging and captive box turtles and tortoises in the United States. Journal of Wildlife Diseases, 44:851-863, Oct 2008. • DebRoy C, Roberts E, Jayarao BM, Brooks JW. Bronchopneumonia associated with Extraintestinal Pathogenic Escherichia coli in a horse. Journal of Veterinary Diagnostic Investigation, 20:661-664, Sept 2008. • Brooks JW, Jayarao BM. Management practices used by white-tailed deer farms in Pennsylvania and herd health problems. JAVMA, 232:98-104, Jan 2008. • Brooks JW, Key DW, Hattel AL, Hovingh EP, Peterson R, Shaw DP, Fisher JS. Failure to detect bovine viral diarrhea virus in necropsied farm-raised white-tailed deer (Odocoileus virginianus) in Pennsylvania. Journal of Veterinary Diagnostic Investigation, 19:298-300, May 2007. • Hattel AL, Shaw DP, Fisher JS, Brooks JW, Love BC, Drake TR, Wagner DC. Mortality in Pennsylvania captive elk (Cervus elaphus): 1998-2006. Journal of Veterinary Diagnostic Investigation, 19:334-337, May 2007. • Hattel AL, Pokorny S, Brooks JW, Fisher JS. Arnold-Chiari type II malformation in two captive African lion littermates. Poster presentation: ACVP Annual Meeting. Tucson, AZ. Dec 2006. • Brooks JW, Whary MT, Hattel AL, Shaw DP, Ge Z, Fox JG, Poppenga RH. Clostridium piliforme infection in two farm-raised white-tailed deer fawns (Odocoileus virginianus) and association with copper toxicosis. Veterinary Pathology, 43:765-768, Sept 2006. • Hattel AL, Shaw DP, Love BC, Wagner DC, Drake TR, Brooks JW. A retrospective study of mortality in Pennsylvania captive white-tailed deer (Odocoileus virginianus): 2000- 2003. Journal of Veterinary Diagnostic Investigation 16:515-521, Nov 2004. • Hattel AL, Shaw DP, Love BC, Brooks JW, Fisher JS. Systemic mycoses in three captive juvenile reindeer. Poster presentation: ACVP Annual Meeting. Orlando, FL. Nov 2004.