CAMPYLOBACTER SPP. IN CONVENTIONAL AND ORGANIC POULTRY OPERATIONS
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctoral of Philosophy in the Graduate School of The Ohio State University
By
Taradon Luangtongkum, D.V.M.
*****
The Ohio State University 2005
Dissertation Committee: Dr. Teresa Y. Morishita, Advisor Approved by Dr. Yehia M. Saif Dr. Charles A. Neer ______Dr. Qijing Zhang Advisor Graduate Program in Veterinary Preventive Medicine
ABSTRACT
Campylobacter spp. particularly C. jejuni has been recognized as one of the most important causes of foodborne bacterial diarrhea in humans worldwide. Since poultry are considered the major reservoir of this foodborne pathogen, the prevalence along with the antimicrobial resistance of Campylobacter spp. in conventional and organic poultry operations is a matter of concern. However, relatively little information on the impact of different food animal production practices on the prevalence of antibiotic-resistant
Campylobacter is available. Hence, a farm-based study was conducted to determine the prevalence and the antimicrobial resistance rates of Campylobacter isolates from both conventionally-raised and organically-raised poultry. Although Campylobacter was highly prevalent in both organic and conventional production systems, the antibiotic resistance rates were significantly higher in conventional operations than in organic operations. These findings clearly indicate the influence of different production practices on the antibiotic resistance rates of Campylobacter spp. on poultry farms, which is likely due to the result of antimicrobial usage in conventional poultry production. However, antimicrobial usage alone may not solely be responsible for the increased antibiotic resistance in Campylobacter isolates because even in the absence of antibiotic exposure, a high level of tetracycline resistance was also observed in Campylobacter isolates from organic poultry farms. Since antibiotics including tetracycline have never been used on these organic poultry farms, a high
ii prevalence of tetracycline resistance observed among Campylobacter isolates from this operation type is interesting. Hence, another part of this study was conducted to investigate the epidemiology of tetracycline resistance of thermophilic Campylobacter on organic poultry farms. The results showed the changes of tetracycline resistance rates in
Campylobacter isolates during the production cycle of one organic broiler flock with 0%,
100%, and 33.33% of tetracycline resistance rates detected at week 3, 6, and 10 of the production period, respectively. Although no tetracycline-resistant Campylobacter strains were isolated from unexposed environmental samples collected from these organic poultry farms except for one organic broiler farm, some of these samples were positive for tetracycline resistance gene [tet(O) gene]. In addition, the agreement between the disk diffusion method and the agar dilution method in identification of antimicrobial resistance of thermophilic Campylobacter was also investigated. The results indicated that the disk diffusion method can be used as a reliable alternative method for susceptibility testing of
Campylobacter species to fluoroquinolone and aminoglycoside antibiotics; however, until the standard resistance breakpoints specific for Campylobacter is established and validated, the agar dilution method should be used to determine antimicrobial resistance of Campylobacter to other classes of antibiotics. Together, this study reveals the complex nature in the spread of antibiotic resistance and further highlights the need for prudent measures to prevent the occurrence and transmission of antibiotic-resistant Campylobacter in the poultry reservoir.
iii
Dedicated to my parents, Songkram and Theraphan Luangtongkum
Without their love and support this dissertation would not have been possible
iv ACKNOWLEDGEMENTS
I wish to thank my advisor, Dr. Teresa Y. Morishita, for providing me the
opportunity and freedom to conduct this research, and for her intellectual guidance,
support, and encouragement, which made this dissertation possible. The experiences that
I have gained from working with her for over the last 5 years here at The Ohio State
University have been invaluable to my future career.
I am also grateful to my dissertation committee members, Dr. Yehia Mo. Saif, Dr.
Charles A. Neer, and Dr. Qijing Zhang for their kindness and willingness to serve on the committee, as well as for their contribution on this dissertation. In this regard, I particularly would like to thank Dr. Qijing Zhang for his invaluable advice, for sharing with me his knowledge and discussions, and for being so patient in answering all the questions that I have had over the last several years, especially while I was writing the manuscripts for publication.
I also wish to thank Dr. Amna B. El-Tayeb, Dr. Meredith F. Davis, Dr. Benjamas
Promsopone, Ms. Elisabeth J. Angrick, Mr. Aaron J. Ison, and all of the many fellow graduate, undergraduate, and veterinary students at the Avian Disease Investigation
Laboratory, both past and present, whom I have had a chance to know and work with, for creating a pleasant environment in which to work, and for offering me their support, especially friendship inside and outside of the laboratory for over the last several years
v that I have been here in the States. The friendship and encouragement that I have received from Dr. William L. Ingalls since the first day that I have been in the Department also deserve a space here.
Many thanks also go to the organic and conventional poultry farms and processing plants for participating in this study. Research funding from the United States
Department of Agriculture (USDA) and the North Central Region program for
Sustainable Agriculture Research and Education (NCR-SARE) are also gratefully acknowledged.
I am indebted to The Anandamahidol Foundation for their financial support throughout my entire study. Without this support, I would not have been able to pursue my doctoral degree in the United States.
I am also very thankful to Ms. Nuttha Thongchul for everything she has done for me and for being so supportive and understanding. The help of Ms. Supaporn
Suwannakham on this dissertation is truly appreciated as well.
Finally, my heartfelt gratitude goes to those who are the most important persons in my life, my parents, Songkram and Theraphan Luangtongkum, for their unconditional love and life-long support, for always encouraging me, believing in me, being my inspiration, and helping me get through the tough, difficult, and stressful times during my studies. Without them, this doctoral degree would not have been possible.
vi VITA
April 19, 1976 ………………… Born – Bangkok, Thailand 1993 – 1999 …………………... Doctor of Veterinary Medicine (Honors) Chulalongkorn University, Bangkok, Thailand 1999 – present ………………… Instructor, Department of Veterinary Public Health Chulalongkorn University, Bangkok, Thailand 2000 – present ………………… Graduate Student, Department of Veterinary Preventive Medicine The Ohio State University, Columbus, OH
PUBLICATION
1. Huang, S., T. Luangtongkum, T. Y. Morishita, and Q. Zhang. 2005. Molecular typing of Campylobacter strains using the cmp gene encoding the major outer membrane protein. Foodborne Pathog. Dis. 2: 12-23.
FIELDS OF STUDY
Major Field: Veterinary Preventive Medicine Studies in Food Safety and Public Health
vii TABLE OF CONTENTS
Page Abstract…………………………………………………………….………...... …. …ii Dedication…………………………………………………………….…………...... …iv Acknowledgments………………………………………………………………..……..….v Vita…………………………………………………………………………………...... vii List of Tables……………………………………………………………...... ….x List of Figures………………………………………………………………………...…. .xii
Chapters:
1. Literature review ……………….……………………………………………...…....…..1
1.1 Introduction…………………………………………...………………………..…..1 1.2 Campylobacter: The Organism…………………………………………………...... 7 1.2.1 Historical background……………………….…………………………... …..7 1.2.2 General characteristics ….………………………………………………. …..9 1.2.3 Isolation and identification………………...…………………………….…..13 1.3 Campylobacter spp. in Humans…………………………………………..………. ..31 1.3.1 Sources and modes of transmission………….……………………………….31 1.3.2 Infection and clinical manifestation……………………………….....…..…..36 1.3.3 Treatment and control and prevention……….…………………………..…..45 1.4 Campylobacter spp. in Poultry…………………………..……………………..…..56 1.4.1 Prevalence in poultry and other animals……………………………………. ..56 1.4.2 Sources of poultry flock colonization….……………………………………..70 1.4.3 Control and prevention at the farm level……………………………..……..81 1.5 Antimicrobial Resistance of Campylobacter species…………………………..…..86 1.5.1 Antimicrobial susceptibility testing methods……………...………………..86 1.5.2 Occurrence and trends of antimicrobial resistance………...…………….…. ..89 1.5.3 Mechanisms of antimicrobial resistance………………………..………..…..95 1.6 References……………………………………………………………………...…101
2. The effect of different poultry production practices on antibiotic resistance rates of Campylobacter in poultry…………………………………………………….....…150 2.1 Abstract…………………………………………………………………………..150 2.2 Introduction…………………………………………………………………….…151 2.3 Materials and methods……………………………………………………………155
viii 2.4 Results…………………………………………………....…………………….…159 2.5 Discussion…………………………………………………………....………...…164 2.6 Acknowledgement…………………………………………………………………172 2.7 References………………………………………………………..…………….…172
3. Antimicrobial susceptibility testing of Campylobacter by the agar dilution and the agar disk diffusion methods………………………………………………………190 3.1 Abstract…………………………………….……………………………..…….…190 3.2 Introduction…………………………………………………………………….…..191 3.3 Materials and methods………………………………………………………….…194 3.4 Results…………………………………………………………………………....197 3.5 Discussion…………………………………………………………………………. 200 3.6 Acknowledgement…………………………………………………………………207 3.7 References……………………………………………………………………....…208
4. Tetracycline resistance of Campylobacter isolates on organic poultry farms……..…216 4.1 Abstract………………………………………………………………….....…..…216 4.2 Introduction…………………………………………………………………….…217 4.3 Materials and methods…………………………….....……………………………222 4.4 Results………………………………………………………………………….…228 4.5 Discussion……………………………………………………………………...…232 4.6 Acknowledgement……………………………………………………………..…239 4.7 References…………………………………………………………………………239
Bibliography……………………………………………………………….………....…253
ix LIST OF TABLES
Table Page
2.1 Antimicrobial test ranges, MIC quality control ranges, and MIC breakpoints used for antimicrobial susceptibility testing .………...……….179
2.2 MIC distributions of C. jejuni isolated from conventional and organic poultry farms…………………………………………….……………………….….180
2.3 MIC distributions of C. coli isolated from conventional and organic poultry farms………...... ….181
2.4 MICs of nine antimicrobial agents against C. jejuni and C. coli isolated from conventional and organic poultry farms ………………………………………182
2.5 Resistance rates of C. jejuni and C. coli isolated from conventional and organic broiler farms ……………………………………………………………..…183
2.6 Resistance rates of C. jejuni and C. coli isolated from conventional and organic turkey farms …………………………………………………...………...…184
2.7 Resistance rates of C. jejuni and C. coli isolated from conventional and organic poultry farms……………………………………………………………185
2.8 Resistance rates of Campylobacter strains isolated from different poultry production systems……………………………………...……………..……186
2.9 Multidrug resistance of Campylobacter strains isolated from conventional 30° and organic poultry farms ………………………………………………………..…187
2.10 Major multidrug resistance patterns of C. jejuni and C. coli isolated from conventional and organic poultry farms………………..……………………188
3.1 Breakpoints of the agar dilution method and the disk diffusion method used 0° to determine antimicrobial susceptibility of Campylobacter isolates ………...….…211
×
x 3.2 Antimicrobial susceptibility patterns of Campylobacter spp. identified 30° by the agar dilution method and the disk diffusion method ……………………..…212
3.3 The agreement between the agar dilution method and the disk diffusion method for antimicrobial susceptibility testing of Campylobacter (no intermediate strains were included) ………………………………………….…213
3.4 The agreement between the agar dilution method and the disk diffusion method for antimicrobial susceptibility testing of Campylobacter (intermediate strains were classified as resistant strains)……………………………214
3.5 The agreement between the agar dilution method and the disk diffusion method for antimicrobial susceptibility testing of Campylobacter (intermediate strains were classified as susceptible strains)……………………...…215
4.1 PCR primers and annealing temperature (°C)………………………………………245 …× 4.2 Campylobacter and tetracycline resistance rates on an organic broiler … farm from the first week until the end of the production cycle………………………246
4.3 Campylobacter and tetracycline resistance rates on 5 organic broiler farms….……247
4.4 Campylobacter and tetracycline resistance rates on 3 organic turkey farms…….…248
4.5 The presence of Campylobacter and tet(O) gene in environmental samples before coming in contact with the organically-raised broilers and turkeys…………249
4.6 Age of the birds when environmental and fecal samples and intestinal samples were collected from each organic poultry farm………………………....…250
4.7 Prevalence of Campylobacter and tetracycline resistance in organically-raised broilers and turkeys between our previous and recent studies……………………………………………………………………251
xi LIST OF FIGURES
Figure Page
2.1 PFGE patterns of Campylobacter strains isolated from conventional broiler and turkey farms and from organic broiler and turkey farms.……………………….189
4.1 Tetracycline resistance gene [tet(O) gene] of Campylobacter strains isolated from organically-raised broilers and turkeys.………….………………..….252
xii CHAPTER 1
LITERATURE REVIEW
1.1 Introduction
Campylobacter jejuni has been recognized as an important cause of food-borne
illness in humans since the late 1970s (Butzler, 2004; Skirrow, 1977). Campylobacter
species are gram-negative, motile, nonspore-forming, curved-rod shaped bacteria that are
approximately 0.2 to 0.5 µm wide and about 0.5 to 5 µm long (Doyle, 1990; Kaplan and
Weissfeld, 1994; Mahon and Manuselis, 2000; Penner, 1988; Rowe and Madden, 2000;
Shane and Montrose, 1985). Unlike other recognized food-borne pathogens,
Campylobacter spp. particularly C. jejuni is strictly microaerophilic. This means they require low level of oxygen for growth (Doyle, 1990; Rowe and Madden, 2000). C. jejuni
and related organisms grow best in an atmosphere containing approximately 3 to 8% O2 and 5 to 15% CO2 (Doyle, 1990; Nachamkin, 1999; Rowe and Madden, 2000). Although
most Campylobacter spp. grow at 37 °C, C. jejuni and C. coli show optimal growth at 42
°C (Blaser, 2000; Doyle, 1990; Shane and Montrose, 1985). In order to successfully
isolate Campylobacter spp., appropriate selective media and optimal incubation
conditions should be used (Forbes et al., 1998). As mentioned earlier that most
1 Campylobacter species are strictly microaerophilic, the the ideal environment for optimal recovery of Campylobacter spp. is an atmosphere containing approximately 5% O2, 10%
CO2, and 85% N2. Therefore, the selective agar plates should be incubated under this
microaerophilic condition at 37 °C or even better at 42 °C for 24 to 48 hours. Currently,
new techniques such as latex agglutination tests, enzyme immunoassays (EIA), enzyme-
linked immunosorbent assay (ELISA), DNA hybridization assays, and polymerase chain
reaction (PCR) have been developed for detection and identification of Campylobacter
species (Sahin et al., 2003c); however, these methods should be used with the traditional
culture method for the best isolation and identification results.
Most Campylobacter infections in humans usually occur as sporadic cases and
consumption of undercooked poultry and/or other foods that are cross-contaminated with
raw poultry meat during food preparation is considered to be a major risk factor of this
food-borne campylobacteriosis (Allos, 2001; Altekruse et al., 1999; Altekruse and
Tollefson, 2003; Blaser, 1997; Harris et al., 1986b; Kapperud et al., 1992; Norkrans and
Svedhem, 1982; Pearson et al., 2000; Skirrow, 1990). Generally, clinical symptoms of C.
jejuni infection including diarrhea, abdominal pain, high fever, and malaise commonly
occur within 2 – 5 days after ingestion of contaminated foods or water (Blaser, 1997;
Blaser, 2000; Butzler, 2004; Butzler and Skirrow, 1979; Matthews, 1999; Skirrow, 1990).
Since the symptoms of Campylobacter gastroenteritis are indistinguishable from those
caused by Salmonella, Shigella, or other enteric bacterial pathogens, a definitive
diagnosis of food-borne Campylobacter infection requires isolating the organism from
the stools of infected persons (Allos, 1998; Allos, 2001; Allos and Taylor, 1998; Blaser,
2 1997; Skirrow, 1990). In addition to an important cause of bacterial gastroenteritis in
humans, C. jejuni has also been associated with Guillane-Barré Syndrome (GBS), an
acute immune-mediated demyelinating disorder of the peripheral nervous system (Allos,
1997; Allos, 1998; Allos, 2001; Allos and Taylor, 1998; Blaser, 1997; Butzler, 2004;
Nachamkin et al., 1998; Nachamkin, 1999; Wassenaar and Blaser, 1999). Although most
Campylobacter infections are mild, self-limiting, and usually resolve within a few days
without any antibiotic treatment, severe or prolonged infections can also occur under
certain circumstances, in which antimicrobial therapy is usually warranted (Allos, 1998;
Allos, 2001; Allos and Blaser, 1999; Blaser, 1997; Blaser, 2000; Butzler, 2004; Skirrow,
1990). Nevertheless, supportive treatment consisting of fluid and electrolyte replacement seems to be the most important management of Campylobacter infections in humans
(Allos, 1998; Allos, 2001; Blaser, 2000; Butzler, 2004; Hoeprich et al., 1994; Perez-Perez
and Blaser, 1996). If antimicrobial therapy is indicated, erythromycin and
fluoroquinolones such as ciprofloxacin are usually considered the drugs of choice for
treatment of food-borne campylobacteriosis (Allos, 1998; Allos, 2001; Allos and Blaser,
1999; Allos and Taylor, 1998; Blaser, 1997; Blaser, 2000; Butzler, 2004; Butzler and
Skirrow, 1979; Nachamkin, 1999). Since the major risk factor of Campylobacter
infections in humans is consumption of undercooked poultry and/or other foods that are
cross-contaminated with raw poultry meat during food preparation, proper handling and
cooking of foods of animal origins particularly poultry meat as well as prevention of
cross-contamination of cooked or ready-to-eat foods with raw poultry and other meats
seem to be one of the most important actions that can help reduce the risk of food-borne
3 campylobacteriosis. Other actions that can also provide protection against the risk of
Campylobacter infections include avoiding drinking untreated water and unpasteurized
milk as well as handwashing after contact with animals (Allos, 1998; Allos, 2001; Blaser,
1986).
Although thermophilic Campylobacter is highly prevalent in poultry particularly broilers and turkeys, this organism is rarely detected in commercial broiler flocks under the age of 2 - 3 weeks old (Berndtson et al., 1996b; Engvall et al., 1986; Evans and
Sayers, 2000; Jacobs-Reitsma, 1997; Jacobs-Reitsma et al., 1995; Newell and Fearnley,
2003; Pokamunski et al., 1986; Sahin et al., 2002; Shane, 1992; Shane, 2000; Stern,
1992). Interestingly, once Campylobacter organisms are isolated from the flock, which is probably around 3 weeks of age, most of the birds in that particular flock will become colonized and environment within or around the poultry house seems to be contaminated with Campylobacter spp. (Jacobs-Reitsma, 1997; Sahin et al., 2002; Saleha et al., 1998).
Although many studies suggest that horizontal transmission from contaminated farm environment is the major mode of Campylobacter colonization in broiler flocks (Jacobs-
Reitsma et al., 1995; Newell and Fearnley, 2003; Pearson et al., 1993; Stern, 1992), several findings indicate that vertical transmission from breeders may also play a role on
Campylobacter infections in broilers (Chuma et al., 1994; Chuma et al., 1997b; Doyle,
1984; Pearson et al., 1996; Shane et al., 1986; Shanker et al., 1986). Even though the sources of poultry flock colonization are still unclear, it is unlikely that the vertical transmission will be the major source (Altekruse and Tollefson, 2003; Newell and
Wagenaar, 2000). Since the horizontal transmission seems to be the primary route for
4 introduction of Campylobacter spp. to broiler flocks, the current intervention measures are mainly focused on strict biosecurity along with other supportive measures such as
competitive exclusion or vaccination in order to control and prevent the introduction of
Campylobacter species from contaminated farm environment to the poultry flock via the
horizontal transmission (Altekruse and Tollefson, 2003; Newell and Davison, 2003;
Newell and Wagenaar, 2000; Rowe and Madden, 1999).
Over the last decade, the emergence of antimicrobial resistance has increased
dramatically and this problem is likely due to the widespread use of antimicrobial agents
in both humans and animals (Khachatourians, 1998; Threlfall et al., 2000; White et al.,
2002). In general, antimicrobial agents have been used in livestock and poultry for
treatment, control, and prevention of the diseases as well as for improving growth and
feed efficiency in animals for long period of time (Aarestrup and Wegener, 1999;
Khachatourians, 1998; McEwen and Fedorka-Cray, 2002; Threlfall et al., 2000; Van den
Bogaard and Stobberingh, 1999). Recently, a rapid increase in the proportion of
Campylobacter strains resistant to antimicrobial agents particularly to fluoroquinolones has been reported from many countries around the world (Engberg et al., 2001;
Nachamkin et al., 2000b; Smith et al., 2000). Because fluoroquinolones are effective against a wide range of enteric pathogens, this class of antibiotics is not only considered the drug of choice for treatment of food-borne campylobacteriosis, but it is also considered the drug of choice for treatment of gastroenteritis caused by other bacteria as well as for bacterial gastroenteritis that a cause of diarrheal illness has not yet been identified (Allos, 1998; Allos, 2001; Blaser, 2000; Hoeprich et al., 1994; Nachamkin,
5 1999). Thus, a high prevalence of fluoroquinolone resistance among food-borne pathogens including Campylobacter spp. is undesirable. There is a direct evidence suggested that the emergence of fluoroquinolone resistance in Campylobacter spp. could occur very rapidly after the chickens were treated with fluoroquinolones (Griggs et al.,
2005; Jacobs-Reitsma et al., 1994b; Luo et al., 2003; McDermott et al., 2002). In addition, several studies also revealed the association between the licensing of fluoroquinolones for use in food animals and the increasing numbers of fluoroquinolone resistance in Campylobacter isolates from both humans and animals (Endtz et al., 1991;
Gaunt and Piddock, 1996; Sanchez et al., 1994; Smith et al., 1999; Velazquez et al.,
1995). Since no antibiotics have been used in the organic poultry production system, the differences in antimicrobial susceptibility patterns of Campylobacter isolates from conventional and organic poultry operations should reveal the relationship between antimicrobial usage in animal agriculture and the development of antimicrobial resistance in food-borne bacterial pathogens.
Recently, organic poultry are becoming an increasingly important sector of the retail chicken market in many industrialized countries (El-Shibiny et al., 2005). In addition to the strict rules regarding the use of antimicrobial substances, the organic birds must be fed on organically-produced feed and they also need to have access to the outside environment (El-Shibiny et al., 2005). When compared to the conventional production system, the organic production system usually has significantly lower density of the birds in each flock but longer rearing cycle. The organic broiler chickens are generally raised on the farm for 8 to 12 weeks before they are sent to the processing plant, while the
6 conventional broiler chickens are commonly sent to the processing plant at 6 weeks of age. Interestingly, since there is no regulation regarding the source of the birds, all organic birds are usually obtained from conventional breeder flocks.
1.2 Campylobacter: The Organism
1.2.1 Historical Background
The pathogenic bacteria that are now known as Campylobacter were first recognized as a cause of infectious abortion in sheep during the early 1900s by McFadyean and
Stockman (Butzler, 1984). At that time, these bacteria were classified as “Vibrio fetus”
(V. fetus) because their morphology was similar to those of Vibrio species (Smith and
Taylor, 1919). Several years later the first document indicated that a “vibrio” organism could also cause enteric infection and diarrhea was reported by Jones et al. (1931). They revealed that a “vibrio” called “V. jejuni” was associated with enteritis and dysentery in calves and cattle (Jones et al., 1931). Similarly during the mid-1940s, Doyle isolated another microaerophilic “vibrio”, which he called “V. coli”, from swine with diarrhea. He proposed that this organism was the cause of swine dysentery (Doyle, 1944). However, the association between microaerophilic “vibrio” and human infections was not reported until 1946 when Levy isolated the organisms that were similar to “V. jejuni” from blood cultures of patients suffered from acute diarrheal illness during a milk-borne outbreak in
Illinois (Levy, 1946). In 1947, Vinzent et al. reported the first isolation of “V. fetus” from blood culture of a woman who had septic abortion (Butzler, 1984). During the late 1940s and 1950s, the clinical spectrum of infectious abortion due to “V. fetus” in sheep and
7 cattle was becoming better understood (Bryner et al., 1964; Butzler, 1984; Clark, 1971;
Miller et al., 1959; Plastridge et al., 1947). As a result, two different varieties of “V.
fetus”, V. fetus var. intestinalis and V. fetus var. venerealis, were identified (Butzler,
1984).
One of the most important studies of Campylobacter occurred in 1957 when
Elizabeth King found that microaerophilic “vibrio” could be distinguished into two major
groups based on their ability to grow at different temperatures (King, 1957). The first
group, which had been known as “V. fetus”, was the organisms that were able to grow at
25 and 37 °C but not at 42 °C, whereas the other group, which she called “related
vibrios”, was the organisms that could grow at 37 °C and even better at 42 °C but not at
25 °C (King, 1957). In addition, Elizabeth King also described that these “related vibrios” were identical to “V. jejuni” and “V. coli” identified by Jones et al. in 1931 and
Doyle in 1944, respectively. Since these “related vibrios” most of the time were isolated
from bloodstream of the patients with diarrhea, she suggested that these “related vibrios”
might be an important cause of acute diarrheal illness in humans even though these
organisms could not be isolated from fecal samples during that time (King, 1957). In the
early 1970s, a major progress in Campylobacter studies occurred when the technique for
culturing Campylobacter from fecal specimens were available (Butzler et al., 1973;
Dekeyser et al., 1972). During the same period of time, a major taxonomic study of
Campylobacter was also reported after Veron and Chatelain found that the biochemical characteristics of “V. fetus” and “related vibrios” were different from those of the
Vibrionaceae. As a result, a new genus, Campylobacter, was proposed (Veron and
8 Chatelain, 1973). Under this scheme, King’s “related vibrios” (“V. jejuni” and “V. coli”)
became Campylobacter jejuni (C. jejuni) and Campylobacter coli (C. coli), while V. fetus
var. intestinalis and V. fetus var. venerealis became Campylobacter fetus subsp. fetus (C. fetus subsp. fetus) and Campylobacter fetus subsp. venerealis (C. fetus subsp. venerealis), respectively (Veron and Chatelain, 1973). A significant development of Campylobacter studies occurred once again during the late 1970s when selective media for culturing
Campylobacter was developed (Skirrow, 1977). The use of antibiotic-containing media in combination with simple methods for obtaining a microaerophilic environment not only made a routine diagnosis of Campylobacter infections possible, but also opened up the study of Campylobacter on a worldwide scale (Allos and Taylor, 1998).
1.2.2 General Characteristics
Campylobacter species belong to the family Campylobacteraceae (Vandamme,
2000). The genus name Campylobacter is derived from the Greek words “campylos”, which means “curved” and “baktron”, which means “rod”. The name Campylobacter or
“curved rod” therefore describes the appearance of this organism under the microscope
(Blaser, 1986). Currently, 14 Campylobacter species are classified in the genus
Campylobacter including C. jejuni and C. coli, the major cause of food-borne
Campylobacter infection in humans, C. lari, C. fetus, C. hyointestinalis, C. upsaliensis,
C. helveticus, C. mucosalis, C. concisus, C. curvus, C. rectus, C. showae, C. sputorum, and C. gracilis (Vandamme, 2000). The main characteristic of this genus is the low level of the G + C content of the DNA ranging from 28 to 47 mol% (Parkhill et al., 2000;
9 Penner, 1988; Vandamme, 2000). In addition, Campylobacter species also have a small
genome with approximately 1,600 – 1,700 kilobases (kb) in size, which is about one-third
the size of the E. coli genome (Chang and Taylor, 1990; Nachamkin, 1997; Parkhill et al.,
2000).
Campylobacter spp. are gram-negative, nonspore-forming, curved-rod shaped
bacteria that are approximately 0.2 to 0.5 µm wide and about 0.5 to 5 µm long (Doyle,
1990; Kaplan and Weissfeld, 1994; Mahon and Manuselis, 2000; Penner, 1988; Rowe
and Madden, 2000; Shane and Montrose, 1985). In addition to curved-rod shape, other
forms of Campylobacter such as S shapes, seagull-wing shapes, spiral rods, commas, and
coccoid shapes have also been reported (Blaser, 1986; Doyle, 1990; Kaplan and
Weissfeld, 1994; Mahon and Manuselis, 2000). However, spherical or coccoid forms of
Campylobacter seem to occur only when the culture is old or exposed to the air for a long
period of time (Blaser, 1986; Doyle, 1990; Kaplan and Weissfeld, 1994; Mahon and
Manuselis, 2000). Although most Campylobacter are motile with a distinctive corkscrew darting type motility using a single polar flagellum at one or both ends (Blaser, 1986;
Penner, 1988; Rowe and Madden, 2000; Shane and Montrose, 1985), some
Campylobacter species such as C. gracilis is not motile, while other species such as C.
showae are motile by multiple flagella (Vandamme, 2000).
The colony morphology of thermophilic Campylobacter has been reported in two major types (Kaplan and Weissfeld, 1994; Shane and Montrose, 1985; Skirrow and
Benjamin, 1980). The first type of C. jejuni and C. coli colonies is flat, mucoid, or wet appearing, and usually formed large islands of growth. Spreading along the streak line or
10 swarming on the agar is also commonly observed. The other type of these C. jejuni and
C. coli colonies is round, convex, or raised, and has a discrete margin. Occasionally, the
simultaneous occurrence of both types of Campylobacter colony in one culture on the agar plate has been reported (Skirrow and Benjamin, 1980). In addition, Skirrow and
Benjamin (1980) also described that the swarming growth of Campylobacter colony seemed to be more characteristic of C. jejuni strain than C. coli strain. In general, the transparent colonies like droplets of water sprayed on the medium or the grayish translucent Campylobacter colonies are usually present on the agar plate after 18 to 24 hours of incubation (Shane and Montrose, 1985; Skirrow and Benjamin, 1980). If the incubation is continued for 24 to 48 hours, the colonies will be thickened and appear in gray or pinkish and yellowish gray or even tan or slightly pink and orange in color (Allos,
1998; Forbes et al., 1998; Kaplan and Weissfeld, 1994; Shane and Montrose, 1985;
Skirrow and Benjamin, 1980). Interestingly, a “metallic” surface sheen is frequently observed in mature cultures of C. jejuni and C. jejuni-like strains, while this feature is absent or much less evident in C. coli and C. coli-like strains (Skirrow and Benjamin,
1980).
Although almost every strain of Campylobacter is oxidase- and catalase-positive, a wide range of oxidase and catalase activity can be noticed especially among C. jejuni and C. coli strains. On average, C. jejuni-like strains were reported to be less active than
C. coli-like strains and on a few occasions these C. jejuni-like strains were described as catalase-negative Campylobacter spp. (Skirrow and Benjamin, 1980). Besides oxidase- and catalase-positive, thermophilic Campylobacter strains can also reduce fumarate to
11 succinate, reduce nitrate to nitrite as well as reduce selenite, but they cannot hydrolyze
urea or grow in the presence of 3.5% NaCl and they are acetoin- and indole-negative
(Forbes et al., 1998; Penner, 1988; Shane and Montrose, 1985; Skirrow and Benjamin,
1980; Vandamme, 2000). The ability to grow in 1% glycine is variable among species
(Forbes et al., 1998). Although C. jejuni, C. coli, and C. lari are closely related, only C.
jejuni can hydrolyze hippurate. The ability to hydrolyze hippurate helps distinguish C.
jejuni from other Campylobacter species; however, hippurate-negative C. jejuni can also
occur (Blaser, 2000; Doyle, 1990; Kaplan and Weissfeld, 1994; Nachamkin, 1999). Since
thermophilic Campylobacter strains do not ferment or oxidize carbohydrates, e.g.
glucose, their energy sources for growth are mainly obtained from either amino acids
such as aspartate and glutamate or tricarboxylic acid (TCA) cycle intermediates (Blaser,
1986; Penner, 1988; Rowe and Madden, 2000; Shane and Montrose, 1985; Skirrow and
Benjamin, 1980; Vandamme, 2000).
Unlike other recognized food-borne pathogens, Campylobacter spp. particularly
C. jejuni is strictly microaerophilic. This means they require low level of oxygen for
growth (Doyle, 1990; Rowe and Madden, 2000). C. jejuni and related organisms grow
best in an atmosphere containing approximately 3 to 8% O2 and 5 to 15% CO2 (Doyle,
1990; Nachamkin, 1999; Rowe and Madden, 2000). Although most Campylobacter grow at 37 °C, C. jejuni and C. coli show optimal growth at 42 °C (Blaser, 2000; Doyle, 1990;
Shane and Montrose, 1985). In contrast to the thermophilic Campylobacter, C. fetus grow
12 well at 25 °C (Skirrow and Benjamin, 1980). This characteristic helps differentiate C.
fetus from other Campylobacter species (Shane and Montrose, 1985; Skirrow and
Benjamin, 1980).
1.2.3 Isolation and Identification
Isolation of Campylobacter spp.
In order to isolate Campylobacter from the samples that may contain different
varieties of bacteria such as fecal samples, the selective media should be used to inhibit
the growth of more rapidly growing components of the enteric bacterial flora because
Campylobacter species multiply much more slowly than other enteric bacteria (Allos,
1998; Blaser, 2000). Since the late 1970s a number of Campylobacter selective media have been described including blood-free media such as modified charcoal cefoperazone deoxycholate agar (mCCDA), charcoal-based selective medium (CSM), and Karmali agar; semi-solid blood-free selective motility medium; and blood-containing media such as Skirrow’s medium, Butzler’s medium, Blaser’s medium, Campy-BAP medium,
Preston agar, and Campy CVA agar (Bolton and Robertson, 1982; Bolton et al., 1983;
Bolton et al., 1984; Corry et al., 1995; Corry, 2000; Forbes et al., 1998; Goossens et al.,
1989; Gun-Munro et al., 1987; Karmali et al., 1986; Kaplan and Weissfeld, 1994; Mahon and Manuselis, 2000; Nachamkin, 1999; Sahin et al., 2003c; Skirrow, 1977). These selective media usually contain combinations of antibiotics to which thermophilic
Campylobacter are intrinsically resistant but other bacteria particularly enteric microbial flora are susceptible (Corry et al., 1995; Sahin et al., 2003c). Antimicrobial agents that
13 are usually used in Campylobacter selective media include a combination of cephalosporins such as cephalothin or cefoperazone and vancomycin or bacitracin.
Although both cephalothin and cefoperazone have been used in Campylobacter selective media, cefoperazone is now more preferred because some strains of C. jejuni such as C. jejuni subsp. doylei and C. coli as well as C. fetus and C. upsaliensis are inhibited by
cephalothin (Corry et al., 1995; Nachamkin, 1999; Ng et al., 1988). Since these
antibiotics are mainly active against gram-positive bacteria, other antibiotics that active
against gram-negative rod-shaped bacteria such as polymyxin B or closely related
antibiotic, polymyxin E (colistin), are often added to the selective media (Corry et al.,
1995; Corry, 2000). Although most gram-negative bacteria are inhibited by polymyxin,
Proteus spp. seem to be resistant to this antibiotic. Because trimethoprim generally inhibits Proteus spp. as well as other gram-negative bacteria and because it does not have inhibitory effect on some strains of Campylobacter as polymyxin B or colistin, it has
been used in several selective media (Corry et al., 1995; Corry, 2000). Rifampicin is
another antibiotic that is used in Campylobacter selective media. This antibiotic is active
against both gram-positive and particularly gram-negative bacteria (Corry, 2000). Other
antimicrobials that are effective against yeasts and molds such as cycloheximide
(actidione), amphotericin B, or nystatin are also present in many Campylobacter selective
media (Corry et al., 1995; Corry, 2000). The detail formulation of each Campylobacter
selective medium has been previously summarized by Corry et al. (1995). Since each
selective medium contains different combination and amount of antibiotics, a
combination of media should be used in order to increase the success rates of
14 Campylobacter isolation (Nachamkin, 1999). In addition, since Campylobacter spp. are sensitive to oxygen, most selective media for thermophilic Campylobacter usually contain substances that help protect Campylobacter from the toxic effect of oxygen derivatives (Corry et al., 1995; Corry, 2000; Sahin et al., 2003c). The most commonly used substances for neutralizing these toxic oxygen derivatives include whole, lysed, or defibrinated blood; charcoal; a combination of ferrous sulfate, sodium metabisulfite, and sodium pyruvate; and haemin or haematin (Corry et al., 1995; Corry, 2000; Sahin et al.,
2003c).
Besides Campylobacter selective media, a number of enrichment broths and transport media have also been formulated to enhance the recovery rates of thermophilic
Campylobacter. The most widely used enrichment broths for Campylobacter spp. include
Preston broth, Doyle and Roman broth, Park and Sanders broth, Hunt and Radle broth, and Exeter broth (Bolton et al., 1982; Corry et al., 1995; Corry, 2000; Doyle and Roman,
1982). These enrichment broths seem to be very beneficial especially when low numbers of Campylobacter in the samples are expected. In addition, if the samples cannot be processed within 2 or 3 hours of collection, the transport media such as Cary-Blair or
Campy-thioglycollate broth should be used to maintain the viability of thermophilic
Campylobacter in the samples (Allos and Taylor, 1998; Forbes et al., 1998; Kaplan and
Weissfeld, 1994; Martin et al., 1983; Rubin et al., 1983; Wang et al., 1983).
Campylobacter isolation can be obtained by a direct plating method or a selective enrichment method depending on the types of sample and the numbers of Campylobacter present in those samples (Sahin et al., 2003c). For example, feces or intestinal/cecal
15 contents from chickens usually contain high numbers of Campylobacter organisms.
Therefore, these types of sample can be directly plated onto Campylobacter selective agar (Musgrove et al., 2001; Sahin et al., 2003c). On the other hand, Campylobacter species are unlikely to be present in high numbers in food or environmental samples such as water samples. Therefore, the selective enrichment method should be performed prior to plating these samples directly onto selective media (Sahin et al., 2003c). For the optimal recovery, samples should be enriched in the enrichment broth no longer than 24 hours because a prolonged enrichment can decrease the isolation rate to a level even below the recovery rate obtained from the direct plating method (Sahin et al., 2003c).
Interestingly, the selective enrichment method may not always be superior for
Campylobacter isolation especially when the samples containing high numbers of fast- growing background flora such as fecal samples are enriched. This is because the background flora may overgrow thermophilic Campylobacter during the enrichment step leading to the reduction of Campylobacter recovery rates (Sahin et al., 2003c). Because of their size and motility, Campylobacter species can be isolated by the filtration method as well (Blaser, 2000; Corry, 2000; Forbes et al., 1998). In fact, the filtration method in conjunction with a nonselective medium such as blood agar has been used to isolate
Campylobacter species from fecal samples since 1970s (Kaplan and Weissfeld, 1994;
Shane and Montrose, 1985). This method is based on the principle that Campylobacter can pass through the 0.45 µm or 0.65 µm pore-size membrane filter relatively easy during the short processing time (about 30 to 60 minutes), while other enteric bacteria cannot
(Allos and Taylor, 1998; Blaser, 2000; Corry, 2000; Forbes et al., 1998; Nachamkin,
16 1999). However, the filtration method can recover Campylobacter spp. from the samples only when those samples contain at least 105 CFU of Campylobacter microorganisms
(Corry, 2000; Nachamkin, 1999). Although this method is not quite as sensitive as
primary culturing with selective media, it is very useful especially for recovering
Campylobacter spp. such as C. upsaliensis that are susceptible to antibiotics used in the
selective media (Corry, 2000; Forbes et al., 1998; Kaplan and Weissfeld, 1994;
Nachamkin, 1999). Therefore, the filtration method should be used only to complement
the direct plating or the selective enrichment methods but not as a replacement (Corry,
2000; Nachamkin, 1999). A combination of the filtration method and the selective enrichment method has been used successfully for isolation of thermophilic
Campylobacter from water samples (Blaser and Cody, 1986; Pearson et al., 1993; Sahin et al., 2003c).
To successfully isolate Campylobacter spp. from samples, selective media and optimal incubation conditions are very crucial (Forbes et al., 1998). As mentioned earlier that most Campylobacter species are strictly microaerophilic, the ideal environment for optimal recovery of Campylobacter is the atmosphere containing approximately 5% O2,
10% CO2, and 85% N2. However, some Campylobacter species such as C.
hyointestinalis, C. mucosalis, C. concisus, C. curvus, C. rectus, and C. sputorum seem to require higher level of hydrogen (about 6% H2) for growth and for primary isolation, so
these Campylobacter species may not be recovered under the conventional
microaerophilic conditions (Mahon and Manuselis, 2000; Nachamkin, 1999). The
methods used to create the suitable microaerophilic environment for Campylobacter spp.
17 isolation have been summarized by Kaplan and Weissfeld (1994) and by Mahon and
Manuselis (2000). Since different Campylobacter species seem to have different optimal temperatures for growth, the choice of incubation temperature for isolating
Campylobacter species from samples is critical (Nachamkin, 1999). In general, most
laboratories use 42 °C as the primary incubation temperature for Campylobacter isolation
(Nachamkin, 1999). This temperature is the optimal growth temperature of thermophilic
Campylobacter including C. jejuni, C. coli, C. lari, and C. upsaliensis; however, it may
not be suitable for the growth of several Campylobacter species (Corry, 2000;
Nachamkin, 1999). For example, C. fetus, an opportunistic pathogen that is commonly
involved in extraintestinal Campylobacter infections in humans, is unlikely to grow and
most of the time is not recovered at 42 °C (Nachamkin, 1999; Shane and Montrose, 1985;
Skirrow and Benjamin, 1980). Since C. fetus grows well at 25 °C, this temperature
should be used for isolating this organism (Joklik et al., 1992; Nachamkin, 1999; Shane
and Montrose, 1985; Skirrow and Benjamin, 1980). Besides the optimal incubation
conditions that directly affect the recovery rates of Campylobacter, another thing that is
also important when trying to isolate Campylobacter spp. from samples is the
susceptibility of Campylobacter strains to antibiotics used in the selective media. Since
several Campylobacter strains may be inhibited by cephalothin, rifampin, colistin, and
polymyxin B, the isolation of Campylobacter spp. on the selective agar containing these
antimicrobial agents should be performed carefully (Allos and Taylor, 1998; Forbes et
al., 1998; Joklik et al., 1992; Nachamkin, 1999). In addition, because C. upsaliensis and
C. fetus are generally sensitive to cephalothin as well as other cephalosporin antibiotics,
18 these organisms should not be isolated on the selective media containing these types of antimicrobial agents (Forbes et al., 1998; Joklik et al., 1992; Nachamkin, 1999). Finally, in order to isolate thermophilic Campylobacter, the selective agar plates should be incubated at 37 °C or even better at 42 °C for 24 to 48 hours under a microaerophilic condition. If no suspected Campylobacter colonies are present on the plates, the incubation should be extended to 72 to 96 hours before being reported as negative; however, Campylobacter strains that cause septicemia or other extraintestinal infections such as C. fetus may require as long as 2 weeks before their growth can be detected
(Forbes et al., 1998; Nachamkin, 1999).
Identification of Campylobacter spp.
As mentioned earlier, Campylobacter colonies may appear differently depending on the media used; however, colonies that appear gray to pinkish gray in color, nonhemolytic, flat, and slightly mucoid should be suspected of being Campylobacter species (Forbes et al., 1998; Kaplan and Weissfeld, 1994; Nachamkin, 1999; Sahin et al.,
2003c). When Campylobacter spp. are subcultured onto freshly prepared moist media, spreading of the colonies along the streak line or swarming of the colonies on the agar plate is commonly noticed. As the moisture content decreases, Campylobacter colonies may become round, convex, and glistening with little spreading (Nachamkin, 1999; Sahin et al., 2003c; Shane and Montrose, 1985). Besides colony morphology, microscopic morphology can help identify Campylobacter species as well. Campylobacter spp. are gram-negative, curved-rod shaped bacteria that are approximately 0.2 to 0.5 µm wide and
19 about 0.5 to 5 µm long (Doyle, 1990; Kaplan and Weissfeld, 1994; Mahon and
Manuselis, 2000; Penner, 1988; Rowe and Madden, 2000; Shane and Montrose, 1985).
Since Campylobacter species are not stained very well and are not easily visualized with the safranin counterstain as commonly used in the standard Gram stain procedure, carbolfuchsin or 0.1% – 1% aqueous basic fuchsin should be used as the counterstain instead. However, if safranin is used, counterstaining should be extended to 2 to 3 minutes (Allos and Taylor, 1998; Kaplan and Weissfeld, 1994; Mahon and Manuselis,
2000; Nachamkin, 1999). In general, when the smears are prepared from fresh, young cultures, Campylobacter spp. may appear in different shapes such as S shapes, seagull- wing shapes, long spirals, short curves, or comma shapes. However, these organisms can also appear as coccoid or coccobacilli especially when the smears are prepared from older cultures (Blaser, 1986; Doyle, 1990; Kaplan and Weissfeld, 1994; Mahon and
Manuselis, 2000).
Although the presumptive identification of Campylobacter spp. can be made by typical colonial and typical microscopic morphology as well as the characteristic rapid darting motility as observed under phase-contrast or darkfield microscope, other phenotypic tests especially oxidase and catalse tests should also be performed to confirm the identification (Kaplan and Weissfeld, 1994; Mahon and Manuselis, 2000;
Nachamkin, 1999; Sahin et al., 2003c). For oxidase test, a heavy inoculum sometimes is required to conduct this phenotypic test (Kaplan and Weissfeld, 1994). Several biochemical tests have been described for identifying Campylobacter spp.; however, the most routinely useful tests for initial identification of thermophilic Campylobacter
20 include ability to grow at 37 or 42 °C but not at 25 °C, oxidase test, catalase test, hippurate hydrolysis test, production of hydrogen sulfide (H2S) in triple sugar iron agar butts, nitrate reduction, urease production, indoxyl acetate hydrolysis test, cephalothin sensitivity test, and nalidixic acid susceptibility test (Doyle, 1990; Forbes et al., 1998;
Kaplan and Weissfeld, 1994; Nachamkin, 1999; Sahin et al., 2003c; Shane and Montrose,
1985; Skirrow and Benjamin, 1980).
As mentioned earlier, hydrolysis of sodium hippurate is the major biochemical criterion that is commonly used to differentiate C. jejuni from other thermophilic
Campylobacter species because only C. jejuni can hydrolyze sodium hippurate and give a positive result to this test. However, some strains of C. jejuni may occasionally be hippurate-negative (Blaser, 2000; Doyle, 1990; Kaplan and Weissfeld, 1994; Nachamkin,
1999; Sahin et al., 2003c). Susceptibility to cephalothin and nalidixic acid is also another important criterion for differentiation among thermophilic Campylobacter species. In general, both C. jejuni and C. coli are resistant to cephalothin, but they both are susceptible to nalidixic acid. On the other hand, C. lari is resistant to both cephalothin and nalidixic acid, while C. upsaliensis is susceptible to both of these antimicrobial agents (Kaplan and Weissfeld, 1994; Nachamkin, 1999). Over the last decade, the emergence of fluoroquinolone resistance among C. jejuni and C. coli isolates has increased drastically (Endtz et al., 1991; Nachamkin et al., 2000). Since fluoroquinolone- resistant C. jejuni and C. coli strains are also cross-resistant to nalidixic acid, the occurrence of nalidixic acid resistance among C. jejuni and C. coli strains can directly lead to problems with identification of thermophilic Campylobacter to the species level
21 (Endtz et al., 1991; Nachamkin, 1999; Nachamkin et al., 2000). Therefore, if the susceptibility to nalidixic acid is going to be used for identification of Campylobacter species, the interpretation of this test should be conducted carefully. Recently, the phenotypic identification of thermophilic Campylobacter also can be performed with commercial kits such as API Campy (Huysmans et al., 1995; Sahin et al., 2003c). The biochemical characteristics of each Campylobacter species have been summarized by
Kaplan and Weissfeld (1994).
At present, new techniques have been developed for detection and identification of Campylobacter species. These techniques are mainly based on antigen-antibody interactions or nucleic acid (DNA) detection (Sahin et al., 2003c). The methods for rapid detection and identification of Campylobacter species include latex agglutination tests, enzyme immunoassays (EIA), enzyme-linked immunosorbent assay (ELISA), immunoblotting technique, colony blotting technique, colony lift immunoassay (CLI), immunomagnetic separation (IMS), DNA hybridization assays, and polymerase chain reaction (PCR) (Sahin et al., 2003c).
Latex agglutination tests are mainly used for rapid identification and confirmation of Campylobacter colonies (Hazeleger and Beumer, 2000; Hazeleger et al., 1992;
Hodinka and Gilligan, 1988; Kaplan and Weissfeld, 1994; Mahon and Manuselis, 2000;
Nachamkin and Barbagallo, 1990; Sahin et al., 2003c; Sutcliffe et al., 1991). These tests help identify whether the suspected colonies are Campylobacter or not, but they cannot differentiate between Campylobacter species; i.e., the latex agglutination tests can identify Campylobacter spp. at the genus level not at the species level (Hazeleger and
22 Beumer, 2000; Hodinka and Gilligan, 1988; Kaplan and Weissfeld, 1994; Mahon and
Manuselis, 2000; Nachamkin, 1999; Nachamkin and Barbagallo, 1990). Latex
agglutination tests are usually performed after the primary isolation of thermophilic
Campylobacter on selective agar plates and these tests are not intended to use for direct
detection of Campylobacter spp. from field samples (Hazeleger and Beumer, 2000;
Hazeleger et al., 1992; Mahon and Manuselis, 2000; Sahin et al., 2003c). Examples of the
commercially available latex agglutination tests include Campyslide (BBL Microbiology
Systems, Cockeysville, MD), Meritec Campy [jcl] (Meridian Diagnostics, Cincinnati,
OH), ID Campy (Integrated Diagnostics, Baltimore, MD), INDX-Campy [jcl] (PanBio
INDX, Inc., Baltimore, MD), and Microscreen Campylobacter (Mercia Diagnostics,
Guildford, UK) (Hazeleger and Beumer, 2000; Hazeleger et al., 1992; Hodinka and
Gilligan, 1988; Kaplan and Weissfeld, 1994; Mahon and Manuselis, 2000; Nachamkin,
1999; Nachamkin and Barbagallo, 1990; Sahin et al., 2003c). Most of these tests can
detect C. jejuni, C. coli, and C. lari (Hazeleger and Beumer, 2000; Hazeleger et al., 1992;
Hodinka and Gilligan, 1988; Kaplan and Weissfeld, 1994; Mahon and Manuselis, 2000;
Nachamkin, 1999; Nachamkin and Barbagallo, 1990; Sutcliffe et al., 1991), while some
of these tests can also detect C. fetus and C. upsaliensis (Hazeleger and Beumer, 2000;
Kaplan and Weissfeld, 1994; Mahon and Manuselis, 2000; Nachamkin and Barbagallo,
1990; Sutcliffe et al., 1991). More details on latex agglutination tests can be obtained
from the previously published articles by Hazeleger and Beumer (2000) and On (1996).
When compared to the latex agglutination tests, the enzyme immunoassays (EIA) seem to
have better sensitivity and specificity and they can also be used for direct detection of
23 Campylobacter spp. in field samples (Hindiyeh et al., 2000; Sahin et al., 2003c).
Although these methods provide rapid detection and identification of Campylobacter species, they are not as sensitive as the traditional culture methods; hence, they should not be used for detection of Campylobacter spp. from samples in which the organisms are suspected to be low in numbers (Sahin et al., 2003c). Examples of the commercially available EIA kits include VIDAS Campylobacter kit (bioMerieux, France), EIA-Foss
Campylobacter enzyme-linked immunosorbent assay kit (Foss Electric, Denmark), and
ProSpecT Campylobacter microplate assay (Alexon-Trend, Inc., Ramsey, MN) (Hindiyeh et al., 2000; Hoorfar et al., 1999; Sahin et al., 2003c). In addition, a commercial enzyme- linked immunosorbent assay (Alexon-Trend, Inc., Minneapolis, MN) for direct detection of Campylobacter antigen in field samples; e.g., fecal samples was also recently introduced (Nachamkin et al., 2000).
The use of nucleic acid or DNA probes for detection and identification of
Campylobacter species has increased in recent years (On, 1996). These DNA probes may be genus or species specific (On, 1996). In general, the majority of DNA probes used for hybridization assays of Campylobacter species are designed from 16 S rRNA genes of these organisms (On, 1996; Sahin et al., 2003c). The major advantage of DNA probes over conventional methods for identification of Campylobacter spp. is that
Campylobacter colonies can be identified relatively quickly and efficiently. Moreover, certain probes can also be used for direct detection of thermophilic Campylobacter from field samples (On, 1996; Sahin et al., 2003c). However, the sensitivity and specificity of
DNA probes seem to be lower than those of traditional culture methods especially when
24 the target gene sequence is mutated or absent. In addition, these methods also are costly and require specialized reagents and equipment (Olsen et al., 1995; On, 1996; Sahin et al., 2003c; Stucki et al., 1995). Generally, DNA probes are often used for culture confirmation and usually combined with culture methods to increase detection efficiency
(Nachamkin, 1999; Sahin et al., 2003c). Examples of the commercially available DNA
probes include SNAP (Syngene, San Diego, CA), DNA Probe System (Enzo
Biochemicals, New York, NY), and AccuProbe (Gen-Probe, Inc., San Diego, CA). These
DNA probes are commonly used for identification of thermophilic Campylobacter
including C. jejuni, C. coli, and C. lari (Kaplan and Weissfeld, 1994; Nachamkin, 1999;
On, 1996; Sahin et al., 2003c).
Recently, polymerase chain reaction (PCR)-based methods have been developed
for rapid detection, identification, and confirmation of Campylobacter species as well as
for typing of Campylobacter strains (Blaser, 2000; Englen et al., 2003; Sahin et al.,
2003c). When compared to DNA hybridization assays, PCR-based methods are more
commonly used than DNA probes not only for detection and identification of
Campylobacter spp. but also for detection and identification of other food-borne pathogens (Sahin et al., 2003c). One of the most important components of PCR-based method is oligonucleotide primers (PCR primers), which can be designed from either variable or conserved gene sequences of Campylobacter species. Depending on the primers used, PCR methods can be used for detection and identification of
Campylobacter genus as well as for differentiation between Campylobacter species. In general, PCR primers designed from conserved regions; for example, 16S rRNA genes or
25 23S rRNA genes are commonly used for general or genus detection, while primers designed from variable regions such as hipO gene, flaA gene, mapA gene, ceuE gene, glyA gene, cadF gene, or lpxA gene are normally used for differentiation of species or strains (Al Rashid et al., 2000; Bang et al., 2002; Cloak and Fratamico, 2002; Englen et al., 2003; Eyers et al., 1993; Fermer and Engvall, 1999; Giesendorf et al., 1992; Gonzalez et al., 1997; Hiett et al., 2002a; Houng et al., 2001; Itoh et al., 1995; Kirk and Rowe,
1994; Klena et al., 2004; Konkel et al., 1999; Kulkarni et al., 2002; Linton et al., 1997;
Logan et al., 2001; Lubeck et al., 2003; Metherell et al., 1999; O’Sullivan et al., 2000;
Oyofo et al., 1992; Perelle et al., 2004; Rasmussen et al., 1996; Sahin et al., 2003c; Stucki et al., 1995; Waage et al., 1999; Waegel and Nachamkin, 1996; Wang et al., 2002). Over the last several years, PCR methods targeting genus-specific sequences or species- specific sequences have been developed and used for detection and identification of
Campylobacter spp. in different types of samples (Bang et al., 2002; Bolton et al., 2002;
Chuma et al., 1997b; Collins et al., 2001; Denis et al., 2001; Docherty et al., 1996;
Engvall et al., 2002; Fermer and Engvall, 1999; Giesendorf et al., 1992; Hernandez et al.,
1995; Hiett et al., 2002a; Inglis and Kalischuk, 2003; Itoh et al., 1995; Josefsen et al.,
2004; Kirk and Rowe, 1994; Lawson et al., 1998; Linton et al., 1997; Lund et al., 2003;
Magistrado et al., 2001; Metherell et al., 1999; O’Sullivan et al., 2000; Oyofo et al.,
1992; Padungtod et al., 2002; Rasmussen et al., 1996; Sails et al., 2002; Sails et al., 2003;
Sahin et al., 2003c; Steinbrueckner et al., 1999; Studer et al., 1999; Vanniasinkam et al.,
1999; Waage et al., 1999; Waegel and Nachamkin, 1996; Waino et al., 2003; Waller and
Ogata, 2000; Wang et al., 2002; Winters and Slavik, 2000).
26 Since PCR methods can amplify targeted or specific genetic sequences over a billion fold within a short period of time, these methods definitely improve detection and identification of Campylobacter spp. from samples containing low numbers of
Campylobacter species (Englen et al., 2003; Sahin et al., 2003c). In addition, when species-specific primer sets are used, PCR can efficiently distinguish C. jejuni from C. coli. Therefore, it is very useful for differentiating between hippurate-negative C. jejuni and C. coli, which cannot be differentiated by phenotypic methods (Nachamkin et al.,
2000; Rautelin et al., 1999). Several specific genes such as hipO gene, flaA gene, mapA gene, ceuE gene, and glyA gene have been reliably used to distinguish between C. jejuni and C. coli by PCR methods over the last several years (Al Rashid et al., 2000; Bang et al., 2002; Cloak and Fratamico, 2002; Englen et al., 2003; Fermer and Engvall, 1999;
Gonzalez et al., 1997; Hiett et al., 2002; Houng et al., 2001; Itoh et al., 1995; Kirk and
Rowe, 1994; Linton et al., 1997; Nachamkin et al., 2000; Oyofo et al., 1992; Rasmussen et al., 1996; Rautelin et al., 1999; Stucki et al., 1995; Waage et al., 1999; Waegel and
Nachamkin, 1996; Wang et al., 2002). Although PCR method provides rapid detection as well as high sensitivity and specificity in identification of Campylobacter species, it usually works well with bacterial isolates and its performance on direct testing of field samples seems to be reduced drastically due to PCR inhibitors that may be present in some types of field samples such as in fecal samples (Sahin et al., 2003c). Another drawback of PCR method is that it is unable to discriminate between viable and non- viable Campylobacter cells, which may be essential in some epidemiological studies
(Sahin et al., 2003c).
27 A variety of PCR assays such as multiplex PCR (Cloak and Fratamico, 2002;
Denis et al., 1999; Harmon et al., 1997; Houng et al., 2001; Klena et al., 2004; Korolik et al., 2001; Wang et al., 2002; Winters and Slavik, 2000), real-time PCR (Logan et al.,
2001; Lund et al., 2004; Nogva et al., 2000; Perelle et al., 2004; Rudi et al., 2004; Sails et al., 2003), reverse transcriptase-PCR (RT-PCR) (Sails et al., 1998) have been designed and used in order to improve detection and identification of Campylobacter species
(Sahin et al., 2003c). For multiplex PCR, multiples sets of primers such as genus-specific and species-specific primers or several species-specific primers are included in one PCR reaction so that detection and identification can be simultaneously accomplished in a single test (Sahin et al., 2003c). In terms of real-time PCR, although this method can accurately measure the quantity of template DNA and consequently provide approximate numbers of organisms present in the sample, it requires special equipment and reagents that usually are more expensive than conventional PCR method (Sahin et al., 2003c).
Reverse transcriptase-PCR (RT-PCR) has been used to differentiate between viable and non-viable Campylobacter cells by amplifying mRNA, which is regarded as a marker for cell viability because it is much less stable than DNA and usually disappears quickly after cells die (Sahin et al., 2003c; Sails et al., 1998). However, this method requires pure
RNA preparations because DNA contamination in the reaction can lead to unreliable results (Sahin et al., 2003c; Sails et al., 1998). Besides different PCR assays previously mentioned, PCR can also be combined with other identification or typing techniques such as ELISA (Bolton et al., 2002; Kulkarni et al., 2002; Lawson et al., 1999; Metherell et al.,
1999; Sails et al., 2001; Sails et al., 2002; Waller and Ogata, 2000), hybridization assays;
28 e.g., Southern blot (Al Rashid et al., 2000; Collins et al., 2001; Geisendorf et al., 1993;
Konkel et al., 1999; Lawson et al., 1998; O’Sullivan et al., 2000), or reverse
hybridization assay (O’Sullivan et al., 2000; van Doorn et al., 1999), restriction fragment
length polymorphism (RFLP) (Fermer and Engvall, 1999; Nachamkin et al., 2000;
Steinhauserova et al., 2001; Steinhauserova et al., 2002), and restriction enzyme assay
(REA) (Comi et al., 1996; Engvall et al., 2002) to enhance detection efficiency (Sahin et
al., 2003c). Other molecular detection and identification methods for Campylobacter
species have been reviewed by On (1996) and Sahin et al. (2003c).
As mentioned earlier, nucleic acids (DNA)-based methods such as PCR are not
only useful for rapid detection and identification of Campylobacter species, but they are also useful for epidemiological studies of Campylobacter infections in both humans and animals (Sahin et al., 2003c; Sails et al., 1998). Several typing systems have been developed to study the epidemiology of Campylobacter infections (Nachamkin, 1999;
Nachamkin et al., 2000). These typing systems can be divided into two major groups, which are phenotypic typing methods and genotypic typing methods (Farber et al., 2001).
These methods seem to vary in complexity and ability to discriminate between
Campylobacter strains (Nachamkin, 1999; Nachamkin et al., 2000). The methods for epidemiological typing of Campylobacter species include serotyping, biotyping, phage typing, auxotyping, lectin binding, bacteriocin sensitivity testing, detection of preformed enzymes, multilocus enzyme electrophoresis (MEE) and molecular typing methods such as pulsed-field gel electrophoresis (PFGE), random amplified polymorphic DNA
(RAPD), amplified fragment length polymorphism (AFLP), PCR-restriction fragment
29 length polymorphism (PCR-RFLP), restriction endonuclease enzyme analysis (REA), ribotyping, plasmid typing, and flagellin typing (fla typing) (Farber et al., 2001; Kaplan and Weissfeld, 1994; Nachamkin, 1999; Nachamkin et al., 2000; Newell et al., 2000;
Sahin et al., 2003c; Wassenaar and Newell, 2000). Among the phenotypic typing methods, serotyping is the most frequently used method (Nachamkin, 1999). The two most widely used serotyping systems for Campylobacter species particularly for C. jejuni and C. coli are the Penner system and the Lior system (Allos and Taylor, 1998;
Nachamkin et al., 2000). The Penner system identifies soluble heat-stable (somatic, O) antigens; e.g. lipopolysaccharide, by passive or indirect hemagglutination technique, whereas the Lior system measures heat-labile (flagellar, H) antigens by slide agglutination technique (Allos and Taylor, 1998; Joklik et al., 1992 Nachamkin et al.,
2000). Recently, there are more than 90 reference serotypes defined by the Penner system and 112 serotypes defined by the Lior system (Allos and Taylor, 1998). Both systems identify more than 90% of Campylobacter species isolated from humans and nonhuman sources (Allos and Taylor, 1998; Jones et al., 1985; Patton et al., 1985). Although serotyping method is a useful tool for epidemiological studies of Campylobacter infections in both humans and animals, it is performed only in a few reference laboratories because of the time and expense needed to maintain quality antisera
(Nachamkin, 1999). In terms of genotypic typing methods, several methods have been used and each method seems to have its own advantages and disadvantages. More details
30 of molecular typing methods of Campylobacter species can be obtained from comprehensive reviews previously published by Newell et al. (2000), and Wassenaar and
Newell (2000).
1.3 Campylobacter spp. in Humans
1.3.1 Sources and Modes of Transmission
Campylobacter species particularly C. jejuni and C. coli have been recognized as important causes of bacterial gastrointestinal infections in humans since the late 1970s
(Butzler, 2004). Human campylobacteriosis is a zoonosis and is mainly a food-borne infection in which foods of animal origin particularly poultry seem to play an important role (Allos, 2001; Altekruse et al., 1999; Altekruse and Tollefson, 2003; Butzler, 2004;
Butzler and Skirrow, 1979; Skirrow, 1982). Most Campylobacter infections in humans usually occur as sporadic cases and consumption of undercooked poultry and/or other foods that are cross-contaminated with raw poultry meat during food preparation is considered to be a major risk factor of this food-borne campylobacteriosis (Allos, 2001;
Altekruse et al., 1999; Altekruse and Tollefson, 2003; Blaser, 1997; Harris et al., 1986b;
Kapperud et al., 1992; Norkrans and Svedhem, 1982; Pearson et al., 2000; Skirrow,
1990). Since Campylobacter species particularly C. jejuni are found as commensals in the intestinal tracts of warm-blooded animals especially poultry, it is not surprising that the majority of retail chicken meats will be contaminated with these organisms (Butzler and
Skirrow, 1979). Several survey studies revealed that at least 70% of retail chickens sold in the United States were contaminated with Campylobacter spp., while less than 2% of
31 pork and beef were contaminated with these organisms (Smith et al., 1999; Zhao et al.,
2001). Likewise, about 40% of poultry meats sold in Europe and Asia were contaminated
with Campylobacter spp., which was 10 times higher than that of pork meats sold in the
same area (Ono and Yamamoto, 1999; Zanetti et al., 1996). Although other risk factors
such as eating barbequed pork or sausage, drinking raw milk or untreated water, and
traveling abroad have been reported to be associated with sporadic Campylobacter
infections (Altekruse et al., 1999; Hopkins et al., 1984; Kapperud et al., 1992; Norkrans
and Svedhem, 1982; Schmid et al., 1987), a majority (50% - 70%) of Campylobacter
infections in humans in many parts of the United States, Europe, and Australia have been attributed to consumption of chickens (Allos, 2001).
Besides poultry meats, raw or unpasteurized milk is also recognized as another
important source of Campylobacter infections in humans (Skirrow, 1982). Unlike
sporadic infections, most outbreaks of food-borne campylobacteriosis are associated with
consumption of raw or improperly pasteurized milk (Allos, 2001; Friedman et al., 2000;
Robinson and Jones, 1981; Skirrow, 1982). Since Campylobacter spp. are commonly
found in cow feces, it is possible that these organisms may probably be introduced into
the milk by fecal contamination at the time of milking (Butzler and Skirrow, 1979;
Skirrow, 1990). Occasionally, Campylobacter spp. may be excreted into the milk from cows that have Campylobacter mastitis as well (Skirrow, 1990).
Another important source of outbreaks of Campylobacter gastroenteritis is improperly treated or untreated water (Friedman et al., 2000; Skirrow, 1982). Large waterborne outbreaks have occurred in municipal water system as a result of a break in
32 chlorination or when a non-chlorinated ground water supply becomes contaminated
(Friedman et al., 2000). In addition, consumption of untreated natural water from lakes, rivers, streams, or even the sea that may be contaminated with feces from wild birds or
waterfowl has also been described in several Campylobacter gastroenteritis cases
(Butzler and Skirrow, 1979; Skirrow, 1982; Skirrow, 1990).
The transmission of Campylobacter spp. from food producing animals to humans usually occurs as indirect transmission via contaminated meats and meat products
(Skirrow, 1982; Skirrow, 1990). Since food producing animals such as cattle, sheep, and
pigs commonly carry Campylobacter spp. in their intestinal tracts, it is not unusual that
carcasses of these animals will regularly become contaminated with Campylobacter spp.
at slaughter (Skirrow, 1982; Skirrow, 1990). Fortunately, the process of carcass dressing
particularly at air chilling step greatly helps reduce numbers of Campylobacter spp. on the carcass by means of surface drying (Skirrow, 1990). Thus, contamination rates of red meats at retail outlets are usually low as previously mentioned (Ono and Yamamoto,
1999; Skirrow, 1982; Skirrow, 1990; Zanetti et al., 1996; Zhao et al., 2001).
In general, people can become infected with Campylobacter spp. from contaminated meats by three different ways: (i) by handling raw meats and meat products, which is commonly reported in persons preparing the food or in inexperienced workers in processing plants; (ii) by consumption of raw or undercooked beef, hamburgers, and sausages; salted, smoked, or slightly cooked pork; as well as raw fish and shellfish; e.g. clams; (iii) by consumption of foods that are usually eaten raw or without further cooking such as salads and breads, which may become cross-
33 contaminated with raw meats in the kitchen (Skirrow, 1982; Skirrow, 1990). Since the infectious dose of Campylobacter species can be as low as a few hundred bacteria such as
500 microorganisms, the cross-contamination of other foods in the kitchen, which is
probably one of the most important routes of food-borne campylobacteriosis in humans,
should be concerned (Robinson and Jones, 1981; Skirrow, 1982; Skirrow, 1990). In
addition, it has been reported that cooking methods also have an effect on the risk of
Campylobacter infections in humans. Generally, the methods that require shorter cooking times such as fondue or barbecue seem to carry an increased risk of infections and seem
to be more often associated with Campylobacter infections than the methods that require
longer cooking times such as roasting, baking, or boiling (Skirrow, 1982; Skirrow, 1990).
Besides the foods mentioned earlier, non-meat products including mushrooms and
other vegetables can be the sources of Campylobacter infections in humans as well.
These products may be contaminated with Campylobacter spp. by several routes such as
the application of fertilizers or contaminated surface waters (Allos and Taylor, 1998).
Although there have been some reported outbreaks due to sources of raw foods such as
lettuce or salad, it has been suggested that these foods were not the original source but
rather the vehicle, which became contaminated during preparation (Jacobs-Reitsma,
2000). Unlike other food-borne pathogens including Salmonella and Staphylococcus,
Campylobacter species do not multiply in foods. This may be the reason why outbreaks
of food-borne campylobacteriosis do not occur frequently (Skirrow, 1990; Skirrow,
1997).
34 In addition to the indirect transmission, Campylobacter infections in humans can
also be acquired through direct contact with infected animals (Skirrow, 1982). Direct
transmission is mainly occupational and individuals who are regularly exposed to
infected animals seem to develop immunity to Campylobacter species (Altekruse and
Tollefson, 2003; Butzler, 2004; Skirrow, 1982; Skirrow, 1990). Veterinarians, farmers,
abattoir workers, and those engaged in meat processing are likely to be at an increased
risk of exposure to Campylobacter species (Altekruse and Tollefson, 2003; Butzler,
2004; Skirrow, 1982; Skirrow, 1990). In addition, contact with household pets, especially
the ones with diarrhea or puppies and kittens as well as pet chickens has also been
identified as risk factor for human campylobacteriosis, accounting for perhaps 5% of
Campylobacter infections in humans (Altekruse et al., 1999; Altekruse and Tollefson,
2003; Butzler, 2004; Hopkins et al., 1984; Kapperud et al., 1992; Norkrans and Svedhem,
1982; Skirrow, 1981; Skirrow, 1982; Skirrow, 1990). Since Campylobacter spp. has been
found in sand from bathing beaches, direct contact with this type of environment may be
considered as a risk for Campylobacter infections in humans as well although it is
probably infrequent. Shedding of Campylobacter spp. in the feces of wild animals, particularly wild birds seem to be an important source of such environmental contamination (Allos and Taylor, 1998).
Like other enteric pathogens, person-to-person transmission of Campylobacter
spp. can occur although it is unlikely to happen (Altekruse et al., 1999). In addition,
although transmission from infected food handlers who are symptomatic or asymptomatic
is uncommon, an outbreak resulted from eating food contaminated with Campylobacter
35 spp. by a food handler may occur (Allos and Taylor, 1998). However, if person-to-person spread does occur, it is mainly observed among young children (Butzler, 2004; Butzler
and Skirrow, 1979; Skirrow, 1991). The transmission of Campylobacter species from
small children with diarrhea to their siblings or their parents taking care of them has been
documented (Butzler and Skirrow, 1979). There have been several reports of vertical
transmission of Campylobacter species from mothers who have Campylobacter enteritis to their neonates (Butzler and Skirrow, 1979). Perinatal transmission from a mother, who is not necessarily symptomatic, may occur following exposure in utero, during passage through the birth canal, or during the first days of life (Butzler, 2004).
1.3.2 Infection and Clinical Manifestation
Infection with Campylobacter spp. does not always produce clinical symptoms.
Clinical manifestation due to Campylobacter infections can vary from asymptomatic to severe illnesses (Allos, 1998; Allos and Taylor, 1998; Blaser, 2000; Butzler and Skirrow,
1979). Although the factors responsible for this phenomenon are unclear, it appears that the numbers of Campylobacter spp. reaching the small intestinal tract and the host’s immunity seem to play important roles (Blaser, 2000). Some experimental studies have shown that ingestion of as low as 500 organisms can cause illness in some persons, while in other studies, higher numbers, e.g. 106 organisms, are required to cause clinical
symptoms. This information suggests that host susceptibility also dictates infectious dose
to some degree (Allos and Taylor, 1998; Black et al., 1988; Blaser, 2000; Matthews,
1999). In general, a dose of less than 104 organisms does not seem to cause illness very
36 frequently (Black et al., 1988; Blaser, 2000). Although the exact number of
Campylobacter cells needed to cause the disease in humans is unknown, a high dose is probably required because Campylobacter spp. are susceptible and likely to be destroyed by acid in the stomach (Blaser, 2000). However, vehicles such as milk or fatty foods that favor passage through the gastric acid barrier may allow some infections to occur at relatively low doses (Blaser, 2000).
Infection with Campylobacter spp. particularly C. jejuni produces acute inflammatory enteritis affecting both small and large intestines (Allos, 1998). Generally, jejunum and ileum are the first sites to become colonized by Campylobacter organisms and then the infection extends distally to affect terminal ileum and usually colon and rectum (Blaser, 2000; Skirrow, 1997). The signs and symptoms of Campylobacter infections suggest an invasive mechanism of the disease. Campylobacter first penetrates and colonizes the intestinal mucous layer and then invades and/or translocates through the epithelial surface to the underlying tissue (Nachamkin, 1999). Penetration and colonization of the mucous lining of the intestinal tract is an important ability of
Campylobacter to produce the disease. A polar flagellum at one or both ends in combination with a rapid motility gives Campylobacter a selective advantage in penetrating and colonizing the thick mucous layer covering the gut surface (Rowe and
Madden, 2000). The pathologic lesion of Campylobacter enteritis is a mucosal invasion characterized by ulceration of the mucosal epithelium and destruction of epithelial glands with crypt abscess formation leading to degeneration, atrophy, and loss of mucus as well as inflammatory infiltration of the lamina propria with acute and chronic inflammatory
37 cells including neutrophils, eosinophils, and mononuclear cells (Allos, 1998; Allos and
Taylor, 1998; Blaser, 2000; Perez-Perez and Blaser, 1996; Skirrow, 1997). These changes; however, are not pathognomonic lesions and are indistinguishable from those seen in ulcerative colitis, Crohn’s disease, or Salmonella, Shigella, or Yersinia infections
(Allos, 1998; Allos and Taylor, 1998; Blaser, 2000; Hoeprich et al., 1994; Skirrow,
1997). In addition to an invasive mechanism, some Campylobacter strains can also produce enterotoxin and cytotoxin. A heat-labile enterotoxin produced by some
Campylobacter strains is similar to the toxin produced by E. coli or cholera toxin, which is commonly detected from the patients with watery diarrhea. This enterotoxin causes a secretory diarrhea by stimulating adenylate cyclase activity in the intestinal mucosa and disrupting the normal ion transport in the enterocytes. Cytotoxin, verotoxin, or Shiga-like toxin is also produced by some strains of Campylobacter species. This toxin is similar to the toxins produced by Shigella species and E. coli O157:H7. The roles of these toxins in the pathogenesis of Campylobacter infections are unclear (Allos and Taylor, 1998;
Blaser, 2000; Hoeprich et al., 1994; Perez-Perez and Blaser, 1996).
The incubation period of Campylobacter infections is commonly 2 – 5 days after ingestion of contaminated foods or water, but the range can extend from 1 - 7 days and perhaps up to 10 days (Blaser, 1997; Blaser, 2000; Butzler, 2004; Butzler and Skirrow,
1979; Matthews, 1999; Skirrow, 1990). In general, the length of the incubation period seems to be reversely related to the dose ingested (Allos and Taylor, 1998; Blaser, 2000).
The typical symptom of Campylobacter infection is characterized by an acute diarrheal illness, which is indistinguishable from gastroenteritis caused by Salmonella, Shigella, or
38 other enteric bacterial pathogens (Allos, 1998; Allos, 2001; Allos and Taylor, 1998;
Blaser, 1997; Skirrow, 1990). Symptoms may range from mild gastrointestinal distress
lasting 24 hours to severe relapsing colitis lasting several weeks (Allos, 1998; Allos and
Taylor, 1998; Blaser, 1997). Although Campylobacter enteritis may persist from 1 day to a few weeks, most patients recover from illness within a week without any antibiotic treatment (Allos, 1998; Skirrow, 1990).
Clinical manifestation of human campylobacteriosis usually starts with abdominal pain; however, in some patients, an influenza-like prodrome consisting of fever, headache, myalgia, malaise, and dizziness may happen (Allos, 1998; Allos and Taylor,
1998; Butzler and Skirrow, 1979). Although the predromal symptoms commonly occur
12 to 24 hours before the onset of intestinal symptoms, these symptoms may coincide with the intestinal phase or, less often, may follow it (Blaser, 2000). The most common symptoms of Campylobacter enteritis include diarrhea, abdominal pain, fever, and malaise (Blaser, 2000). Nausea is also frequently reported among the patients with
Campylobacter infections, while vomiting is uncommon (Allos, 1998).
Diarrhea due to Campylobacter infections may range in severity from loose stools to massive watery or even bloody stools with 8 or more bowel movements within one day
(Allos, 1998; Blaser, 1997; Blaser, 2000). This acute diarrhea usually lasts for 2 to 3 days; however, it may persist for 1 week or longer particularly in patients with an acute colitis (Blaser, 2000; Butzler and Skirrow, 1979). Occasionally, acute abdominal pain may be the major and only symptom of Campylobacter infections. Since the abdominal pain is usually cramping in nature and most often occurs at the right lower quadrant of
39 the abdomen, severe abdominal pain may mimic acute appendicitis (Allos, 1998; Allos
and Taylor, 1998; Blaser, 1997; Blaser, 2000; Butzler, 2004). Other common symptoms
of Campylobacter enteritis include tenesmus and high fever (40 °C) (Blaser, 2000). Like
acute abdominal pain, high and persistent fever may be the sole manifestation of
Campylobacter infections (Blaser, 2000).
In general, gastrointestinal complications due to Campylobacter infections are
rarely occur. However, Campylobacter spp. may infect the biliary tract and lead to
cholecystitis, pancreatitis, or obstructive hepatitis (Allos, 1998; Allos and Taylor, 1998;
Blaser, 2000; Butzler, 2004). Massive gastrointestinal hemorrhage and toxic megacolon
may also occur in some severe cases (Allos, 1998; Blaser, 2000). In patients undergoing
peritoneal dialysis, Campylobacter infections may cause peritonitis although it rarely
occurs (Allos, 1998; Allos and Taylor, 1998; Butzler, 2004). Unlike gastrointestinal
complications that seem to be the result of local invasion, extraintestinal complications
such as bacteremia, meningitis, endocarditis, septic arthritis, and osteomyelitis are usually
the result of systemic spread (Allos, 1998; Allos, 2001; Blaser, 2000; Butzler, 2004).
Although bacteremia has been noted in less than 1% of patients with Campylobacter
infections, it seems to be more common in immunocompromised or very young or very
old persons (Allos, 1998; Allos, 2001; Blaser, 2000; Butzler, 2004). In addition,
immunodeficient persons also often develop prolonged, severe, and recurrent
Campylobacter infections, especially with bacteremia and other extraintestinal
manifestations (Blaser, 2000). In terms of Campylobacter infections in pregnant women,
40 although the symptoms are likely to be mind and self-limited, neonatal sepsis and death
can occur particularly if the woman is infected during her trimester (Allos, 1998; Allos
and Taylor, 1998).
The most important postinfectious complication of C. jejuni infection is Guillane-
Barré Syndrome (GBS), an acute immune-mediated demyelinating disorder of the peripheral nervous system (Allos, 1998; Allos, 2001; Allos and Taylor, 1998; Blaser,
1997; Butzler, 2004; Nachamkin et al., 1998; Nachamkin, 1999; Wassenaar and Blaser,
1999). Generally, about 20 – 40% of GBS patients are infected with C. jejuni (Allos and
Taylor, 1998). Symptoms of GBS usually occur 1 - 3 weeks after the onset of
Campylobacter enteritis (Allos, 1998; Allos and Taylor, 1998; Butzler, 2004). Although
C. jejuni infection seems to be a common trigger of GBS, the risk of developing GBS after C. jejuni infection is actually quite small (less than 1 case of GBS per 1000 cases of
C. jejuni infection) (Allos, 1997; Allos, 1998; Allos, 2001; Butzler, 2004; Nachamkin,
2002; Nachamkin et al., 2000a). In addition, the development of GBS after C. jejuni infections does not appear to be related to the severity of gastrointestinal symptoms. In fact, GBS can follow asymptomatic C. jejuni infections (Allos, 1998; Allos, 2001; Allos and Taylor, 1998). Although different C. jejuni strains may involve with GBS, 30 – 80% of C. jejuni isolates from GBS patients belong to Penner serotype O:19 (Allos, 1998;
Allos, 2001; Allos and Taylor, 1998; Nachamkin, 1999). Early symptoms of GBS generally include burning sensation and numbness that can progress to flaccid analysis
(Allos, 1997; Nachamkin et al., 1998; Nachamkin, 2002; Nachamkin et al., 2000a).
Interestingly, GBS that occurs after C. jejuni infection is usually more severe, associated
41 with an irreversible neurological damage, and may require intensive treatment with
possible long-term disability (Allos, 2001; Butzler, 2004). Because the neurological symptoms of GBS that follow C. jejuni infection typically occur 1 – 3 weeks after the onset of diarrheal illness, humoral immunopathogenic mechanisms seem to play important roles in the pathogenesis of Campylobacter-induced GBS (Allos, 1998; Allos,
2001; Butzler, 2004; Nachamkin, 2002; Nachamkin et al., 2000a). It is possible that antibodies and/or T-cells are induced by the infection and are initially directed against
Campylobacter, leading to eradication of the organism. However, because peripheral nerve glycolipids or myelin proteins (self-antigens) are similar to structures on the lipopolysaccharides of some Campylobacter strains (microbial antigens), the self- antigens; e.g. peripheral nerve cells or tissues, may be destroyed by antibodies and/or T- cells as well, leading to GBS (Allos, 1998; Allos, 2001; Butzler, 2004; Nachamkin, 2002;
Nachamkin et al., 2000a). More details on Guillane-Barré Syndrome and Campylobacter infections have been reviewed by Allos (1997), Nachamkin et al. (1998), and Nachamkin et al. (2000a).
In addition to GBS, C. jejuni infection has been implicated as a potential cause of acute motor axon neuropathy (AMAN), which is clinically indistinguishable from GBS
(Allos and Taylor, 1998) and Miller Fisher syndrome (MFS), which is another related neurological syndrome that can follow C. jejuni infection (Endtz et al., 2000; Takahashi et al., 2005). MFS is characterized by acute onset of opthalmoplegia, ataxia, and areflexia. This complication is considered a variant of GBS because some patients who have MFS can progress to GBS (Endtz et al., 2000; Takahashi et al., 2005). Another
42 postinfectious complication of C. jejuni infection is reactive arthritis, which may occur up
to several weeks after Campylobacter infections. Reactive arthritis most commonly
affects large weight-bearing joints such as the knees and the lower back. Like GBS, reactive arthritis can follow symptomatic or asymptomatic C. jejuni infection and both are thought to be autoimmune responses triggered by the infection. However, reactive arthritis can occur after Salmonella, Shigella, and Yersinia infections, while GBS is associated only with Campylobacter infections (Allos, 1998; Allos, 2001; Blaser, 2000).
When reactive arthritis occurs as a part of a triad of symptoms with inflammation of the urethra and conjunctiva, it is referred to as Reiter’s syndrome (Altekruse et al., 1998).
Other reported but rare postinfectious complications of Campylobacter infections include uveitis, hemolytic anemia, hemolytic uremic syndrome, IgA nephropathy, interstitial
nephritis, hepatitis, and encephalopathy (Allos, 1998; Allos, 2001; Allos and Taylor,
1998; Blaser, 2000).
The clinical manifestations of infections due to other Campylobacter species
overlap substantially with those of C. jejuni infections. In general, C. coli infections seem
to produce more mild diseases than C. jejuni infections (Blaser, 2000; Nachamkin, 1999).
Although C. fetus can cause intermittent diarrhea with or without nonspecific abdominal
pain, it is primarily associated with bacteremia and extraintestinal infections, especially
in patients with underlying diseases such as acquired immunodeficiency syndrome
(AIDS) (Blaser, 2000; Nachamkin, 1999). Clinical aspects of Campylobacter infections
by C. jejuni and other Campylobacter species have been summarized by Skirrow and
Blaser (2000) and Lastovica and Skirrow (2000).
43 In general, specific humoral antibodies IgG, IgM, and IgA and specific intestinal antibody IgA are usually developed after Campylobacter infections (Allos, 1998; Black et al., 1988; Black et al., 1992; Blaser, 2000; Perez-Perez and Blaser, 1996; Skirrow,
1997). In addition, serum antibodies developed in response to infection with one
Campylobacter strain are also cross-reactive with other Campylobacter strains (Allos and
Taylor, 1998). Although the specific humoral antibodies particularly IgA and IgM antibodies commonly appear within 10 days and peak within 2 to 4 weeks after the infection, they decline rapidly (Allos, 1998; Skirrow, 1997). Immunoglobulin G (IgG), on the other hand, may not peak until 3 – 4 weeks after the onset of illness; however, it can persist for several weeks or months (Blaser, 2000; Skirrow, 1997). It has been reported that repeated infections may stimulate specific IgA production, but these seem to have little effect on systemic IgG production (Allos and Taylor, 1998). In addition, high exposure to Campylobacter can also lead to the development of solid gut immunity
(Allos and Blaser, 1999). Currently, it is not known whether the antibody response helps eliminate the infection or helps protect against reinfection (Perez-Perez and Blaser,
1996). However, it has been shown that persons with elevated serum and intestinal antibody levels were likely to develop asymptomatic infection with a brief duration of pathogen excretion after they were challenged with C. jejuni (Allos and Taylor, 1998;
Black et al., 1988; Nachamkin, 1997; Perez-Perez and Blaser, 1996). Likewise, persons who regularly drink raw milk seem to have specific antibody against Campylobacter species and seem to be protected from illness when exposed to contaminated raw milk that led to illness in other persons (Allos and Taylor, 1998; Jones et al., 1981; Blaser et
44 al., 1987; Nachamkin, 1997; Perez-Perez and Blaser, 1996; Skirrow, 1982). Based on the above information, it looks like specific serum and intestinal antibodies can help protect against illness but not necessarily against Campylobacter infection colonization (Black et al., 1988; Nachamkin, 1997). Although the role of the cellular immune response in the control of Campylobacter infections is unclear, the increased incidence, severity, and duration of Campylobacter infections in HIV-infected persons suggest that cell-mediated immunity seem to play an important role as well (Allos, 1998; Allos and Taylor, 1998;
Allos and Blaser, 1999; Blaser, 2000).
1.3.3 Treatment and Control and Prevention
Treatment
The most important intervention in treatment of Campylobacter infections or any other diarrheal illnesses is maintenance of proper rehydration and electrolyte balance, which in most cases can be accomplished by encouraging oral intake of water and other fluids such as glucose and electrolyte solutions (Allos, 1998; Allos, 2001; Blaser, 2000;
Butzler, 2004; Hoeprich et al., 1994; Perez-Perez and Blaser, 1996). Although oral rehydration solutions are the best method of maintaining fluid and electrolyte balance, intravenous fluids may be needed especially in the persons who are severely dehydrated or in the very young and very old persons (Allos, 1998; Allos and Blaser, 1999; Blaser,
2000). Since most clinical symptoms of Campylobacter infections are mild, self-limiting, and usually resolve within a few days without any antimicrobial therapy, treatment with antibiotics seems to be unnecessary (Allos, 2001; Allos and Blaser, 1999; Blaser, 1997;
45 Butzler, 2004). However, in some certain circumstances such as in severe infections
(persistent high fever, bloody diarrhea, or more than eight bowel movements within a
day), prolonged illness (symptoms last longer than 1 week), or systemic infections
particularly in infants, elderly, or immunocompromised persons, antimicrobial treatment
is usually warranted (Allos, 1998; Allos, 2001; Allos and Blaser, 1999; Blaser, 1997;
Blaser, 2000; Butzler, 2004). If possible, antibiotic treatment should be initiated on the
first day of illness. Several studies have shown shortened duration of symptoms and
Campylobacter excretion in patients who received antibiotics at the onset of their illness,
while other studies in which initiation of treatment was delayed for several days after the
onset of symptoms did not show a therapeutic effect (Allos, 1998; Blaser, 2000).
In vitro, Campylobacter spp. particularly C. jejuni are susceptible to a wide
variety of antimicrobial agents including macrolides (e.g. erythromycin),
fluoroquinolones (e.g. ciprofloxacin), aminoglycosides (e.g. gentamicin), and
chloramphenicol, clindamycin and imipenem. In contrast, they are generally resistant to
cephalosporins (e.g. cephalothin), penicillin, vancomycin, bacitracin, lincomycin, and
trimethoprim. Although a majority of Campylobacter isolates are susceptible to
tetracycline, resistance to this antibiotic has been reported. For ampicillin and
trimethoprim-sulfamethoxazole, susceptibility to these antimicrobials is variable (Allos,
1998; Allos, 2001; Allos and Blaser, 1999; Allos and Taylor, 1998; Blaser, 2000; Butzler and Skirrow, 1979; Nachamkin, 1999; Skirrow, 1997). When antimicrobial therapy is indicated, erythromycin is commonly recognized as the drug of choice for treatment of
Campylobacter infections because of its efficacy, safety (low toxicity), ease of
46 administration, low cost, and relatively narrow spectrum of activity (less inhibitory effect
on fecal flora) (Allos, 1998; Allos, 2001; Allos and Blaser, 1999; Allos and Taylor, 1998;
Blaser, 1997; Blaser, 2000; Butzler, 2004; Butzler and Skirrow, 1979; Nachamkin, 1999).
Unlike other antibiotics such as fluoroquinolones or tetracyclines, erythromycin can be
used safely with children and pregnant women and the resistance rate of Campylobacter
spp. particularly C. jejuni to this antibiotic is still low when compared to other antibiotics
(Allos, 2001; Allos and Blaser, 1999; Nachamkin, 1999). In addition to its systemic effects, erythromycin seems to have a local or a contact effect throughout the bowel as well because some forms of erythromycin; e.g. erythromycin stearate, is quite stable and incompletely absorbed (Allos, 2001; Allos and Blaser, 1999). The recommended dosage of erythromycin for adults is 250 mg administered orally 4 times per day or 500 mg administered orally twice a day for 5 to 7 days. For children, the recommended dosage is
30 – 50 mg/kg/day in divided doses for 5 days (Allos, 2001; Blaser, 2000; Butzler and
Skirrow, 1979). The new macrolides such as azithromycin and clarithromycin are also effective in treatment of Campylobacter infections; however, they are more expensive and provide little or no clinical advantage over erythromycin (Allos, 1998; Allos, 2001;
Allos and Blaser, 1999). Interestingly, Campylobacter strains that are resistant to erythromycin seem to be resistant to these new extended-spectrum macrolides as well
(Allos and Blaser, 1999).
In addition to erythromycin, fluoroquinolones such as ciprofloxacin is considered the treatment of choice for Campylobacter enteritis as well (Allos, 1998; Allos, 2001;
Allos and Blaser, 1999; Allos and Taylor, 1998; Blaser, 1997, Blaser, 2000; Butzler,
47 2004; Hoeprich et al., 1994; Nachamkin, 1999). Unfortunately, the emergence of fluoroquinolones resistance that has been increased drastically over the last decade in many parts of the world has limited their effectiveness (Allos, 1998; Allos, 2001; Allos and Blaser, 1999; Allos and Taylor, 1998; Blaser, 2000; Butzler, 2004; Nachamkin,
1999). In addition, although the development of fluoroquinolne particularly ciprofloxacin resistance during therapy has been reported, fluoroquinolones have retained the advantage of being effective against a wide range of enteric pathogens (Allos, 1998;
Hoeprich et al., 1994; Nachamkin, 1999). Because symptoms of Campylobacter enteritis are clinically indistinguishable from those of gastroenteritis caused by other enteric bacteria, fluoroquinolones may still be considered the drug of choice especially when bacterial gastroenteritis is suspected and a cause of diarrheal illness has not yet been identified (Allos, 1998; Allos, 2001; Blaser, 2000). The recommended dosage of ciprofloxacin is 500 mg administered orally twice daily for 5 to 7 days (Blaser, 2000;
Hoeprich et al., 1994). Although new fluoroquinolones exhibit considerable activity against Campylobacter species, it is possible that these agents might not be effective against Campylobacter strains already resistant to ciprofloxacin (Allos and Blaser, 1999).
Other antimicrobial agents that can be used as alternative agents for treatment of
Campylobacter infections include tetracycline, clindamycin, chloramphenicol, amoxicillin plus clavulanic acid, third-generation or fourth-generation cephalosporins
(e.g. cefotaxime), imipenem, and gentamicin (Allos, 1998; Allos, 2001; Allos and Blaser,
1999; Allos and Taylor, 1998; Blaser, 1997; Blaser, 2000; Butzler and Skirrow, 1979).
Since small to moderate numbers of Campylobacter isolates are resistant to tetracycline,
48 care should be taken before prescribing this antibiotic. In addition, because tetracycline is contraindicated in children under 9 years of age, clindamycin may be used in such patients (Allos and Blaser, 1999; Blaser, 2000; Butzler and Skirrow, 1979). It has been reported that the activity of clindamycin against C. jejuni is not different from that of erythromycin (Allos, 2001; Allos and Blaser, 1999). In special circumstances, as when
Campylobacter strain has an unusual antimicrobial resistance pattern or when antibiotics need to be prescribed for a patient who is intolerant of many classes of antimicrobial agents, alternative agents such as chloramphenicol should be used; nearly all
Campylobacter strains are susceptible to this antibiotic (Allos, 2001; Allos and Blaser,
1999; Butzler and Skirrow, 1979). Although C. jejuni and C. coli may produce enzyme β-
lactamase causing penicillin and cephalosporin resistance in these Campylobacter
isolates, this enzyme has been reported to be inhibited by clavulanic acid but not by
sulbactam or tazobactam. This is why amoxicillin plus clavulanic acid (but not sulbactam
or tazobactam) appears to be generally effective against Campylobacter infections
(Blaser, 2000; Nachamkin, 1999). For bacteremia and other extraintestinal infections,
gentamicin and imipenem are the most commonly prescribed antibiotics in these
circumstances because less than 1% of Campylobacter isolates are resistant to these
antimicrobial agents (Allos, 1998; Allos and Blaser, 1999; Allos and Taylor, 1998;
Blaser, 2000; Butzler and Skirrow, 1979). However, because gentamicin is ineffective
against Campylobacter infections in the gut, oral therapy with an effective and
absorbable drug should be given as well (Allos, 1998; Allos and Blaser, 1999; Allos and
T
49
Taylor, 1998). Other alternative agents such as cefotaxime or chloramphenicol also have been indicated for treatment of bacteremia or other extraintestinal infections by
Campylobacter species (Blaser, 2000).
Systemic C. fetus infections or other severe Campylobacter infections should be
treated parenterally with erythromycin, gentamicin or other aminoglycosides, and
chloramphenicol, depending upon the type of infection. In addition, ampicillin and third-
generation cephalosporins may be listed as alternative agents as well (Blaser, 2000;
Butzler and Skirrow, 1979; Nachamkin, 1999). Unlike in Salmonella infections,
treatment with antibiotics does not prolong carriage of Campylobacter spp.; in contrast, it
helps eliminate carriage in most patients. For example, treatment with erythromycin can
eliminate carriage of C. jejuni within 72 hours (Blaser, 2000). Anti-intestinal motility
agents should not be used for Campylobacter infections because they do not only prolong duration of symptoms, but they also have been associated with fatalities (Blaser, 2000;
Smith and Blaser, 1985).
Control and Prevention
Control and prevention of Campylobacter infections in humans depend mainly on
interruption of transmission of Campylobacter from risk factor sources such as farm and
domestic animals, foods of animal origin, or contaminated environment to humans (Allos
and Taylor, 1998; Blaser, 1986). As mentioned earlier, the major risk factor of
Campylobacter infections in humans is consumption of undercooked poultry and/or other
50 foods that are cross-contaminated with raw poultry meat during food preparation.
Therefore, careful food preparation habits in the kitchen seem to be one of the most
important actions that will help reduce the risk of Campylobacter infections in humans
(Allos, 1998; Allos, 2001). In addition, basic hygiene concepts also should be applied in the kitchen and food preparation area (Skirrow, 1982). Raw animal meats and animal products should be kept separately from other foods to avoid cross-contamination of cooked or ready-to-eat foods. All foods derived from animal sources particularly poultry should be thoroughly cooked (Allos, 1998; Allos, 2001; Skirrow, 1982). Since
Campylobacter spp. are sensitive to heat (the decimal reduction time for Campylobacter spp. at 55 °C is about one minute), the organisms should not survive in food products brought to adequate cooking temperature. Thorough cooking of raw chicken and other meats; therefore, provides protection against food-borne campylobacteriosis (Doyle,
1990; Nachamkin, 1997). It is recommended that poultry should be heated to an internal temperature of 82 °C (180 °F) to kill Campylobacter organisms (Altekruse and Tollefson,
2003). So, it seems to be a good idea to use a meat thermometer when cooking meats to
ensure that temperature is adequate to kill these organisms (Allos, 1998; Allos, 2001;
Altekruse and Tollefson, 2003). Prevention of cross-contamination with raw poultry and
other meats will help reduce the risk of Campylobacter infections as well. Cutting boards
and other cooking utensils that are used in handling uncooked poultry or other meats
should be washed with hot, soapy water before being used for preparation of salads or
other foods that will be served without subsequent cooking (Allos, 1998; Allos, 2001).
Like
51
Likewise, hands should be washed and food contact surfaces in the kitchen should be
cleaned and disinfected every time after contact with raw meats and poultry (Altekruse
and Tollefson, 2003).
In addition to proper handling and cooking of foods, reduction of Campylobacter
contamination on the carcasses is also another important step that can help reduce the risk
of food-borne campylobacteriosis (Altekruse and Tollefson, 2003). It has been reported
that the microbial quality of broiler carcasses has been associated with the abattoir where
the carcasses were processed (Altekruse and Tollefson, 2003; Wedderkopp et al., 2000).
In general, the numbers of Campylobacter organisms on poultry carcasses seem to
increase during defeathering and evisceration processes, but then decrease during
scalding, washing, and chilling processes (Acuff et al., 1986; Altekruse et al., 1998;
Berrang et al., 2000a; Berrang et al., 2000b; Berrang et al., 2001a; Berrang et al., 2001b;
Whyte et al., 2001). Treatment of wash water with active chlorine, organic acids, sodium
chloride, and tri-sodium phosphate seems to be an important control step that can help
reduce carcass contamination by Campylobacter species (Altekruse et al., 1999;
Altekruse and Tollefson, 2003). In one study, the use of electrolyzed water for washing poultry carcasses resulted in a 3 log10 reduction of C. jejuni on chicken carcasses
(Altekruse and Tollefson, 2003; Park et al., 2002). Another study also revealed that 10%
oleic acid significantly reduced the numbers of Campylobacter on poultry skin (Altekruse
and Tollefson, 2003; Hinton and Ingram, 2000). Since Campylobacter spp. are sensitive
to active chlorine, the chlorination of carcass wash water definitely helps reduce the risk
52 of carcass contamination by Campylobacter species (Altekruse and Tollefson, 2003;
Yang et al., 2001). In one processing plant, use of chlorinated sprays on equipment and
working surfaces in combination with maintenance of clean working surfaces and
reduction of carcass contact surfaces resulted in significant reduction of Campylobacter
counts on packaged poultry (Altekruse et al., 1998; Altekruse et al., 1999; Mead et al.,
1995). In addition, treatment of poultry chiller water containing sodium chloride or
trisodium phosphate with an electrical current can reduce C. jejuni contamination on
poultry carcasses as well (Altekruse et al., 1998; Altekruse et al., 1999; Li et al., 1995).
Because Campylobacter species are sensitive to freezing at temperatures between – 20 °C
to – 5 °C, it is not surprising that freezing poultry carcasses to – 20 °C would reduce
Campylobacter counts by 2 log10 (Altekruse and Tollefson, 2003; Doyle, 1990; Stern et
al., 1985). Recent studies suggest that Campylobacter species are more radiation sensitive than other food-borne pathogens such as Salmonella or Listeria monocytogenes and the irradiation treatment that is effective against the latter organisms should be sufficient to kill Campylobacter spp. as well (Nachamkin, 1997). Currently, gamma irradiation and
electron beam irradiation have been successfully used for elimination of Campylobacter
spp. from poultry products; however, consumer acceptance rate for control of
Campylobacter contamination on poultry carcasses and products by the irradiation
method seems to be limited (Altekruse et al., 1998; Altekruse and Tollefson, 2003; Lewis
et al., 2002; Nachamkin, 1997; Patterson, 1995; Skirrow, 1997). Since Campylobacter
spp., particularly C. jejuni and C. coli are widespread in the intestinal tract of poultry and other food-producing animals, the reduction or elimination of colonization of poultry or
53 other food animals with Campylobacter spp. on the farm will definitely help reduce the risk of carcass contamination by Campylobacter spp. at the processing plant and in the
kitchen, which unquestionably benefits consumers. Information on control and
prevention of Campylobacter spp. at the farm level is discussed under the
“Campylobacter spp. in poultry” section.
Because several outbreaks and sporadic cases of Campylobacter infections are caused by consumption of contaminated raw or unpasteurized milk and dairy products as well as untreated surface water, prevention of these outbreaks and sporadic cases can be easily accomplished by consuming only pasteurized milk and dairy products and treated water (Allos, 1998; Allos, 2001; Allos and Taylor, 1998; Blaser, 1986; Hoeprich et al.,
1994). In addition, avoiding consumption of unpasteurized milk and dairy products and untreated water also should be emphasized to pregnant women, the elderly, immunocompromised persons, or other persons in whom Campylobacter infections may
have serious consequences (Allos, 1998; Allos, 2001). As mentioned earlier,
Campylobacter spp. are sensitive to heat and chlorine. Hence, they should be effectively
destroyed by pasteurization and chlorination of milk and water, respectively (Doyle,
1990; Jay, 2000; Nachamkin, 1997; Skirrow, 1982). Because overflow of waste from
farms and processing plants, entering rivers, has the potential to contaminate public water
supplies, special care should also be taken to prevent contamination of community drinking water supplies from those sources as well as from fecal contamination by wild birds, domestic animals, or sewage effluent (Allos and Taylor, 1998; Jacobs-Reitsma,
2000; Skirrow, 1982). The safe disposal of sewage and the protection and purification of
54 water supplies are fundamental to control of most diseases due to enteric pathogens including Campylobacter spp. (Allos and Blaser, 1999). Although seafood such as
shellfish may not be considered as an important source of Campylobacter infections in
humans, avoiding consumption of raw or uncooked shellfish should help reduce the risk
of food-borne campylobacteriosis at least in immunocompromised persons (Jacobs-
Reitsma, 2000).
Since direct contact with animals especially household pets such as puppies and
kittens as well as diarrheic animals has been identified as the risk factor for human
campylobacteriosis, people who handle pets or other animals should wash their hands
before eating or involving with any food activities (Allos, 1998; Allos, 2001; Altekruse
and Tollefson, 2003). Young children in particular should be encouraged to wash their
hands every time after playing with pets particularly before eating. Hand-washing after
contact with animals is a prudent step in prevention of zoonotic transmission of
Campylobacter spp. in household and occupational settings (Altekruse and Tollefson,
2003).
Although person-to-person transmission of Campylobacter infections is
uncommon and has not been demonstrated, persons with any acute diarrheal illness
should avoid preparation and handling of foods until their illness resolves (Allos, 1998;
Allos, 2001; Hoeprich et al., 1994). As part of good general hygiene, all persons should
wash their hands after using the bathroom, especially if they have diarrhea (Allos, 1998;
Allos, 2001). Currently, there is no proven benefit of antibiotic prophylaxis to prevent
Campylobacter infections in travelers, particularly the ones who travel to tropical
55 countries and in fact the routine use of antibiotic prophylaxis to prevent Campylobacter infections is not recommended. One thing that travelers can do to reduce the risk of
Campylobacter infections is that they should be cautioned against drinking untreated water (Allos, 1998; Allos, 2001; Allos and Blaser, 1999). In addition, since there is no vaccine for prevention of Campylobacter infections at this stage, non-specific methods for prevention as mentioned above are very important (Allos and Blaser, 1999).
1.4 Campylobacter spp. in Poultry
1.4.1 Prevalence in Poultry and Other Animals
Commercial poultry such as broilers, layers, turkeys, and ducks as well as free- living birds are considered to be natural reservoirs of thermophilic Campylobacter
(Shane, 1992; Shane, 2000). However, the prevalence of Campylobacter spp. in different species of the birds seems to be different and this prevalence also varies among countries
(Newell and Fearnley, 2003). In the United States, the prevalence of Campylobacter spp. in commercial broilers ranged from 20% to nearly 90% (Aho and Hirn, 1988; Harris et al., 1986a; Hiett et al., 2002b; Jones et al., 1991; Luangtongkum et al., 2005; Munroe et al., 1983; Saleha et al., 1998; Shane, 2000; Stern et al., 2001b), while the prevalence of this organism in other countries in North America such as Canada and in South America was around 45% to 48% and 20% to 96%, respectively (Aho and Hirn, 1988; Newell and
Fearnley, 2003; Shane, 2000). In Europe, the proportion of commercial broiler flocks colonized with Campylobacter spp. varied from 2.9% to more than 92% (Aho and Hirn,
1988; Atanassova and Ring, 1999; Berndtson et al., 1996a; Bouwknegt et al., 2004;
56 Engvall et al., 1986; Evans and Sayers, 2000; Hald et al., 2000; Heuer et al., 2001;
Humphrey et al., 1993; Jacobs-Reitsma et al., 1994a; Jacobs-Reitsma et al., 1995;
Kapperud et al., 1993; Newell and Fearnley, 2003; Pearson et al., 1996; Perko-Makela et al., 2002; Refregier-Petton et al., 2001; Saleha et al., 1998; Shane, 2000; Stern et al.,
2005; Van de Giessen et al., 1996; Wedderkopp et al., 2000; Wedderkopp et al., 2001) with the lowest flock prevalence (2.9%) observed in Finland (Perko-Makela et al., 2002).
Likewise, the variation in the prevalence of thermophilic Campylobacter among
commercial broiler flocks was also observed in other regions of the world ranging from
13.6% to 87% in Africa (Adekeye et al., 1989; Aho and Hirn, 1988; Cardinale et al.,
2004; Kazwala et al., 1993; Sackey et al., 2001), 24% to 54% in Asia (Newell and
Fearnley, 2003), and about 42% in Australia (Saleha et al., 1998; Shanker et al., 1982).
In addition to the prevalence of thermophilic Campylobacter in commercial
broiler flocks, the prevalence of Campylobacter spp. on broiler chicken carcasses also
varies among studies conducted in different countries. Several studies in the United
States revealed that 22% to 98% of retail broiler chickens sold in the U.S. were
contaminated with Campylobacter spp. (Baker et al., 1987; Berrang et al., 2001b; Harris
et al., 1986a; Saleha et al., 1998; Shane, 2000; Smith et al., 1999; Willis and Murray,
1997; Zhao et al., 2001), while 38% of broiler carcasses sold in Canada and 79% to 84%
of retail broiler chickens sold in South America were contaminated with this organism
(Aho and Hirn, 1988; Shane, 2000). Likewise, the prevalence of Campylobacter spp. on
broiler chicken carcasses sold in different European countries also varied from 14% to
88% with the lowest rates (14%) in Norway and the highest rates (88%) in the United
57 Kingdom (Aho and Hirn, 1988; Atanassova and Ring, 1999; Jorgensen et al., 2002;
Shane, 2000; Wilson, 2002; Zanetti et al., 1996). In Asia, the prevalence of
Campylobacter spp. on retail chicken meats sold in Japan, Malaysia, India, and Taiwan ranged from 46% to 100% (Ono and Yamamoto, 1999; Padungton and Kaneene, 2003;
Saleha et al., 1998; Shane, 2000; Shih, 2000), whereas 45% and 77% of broiler chicken carcasses sold in Australia and Africa were contaminated with Campylobacter spp., respectively (Padungton and Kaneene, 2003; Shanker et al., 1982).
A high prevalence of Campylobacter spp. is observed not only in broilers, but it is also observed in other poultry species. Generally, the prevalence of thermophilic
Campylobacter in commercial turkeys is quite high ranging from 50% to 100% (Acuff et
al., 1986; Luangtongkum et al., 2005; Luechtefeld and Wang, 1981; Shane, 2000; Smith
et al., 2004; Wallace et al., 1998; Wempe et al., 1983). In addition, the prevalence of this
organism on turkey carcasses was also reported in a wide range from 14% to 94% (Acuff
et al., 1986; Logue et al., 2003; Luechtefeld and Wang, 1981; Wallace et al., 1998;
Zanetti et al., 1996; Zhao et al., 2001). Likewise, commercial layers are also frequently
infected with thermophilic Campylobacter (Shane, 1992). The prevalence rates of
commercial layer flocks colonized by Campylobacter spp. ranged from 13% to 62%
(Doyle, 1984; Shane, 1992; Shane et al., 1986). A high prevalence of thermophilic
Campylobacter is also observed in commercial ducks and geese (Aydin et al., 2001;
Kasrazadeh and Genigeorgis, 1987; Tsai and Hsiang, 2004). Campylobacter spp. were
isolated from 43.5% and 92% of ducks and duck farms, respectively (Tsai and Hsiang,
2004), whereas 100% of geese carried thermophilic Campylobacter in their intestinal
58 tracts (Aydin et al., 2001). For commercial pheasant, guinea fowl, and squab (young
pigeon) farms, the isolation rates of Campylobacter organisms from these pheasants,
guinea fowl, and squabs were 25.9%, 35.7%, and 3.9%, respectively (Adekeye et al.,
1989; Atanassova and Ring, 1999; Jeffrey et al., 2001). Moreover, Adekeye et al. (1989) also reported that 66.7% of guinea fowl flocks were positive for C. jejuni (Adekeye et al.,
1989). In addition, thermophilic Campylobacter can be isolated from quails (Minakshi and Ayyagari, 1988; Shane, 1992) and ostriches (Ley et al., 2001; Stephens et al., 1998) as well.
As mentioned earlier, a wide range of free-living and migrating birds and waterfowl can serve as natural reservoirs of thermophilic Campylobacter (Shane, 1992;
Shane, 2000). Although Campylobacter spp. can be isolated from free-living birds including sparrows, pigeons, crows, blue magpies, gray starlings, blackbirds, bulbuls, eastern turtledoves, owl goldeneye, and reed bunting, the highest prevalence of
Campylobacter species was found in crows, which 34% to 89.8% of the birds were
Campylobacter positive, followed by blue magpies (20%), gray starlings (14%), bulbuls
(11%), and eastern turtledoves (2%) (Chuma et al., 2000; Fukuyama et al., 1986; Ito et al., 1988; Kapperud and Rosef, 1983; Kinjo et al., 1983; Luechtefeld et al., 1981; Shane,
1992; Yogasundram et al., 1989). Likewise, the prevalence of these Campylobacter organisms in pigeons has also been reported ranging from 4.2% to 26% (Ito et al., 1988;
Kapperud and Rosef, 1983; Kinjo et al., 1983; Luechtefeld et al., 1981; Shane, 1992). In addition to the avian species mentioned above, other wild birds as well as free-living and migratory waterfowl including passerines, Canada geese, mallards, shovelers, pintails,
59 American widgeons, green-winged teals, gadwalls, common gulls or seagulls, blacked-
headed gulls, herring gulls, puffin, dunlins, sandpipers, and common terns also harbor thermophilic Campylobacter (Broman et al., 2002; Broman et al., 2004; Craven et al.,
2000; Fallacara et al., 2001; Kapperud and Rosef, 1983; Kapperud et al., 1983; Kaneuchi
et al., 1987; Luechtefeld et al., 1980; Shane, 1992; Waldenström et al., 2002;
Yogasundram et al., 1989). The prevalence of Campylobacter spp. in Canada geese was
about 52%, while the prevalence of this organism in mallards ranged between 34% and
40% (Fallacara et al., 2001; Luechtefeld et al., 1980). For other migratory waterfowl, the
Campylobacter isolation rates were as the followings: 66% in shovelers, 50% in pintails,
42% in American widgeons, 16% in green-winged teals, and 15% in gadwalls
(Luechtefeld et al., 1980). Similarly, a high frequency of Campylobacter spp. was also
observed in shorebirds particularly in puffins. Campylobacter spp. were isolated from
51% to 78% of puffins, whereas only 5.6% of common terns were positive for
Campylobacter species (Kapperud and Rosef, 1983; Kapperud et al., 1983). Among various species of gulls, the prevalence of thermophilic Campylobacter varied from 4.2%
in herring gulls to 18.9% in common gulls and to 13.2% - 41.4% in black-headed gulls
(Broman et al., 2002; Kapperud and Rosef, 1983; Kaneuchi et al., 1987).
In terms of the prevalence of thermophilic Campylobacter in free-range and
organic poultry production systems, several studies revealed a high frequency of
Campylobacter isolation from these free-range and organic broilers and turkeys (Avrain
et al., 2003; El-Shibiny et al., 2005; Heuer et al., 2001; Kazwala et al., 1993;
Luangtongkum et al., 2005; Saleha et al., 1998). In general, the prevalence of
60 Campylobacter spp. in free-range and organic broilers ranged from 68.5% to 100%
(Avrain et al., 2003; El-Shibiny et al., 2005; Heuer et al., 2001; Kazwala et al., 1993;
Luangtongkum et al., 2005; Saleha et al., 1998), while about 87% of organic turkeys
were reported to be colonized by thermophilic Campylobacter (Luangtongkum et al.,
2005).
Among Campylobacter-positive flocks, C. jejuni was the predominant species in
poultry especially in commercial broilers. Many studies on the prevalence of
Campylobacter spp. in commercial broilers have shown that 85% to 98% of commercial
broiler flocks were colonized by C. jejuni, while about 2% - 11% and 1% - 5% were
colonized by C. coli and C. lari, respectively (Avrain et al., 2003; Berndtson et al.,
1996a; Berndtson et al., 1996b; Evans and Sayers, 2000; Hald et al., 2000; Heuer et al.,
2001; Jorgensen et al., 2002; Luangtongkum et al., 2005; Nielsen et al., 1997; Refregier-
Petton et al., 2001; Saleha et al., 1998; Wedderkopp et al., 2000). Interestingly, although
several studies (Heuer et al., 2001; Luangtongkum et al., 2005; Saleha et al., 1998)
indicated that C. jejuni was the major Campylobacter species isolated from free-range or
organic broilers with the prevalence ranging from 72% to 91%, other studies (Avrain et
al., 2003; El-Shibiny et al., 2005), on the other hand, reported that the majority (43% to
92%) of thermophilic Campylobacter colonized free-range or organic broilers were C.
coli.
Unlike commercial broilers, the prevalence of C. jejuni and C. coli in commercial turkeys varies drastically among studies. Some studies (Lee et al., 2005; Smith et al.,
2004) showed that 80% - 90% of Campylobacter strains isolated from turkeys were C.
61 coli, while other studies (Wallace et al., 1998; Van Looveren et al., 2001) found that
almost 100% of Campylobacter isolated from commercial turkeys were C. jejuni. In one
survey study, the difference in the prevalence of C. coli in turkey samples collected from
2 processing plants located in the Midwestern region of the United States was reported.
The prevalence of C. coli in commercial turkeys collected from one processing plant was
40.5%, while the prevalence of this organism in commercial turkeys collected from
another processing plant located in the same region was 14.6% (Logue et al., 2003). In
addition, the recent study on the prevalence of thermophilic Campylobacter in
commercial and organic turkeys revealed that about 46% and 54% of Campylobacter
isolates from commercial turkeys were C. jejuni and C. coli, respectively; whereas about
66% of Campylobacter isolates from organic turkeys were identified as C. jejuni and
about 34% of these isolates were identified as C. coli (Luangtongkum et al., 2005).
Among Campylobacter spp. isolates from other poultry species, C. jejuni was the
predominant Campylobacter species isolated from ducks and geese (Aydin et al., 2001;
Tsai and Hsiang, 2004). About 95% of Campylobacter isolates from ducks were
identified as C. jejuni and 5.2% of these isolates were classified as C. coli (Tsai and
Hsiang, 2004). Likewise, all Campylobacter isolates from geese were identified as C.
jejuni (Aydin et al., 2001). C. jejuni and C. coli were found at a rate of 28% and 21% in
pheasants, respectively (Atanassova and Ring, 1999). In addition, only C. jejuni was
isolated from squabs and quails (Jeffrey et al., 2001; Minakshi and Ayyagari, 1988). For
free-living and migrating birds and waterfowl, C. jejuni was the main Campylobacter
species isolated from these birds (Broman et al., 2002; Broman et al., 2004; Chuma et al.,
62 2000; Craven et al., 2000; Fallacara et al., 2001; Ito et al., 1988; Kaneuchi et al., 1987;
Kapperud and Rosef, 1983; Kapperud et al., 1983; Kinjo et al., 1983; Luechtefeld et al.,
1980; Shane, 1992; Waldenström et al., 2002; Yogasundram et al., 1989), while a few
Campylobacter isolates from pigeons and black-headed gulls were identified as C. coli
(Broman et al., 2002; Kaneuchi et al., 1987; Kinjo et al., 1983). In addition, besides C.
jejuni and C. coli, C. lari was also isolated from gulls; however, the isolation rate of this
Campylobacter species was much lower than that of C. jejuni (less than 5% versus almost
95%) (Broman et al., 2002; Kaneuchi et al., 1987).
The prevalence of Campylobacter spp. in poultry particularly in broilers has been
shown to be associated with the age of the birds at slaughter. In general, Campylobacter
infection rates in commercial broiler flocks increase when age of the birds increases
(Berndtson et al., 1996a; Berndtson et al., 1996b; Evans and Sayers, 2000; Genigeorgis et al., 1986; Lindblom et al., 1986; Luangtongkum et al., 2005; Newell and Fearnley, 2003;
Newell and Wagenaar, 2000; Northcutt et al., 2003; Sahin et al., 2002; Willis and
Murray, 1997). Colonization of broiler intestinal tract usually begins after the first or the second week, then this colonization rate increases with age and remains high or even
peaks at the market age, which is normally about 6 to 7 weeks (Genigeorgis et al., 1986;
Jacobs-Reitsma, 1997; Jacobs-Reitsma et al., 1995; Lindblom et al., 1986; Padungton and
Kaneene, 2003). A study under commercial conditions revealed that the Campylobacter
colonization rate of the birds increased from 2.3% in 10 days old birds to 81.8% in
slaughter-age birds (Genigeorgis et al., 1986). Likewise, another study in the United
Kingdom also found that 40% of commercial broiler flocks were colonized with
63 Campylobacter spp. by four weeks and up to 90% by seven weeks (Evans and Sayers,
2000). Similarly, turkeys also rapidly acquire Campylobacter infections (Kaneene and
Potter, 2003). In one study, colonization of turkey chicks began at seven days after the birds arrived at the farm and this colonization rate reached 100% within three weeks
(Wallace et al., 1998). In addition to the age of the birds at slaughter, the prevalence of
Campylobacter spp. in broilers is also affected by season (Jacobs-Reitsma, 1997; Jacobs-
Reitsma et al., 1994a; Kaneene and Potter, 2003; Kapperud et al., 1993; Newell and
Wagenaar, 2000; Patrick et al., 2004; Refregier-Petton et al., 2001; Sahin et al., 2002;
Shane, 2000; Wallace et al., 1997; Wedderkopp et al., 2001; Wedderkopp et al., 2000;
Willis and Murray, 1997). Seasonal variation in presence of Campylobacter spp. in broiler flocks or broiler carcasses has been reported by several studies. Generally, the highest Campylobacter isolation rate was observed during summer/autumn months, while the lowest rate was observed during winter months (Jacobs-Reitsma, 1997; Jacobs-
Reitsma et al., 1994a; Kaneene and Potter, 2003; Kapperud et al., 1993; Meldrum et al.,
2005; Newell and Wagenaar, 2000; Patrick et al., 2004; Refregier-Petton et al., 2001;
Sahin et al., 2002; Shane, 2000; Wallace et al., 1997; Wedderkopp et al., 2001;
Wedderkopp et al., 2000; Willis and Murray, 1997). The seasonal variation in
Campylobacter prevalence in broilers may correlate directly with temperature, relative humidity, or sun light hours (Jacobs-Reitsma et al., 1994a; Kaneene and Potter, 2003;
Patrick et al., 2004; Wallace et al., 1997; Willis and Murray, 1997). In addition, other sources of contamination within the farm environment that are temperature dependent
such
64 such as migratory birds, rodents, and insects, e.g. darkling beetles, may play some roles
on the increased Campylobacter carriage in broilers during the summer/autumn months as well (Jacobs-Reitsma, 1997; Jacobs-Reitsma et al., 1994a; Patrick et al., 2004).
In terms of the prevalence of Campylobacter spp. in other animals, thermophilic
Campylobacter have been detected in many domestic and wild animals including cattle, sheep, swine, wildlife, and pet. Campylobacter species often inhabit the bovine intestinal
tract (Altekruse and Tollefson, 2003). Although prevalence rates of Campylobacter spp. in cattle vary from less than 5% to more than 89% (Altekruse and Tollefson, 2003;
Atabay and Corry, 1998; Beach et al., 2002; Busato et al., 1999; Cabrita et al., 1992;
Giacoboni et al., 1993; Harvey et al., 2004; Nielsen, 2002; Rosef et al., 1983; Stanley et al., 1998c; Turkson et al., 1988; Wesley et al., 2000), a wide range of Campylobacter prevalence is mainly observed in beef cattle, while the prevalence of this organism in dairy cows is usually less than 40% (Atabay and Corry, 1998; Harvey et al., 2004;
Wesley et al., 2000). In addition, although cattle can be colonized by several
Campylobacter species, C. jejuni is the most common thermophilic Campylobacter species isolated from cattle (Busato et al., 1999; Harvey et al., 2004; Stanley et al.,
1998c; Wesley et al., 2000). In general, C. jejuni comprised 70% to 100% of thermophilic Campylobacter in cattle (Harvey et al., 2004; Hoofar et al., 1999; Manser and Dalziel, 1985; Munroe et al., 1983; Nielsen et al., 1997; Stanley et al., 1998c; Wesley et al., 2000) except for one study, which reported that C. jejuni comprised only 7% of the
Campylobacter isolates (Atabay and Corry, 1998). Besides C. jejuni, C. hyointestinalis is another Campylobacter species that is commonly found in cattle (Nielsen et al., 1997).
65 The prevalence of this organism in intestinal tract of cattle could range from 24% to 88%
(Grau, 1988; Nielsen et al., 1997). Risk factors that influence the prevalence of
thermophilic Campylobacter in cattle include age of the animals and season of the year
(Busato et al., 1999; Harvey et al., 2004; Stanley et al., 1998c; Wesley et al., 2000). New- born calves were reported to be Campylobacter free at birth but became colonized rapidly within a few days via horizontal transmission from farm environment (Stanley et al.,
1998c). It has been reported that high numbers of Campylobacter could be found in fecal
samples of calves by 1 – 2 months of age and the overall prevalence of thermophilic
Campylobacter in calves during the first 3 months of age was about 39% (Altekruse and
Tollefson, 2003; Busato et al., 1999; Stanley et al., 1998c). In addition, calves are more frequently infected with Campylobacter spp. than adult cattle especially in dairy herds where young animals had a higher prevalence of Campylobacter than older animals
(Altekruse and Tollefson, 2003; Nielsen 2002). Moreover, feedlot cattle are also more likely to be colonized by Campylobacter species than the cattle raised on pasture (Garcia et al., 1985; Grau, 1988; Stanley et al., 1998c). The significance of thermophilic
Campylobacter colonization in cattle relates not only to the potential for contamination of
milk at the farm or the carcass at slaughter, but also environmental contamination
especially water contamination during disposal of abattoir effluents (Stanley et al., 1998a;
Stanley et al., 1998c).
The prevalence of thermophilic Campylobacter in sheep and lambs varies significantly between studies. Although several studies reported that the prevalence of
Campylobacter species in sheep and lambs at slaughter was about 92% (Altekruse and
66 Tollefson, 2003; Jones et al., 1999; Stanley et al., 1998b), other studies showed that the
level of carriage of Campylobacter in sheep as well as the level of carcass contamination
at slaughter was very low with 2% Campylobacter isolation rate (Turkson et al., 1988).
Almost 90% of thermophilic Campylobacter isolated from sheep and lambs were identified as C. jejuni, while C. coli accounted for 8% to 10% of the isolates (Jones et al.,
1999; Stanley et al., 1998b). In addition, a low frequency of other Campylobacter species such as C. hyointestinalis and C. lari could be isolated from sheep and lambs as well
(Stanley et al., 1998b). Like poultry and cattle, seasonal variation in the numbers of thermophilic Campylobacter in the intestinal tract was also observed in sheep and lambs
(Stanley et al., 1998b). When compared to other animals, the prevalence of
Campylobacter spp. in goats is relatively low with the prevalence around 6% (Turkson et
al., 1988).
Intestinal carriers of thermophilic Campylobacter are very common in swine. The
isolation rate of Campylobacter spp. from swine generally ranged from 44% to 100%
(Manser and Dalziel, 1985; Nielsen et al., 1997; Oosterom et al., 1985; Rosef et al., 1983;
Turkson et al., 1988; Weijtens et al., 1993). Unlike other animals mentioned earlier that
were mainly colonized by C. jejuni, C. coli is the predominant Campylobacter species
that colonizes swine intestinal tract (Altekruse and Tollefson, 2003; Nielsen et al., 1997;
Rosef et al., 1983; Turkson et al., 1988; Van Looveren et al., 2001). Since most studies have reported high carriage rates (almost 100%) of C. coli among healthy swine, this information provides evidence that C. coli may be a normal component of the intestinal
flora
67 flora of this animal (Rosef et al., 1983; Sticht-Groh, 1982; Van Looveren et al., 2001).
However, other Campylobacter species such as C. jejuni can be isolated from swine as
well (Turkson et al., 1988).
As mentioned earlier, Campylobacter species are naturally found in domestic animals and pets (Svedhem and Kaijser, 1981). Since Campylobacter carriage in healthy pets is very common, it is not surprising that household pets especially puppies will be responsible for the transmission of these organisms to humans (Blaser et al., 1978).
Several studies revealed that the prevalence of thermophilic Campylobacter in dogs ranged from 4.6% to 76.2% (Baker et al., 1999; Bruce et al., 1980; Burnens et al., 1992;
Gondrosen et al., 1985; Hald and Madsen, 1997; Hald et al., 2004a; Olson and Sandstedt,
1987; Sandberg et al., 2002; Steinhauserova et al., 2000; Torre and Tello, 1993; Wright,
1982), while the prevalence of this organism in cats varied from 5% to 45% (Baker et al.,
1999; Bruce et al., 1980; Burnens et al., 1992; Gondrosen et al., 1985; Hald and Madsen,
1997; Sandberg et al., 2002; Steinhauserova et al., 2000). Interestingly, the species
distribution of Campylobacter isolates from household pets especially dogs differs considerably between studies; however, C. jejuni and C. upsaliensis are the two major species that account for more than 95% of thermophilic Campylobacter isolated from these animals (Burnens et al., 1992; Hald and Madsen, 1997; Hald et al., 2004a; Olson and Sandstedt, 1987; Sandberg et al., 2002). Several studies reported that the ratios of C. upsaliensis to C. jejuni can be either high; for example, 80% to 19% or low; e.g., 15% to
82% (Baker et al., 1999; Burnens et al., 1992; Hald and Madsen, 1997; Hald et al.,
2004a; Sandberg et al., 2002). Unlike C. jejuni, C. upsaliensis has been reported to occur
68 only in dogs and cats (Hald and Madsen, 1997; Hald et al., 2004a; On, 1996). Other
Campylobacter species that can be isolated from household pets include C. coli and C. lari (Hald and Madsen, 1997; Hald et al., 2004a). In addition, Hald et al. (2004a) revealed that the prevalence of thermophilic Campylobacter in dogs increased from 60% at 3 months of age to nearly 100% at 1 year of age, then the carrier rate decreased to 67% at 2 years of age. Moreover, Hald et al. (2004a) also reported that C. jejuni was isolated more frequently in dogs under 1 year of age than in the older dogs. This finding may help explain the higher prevalence of C. jejuni than that of C. upsaliensis (76% versus 19%) observed in puppies (Hald and Madsen, 1997).
Wildlife and zoo animals including bobcat, cheetah, capybaras, coatimundis, hares, polar bear, red panda, tapir, llama, reindeer, roan antelope, wallaroos, wolves, and several species of primates have been documented to carry thermophilic Campylobacter
(Luechtefeld et al., 1981; Rosef et al., 1983). However, the isolation rate of
Campylobacter spp. from these animals is quite low with the prevalence less than 10% and C. jejuni is considered the main organism isolated from these animals (Luechtefeld et al., 1981; Rosef et al., 1983). In addition to wildlife and zoo animals, Campylobacter spp. can also be isolated from laboratory and wild rodents such as rats and bank voles, but not from rabbits, laboratory mice, hamsters, guinea-pigs, field mice or field voles (Fernie and
Park, 1977).
69 1.4.2 Sources of Poultry Flock Colonization
In general, Campylobacter species are rarely detected in commercial broiler
flocks under the age of 2 - 3 weeks old (Berndtson et al., 1996b; Engvall et al., 1986;
Evans and Sayers, 2000; Jacobs-Reitsma, 1997; Jacobs-Reitsma et al., 1995; Newell and
Fearnley, 2003; Pokamunski et al., 1986; Sahin et al., 2002; Stern, 1992; Stern et al.,
1998). The explanation for this phenomenon is unclear; however, it is possible that the presence of Campylobacter-specific maternal antibodies in young chicks (Jacobs-
Reitsma, 1997; Newell and Fearnley, 2003; Rice et al., 1997; Sahin et al., 2001; Sahin et al., 2002; Sahin et al., 2003b; Stern et al., 1998) or the presence of unique microbial flora in the intestinal tract especially in the cecum of these young chicks (competitive cecal microflora) (Humphrey et al., 1989; Sahin et al., 2002) may have an inhibitory effect or play a role on Campylobacter colonization during the first 2 weeks of life of these commercial broilers. Interestingly, once Campylobacter spp. are introduced into the flock, they tend to spread very rapidly and once these organisms are isolated from the flock, which is probably around 3 weeks of age, most or even all of the birds in that particular flock will become colonized and all environmental sources will be contaminated with Campylobacter spp. (Jacobs-Reitsma, 1997; Sahin et al., 2002; Saleha et al., 1998). In addition, Campylobacter isolation rates are also likely to increase and remain high until the birds are sent to the processing plant (Jacobs-Reitsma, 1997). Many studies suggest that horizontal transmission from environmental sources is the major mode of Campylobacter colonization in broiler flocks (Jacobs-Reitsma et al., 1995;
Newell and Fearnley, 2003; Pearson et al., 1993; Stern, 1992); however, several findings
70 suggest that vertical transmission from breeders may also play a role in Campylobacter infection in broilers (Chuma et al., 1994; Chuma et al., 1997b; Doyle, 1984; Pearson et al., 1996; Shane et al., 1986; Shanker et al., 1986).
Vertical transmission. Several studies have shown that Campylobacter spp. can be isolated from the intestinal tracts and reproductive tracts of healthy laying hens and broiler breeder hens (Buhr et al., 2002; Camarda et al., 2000; Cox et al., 2002a; Jacobs-
Reitsma, 1995; Pearson et al., 1996; Sahin et al., 2003a; Shanker et al., 1986) as well as from semen of broiler breeder roosters (Cox et al., 2002b; Hiett et al., 2003). In addition, some molecular studies also demonstrated that the genotypes of Campylobacter strains isolated from breeder flocks were similar to those of the isolates from their progeny flocks (Cox et al., 2002a; Pearson et al., 1996). This information in combination with the positive detection of Campylobacter DNA from cecal contents of newly hatched chicks
(Chuma et al., 1994; Chuma et al., 1997b) suggest that vertical transmission of
Campylobacter spp. from breeder flocks to broiler flocks through the egg may occur and this vertical transmission might be the source of Campylobacter colonization in broiler chickens (Jacobs-Reitsma, 1997; Newell and Fearnley, 2003; Sahin et al., 2002; Saleha et al., 1998). Although the findings mentioned above seem to support a vertical transmission, other studies suggest that vertical transmission of Campylobacter spp. via the egg is considered unlikely, mainly because of the difficulty or inability to culture
Campylobacter spp. from naturally or experimentally infected eggs as well as from newly hatched chicks originating from infected breeder flocks (Acuff et al., 1982; Baker et al.,
1987; Doyle, 1984; Hiett et al., 2002a; Jacobs-Reitsma, 1995; Jacobs-Reitsma et al.,
71 1995; Jacobs-Reitsma, 1997; Jones et al., 1991; Newell and Fearnley, 2003; Sahin et al.,
2002; Sahin et al., 2003a; Saleha et al., 1998; Shane et al., 1986; Shanker et al., 1986).
Even though the detection of Campylobacter DNA in eggs, embryos, and cecal contents
of newly hatched chicks has been shown in several investigations, none of these studies has been able to detect any live Campylobacter organisms from those samples (Chuma et al., 1994; Chuma et al., 1997b; Hiett et al., 2002a). In addition, several researchers also showed that isolation of Campylobacter species from broiler flocks before 2 or 3 weeks of age was hardly accomplished, even though the chicks were hatched from eggs obtained from infected parent flocks (Berndtson et al., 1996a; Jacobs-Reitsma, 1997;
Newell and Fearnley, 2003; Sahin et al., 2002; Shanker et al., 1986; Van de Giessen et al., 1992). Moreover, broilers from the same parent flocks were also found to be colonized in one production cycle but Campylobacter-free in another cycle (Jacobs-
Reitsma, 1995; Jacobs-Reitsma et al., 1995; Jacobs-Reitsma, 1997; Newell and Fearnley,
2003; Sahin et al., 2002). When Campylobacter strains from breeder flocks and broiler
flocks were investigated, several studies revealed that Campylobacter strains infected
broiler flocks were frequently found to be different from those infected breeder flocks
(Chuma et al., 1997a; Petersen et al., 2001b; Sahin et al., 2002). In addition,
Campylobacter strains isolated from different broiler flocks originating from the same
breeder flocks did not always have the same serotypes, while Campylobacter strains
isolated from broilers originating from different hatcheries but were raised in the same
farm may be colonized with the same Campylobacter strains, indicating that the serotype
pattern was more associated with the farm than with the origins of the birds (Berndtson et
72 al., 1996b; Jacobs-Reitsma, 1997; Newell and Fearnley, 2003; Petersen and Wedderkopp,
2001; Sahin et al., 2002). Based on all information available currently, it is possible that
vertical transmission of Campylobacter spp. through the egg may occur; however, it is
probably a rare event and does not seem to play an important role in the introduction of
Campylobacter species to broiler flocks (Sahin et al., 2002; Sahin et al., 2003a).
Horizontal transmission. Many studies have suggested that horizontal
transmission of Campylobacter spp. from the rearing environment is likely to be the
major source of Campylobacter infection in broiler flocks (Newell and Fearnley, 2003;
Sahin et al., 2002; Saleha et al., 1998). Although many environmental sources including
feed, litter, untreated water, insects, flies, rodents, other farm animals, domestic pets,
wildlife species particularly wild birds, equipment and transport vehicles, and farm
workers have been suspected to be the source of Campylobacter infection in broiler
farms, none of these suspected sources has been identified conclusively as the actual source of infection and mode of transmission of this organism to broilers in the farms has been unclear (Sahin et al., 2002; Saleha et al., 1998). In general, Campylobacter spp. were usually detected from environmental sources after the broilers had become infected, suggesting that the birds might be the source of environmental contamination instead of being infected from environmental sources (Berndtson et al., 1996a; Jacobs-Reitsma et al., 1995; Kazwala et al., 1990; Sahin et al., 2002; Stern et al., 2001b).
As mentioned earlier, Campylobacter spp. usually have not been isolated from fresh litter and feed samples before the flock had been colonized with Campylobacter
organisms (Gregory et al., 1997; Humphrey et al., 1993; Jacobs-Reitsma et al., 1995;
73 Pearson et al., 1993; Sahin et al., 2002). Because of the low water content of feed and
litter, Campylobacter spp. are unlikely to survive for a long period of time in these
environmental sources (Berndtson et al., 1996b; Evans, 1992; Jacobs-Reitsma, 1997;
Jacobs-Reitsma et al., 1995; Kaneene and Potter, 2003; Newell and Fearnley, 2003;
Pokamunski et al., 1986; Sahin et al., 2002; Shane, 2000). Therefore, feed and litter are
not regarded as the major sources of Campylobacter infection in broiler flocks (Newell
and Fearnley, 2003; Sahin et al., 2002; Shane, 2000). Although used litter that became
contaminated with Campylobacter spp. may play a role in maintaining Campylobacter
spp. in the farm environment and transmitting this organism to other broiler flocks
(Montrose et al., 1985; Shane, 1992), a study by Payne et al. (1999) showed that the role
of used litter in the transmission of Campylobacter species to successive flocks in the same poultry house was insignificant.
Several studies indicated that the use of untreated water such as groundwater or well water seemed to be an important risk factor for Campylobacter colonization in broiler flocks (Jacobs-Reitsma, 1997; Kapperud et al., 1993; Pearson et al., 1993). This is mainly because untreated water could be comtaminated with Campylobacter species from other environmental sources, particularly from livestock or wild birds (Jones, 2001;
Kaneene and Potter, 2003; Sahin et al., 2002; Stanley et al., 1998a). Contaminated or non-chlorinated water supplied to broilers has been demonstrated or implicated as a source of Campylobacter infection as well as a vehicle for transmission of
Campylobacter spp. to broiler flocks in several countries (Engvall et al., 1986; Kaneene and Potter, 2003; Kapperud et al., 1993; Pearson et al., 1993; Sahin et al., 2002; Shane,
74 2000). In general, fresh tap water from broiler farms was not found to be contaminated with Campylobacter spp. (Jacobs-Reitsma, 1997). However, water samples from the farms, particularly the ones collected from bell-shaped drinkers inside the broiler houses, could become Campylobacter positive especially after or at the same time that broilers in the houses were colonized by Campylobacter species (Berndtson et al., 1996a; Jacobs-
Reitsma, 1997; Jacobs-Reitsma et al., 1995; Newell and Fearnley, 2003). This means water contamination usually follows rather than precedes Campylobacter colonization of a flock (Engvall et al., 1986; Kazwala et al., 1990; Linblom et al., 1986; Newell and
Fearnley, 2003). In addition, the occurrence of viable but non-culturable form of
Campylobacter species in water may be significant in introducing infection into broiler flocks as well (Newell and Fearnley, 2003; Shane, 2000). Since contaminated water can introduce Campylobacter infections into broiler houses and lead to subsequent dissemination of this organism within the flocks, water provided for the birds should be chlorinated and water supply system should be cleaned thoroughly and disinfected after the previous flock has been removed and before the new flock is introduced to the house
(Engvall et al., 1986; Kapperud et al., 1993; Pearson et al., 1993; Saleha et al., 1998;
Shane, 1992; Shanker et al., 1990; Smitherman et al., 1984).
Insects including flies (e.g. house flies, filth flies), darkling beetles, cockroaches, and mealworms in and around broiler houses can serve as mechanical vectors and may transmit or carry Campylobacter spp. from one location to another within or between the houses or farms as well as between successive broiler flocks (Bates et al., 2004;
Berndtson et al., 1996a; Gregory et al., 1997; Hald et al., 2004b; Jacobs-Reitsma, 1997;
75 Jacobs-Reitsma et al., 1995; Kaneene and Potter, 2003; Newell and Fearnley, 2003;
Rosef and Kapperud, 1983; Sahin et al., 2002; Shane, 2000; Shane et al., 1985; Skove et al., 2004; Szalanski et al., 2004). Interestingly, serotypes and genotypes of
Campylobacter strains isolated from insects and broilers within the same broiler house were found to be identical; therefore, it was possible that an infection route from insects to broilers might occur (Berndtson et al., 1996a; Jacobs-Reitsma, 1997; Jacobs-Reitsma et al., 1995; Rosef et al., 1985; Sahin et al., 2002; Stern et al., 1997). However, since insects in broiler houses were usually not positive for Campylobacter spp. until the organism was isolated from the broilers (Berndtson et al., 1996a; Jacobs-Reitsma, 1997;
Jacobs-Reitsma et al., 1995; Nesbit et al., 2001; Sahin et al., 2002), this finding suggested that an infection route was likely to occur from broilers to insects rather than from insects to broilers and the possibility that insects were an original source of Campylobacter infection in broiler flocks seemed to be very small (Sahin et al., 2002).
Although the presence of rodents such as rats and mice on a farm can lead to an increased risk of flock colonization with Campylobacter spp., this risk factor is considered unlikely to be significant especially for the farm that has an effective vermin control program (Berndtson et al., 1996b; Kapperud et al., 1993; Newell and Fearnley,
2003; Sahin et al., 2002). In addition, several studies also reported that they could not isolate Campylobacter species from trapped rats and mice on broiler farms with infected flocks (Gregory et al., 1997; Jones et al., 1991; Shane, 2000). Therefore, the role of rodents as a common source of Campylobacter infection in broiler flocks seems to be questionable (Evans and Sayers, 2000; Gregory et al., 1997; Sahin et al., 2002).
76 In addition to rodents, Campylobacter spp. can also be recovered from other wild animals including hares, raccoons, badgers, foxes, and deer (Newell and Fearnley, 2003;
Sahin et al., 2002). And as mentioned earlier, a wide range of free-living and migrating birds and waterfowl can also serve as natural reservoirs of thermophilic Campylobacter
(Broman et al., 2002; Broman et al., 2004; Craven et al., 2000; Fallacara et al., 2001;
Kapperud and Rosef, 1983; Kapperud et al., 1983; Kaneuchi et al., 1987; Luechtefeld et al., 1980; Shane, 1992; Shane, 2000; Waldenström et al., 2002; Yogasundram et al.,
1989). Therefore, it is not surprising that Campylobacter spp. are frequently isolated from feces of wild birds around broiler houses (Newell and Fearnley, 2003). Although
Campylobacter strains isolated from wild birds around broiler houses were usually found to be different from those of broiler chickens, some epidemiological studies reported that these Campylobacter strains could occasionally be recovered from the intestinal tracts of of broilers in those houses (Gregory et al., 1997; Hiett et al., 2002b; Nesbit et al., 2001;
Newell and Fearnley, 2003; Petersen et al., 2001a; Rosef et al., 1985; Sahin et al., 2002;
Stern et al., 1997). Since wild animals, particularly wild birds seem to have a high carriage rate of Campylobacter species in their intestines, they should therefore be considered as a potential source of Campylobacter infection for broiler flocks as well
(Craven et al., 2000; Sahin et al., 2002).
The presence of domestic livestock including cattle, sheep, and pigs as well as domestic pets such as dogs and cats on broiler farms has been found to be associated with an increased risk of Campylobacter infection in broiler flocks especially in the farms that do not have good or appropriate biosecurity procedures (Berndtson et al., 1996b; Gregory
77 et al., 1997; Kapperud et al., 1993; Newell and Fearnley, 2003; Rosef et al., 1985; Sahin
et al., 2002; Shane, 2000; Stern et al., 1997; Van de Giessen et al., 1992; Van de Giessen et al., 1998). Farm animals can excrete substantial numbers of Campylobacter spp. in
their feces and this can result in contamination of boots, other external clothing, and
equipment used in the farm, which can lead to contamination of broiler flocks (Newell and Fearnley, 2003). Hence, it is not surprising that a mode of transmission of
Campylobacter organisms from farm animals to broilers will be frequently associated with indirect mechanical transmission through farm personnel or farm equipment (Shane,
2000). In addition, farm animals especially cattle and sheep also have potential to contaminate pastures and surface waters, which in turn may act as a source of broiler flock colonization (Jones et al., 1999; Sahin et al., 2002; Stanley et al., 1998b; Stanley et al., 1998c). Like other environmental sources, some studies found that Campylobacter
strains isolated from farm animals were similar to the isolates from broilers, while other
studies repored that Campylobacter strains isolated from farm animals were different from those isolates recoverd from broilers raised on the same farm (Gregory et al., 1997;
Jacobs-Reitsma, 1997; Jacobs-Reitsma et al., 1995; Nesbit et al., 2001; Petersen et al.,
2001a; Rosef et al., 1985; Stern et al., 1997; Van de Giessen et al., 1998). Likewise,
several studies indicated the association between Campylobacter strains colonized broiler flocks and the presence of pigs on the same farm, whereas other studies did not observe this association (Jacobs-Reitsma, 1997; Jacobs-Reitsma et al., 1994; Jacobs-Reitsma et al., 1995; Kapperud et al., 1993; Rosef et al., 1985; Stern et al., 1997; Van de Giessen et
al., 1992; Van de Giessen et al., 1998). However, since C. coli is mainly associated with
78 pigs, an increased proportion of C. coli in broiler flocks might indicate the transmission of these Campylobacter organisms from pigs to broilers (Jacobs-Reitsma, 1997).
Although other domestic animals such as horses can also be infected with Campylobacter
spp. (Stern, 1992), the potential role of these animals as a source of broiler flock infection
has not yet been established (Sahin et al., 2002).
Since healthy carriers of Campylobacter spp. in humans are unlikely, farm
workers are usually not regarded as direct sources of transmission (Jacobs-Reitsma,
1997). However, farm workers may carry and spread Campylobacter spp. from one flock to another by indirect mechanical transmission via boots/footwear, clothing, or
equipment, which seems to occur easily (Berndtson et al., 1996b; Jacobs-Reitsma, 1997;
Kazwala et al., 1990; Lindblom et al., 1986; Sahin et al., 2002; Saleha et al., 1998).
Furthermore, farm workers taking care of broilers or other animals or loading birds for
transport to slaughter can also increase the risk of Campylobacter infection in broiler
flock especially if they enter the broiler house without changing clothes or boots
(Berndtson et al., 1996b; Kapperud et al., 1993; Sahin et al., 2002; Saleha et al., 1998).
Since Campylobacter spp. have been isolated from footbath water, boots, and transport
crates (Hiett et al., 2002b; Jacobs-Reitsma, 1997; Kazwala et al., 1990; Newell et al.,
2001; Slader et al., 2002; Stern et al., 2001b; Van de Giessen et al., 1998), movement of
personal and equipment between flocks and farms, which seems to be associated with
modern integrated production, is likely to contribute to introduction of Campylobacter
infection in broiler flocks especially if clothing, boots, and equipment are contaminated with fresh fecal material from a flock excreting Campylobacter organisms (Newell and
79 Fearnley, 2003; Shane, 2000). However, a recent study showed that two adjacent broiler houses could be colonized with different Campylobacter genotypes even though these houses shared the same equipment and the farmer worked in both houses without changing any boots or clothes (Nesbit et al., 2001).
Other possible environmental sources of Campylobacter colonization in broiler flocks include aerosols, standing water, and soils around broiler houses (Jacobs-Reitsma,
1997; Newell and Fearnley, 2003). Although conditions for survival of Campylobacter species in air do not seem to be favorable, several studies could isolate Campylobacter spp. from air samples on broiler farms (Berndtson et al., 1996a; Engvall et al., 1986;
Kazwala et al., 1990). Likewise, recent studies have shown that Campylobacter spp.
could also be isolated from standing water around broiler houses before flock
colonization and the same Campylobacter genotypes were subsequently recovered from
broiler flock (Hiett et al., 2002b; Newell and Fearnley, 2003). However, the role of these
environmental sources in introduction of Campylobacter spp. to broiler flocks is still
unclear (Jacobs-Reitsma, 1997; Newell and Fearnley, 2003).
Based on information currently available, the source of broiler flock colonization
is most likely mediated by multiple sources rather than specific single source. More
details in sources of Campylobacter colonization in broiler flocks as well as information
on horizontal and vertical transmission can be obtained from the review articles
previously published by Newell and Fearnley (2003) and Sahin et al. (2002).
80 1.4.3 Control and Prevention at the Farm Level
The control and prevention of Campylobacter infection at the farm level are
considered important factors in the reduction of Campylobacter spp. on poultry carcasses
and even in the elimination of these organisms from the human food chain (Kapperud et
al., 1992; Newell and Davison, 2003; White et al., 1997). However, since the sources of
broiler flock colonization are unclear at this stage, the control and prevention of
Campylobacter organisms at the farm level seem to be very difficult. The transmission of
Campylobacter spp. in broiler flocks can occur very rapidly and once the flocks are
infected, the incidence and levels of Campylobacter colonization in those flocks seem to remain similar for farms with both good and poor hygiene (Altekrause et al., 1998;
Humphrey et al., 1993). Therefore, prevention of the first bird becoming infected with this organism is very important for successful control (Newell and Davison, 2003).
Currently, the measures for control and prevention of Campylobacter infection at the
farm level can be classified into two major strategies: (i) biosecurity to exclude the
organisms from the flock, and (ii) reduction or prevention of Campylobacter colonization of the birds by methods such as competitive exclusion and vaccination (Newell and
Davison, 2003; Newell and Wagenaar, 2000; Rowe and Madden, 1999).
Although the sources of Campylobacter infection for broiler flocks are still unclear, it appears that the horizontal transmission of Campylobacter species from contaminated environment within or around the broiler house is likely to be the primary route for introduction of Campylobacter spp. to broiler flocks (Altekruse and Tollefson,
2003; Newell and Wagenaar, 2000). Some of the common risk factors include
81 contaminated water supplies, poor broiler house maintenance leading to insect or rodent infestation, insufficient cleaning and disinfection of broiler house between flocks, and poor hygienic practices such as lack of using boot dips and change of outer clothes before entering broiler house (Newell and Wagenaar, 2000). Since poor biosecurity is likely to
be the main source of poultry flock colonization, the intervention strategy is primarily
focused on maintaining and improving biosecurity measures in an attempt to prevent
introduction of Campylobacter organisms into the broiler house (Altekruse and
Tollefson, 2003; Gibbens et al., 2001; Shane, 1992; Van de Gissen et al., 1996). Good
farm management practices including use of an all-in all-out system, cleansing and
disinfection of building between flocks, control and keep buildings free of vermin such as
insects, rodents, and wild birds, restricting other domestic animals on the farms as well as
personnel contact with those animals, along with change of outer pretective clothing and
footwear at the entrance to broiler house, use of disinfectant footbaths and hand washing
facilities, restriction on the staff and equipment entering broiler house, complete
replacement of litter, and proper disposal of dead birds may help diminish introduction of
Campylobacter spp. to broiler flocks (Altekruse et al., 1998; Altekruse and Tollefson,
2003; Gibbens et al., 2001; Humphrey et al., 1993; Kapperud et al., 1993; Kazwala et al.,
1990; Newell and Davison, 2003; Newell and Wagenaar, 2000; Shane, 1992; Shane,
2000; Van de Gissen et al., 1996; White et al., 1997). As water has been demonstrated to
be an important source of Campylobacter infection in broiler flocks, chlorination of
drinking water, removal of biofilm in water supply lines, tanks, and drinkers using
frequent cycles of flushing, and regular cleaning and disinfection of water supply system
82 can also reduce the proportion of birds colonized with Campylobacter spp. significantly although it may not totally eliminate carriage (Altekruse et al., 1998; Altekruse and
Tollefson, 2003; Gibbens et al., 2001; Humphrey et al., 1993; Kapperud et al., 1993;
Newell and Davison, 2003; Newell and Wagenaar, 2000; Pearson et al., 1993; Rowe and
Madden, 1999; Shane, 2000; Stern et al., 2002; White et al., 1997). In addition, nipple drinkers rather than bell-shaped drinkers should be used in broiler production system in order to avoid fecal contamination of water (White et al., 1997). The association between chlorination of water supply and reduction in the colonization rates of broilers with
Campylobacter spp. was revealed by Pearson et al. (1993). Because the hatchery is considered as a potential link in transmission of Campylobacter spp. from breeder flock to broilers, intensification of biosecurity procedures and decontamination of incubators at the hatchery should be emphasized as well (Shane, 2000).
As mentioned earlier, intensifying biosecurity can reduce but not eliminate the possibility of introduction of Campylobacter infection in broiler flocks (Shane, 2000). In addition, total prevention of flock exposure to Campylobacter spp. is unlikely to be accomplished by strict biosecurity measures alone (Newell and Davison, 2003).
Therefore, other supportive measures such as competitive exclusion or vaccination seem to be required for control and prevention of Campylobacter colonization in broiler flocks
(Newell and Davison, 2003; Shane, 2000). Although competitive exclusion has been a relatively successful approach for the control of Salmonella, its efficacy against
Campylobacter colonization seems to be variable and unpredictable (Aho et al., 1992;
Newell and Wagenaar, 2000). However, several studies reported that competitive
83 exclusion could help reduce the level and the prevalence of Campylobacter colonization
in broilers and it could partly protect the birds from Campylobacter colonization in some
experiments as well (Humphrey et al., 1989; Mead et al., 1996; Morishita et al., 1997;
Newell and Wagenaar, 2000; Saleha et al., 1998; Schoeni and Doyle, 1992; Schoeni and
Wong, 1994; Soerjadi et al., 1982; Soerjadi-Liem et al., 1984; Stern, 1994). In general,
competitive exclusion materials are obtained from adult donor birds and gave to the
chicks by oral administration. These competitive exclusion materials can be fecal
suspension, cecum-colonizing bacteria that produce anti-Campylobacter metabolites, or
anaerobically harvested mucus from the cecal mucosa, either cultured in a laboratory
medium or used as a diluted suspension (Humphrey et al., 1989; Mead et al., 1996;
Newell and Wagenaar, 2000; Saleha et al., 1998; Schoeni and Doyle, 1992; Schoeni and
Wong, 1994; Soerjadi et al., 1982; Soerjadi-Liem et al., 1984; Stern, 1994; Stern et al.,
2001a). However, because these materials are derived from live birds, it is not surprising
that they will be inconsistent and are mainly influenced by the status of the donor birds
and the site from which the starter materials are collected (Laisney et al., 2004; Mead et
al., 1996; Newell and Wagenaar, 2000; Schoeni and Doyle, 1992; Schoeni and Wong,
1994). Recently, several commercial probiotic preparations have been available such as
Avian Pac Soluble, which is a mixture of Lactobacillus acidophilus and Streptococcus faecium (Chang and Chen, 2000; Morishita et al., 1997; Saleha et al., 1998). This avian- specific probiotic containing L. acidophilus and S. faecium has been found to be able to reduce the colonization and frequency of fecal shedding of Campylobacter spp. in broilers as well (Morishita et al., 1997). Besides L. acidophilus and S. faecium, other
84 microorganisms including a mixture of Klebsiella pneumoniae, Citrobacter diversus, and
Escherichia coli (O13:H-) and yeast Saccharomyces boulardii could also protect or at
least reduce the level of Campylobacter colonization in cecum of broilers (Line et al.,
1997; Line et al., 1998; Newell and Wagenaar, 2000; Rowe and Madden, 1999; Schoeni and Doyle, 1992). In addition, Campylobacter strains, which have a high colonizing potential but are nonpathogenic may be another interesting alternative to use as competitive excluders of pathogenic Campylobacter strains (Barrow and Page, 2000;
Chen and Stern, 2001; Newell and Wagenaar, 2000). Although most of competitive exclusion materials mentioned above seem to have good results under experimental conditions, their abilities in control and prevention of Campylobacter colonization in broiler flocks under commercial field conditions still need to be investigated. In addition
to competitive exclusion, vaccination seems to be another possible alternative approach
for control and prevention of Campylobacter colonization; however, effective vaccine
strategies directed against infection with Campylobacter spp. in broilers have yet to be
developed (Newell and Wagenaar, 2000). Passive immunization by oral administration of
anti-Campylobacter antibodies has also been reported to have both therapeutic and
prophylactic properties in broilers (Newell and Wagenaar, 2000; Stern et al., 1990b;
Tsubokura et al., 1997). Therefore, it may be possible to immunize parent flocks to
produce passively protected chickens (Newell and Wagenaar, 2000). Based on the
information currently available, vaccination of chickens is likely to help reduce rather
than prevent colonization (Newell and Wagenaar, 2000). Other intervention strategies
including use of inbred chicken lines, which are resistant to Campylobacter infection, use
85 of enzyme-supplemented or other supplemented diet, or administration of
Campylobacter-specific bacteriophages may be useful for control and prevention of
Campylobacter colonization in broiler flocks as well (Altekruse and Tollefson, 2003;
Bailey, 1993; Connerton et al., 2004; El-Shibiny et al., 2005; Fernandez et al., 2000b;
Heres et al., 2004; Hinton et al., 2002; Kassaify and Mine, 2004; Newell and Davison,
2003; Newell and Wagenaar, 2000; Rowe and Madden, 1999; Shane, 2000; Stern et al.,
1990a).
1.5 Antimicrobial Resistance of Campylobacter Species
1.5.1 Antimicrobial Susceptibility Testing Methods
In general, antimicrobial susceptibility testing of Campylobacter species can be classified into two major tests, which are dilution and diffusion methods (Aarestrup and
Engberg, 2001; Nachamkin et al., 2000b). Although a number of different diffusion and dilution methods have been used to measure susceptibility of bacterial species to various antimicrobial agents, the most commonly used diffusion and dilution methods for antimicrobial susceptibility testing of Campylobacter spp. include disk diffusion test, epsilometer test (E-test), agar dilution test, and broth microdilution test (Aarestrup and
Engberg, 2001; Alfredson et al., 2003; Engberg et al., 1999; Fernandez et al., 2000a;
Frediani-Wolf and Stephan, 2003; Gaudreau and Gilbert, 1997; Ge et al., 2002; Huang et al., 1992; Huysmans and Turnidge, 1997; Luber et al., 2003a; McDermott et al., 2004;
Nachamkin et al., 2000b; Oncul et al., 2003).
86 The agar diffusion test (disk diffusion and E-test) is relatively easy to perform and is very useful especially when several antimicrobial agents need to be tested against a few isolates; however, in order to be reproducible, it requires a high level of standardization and quality control. In addition, the results (the zone of inhibition) of this test are directly influenced by the agar depth and the inoculum size (Acar and Goldstein, 1996; Caprioli et al., 2000; McDermott et al., 2004; Potz et al., 2004). Because of its convenience and low cost, the disk diffusion test has been widely used and probably be the most commonly used antimicrobial susceptibility testing method. However, only qualitative data can be obtained from this test (Acar and Goldstein, 1996; Caprioli et al., 2000;
McDermott et al., 2004; Potz et al., 2004). The E-test, on the other hand, can provide quantitative minimal inhibitory concentration (MIC) values although its cost is much higher than that of the disk diffusion method (Acar and Goldstein, 1996). In terms of the dilution test (agar dilution and broth microdilution), although this method is very reliable, highly reproducible, and provides quantitative MIC values, it is labor-intensive, time- consuming, and quite expensive especially when compared to the disk diffusion method
(Caprioli et al., 2000). Nevertheless, the agar dilution method has been recognized as a standard antimicrobial susceptibility testing method for Campylobacter species
(McDermott et al., 2004; NCCLS, 2002a). The agreement between different antimicrobial susceptibility testing methods particularly between the agar dilution method and the E-test in determining antimicrobial resistance of Campylobacter spp. has been
87 reported by several studies (Alfredson et al., 2003; Engberg et al., 1999; Fernandez et al.,
2000a; Frediani-Wolf and Stephan, 2003; Gaudreau and Gilbert, 1997; Ge et al., 2002;
Huang et al., 1992; Huysmans and Turnidge, 1997; Luber et al., 2003a; Oncul et al.,
2003).
Although antimicrobial susceptibility test of thermophilic Campylobacter by the
agar dilution method has not been standardized yet, it is recommended that Mueller-
Hinton agar supplemented with 5% defibrinated sheep blood should be used and these
agar plates should be incubated at 42 °C for 24 hours or at 36 °C for 48 hours under a
microaerophilic atmosphere (approximately 5% O2, 10% CO2, and 85% N2). In addition,
Campylobacter jejuni ATCC 33560 should also be used as a quality control organism
(McDermott et al., 2004; NCCLS, 2002a). Since there are no internationally accepted standard resistance breakpoints specific for Campylobacter species available currently, the resistance breakpoints of enteric bacteria in the family Enterobacteriaceae have been used to determine antimicrobial resistance of thermophilic Campylobacter (Ge et al.,
2002; Luber et al., 2003a; Nachamkin et al., 2000b). Due to the lack of a standardized
method and the validated resistance breakpoints specific for thermophilic Campylobacter, different antimicrobial susceptibility tests and different resistance breakpoints have been used between studies; therefore, antimicrobial resistance of Campylobacter species reported from different countries in different studies should be analysed with caution.
88 1.5.2 Occurrence and Trends of Antimicrobial Resistance
A rapid increase in the proportion of Campylobacter strains resistant to
antimicrobial agents particularly to fluoroquinolones has been observed in every region
of the world. In the United States, the emergence of fluoroquinolone resistance among
Campylobacter isolates was first noticed about 10 years ago. Prior to 1992 there were no reports of fluoroquinolone-resistant Campylobacter strains isolated from humans in the
U.S. (Nachamkin et al., 2002; Wang et al. 1984). However, the prevalence of fluoroquinolone resistance among these Campylobacter isolates increased significantly
from 1.3% in 1992 to 8% - 13% during 1996 - 1998 and this resistance trend has increased steady since 1998 (Gupta et al., 2004; Nachamkin et al., 2002; Smith et al.,
1999). In 2001, the National Antimicrobial Resistance Monitoring System (NARMS) and
Nachamkin et al. (2002) found that about 19% - 40% of Campylobacter strains isolated
from humans were resistant to ciprofloxacin (Gupta et al., 2004; Nachamkin et al., 2002).
In Canada, no fluoroquinolone resistance was observed in Campylobacter isolates during
1980 to 1986 (Gaudreau and Gilbert, 1998; Lariviere et al., 1986). However, in 1992 -
1997, about 3.5% - 13.6% of these Campylobacter isolates became resistant to fluoroquinolones (Gaudreau and Gilbert, 1998; Harnett et al., 1995). In 1998 - 2000, the rate of ciprofloxacin resistance among C. jejuni in Canada rose to 10% - 27% and
dramatically increased to 47% in 2001 (Gaudreau and Gilbert, 2003). In South America,
18.2% - 25% of Campylobacter strains isolated from humans and animals in Brazil were
also resistant to fluoroquinolones (Aquino et al., 2002).
89 In Europe, particularly in the Netherlands where fluoroquinolone-resistant
Campylobacter isolates were first reported, the prevalence of Campylobacter strains resistant to this antimicrobial agent increased from 0% in 1982 to 11% in 1989 for human isolates and to 14% for the isolates from poultry products (Endtz, 1991). In 1992, five years after enrofloxacin was licensed for use in poultry in the Netherlands, Jacobs-
Reitsma et al. (1994c) found that about 31% of Campylobacter isolates from broilers were resistant to fluoroquinolones. Similarly, 29% of human Campylobacter isolates in
1997 were also resistant to these antibiotics (Talsma et al., 1999). Besides the
Netherlands, the emergence of fluoroquinolone resistance was found in other European countries as well. In Belgium, 44%, 28%, and 35% of C. jejuni isolated from broilers, layers, and turkeys and 62%, 41%, and 28% of C. coli isolated from broilers, layers, and swine in 1998 were resistant to fluoroquinolones, respectively (Van Looveren et al.,
2001). Likewise, 17% of C. jejuni and 40% C. coli isolated from broilers in France in
1999 were also found to be resistant to enrofloxacin (Avrain et al., 2001). In addition, a recent study in Italy showed that 42%, 53%, and 38% of C. jejuni isolated from broilers, chicken meat, and humans and 75%, 79%, and 55% of C. coli isolated from the same sources during 2000 - 2001 were resistant to fluoroquinolones, respectively (Pezzotti et al., 2003) In Switzerland, the prevalence of fluoroquinolone resistance in Campylobacter isolates from poultry carcasses increased from 0% in 1997 - 1998 (Frei et al., 2001) to
29% in 2002 (Ledergerber et al., 2003). Similarly, recent studies in Germany also revealed significant increases in the rates of fluoroquinolone resistance in both human and poultry isolates (Krausse and Ullmann, 2003; Luber et al., 2003b). Krausse and
90 Ullmann (2003) and Luber et al. (2003b) reported that fluoroquinolone-resistant
Campylobacter strains isolated from humans in Germany rose from 0% in 1980 - 1982 to
4.9% in 1991 and to 31% - 45.1% during 2000 - 2002, while the prevalence of fluoroquinolone resistance in Campylobacter strains isolated from chickens and turkeys in Germany increased from 27.3% and 25.8% in 1991 to 45.6% and 33.3% in 2001 -
2002, respectively (Luber et al., 2003b). A significant increase in fluoroquinolone
resistance among human Campylobacter isolates was observed in Greece as well. During
1987 - 1989, no fluoroquinolone-resistant C. jejuni isolates was reported in Greece;
however, 30.6% of Campylobacter isolates in 1998 - 2000 were resistant to these antimicrobial agents (Chatzipanagiotou et al., 2002). In Spain, the resistance rates to fluoroquinolone antibiotics among Campylobacter isolates increased rapidly from 0% -
3% before 1988 to 7% - 14% in 1989 to 30% in 1990 - 1991 and to 31% - 57% in 1992 -
1993 (Navarro et al., 1993; Prats et al., 2000; Reina, 1992; Reina et al., 1992; Reina et al.,
1994, Sanchez et al., 1994; Velazquez et al., 1995). During 1995 and 1998, an extremely high prevalence of fluoroquinolone resistance among Campylobacter isolates in Spain was reported. Approximately 99% of Campylobacter strains isolated from broilers were resistant to fluoroquinolones, while about 71% - 90% of human isolates were resistant to these antibiotics (Mirelis et al., 1999; Prats et al., 2000; Saenz et al., 2000). The emergence of fluoroquinolone resistance among Campylobacter isolates also occurred in the United Kingdom. Until the end of 1990, all Campylobacter isolates in the U.K. were susceptible to ciprofloxacin; however, since 1991 a steady increase in fluoroquinolone resistance in Campylobacter isolates in this country has been reported (Bowler, 1992;
91 Bowler et al., 1996; Gaunt and Piddock, 1996; McIntyre, 1993; Sam et al., 1999;
Thwaites and Frost, 1999). Similarly, Campylobacter strains isolated from humans and poultry in Ireland and Northern Ireland were also resistant to fluoroquinolones (Fallon et al., 2003; Lucey et al., 2000; Lucey et al., 2002; Moore et al., 2001; Oza et al., 2003;
Wilson 2003). In Ireland, fluoroquinolone resistance among Campylobacter isolates from humans increased from 0% in 1996 - 1998 to 34% in 2000, while fluoroquinolone resistance among Campylobacter isolates from poultry increased from 3.1% in 1996 -
1998 to 18% - 29% in 1999 - 2000 (Fallon et al., 2003; Lucey et al., 2000; Lucey et al.,
2002). Prior to 1992, no Campylobacter isolates from humans in Northern Ireland was resistant to fluoroquinolones; however, fluoroquinolone resistance among these
Campylobacter isolates increased to 10% in 1993 and to 17.4% - 23% in 1999 - 2000
(Moore et al., 2001). Interestingly, during the same period of time about 3% of
Campylobacter isolates from broilers and 10% of the isolates from chicken meat were resistant to these antibiotics (Oza et al., 2003; Wilson 2003). Similar observation was also reported in Denmark, where 21% of human isolates, 6% of broiler isolates, and 13% of chicken meat isolates were resistant to ciprofloxacin (Anonymous, 2001). A high frequency of fluoroquinolone resistance among Campylobacter isolates was found not only in Denmark, but also in other Scandinavian countries (Aarestrup et al., 1997;
Andreasen, 1987; Anonymous, 2001; Hakanen et al., 2003; Pedersen and Wedderkopp,
2003; Rautelin et al., 1991; Rautelin et al., 1993; Sjogren et al., 1992; Sjogren et al.,
1997; Svedhem et al., 1981). In Sweden, the trend of fluoroquinolone resistance among
Campylobacter isolates rose from 0% in 1981 to 6.1% in 1992 - 1995 (Sjogren et al.,
92 1992; Sjogren et al., 1997; Svedhem et al., 1981), whereas the prevalence of
ciprofloxacin resistance among Campylobacter strains isolated from humans in Finland
increased dramatically from 0% in 1978 - 1980 to 17% in 1993 and to 46% in 1995 -
2000 (Hakanen et al., 2003; Rautelin et al., 1991; Rautelin et al., 1993). Similar trends of
antimicrobial resistance in Campylobacter strains isolated from humans and poultry were
also observed in other European countries (Adler-Mosca et al., 1991; Engberg et al.,
2001; Frediani-Wolf and Stephan, 2003; Hirschl et al., 1990; Ledergerber et al., 2003;
Lekowska-Kochaniak et al., 1996; Nachamkin et al., 2000b; Smith et al., 2000).
The emergence of fluoroquinolone resistance in Campylobacter isolates was not
restricted only in Europe and North and South America, but this problem also occurred in
other continents of the world. A study in Egypt showed that fluoroquinolone-resistant
Campylobacter strains isolated from humans increased rapidly from 13% in 1995 to 28%
in 1996 to 35% in 1997 and to 48% - 50% in 1998 - 2000 (Putnam et al., 2003). In Asia, particularly in East and Southeast Asia, the high prevalence of fluoroquinolone resistance in Campylobacter isolates was also reported. Approximately 57% and 91% of
Campylobacter strains isolated from humans and poultry in Taiwan during 1994 to 1996 were resistant to ciprofloxacin, respectively (Li et al., 1998). In Japan, no fluoroquinolone resistance among Campylobacter strains isolated from humans was detected in 1981 - 1982; however, since 1993 the frequency of fluoroquinolone-resistant
Campylobacter strains increased remarkably. In addition, 32.4% of Campylobacter strains isolated from broilers in Japan during 1995 - 1999 were also resistant to fluoroquinolones (Chuma et al., 2001; Niwa et al., 2004; Sagara et al., 1987). Likewise,
93 the fluoroquinolone resistance among Campylobacter strains isolated from humans in
Thailand was dramatically increased as well (Hoge et al., 1998; Isenbarger et al., 2002;
Kuschner et al., 1995). Before 1991, all Campylobacter isolates in Thailand were
uniformly susceptible to quinolones. In 1993, 40% - 50% of Campylobacter isolates were found to be resistant to fluoroquinolones (Hoge et al., 1998; Kuschner et al., 1995). By
1994 this resistance rate rose to 76% and during 1995 to 1999 about 77% - 84% of these
Campylobacter isolates in Thailand were resistant to these antimicrobial agents (Hoge et al., 1998; Isenbarger et al., 2002). Similarly, the frequency of fluoroquinolone-resistant
Campylobacter strains isolated from humans in Indonesia also increased from 0% in
1995 - 1997 to 22% - 43% in 1998 - 2001 (Tjaniadi et al., 2003). Although the emergence of fluoroquinolone resistance in Campylobacter isolates was also observed in Australia and New Zealand, the percentage of fluoroquinolone-resistant Campylobacter isolates was very low when compared to the isolates obtained from other countries (Dowling et al., 1998; Sharma et al., 2003).
The development of antimicrobial resistance of Campylobacter strains isolated from humans and animals to erythromycin, tetracycline, as well as other antimicrobial agents was also reported in many countries worldwide (Aarestrup et al., 1997; Alfredson et al., 2003; Avrain et al., 2001; Avrain et al., 2003; Bradbury and Munroe, 1985; Cabrita et al., 1992; Elharrif et al., 1985; Engberg et al., 2001; Fallon et al., 2003; Frediani-Wolf and Stephan, 2003; Gaudreau and Gilbert, 1998; Hoge et al., 1998; Isenbarger et al.,
2002; Jacobs-Reitsma et al., 1994; Karmali et al., 1981; Krausse and Ullmann, 2003;
Lariviere et al., 1986; Ledergerber et al., 2003; Lekowska-Kochaniak et al., 1996; Li et
94 al., 1998; Luber et al., 2003b; Lucey et al., 2000; Michel et al., 1983; Nachamkin et al.,
2000b; Narvarro et al., 1993; Prats et al., 2000; Rautelin et al., 1991; Reina et al., 1992;
Saenz et al., 2000; Sagara et al., 1987; Sanchez et al., 1994; Sjogren et al., 1992; Smith et
al., 2000; Talsma, 1999; Tajada et al., 1996; Taylor et al., 1987; Vanhoof et al., 1982;
Van Looveren et al., 2001; Wang et al., 1984).
1.5.3 Mechanisms of Antimicrobial Resistance
As mentioned in the previous section, the prevalence of antimicrobial resistance
in Campylobacter species has increased drastically over the last decade and this problem
has been reported from many countries in every continent of the world. In addition, a
large number of Campylobacter isolates from both humans and animals were found to be resistant not only to fluoroquinolones, but also to other classes of antibiotics. The mechanisms of antimicrobial resistance in thermophilic Campylobacter to those antibiotics including to fluoroquinolones are summarized in this section. More details on
the mechanisms of antibiotic resistance in Campylobacter species can be obtained from
the previous review articles by Taylor and Courvalin (1988) and Trieber and Taylor
(2000).
Fluoroquinolone resistance in Campylobacter species is mainly due to point
mutations in the gyrA gene, which encodes the A subunit of the DNA gyrase enzyme
(Aarestrup and Engberg, 2001; Engberg et al., 2001; Gibreel et al., 1998; Luo et al.,
2003; Piddock et al., 2003; Wang et al., 1993; Zhang et al., 2003; Zirnstein et al., 1999).
The mutations occurred in the gyrA gene at specific positions such as Thr-86, Asp-90,
95 and Ala-70 were not only responsible for fluoroquinolone resistance in Campylobacter species, but the positions of these mutations also have direct effect on the level of fluoroquinolone resistance in Campylobacter isolates; for example, the Thr-86-Ile mutation was associated with high-level fluoroquinolone resistance, while the mutations occurred at other positions such as Asp-90-Asn, Ala-70-Thr, and Thr-86-Lys mutations were associated with lower level of fluoroquinolone resistance (Aarestrup and Engberg,
2001; Engberg et al., 2001; Gootz and Martin, 1991; Luo et al., 2003; Wang et al., 1993;
Zhang et al., 2003). Besides the point mutations in the gyrA gene, an active multi-drug efflux pump (CmeABC) is also linked to fluoroquinolone resistance in Campylobacter isolates (Charvalos et al., 1995; Lin et al., 2002; Luo et al., 2003; Pumbwe and Piddock,
2002; Pumbwe et al., 2004; Zhang et al., 2003).
Although the mechanism of macrolide resistance can occur through different mechanisms such as target modification by mutation or methylation; enzymatic inactivation by different types of macrolide-inactivating enzymes such as esterases, hydrolases, and transferases; or active efflux system, the mechanism of macrolide resistance in Campylobacter species seems to be associated with an alteration of the target site on the 23S rRNA of Campylobacter ribosomes or mutations at positions 2074 and 2075 of the ribosomal protein genes (23S rRNA) (Aarestrup and Engberg, 2001;
Engberg et al., 2001; Niwa et al., 2001; Payot et al., 2004; Schwarz and Chaslus-Dancla,
2001; Trieber and Taylor, 2000; Yan and Taylor, 1991).
In general, the mechanism of aminoglycoside resistance is mainly due to enzymatic inactivation by aminoglycoside-modifying enzymes. As a result, these
96 aminoglycoside antibiotics are not able to interact with the ribosome, which is the target
site of this antibiotic group (Aarestrup and Engberg, 2001; Schwarz and Chaslus-Dancla,
2001; Shaw et al., 1993; Trieber and Taylor, 2000; Trieu-Cuot and Courvalin, 1986). The
aminoglycoside-modifying enzymes can be divided into three different groups based on the reaction that they mediate (phosphorylation or adenylation/nucleotidylation of a hydroxyl group or acetylation of an amino group). Three different types of these aminoglycoside-modifying enzymes include aminoglycoside phosphotransferases (APH), aminoglycoside adenyltransferases or aminoglycoside nucleotidyltransferases (AAD or
ANT) and aminoglycoside acetyltransferases (AAC) (Aarestrup and Engberg, 2001;
Schwarz and Chaslus-Dancla, 2001; Shaw et al., 1993; Trieber and Taylor, 2000; Trieu-
Cuot and Courvalin, 1986). Each enzyme is also divided into various subgroups depending on the site at which the antibiotic is modified; for example, APHs, which phosphorelate the hydroxyl groups (O-phosphotransferases) at positions 4, 6, 3’, 2”, 3”, and 5” will be recognized as APH(4), APH(6), APH(3’), APH(2”), APH(3”), and
APH(5”), respectively, while AADs, which adenylate the hydroxyl groups (O-
adenyltransferases) at positions 6, 9, 4’, 2”, and 3” will be recognized as AAD(6),
AAD(9), AAD(4’), AAD(2”), and AAD(3”)(9) and AACs, which acetylate the amino groups (N-acetyltransferases) at positions 1, 3, 2’, and 6’ will be recognized as AAC(1),
AAC(3), AAC(2’), and AAC(6’), respectively (Schwarz and Chaslus-Dancla, 2001;
Shaw et al., 1993; Trieber and Taylor, 2000; Trieu-Cuot and Courvalin, 1986). Although several aminoglycoside-modifying enzymes are asoociated with aminoglycoside resistance in Campylobacter species, the most common enzyme that involves kanamycin
97 and structurally related antibiotics such as neomycin resistance in Campylobacter species
is 3’-aminoglycoside phosphotransferase type III [APH(3’)-III], which is encoded by the aphA-3 gene (Aarestrup and Engberg, 2001; Gibreel et al., 2004a; Ouellette et al., 1987;
Papadopoulou and Courvalin, 1988; Saenz et al., 2000; Sagara et al., 1987; Taylor and
Courvalin, 1988; Tenover and Elvrum, 1988; Tenover et al., 1992; Trieber and Taylor,
2000; Trieu-Cuot and Courvalin, 1986). However, the aphA-3 gene is not the only kanamycin resistance gene found in Campylobacter species. Other genes such as aphA-7 and aphA-1 are also associated with a high-level of kanamycin resistance in
Campylobacter and Campylobacter-like organisms, respectively (Gibreel et al., 2004a;
Ouellette et al., 1987; Taylor and Courvalin, 1988; Tenover and Elvrum, 1988; Tenover et al., 1992). The aphA-1 gene, which encodes 3’-aminoglycoside phosphotransferase type I [APH(3’)-I] is chromosomally located, whereas the aphA-3 and aphA-7 genes are generally found on the plasmid. Although both aphA-3 and aphA-7 genes are located on the plasmid, the aphA-3 gene is frequently detected on large plasmids (41 to 132 kb) that also encode other resistance determinants particularly tetracycline resistance determinant
(tetO gene), while the aphA-7 gene seems to be located on smaller plasmids (9.5 to 11.5 kb) that apparently do not encode other resistance traits (Gibreel et al., 2004a; Ouellette et al., 1987; Papadopoulou and Courvalin, 1988; Saenz et al., 2000; Sagara et al., 1987;
Taylor and Courvalin, 1988; Tenover and Elvrum, 1988; Tenover et al., 1992). In addition, kanamycin resistance in Campylobacter species especially in C. coli can be mediated by 3’-aminoglycoside phosphotransferase type IV [APH(3’)-IV] enzyme as well (Aarestrup and Engberg, 2001; Rivera et al., 1986; Trieber and Taylor, 2000).
98 Currently, four different mechanisms associated with tetracycline resistance in bacteria have been reported including (i) an energy-dependent efflux system, which helps reduce the intracellular concentration of tetracycline by an integral membrane protein; (ii) ribosomal protection protein, which protects the ribosomal binding site of tetracycline;
(iii) inactivation of tetracycline by modifying enzymes, which occurs rarely; and (iv) mutation of the 16S rRNA, which found in propionibacteria (Aarestrup and Engberg,
2001; Chopra et al., 1992; Chopra and Roberts, 2001; Roberts, 1996; Ross et al., 1998;
Schnappinger and Hillen, 1996; Speer et al., 1992; Taylor and Chau, 1996; Trieber and
Taylor, 2000). However, only a ribosomal protection protein is associated with tetracycline resistance in Campylobacter species (Aarestrup and Engberg, 2001; Trieber and Taylor, 2000). As mentioned earlier, the mechanism of tetracycline resistance in
Campylobacter is primarily associated with tet(O) gene, which encodes a ribosomal protection protein designated as Tet(O) (Taylor and Courvalin, 1988; Trieber and Taylor,
2000). Tet(O) protein interacts directly with the ribosome, the target of tetracycline, and displaces tetracycline from its primary binding site on the ribosome; thus, protecting the ribosome from the inhibitory effect of tetracycline (Connell et al., 2002; Connell et al.,
2003; Gibreel et al., 2004b; Manavathu et al., 1990; Pratt and Korolik, 2005; Trieber and
Taylor, 2000). Although tet(O) gene is usually carried on transmissible plasmids
(Aarestrup and Engberg, 2001; Gibreel et al., 2004b; Lee et al., 1994; Pratt and Korolik,
2005; Sagara et al., 1987; Taylor and Courvalin, 1988; Trieber and Taylor, 2000), it has been found to be chromosomally-located as well (Gibreel et al., 2004b; Lee et al., 1994;
Pratt and Korolik, 2005).
99 The main resistance mechanism of C. jejuni and C. coli isolates to β-lactam antimicrobial agents such as ampicillin seems to be associated with the production of enzyme β-lactamases, which break the β-lactam ring of β-lactam antibiotics (Aarestrup and Engberg, 2001; Lachance et al., 1991; Lachance et al., 1993; Tajada et al., 1996;
Trieber and Taylor, 2000). Although approximately 90% of C. jejuni and 70% of C. coli produce this type of enzyme, a large number of Campylobacter isolates were found to be susceptible to ampicillin (Lachance et al., 1991; Lachance et al., 1993; Lariviere et al.,
1986; Tajada et al., 1996; Trieber and Taylor, 2000). In addition, several ampicillin- resistant Campylobacter strains, on the other hand, were found to be negative for β- lactamase production (Saenz et al., 2000). This information indicated that other mechanisms of resistance such as alteration of penicillin-binding proteins or decreased permeability of the drug through modification of porins could be involved with ampicillin resistance in Campylobacter species as well (Aarestrup and Engberg, 2001; Saenz et al.,
2000; Tajada et al., 1996; Trieber and Taylor, 2000).
Although the mechanism of multidrug resistance in Campylobacter species has not yet been clearly identified, it appears that the multidrug resistance mechanism in
Campylobacter species seems to be associated with the presence of an active efflux system (Charvalos et al., 1995; Lin et al., 2002; Pumbwe and Piddock, 2002; Pumbwe et al., 2004; Trieber and Taylor, 2000).
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149
CHAPTER 2
THE EFFECT OF DIFFERENT POULTRY PRODUCTION PRACTICES ON
ANTIBIOTIC RESISTANCE RATES OF CAMPYLOBACTER IN POULTRY
2.1 ABSTRACT
Intestinal tracts of broilers and turkeys from 10 conventional poultry farms, where
antibiotics were routinely used, and from 5 organic poultry farms, where antibiotics had
never been used, were collected and cultured for Campylobacter species. A total of 694
Campylobacter jejuni and Campylobacter coli isolates from the conventional and organic
poultry operations were tested for antimicrobial resistance to nine antimicrobial agents by
the agar dilution method. Although Campylobacter spp. were highly prevalent in both
conventional and organic poultry operations, the antibiotic resistance rates of the isolates
were significantly different between the organic operations and the integrated
conventional operations. Less than 2% of Campylobacter strains isolated from organically-raised poultry were resistant to fluoroquinolones, while 46% and 67% of
Campylobacter isolates from conventionally-raised broilers and turkeys were resistant to these antimicrobials. In addition, a high frequency of resistance to erythromycin
(79.60%), clindamycin (64.18%), kanamycin (76.12%), and ampicillin (31.34%) was observed among Campylobacter isolates from conventionally-raised turkeys. None of the
150 Campylobacter isolates obtained in this study was resistant to gentamicin, while a large
number of the isolates from both conventional and organic poultry operations were
resistant to tetracycline. Multidrug resistance was mainly observed among
Campylobacter strains isolated from conventional turkey operation (81.09%). Findings
from this study clearly indicate that different production practices significantly influence
the antibiotic resistance rates of Campylobacter on poultry farms, which is likely due to
the result of antimicrobial usage in conventional poultry production.
2.2 INTRODUCTION
Food-borne campylobacteriosis, a major public health concern in the United
States and many countries worldwide, is caused mainly by Campylobacter jejuni (Mead et al., 1999). It is estimated that over 2 million cases of food-borne bacterial diarrhea that
occur each year in the United States are caused by Campylobacter (Altekruse et al.,
1999). In other industrialized countries, the numbers of Campylobacter infections
exceeded those of Salmonella, Shigella, and E. coli 0157:H7 infections combined (Allos,
2001). Campylobacter jejuni is not only an important cause of bacterial gastroenteritis in
humans, but it has also been associated with Guillane-Barré Syndrome (GBS), an acute
immune-mediated demyelinating disorder of the peripheral nervous system (Blaser, 1997;
Nachamkin et al., 1998).
Most Campylobacter infections in humans usually occur as sporadic cases and are
associated with the ingestion of contaminated or improperly handled/cooked foods as
well as milk or dairy products (Blaser, 1997; Altekruse and Tollefson, 2003). However,
151 consumption of undercooked poultry and/or other foods that are cross-contaminated with raw poultry meat during food preparation is considered a major risk factor for food-borne campylobacteriosis (Altekruse and Tollefson, 2003). As a commensal organism, thermophilic Campylobacter spp. including C. jejuni and C. coli are highly prevalent in chickens and turkeys (Newell and Fearnley, 2003; Newell and Wagenaar, 2000; Sahin et al., 2002). Due to the large number of Campylobacter present in feces, contamination of chicken carcasses by Campylobacter in slaughter houses is extensive and unavoidable, resulting in the potential transmission of Campylobacter from contaminated chicken meats to consumers.
Although most Campylobacter infections are mild, self-limiting, and usually resolve within a few days without any antimicrobial therapy, severe or prolonged infections can also occur under certain circumstances, in which an appropriate antimicrobial treatment is usually warranted (Blaser, 1997; Engberg et al., 2001). For instance, antibiotic therapy may be life saving and is required in invasive and/or systemic infections particularly in infants, elderly, or immunocompromised persons. Among a variety of antibiotics that are generally prescribed for treatment of bacterial gastrointestinal infections in human, macrolides (e.g. erythromycin) and fluoroquinolones
(e.g. ciprofloxacin) are considered to be the drugs of choice for treatment of food-borne campylobacteriosis (Allos, 2001). However, these antibiotics are also used for treatment of diseases in livestock and poultry (Khachatourians, 1998; McEwen and Fedorka-Cray,
2002; Van den Bogaard and Stobberingh, 1999). Because of the widespread use of antimicrobial agents in humans and animals, the emergence of antimicrobial resistance
152 has increased dramatically over the last decade (Khachatourians, 1998; Threlfall et al.,
2000; White et al., 2002). Antimicrobial-resistant Campylobacter strains isolated from humans and animals were reported in many countries worldwide (Engberg et al., 2001;
Nachamkin et al., 2000b; Smith et al., 2000).
In the United States, the emergence of fluoroquinolone resistance among
Campylobacter isolates was first noticed about 10 years ago. Prior to 1992 there were no reports of fluoroquinolone-resistant Campylobacter strains isolated from humans in the
U.S. (Nachamkin et al., 2002). However, the prevalence of fluoroquinolone resistance among these Campylobacter isolates increased significantly from 1.3% in 1992 to 8% -
13% during 1996 - 1998 and this resistance trend has increased steady since 1998 (Gupta et al., 2004; Nachamkin et al., 2002; Smith et al., 1999). In 2001, the National
Antimicrobial Resistance Monitoring System (NARMS) and Nachamkin et al. (2002) found that about 19% - 40% of Campylobacter strains isolated from humans were resistant to ciprofloxacin (Gupta et al., 2004; Nachamkin et al., 2002). In other areas of
North America, such as Canada, no fluoroquinolone resistance was observed in
Campylobacter isolates during 1985 to 1986 (Gaudreau and Gilbert, 1998). However, in
1992 - 1997, 3.5% - 13.6% of Campylobacter isolates tested were resistant to fluoroquinolones (Gaudreau and Gilbert, 1998). In 1998-2000, the rate of ciprofloxacin resistance among C. jejuni in Canada rose to 10% - 27% and drastically increased to 47% in 2001 (Gaudreau and Gilbert, 2003).
In other regions of the world, the prevalence of fluoroquinolone resistance in
Campylobacter strains isolated from humans and animals particularly poultry also
153 increased remarkably over the past decade (Engber et al., 2001; Nachamkin et al.,
2000b). In Europe, the fluoroquinolone resistance rates among Campylobacter isolates
increased from less than 3% before 1988 to more than 95% in some European countries during the late 1990s and the early twenty-first century (Avrain et al., 2001; Engber et al.,
2001; Luber et al., 2003b; Nachamkin et al., 2000b; Saenz et al., 2000; Van Looveren et al., 2001). In Africa, fluoroquinolone-resistant Campylobacter strains isolated from humans increased from 13% in 1995 to 28% in 1996 to 35% in 1997 and to 48% - 50% in
1998 – 2000 (Putnam et al., 2003). In Asia, particularly in East and Southeast Asia, the fluoroquinolone resistance among Campylobacter strains isolated from humans and animals also remarkably increased. Since 1993, approximately 50% to 90% of
Campylobacter isolates from this region were resistant to ciprofloxacin (Engberg et al.,
2001; Li et al., 1998; Nachamkin et al., 2000b). Although the emergence of fluoroquinolone resistance in Campylobacter isolates was also observed in Australia, the percentage of fluoroquinolone-resistant Campylobacter isolates was very low when compared to the isolates obtained from other countries (Sharma et al., 2003).
Despite these advances in understanding the epidemiology of antibiotic-resistant
Campylobacter, relatively little is known about the impact of different food animal
production practices on the prevalence of antibiotic-resistant Campylobacter. Since
contaminated poultry are considered an important source of Campylobacter infection in
humans and the demand for organic animal produce is increasing in the U.S., we
conducted a farm-based study to determine the antimicrobial resistance rates in
Campylobacter isolates from both conventionally-raised and organically-raised broilers
154 and turkeys. It is found that Campylobacter is highly prevalent in both organic and conventional production systems. However, the antibiotic resistance rates are significantly higher in conventional operations than in organic operations. This study also revealed that antibiotic-resistant Campylobacter strains are stable and can persist in the absence of antibiotic usage.
2.3 MATERIALS AND METHODS
Sample collection. This study focused on the prevalence of antibiotic-resistant
Campylobacter in slaughter-age birds. The surveyed farms included 10 integrated conventional broiler operations, 10 integrated conventional turkey operations, 5 organic broiler farms, and 5 organic turkey farms. Intestinal samples were collected from slaughter houses during August 2000 to November 2002. At least 30 intestines from each conventional poultry farm and approximately 60 intestines from each organic poultry farm were collected at each collection period. A total of 345, 360, 355, and 230 intestinal tracts were obtained from conventional broilers, conventional turkeys, organic broilers, and organic turkeys, respectively.
According to the farmers, no antimicrobial agents were used in organic broiler or turkey operations from which the samples were collected. In contrast, antimicrobial agents were used in almost every conventional poultry farm. For conventionally-raised broilers, gentamicin was the most commonly used antibiotic. This antimicrobial agent was given to the birds at the hatchery to prevent early mortality due to E. coli infections. Lincomycin and amprolium were also used to control necrotic enteritis and coccidiosis in some
155 conventional broiler farms, respectively. In addition, bacitracin and virginiamycin, which were supplemented in broiler feed at sub-therapeutic levels in order to promote growth and improve feed efficiency, were also used in the conventional broiler farms in this study. For the conventional broiler flocks surveyed in this study, the birds were not exposed to treatments with fluoroquinolone antibiotics during the production period according to information obtained from the producers. For conventionally-raised turkeys, enrofloxacin was the drug routinely used for flocks with respiratory disease due to E. coli infections, while chlortetracycline was used only in the farms that had fowl cholera problem. Like conventional broiler production system, bacitracin was also used as feed additive for conventionally-raised turkeys.
Bacterial isolation and identification. The intestinal tracts were placed on ice and brought back to the laboratory within 3 hours of collection and cultured for C. jejuni and C. coli.
Each intestine was aseptically opened and the cecal contents were directly streaked onto
Campy CVA agar (BBL Becton Dickinson Microbiology Systems, Cockeysville, MD). The inoculated plates were then incubated at 42 °C for 48 hours under a microaerophilic environment (approximately 5% O2, 10% CO2, and 85% N2) in a CampyPak II anaerobic system jar with CampyPak gas generating system envelopes (BBL Becton Dickinson
Microbiology Systems, Sparks, MD). Suspect Campylobacter colonies were identified by colony morphology characteristics, Gram-stain, oxidase test, catalase test, and
Campylobacter culture-plate latex agglutination confirmation test (INDX-Campy [jcl];
PanBio INDX, Inc., Baltimore, MD). In addition, the hippurate hydrolysis test was also
156 performed to differentiate between C. jejuni and C. coli. From each Campylobacter- positive sample, a single colony was used for antibiotic susceptibity tests. All
Campylobacter isolates were stored in sterile cryovial tubes containing skim milk and 30% glycerol at –85 °C prior to antimicrobial susceptibility test.
Antimicrobial susceptibility testing. A total of 694 C. jejuni and C. coli isolates from conventional and organic poultry farms were tested for antimicrobial resistance to nine antimicrobial agents including ampicillin, tetracycline, gentamicin, kanamycin, clindamycin, erythromycin, ciprofloxacin, norfloxacin, and nalidixic acid by the agar dilution method (NCCLS, 2002a). All antimicrobial agents were obtained from Sigma
Chemical Co., St. Louis, MO except ciprofloxacin (Serologicals Proteins, Inc., Kankakee,
IL). The concentrations of most antimicrobial agents tested in this study ranged from 0.06 to 128 µg/ml except for ciprofloxacin (0.008 to128 µg/ml) and for kanamycin and nalidixic acid (0.25 to 128 µg/ml) (Table 2.1). Briefly, Campylobacter isolates, grown on blood agar
plates for at least 24 hours, were inoculated onto Mueller-Hinton (MH) broth (BBL Becton
Dickinson Microbiology Systems, Cockeysville, MD) and then adjusted to a turbidity
equivalent to 0.5 McFarland standard by a colorimeter (BioMérieux, Inc., Hazelwood,
MO). A multipoint inoculator (a cathra replicator system) with 1 mm pins (Oxoid, Inc.,
Ogdensburg, NY) was used to inoculate approximately 104 CFU of samples onto Mueller-
Hinton agar (BBL Becton Dickinson Microbiology Systems, Cockeysville, MD) containing
a two-fold concentration series of antibiotics and supplemented with 5% defibrinated sheep blood (Cleveland Scientific, Cleveland, OH). Campylobacter jejuni ATCC 33560 was used
157 as the quality control organism. While QC ranges are not currently available for ampicillin,
kanamycin, clindamycin, and norfloxacin, the MIC results for these drugs with C. jejuni
ATCC 33560 were consistent, falling within a three-dilution range throughout the study.
The inoculated plates were incubated in a Forma Series II water-jacketed CO2 incubator
(Thermo Electron Corporation, Marietta, OH) at 42 °C for 24 hours under a microaerobic atmosphere of 5% O2, 10% CO2, and 85% N2. The MIC was defined as the lowest
concentration of antimicrobial agent that completely inhibited the visible growth on the
plates. The resistance breakpoints for each antimicrobial agent were as follows: ≥ 4 µg/ml
for ciprofloxacin and clindamycin; ≥ 8 µg/ml for erythromycin; ≥ 16 µg/ml for tetracycline,
gentamicin, and norfloxacin; ≥ 32 µg/ml for ampicillin and nalidixic acid; and ≥ 64 µg/ml
for kanamycin (Table 2.1) (CDC, 2003; NCCLS, 2002a; NCCLS, 2002b). If an isolate was
resistant to three or more classes of antibiotics, it was defined as multidrug resistance.
Pulsed-field gel electrophoresis (PFGE). To determine the genetic diversity of
Campylobacter isolates from different poultry production systems, sample preparation and
PFGE were performed using the protocol described previously (Huang et al., 2005).
Restriction enzyme KpnI was used for digestion of the gel plugs.
Statistical analysis. A Chi-square (χ2) test at a significance level of P < 0.05 (two-tailed)
with Yates correction for continuity was used for statistical analysis of the data.
158
2.4 RESULTS
Prevelence of Campylobacter. From 345 intestinal tracts of conventionally-raised broilers,
Campylobacter were obtained from 227 (65.80%) intestines. Among these 227
Campylobacter isolates, 220 (96.92%) and 7 (3.08%) were identified as C. jejuni and C.
coli, respectively, on the basis of the hippurate hydrolysis test. For conventionally-raised
turkeys, 299 (83.06%) Campylobacter isolates were obtained from 360 intestines. One
hundred and thirty-seven (45.82%) isolates were classified as C. jejuni, while 162 (54.18%) isolates were classified as C. coli. In terms of the organic poultry production systems, a total of 317 (89.30%) and 201 (87.39%) Campylobacter isolates were obtained from 355 and 230 intestinal tracts of organically-raised broilers and turkeys, respectively. Two hundred and twenty-nine (72.24%) and 88 (27.76%) Campylobacter isolates from organically-raised broilers were identified as C. jejuni and C. coli, respectively, whereas
133 (66.17%) Campylobacter isolates from organically-raised turkeys were identified as C. jejuni and 68 (33.83%) were identified as C. coli. Although Campylobacter spp. could be isolated from every conventional and organic poultry farm, the prevalence of this organism varied among farms. For conventional production systems, the prevalence of
Campylobacter spp. in conventional broiler farms ranged from 44% to 80%, while the prevalence of this organism in conventional turkey farms ranged from 63.33% to 97.78%.
Likewise, the prevalence of Campylobacter spp. ranged from 69.57% to 100% in organic broiler farms and 5.56% to 100% in organic turkey farms.
159
Antimicrobial susceptibility patterns. The MIC distributions of ampicillin, tetracycline, gentamicin, kanamycin, clindamycin, erythromycin, ciprofloxacin, norfloxacin, and nalidixic acid against C. jejuni and C. coli isolates were summarized in Table 2.2 and Table
2.3, respectively. In general, a wider range of MICs were observed with isolates from
conventional farms than from organic farms, except gentamicin, for which the MIC
distributions of Campylobacter strains isolated from both types of productions were
comparable. The MICs at which 50% and 90% of Campylobacter isolates were inhibited
were shown in Table 2.4. The MIC90 of fluoroquinolones (ciprofloxacin and norfloxacin) for the Campylobacter strains isolated from organic poultry farms were 4 to 16 times lower than the resistance breakpoints (Table 2.4). On the contrary, the MIC90 of fluoroquinolones
for Campylobacter strains isolated from conventional poultry farms were 8 times higher
than the resistance breakpoints (Table 2.4). Among Campylobacter strains isolated from
organic poultry farms, the MIC50 and MIC90 of erythromycin against C. coli isolates were
slightly higher than those against C. jejuni isolates, while the MIC50 and MIC90 of clindamycin against C. coli isolates were similar to those against C. jejuni isolates (Table
2.4). In contrast, the MIC50 of erythromycin and clindamycin against C. coli isolated from
conventional poultry farms were more than 32-fold higher than those against C. jejuni
isolates from the same operation types (> 128 µg/ml versus 2 µg/ml for erythromycin and
32 µg/ml versus 1 µg/ml for clindamycin). For the aminoglycoside, Campylobacter strains
isolated from both conventional and organic poultry operations were uniformly susceptible to gentamicin with MIC50 and MIC90 ≤ 1 µg/ml. Interestingly, the MIC50 and MIC90 of
160 kanamycin (8 µg/ml and > 128 µg/ml, respectively) against C. jejuni isolates from both conventional and organic poultry farms were the same, while the MIC50 against C. coli
isolates from conventional poultry farms was more than 16 times higher than that against
the C. coli isolates from organic poultry farms (> 128 µg/ml versus 8 µg/ml). In this study,
tetracycline showed poor activity against Campylobacter strains isolated from both
conventional and organic poultry farms, although the MIC50 of tetracycline against the
Campylobacter isolates (both C. jejuni and C. coli) from conventional farms was 4 times
higher than that against the isolates from organic farms (Table 2.4). In terms of β-lactam
antibiotics, ampicillin showed moderate activity against Campylobacter isolates from
conventionally-raised and organically-raised poultry. For C. jejuni, the MIC50 and MIC90 of
ampicillin against the isolates from conventional farms were two times higher than those
against the isolates from organic farms, while the MIC50 and MIC90 against C. coli isolates
from conventional farms were 2- and 8 - fold higher, respectively, in comparison with the
organic farms.
Antimicrobial resistance rates. The resistance rates of the Campylobacter strains were
tabulated according to Campylobacter species (C. jejuni and C. coli), production types
(conventional and organic), and poultry species (broiler and turkey) and were
summarized in Tables 2.5 – 2.8. One of the most striking findings in this study was the
difference in fluoroquinolone resistance between Campylobacter strains isolated from
conventional poultry farms and organic poultry farms. Approximately 46% of
Campylobacter strains isolated from conventionally-raised broilers and 67% of
161 Campylobacter strains isolated from conventionally-raised turkeys were resistant to
ciprofloxacin, norfloxacin, and nalidixic acid. In contrast, none of Campylobacter strains
isolated from organically-raised broilers and less than 2% of Campylobacter strains
isolated from organically-raised turkeys were resistant to these antibiotics (Table 2.8).
Besides the high resistance rates to fluoroquinolones, Campylobacter strains isolated
from conventional turkey farms were also highly resistant to erythromycin (79.60%) and
clindamycin (64.18%) (Table 2.8). Among C. coli isolated from conventionally-raised
turkeys, 114 (90.48%) and 93 (73.81%) were resistant to erythromycin and clindamycin,
respectively (Table 2.6). When compared to Campylobacter strains isolated from other
poultry production systems, the isolates from conventional turkey operations were
significantly more resistant to erythromycin and clindamycin (P < 0.001). Regardless of
the sources of isolation, none of the Campylobacter strains tested in this study were resistant to gentamicin. However, 11.38% of Campylobacter strains isolated from conventionally-raised broilers and 76.12% of Campylobacter strains isolated from conventionally-raised turkeys were resistant to kanamycin. Interestingly, 16.97% and
31.06% of Campylobacter strains isolated from organically-raised broilers and turkeys were also resistant to this antimicrobial agent (Table 2.8). A high level of tetracycline resistance was observed among the C. jejuni and C. coli isolates, with resistance rates ranging from 50% - 92% in various production types (Table 2.8). For ampicillin, the resistance rates of C. jejuni and C. coli varied among the operation types, with the resistance rate in Campylobacter strains isolated from conventional turkey farms significantly higher (P < 0.001) than the resistance rates found in Campylobacter strains
162 isolated from other production systems (Table 2.8). Overall, the antimicrobial resistance
rates in Campylobacter isolates from organic poultry farms were significantly lower than
those from conventional poultry farms and the highest antibiotic resistance rates were
observed in conventional turkey operation (Table 2.8).
Multidrug resistance. The occurrence of multidrug-resistant Campylobacter strains was mainly observed in strains from conventionally-raised turkeys, with 81.09% of these
isolates showing resistance to three or more classes of antibiotics (Table 2.9). In
comparison with the conventional turkey production, the multidrug resistance rates were
significantly lower (P < 0.001) in Campylobacter strains isolated from other poultry production systems. Among multidrug-resistant Campylobacter strains isolated from conventionally-raised turkeys, 77 isolates (47.24%) were resistant to tetracycline, kanamycin, clindamycin, erythromycin, ciprofloxacin, norfloxacin, and nalidixic acid
(Table 2.10). In addition, 117 (58.21%) of Campylobacter strains isolated from this operation type were also resistant to both erythromycin and ciprofloxacin, the preferred antimicrobials for the treatment of human campylobacteriosis, whereas none of
Campylobacter strains isolated from conventionally-raised broilers and organically-raised broilers and turkeys were concomitantly resistant to both of these antibiotics.
PFGE profiles. Representative C. jejuni and C. coli isolates from different poultry operation types were analyzed by PFGE to determine the genetic diversity of the isolates.
The PFGE patterns generated by KpnI digestion showed great differences among the
163 strains isolated from different farms, while the isolates from a single poultry farm tended
to have similar or identical PFGE patterns (Fig. 2.1). Interestingly, a similar PFGE type
was observed in isolates from two different conventional broiler farms, H (Lane 5; Fig.
2.1) and J (Lane 6 and 7; Fig. 2.1), indicating that both farms harbored a genetically- related Campylobacter strain. This PFGE result revealed diverse genotypes among
Campylobacter species isolated from the conventional and organic poultry operations.
2.5 DISCUSSION
In this study, it is clearly shown that thermophilic Campylobacter is highly
prevalent in both organic and conventional poultry production systems. However, the
antimicrobial resistance rates vary significantly in different production types. In general,
the conventional broiler and turkey farms harbor more antibiotic-resistant Campylobacter
strains than organic poultry farms and the differences are especially obvious with
fluoroquinolones. Little resistance was detected in Campylobacter strains isolated from
organic poultry farms, whereas large numbers (45% - 67%) of the isolates from
conventional broiler and turkey farms were resistant to these antimicrobials. Multidrug
resistance to three or more classes of antibiotics in Campylobacter is high among the
isolates from conventional turkey operations, which may be explained by relatively more
antibiotic usage in turkey operations than in broiler operations due to the long production
cycle of meat turkey and the need for more frequent antibiotic treatments of diseases.
The prevalence of Campylobacter species was significantly higher in organically-
raised broilers than in conventionally-raised broilers (P < 0.001), while the prevalence of
164 this organism in organically-raised turkeys was not significantly different from that in conventionally-raised turkeys (P = 0.19). The high frequency of Campylobacter strains isolated from organic broiler production system seems to be associated with the increased age of the birds at slaughter since the average age of these organically-raised broilers at slaughter were about 8 to 12 weeks old compared to 6 weeks old for conventionally- raised broilers. On the other hand, both conventionally-raised and organically-raised turkeys were sent to the processing plant around the same age (18 to 20 weeks). This may explain why the prevalence of Campylobacter strains isolated from conventional and organic turkey farms was not significantly different. The association between
Campylobacter colonization and age of the birds at slaughter was also noted by other studies (Berndtson et al., 1996b; Evans and Sayers, 2000; Newell and Wagenaar, 2000;
Northcutt et al., 2003), which indicated that prevalence of Campylobacter in poultry elevated when the age of birds at the processing plant increased. In terms of the high prevalence of Campylobacter in organic poultry production systems, similar findings were also noted by other studies conducted in free-range and/or organic poultry operations (Avrain et al., 2003; Heuer et al., 2001; Kazwala et al., 1993).
Among Campylobacter-positive flocks, C. jejuni was the predominant species in both conventional and organic broiler production systems especially in conventional broiler farms, where almost 97% of Campylobacter isolates from our study were C. jejuni. This finding is in accordance with previous studies by other research groups, who reported that 85% to 98% of conventional broiler flocks were colonized by C. jejuni
(Avrain et al., 2003; Berndtson et al., 1996b; Evans and Sayers, 2000; Hald et al., 2000;
165 Heuer et al., 2001; Jorgensen et al., 2002; Nielsen et al., 1997; Wedderkopp et al., 2000).
In this study, the distribution of C. jejuni and C. coli in conventional turkey production system is remarkably different from that in conventional broiler production system.
About 46% of Campylobacter isolates from conventionally-raised turkeys were C. jejuni, whereas 54% of these isolates were C. coli. This finding is different from the previous study by Wallace et al. (1998), who reported that almost 100% of Campylobacter isolates from conventional turkey flocks were C. jejuni. In contrast, Smith et al. (2004) revealed that 80% - 90% of Campylobacter strains that colonized turkey flocks were C. coli. For organic poultry production systems, the proportion of C. jejuni and C. coli among
Campylobacter isolates from organic broiler and organic turkey farms was not significantly different from each other (P = 0.171), with C. jejuni as the predominant
Campylobacter species.
Although no fluoroquinolones were used in the conventional broiler flocks from which the samples were collected, we found that about 46% of Campylobacter isolates from conventionally-raised broilers were resistant to these antimicrobial agents (Table
2.8). The high resistance rate to fluoroquinolones may be associated with the previous use of these antibiotics on the broiler farms. Several studies have shown that
Campylobacter displays a hypermutable phenotype to fluoroquinolone treatment in chickens and the treatment results in rapid occurrence of fluoroquinolone-resistant
Campylobacter in the treated chickens (Griggs et al., 2005; Jacobs-Reitsma et al., 1994b;
Luo et al., 2003; McDermott et al., 2002). Epidemiological surveys also suggested that certain quinolone-resistant clones were stable and able to persist on the farms during
166 several rotations even there had been no selective pressure on that farm for a long period of time (Pedersen and Wedderkopp, 2003; Price et al., 2005). Using clonally-related isolates and isogenic mutants, Luo et al. (2005) recently demonstrated that fluoroquinolone-resistant Campylobacter persisted and outcompeted fluoroquinolone- susceptible Campylobacter in the absence of antibiotic usage. Together, these findings underscore the difficulty in eliminating fluoroquinolone-resistant Campylobacter from poultry production. Since no fluoroquinolones were used in organic poultry operations, it is not surprising that there were little or no resistance to this class of antibiotics in
Campylobacter strains isolated from organic poultry farms. The few fluoroquinolone- resistant Campylobacter isolates in organic turkey operation were likely the result of the transmission of fluoroquinolone-resistant Campylobacter from other sources.
In this study, a high level of erythromycin resistance was mainly observed in
Campylobacter strains isolated from conventional turkey farms (Table 2.8). About 61% of C. jejuni and 90% of C. coli isolates from conventionally-raised turkeys were resistant to erythromycin (Table 2.6). This finding is consistent with the previous study by Ge et al. (2003), who reported that erythromycin resistance rates among Campylobacter strains isolated from processed turkeys were significantly higher than those from chickens.
Consistent with findings in previously published work (Avrain et al., 2003; Ge et al.,
2003; Li et al., 1998; Saenz et al., 2000; Van Looveren et al., 2001), erythromycin resistance was much more common in C. coli than in C. jejuni (Table 2.7). In addition, almost 80% of erythromycin-resistant Campylobacter isolates were also resistant to clindamycin, while all but two clindamycin-resistant Campylobacter strains were
167 resistant to erythromycin. A co-resistance between these two antibiotics was also
observed in previous reports by others (Li et al., 1998; Saenz et al., 2000; Taylor and
Courvalin, 1988). Similar to erythromycin, the prevalence of clindamycin resistance in
our study was mainly observed among C. coli especially the isolates from
conventionally-raised turkeys.
Interestingly, none of C. jejuni and C. coli isolates from both conventionally-
raised and organically-raised broilers and turkeys in our study was resistant to gentamicin. This finding is in agreement with the previous studies by other research groups, who reported that no gentamicin resistance was observed among poultry isolates
(Aarestrup et al., 1997; Li et al., 1998; Luber et al., 2003b). However, in some countries
such as Spain, about 25% of Campylobacter strains isolated from broilers were resistant
to this antimicrobial (Saenz et al., 2000). Although gentamicin was the most commonly
used antibiotic in the conventional broiler production system in this study, it was given to
the birds at the hatchery by subcutaneous injection in the neck region. Since
Campylobacter is usually not present in the intestinal tracts of the birds during the first
week of life, the injection of gentamicin may have very limited impact on selection of
gentamicin resistance in Campylobacter. In terms of other aminoglycoside antibiotics, we
found that Campylobacter isolates particularly the ones from turkeys appeared to be
highly resistant to kanamycin (Table 2.8). In addition, we also noticed that kanamycin
resistance seemed to be more prevalent among C. coli than C. jejuni (Table 2.7).
Although 16.97% and 31.06% of Campylobacter strains isolated from organic broiler and
turkey operations were resistant to kanamycin (Table 2.8), the resistant isolates were
168 limited only two organic poultry operations. Similarly, all kanamycin-resistant
Campylobacter isolates from conventionally-raised broilers were from a single
conventional broiler farm. In contrast, kanamycin-resistant Campylobacter was common on different conventional turkey farms. The reason for this skewed distribution of kanamycin-resistant Campylobacter on conventional broiler and organic poultry farms is unclear at this stage.
High levels of tetracycline resistance among Campylobacter isolates were observed from both conventionally-raised and organically-raised broilers and turkeys
(Table 2.8). Although tetracycline had never been used in those organic farms, tetracycline-resistant Campylobacter strains were present in almost every organic poultry farm surveyed in this study. This widespread distribution of tetracycline resistance is different from that of kanamycin resistance for which only some farms showed resistant strains. Since tetracyclines have been used as feed additives for livestock and poultry for both therapeutic and subtherapeutic purposes for long period of time (Chopra and
Roberts, 2001; Fallon et al., 2003; Trieber and Taylor, 2000), it is possible that
Campylobacter may have evolutionally become resistant to this class of antibiotic, leading to the widespread distribution of tetracycline-resistant Campylobacter in animal reservoirs regardless of the production types. The high prevalence of tetracycline- resistant Campylobacter in poultry was also reported in other studies (Aarestrup et al.,
1997; Avrain et al., 2001; Fallon et al., 2003; Li et al., 1998; Luber et al., 2003b; Saenz et al., 2000; Van Looveren et al., 2001). For human Campylobacter isolates, the frequency
of
169 of tetracycline resistance showed a wide variation in different geographical regions ranging from 11% in Denmark to 95% in Taiwan (Aarestrup et al., 1997; Gaudreau and
Gilbert, 2003; Li et al., 1998; Luber et al., 2003b).
The ampicillin resistance rates reported in this study are in agreement with the resistance rates reported by other studies (Aarestrup et al., 1997; Avrain et al., 2001;
Fallon et al., 2003; Luber et al., 2003b). It is noticed that ampicillin resistance is more prevalent in conventional turkey operations than in other production types (Table 2.8). In addition, it appears that ampicillin resistance is more common in C. jejuni than in C. coli and more prevalent among turkey isolates than broiler isolates (Table 2.5 – 2.6).
This study also revealed a high frequency of multidrug resistance (81.09%) among Campylobacter strains isolated from conventionally-raised turkeys. On the other hand, less than 10% of Campylobacter strains isolated from other poultry production systems were resistant to three or more classes of antibiotics (Table 2.9). Multidrug- resistant strains among Campylobacter isolates from conventionally-raised turkeys were also reported by Lee et al. (2005). Interestingly, none of Campylobacter isolates from conventionally-raised broilers and organically-raised broilers and turkeys were resistant to both macrolides (e.g. erythromycin) and fluoroquinolones (e.g. ciprofloxacin), whereas more than 58% of C. jejuni and C. coli isolates from conventionally-raised turkeys were concomitantly resistant to these antibiotics. If these resistant turkey isolates transmit to humans, the concomitant resistance of the isolates to both macrolide and fluoroquinolone can potentially compromise the efficacy of antibiotic treatment because erythromycin and fluoroquinolones are the drugs of choice for the treatment of food-borne
170 campylobacteriosis in humans (Allos, 2001). Although the exact reason for multidrug
resistance in Campylobacter isolated from conventionally-raised turkeys is unknown, it is
possible that this phenotype is associated with the frequent antibiotic usage in turkey
production.
In summary, this study revealed significant differences in antimicrobial-resistant
Campylobacter in different poultry production systems. The results suggest that the
practice of antimicrobial usage in poultry production systems influences the prevalence
of antibiotic-resistant Campylobacter in poultry. However, antimicrobial usage alone
may not solely be responsible for the increased antibiotic resistance in Campylobacter
because even in the absence of antibiotic exposure, a high level of tetracycline resistance
was observed in organic poultry. Similarly, the resistance rates to fluoroquinolones were also high in the surveyed conventional broiler flocks which did not get exposed to the
class of antibiotics during the entire production period. These observations suggest that
antibiotic-resistant Campylobacter are stable and able to transmit and persist in poultry
even in the absence of selection pressure. Together, these findings reveal the complex
nature in the spread of antibiotic resistance and further highlight the need for prudent
measures to prevent the occurrence and transmission of antibiotic-resistant
Campylobacter in the poultry reservoir.
171
2.6 ACKNOWLEDGEMENT
The authors would like to thank Ms. Sonya M. Bodeis at the Center for Veterinary
Medicine, Food and Drug Administration for her technical assistance in this study. The
authors would also like to thank Dr. Amna B. El-Tayeb, Ms. Elisabeth J. Angrick, and fellow colleagues at the Avian Disease Investigation Laboratory at The Ohio State
University for their help, advice, and technical support.
This work was supported by National Research Initiative Competitive Grants 00 -
51110 - 9741 and 2003 - 35212 – 13316 (to QZ and TYM) from the USDA Cooperative
State Research, Education, and Extension Service and grant 2003 - 38640 - 13225 from the North Central Region program for Sustainable Agriculture Research and Education
(NCR-SARE).
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178
MIC quality Agar e control ranges of MIC breakpoints (µg/ml) Antimicrobial dilution C. jejuni agents test range ATCC 33560c (µg/ml) (µg/ml) S I R
Ampicillina 0.06 - 128 N/Ad ≤ 8 16 ≥ 32 Tetracyclinea 0.06 - 128 1 - 4 ≤ 4 8 ≥ 16 Gentamicina 0.06 - 128 0.5 - 4 ≤ 4 8 ≥ 16 Kanamycina 0.25 - 128 N/A ≤ 16 32 ≥ 64 a Clindamycin 0.06 - 128 N/A ≤ 0.5 1-2 ≥ 4 179 Erythromycina 0.06 - 128 1 - 8 ≤ 0.5 1-4 ≥ 8 Ciprofloxacina 0.008 - 128 0.06 - 0.5 ≤ 1 2 ≥ 4 Norfloxacinb 0.06 - 128 N/A ≤ 4 8 ≥ 16 Nalidixic acida 0.25 - 128 8 - 32 ≤ 16 ≥ 32
a MIC breakpoints for enteric bacteria used by the National Antimicrobial Resistance Monitoring System (NARMS) b MIC breakpoints for Enterobacteriaceae recommended by the National Committee for Clinical Laboratory Standard (NCCLS) c Tentative agar dilution quality control ranges of C. jejuni ATCC 33560 approved by the NCCLS d No data available e S, susceptible; I, intermediate; R, resistant
Table 2.1 Antimicrobial test ranges, MIC quality control ranges, and MIC breakpoints used for antimicrobial susceptibility testing
179
Antimicrobial Operation type Number of isolates inhibited by MIC (µg/ml)a agents 0.06 0.125 0.25 0.5 1 2 4 8 16 32 64 128 >128 Ampicillin conventionalb 0 0 0 0 0 5 98 74 33 22 2 4 2 organicc 0 0 0 0 1 39 80 24 56 4 4 3 0
Tetracycline conventional 0 8 5 2 6 0 2 4 12 60 51 66 24 organic 0 7 35 42 13 4 0 0 7 15 25 28 35
Gentamicin conventional 0 0 23 93 122 2 0 0 0 0 0 0 0 organic 0 0 39 100 72 0 0 0 0 0 0 0 0
Kanamycin conventional - - 1 0 0 9 72 91 3 1 0 1 62 organic - - 0 0 6 20 69 86 1 0 0 0 29
Clidamycin conventional 0 0 1 37 127 38 4 3 1 25 4 0 0 organic 0 0 1 65 110 30 0 0 1 4 0 0 0 180 Erythromycin conventional 0 0 1 3 54 107 29 4 4 1 1 2 34 organic 0 2 1 37 71 78 17 0 0 0 0 0 5
Ciprofloxacin conventional 0 45 37 19 10 1 0 20 80 28 0 0 0 organic 1 51 65 49 38 5 0 0 0 0 2 0 0
Norfloxacin conventional 0 0 9 58 34 8 0 2 0 1 48 75 5 organic 0 5 77 74 41 7 5 0 0 0 0 0 2
Nalidixic acid conventional - - 0 0 2 3 56 29 20 0 1 22 107 organic - - 0 0 0 7 135 60 7 0 0 0 2 a Thin vertical lines indicate the breakpoint between susceptible and intermediate strains. Thick vertical lines indicate the breakpoint between intermediate and resistant strains (except for Nalidixic acid that indicates between susceptible and resistant strains). b C. jejuni isolates from conventional poultry farms (n = 240) c C. jejuni isolates from organic poultry farms (n = 211)
Table 2.2 MIC distributions of C. jejuni isolated from conventional and organic poultry farms
180
Antimicrobial Operation type Number of isolates inhibited by MIC (µg/ml)a agents 0.06 0.125 0.25 0.5 1 2 4 8 16 32 64 128 >128 Ampicillin conventionalb 0 0 0 0 0 0 5 58 32 9 2 19 3 organicc 0 0 0 0 0 0 20 41 50 0 0 4 0
Tetracycline conventional 0 0 0 1 7 4 2 0 1 0 13 63 37 organic 0 0 3 19 21 2 0 0 2 14 1 7 46
Gentamicin conventional 0 0 2 18 107 1 0 0 0 0 0 0 0 organic 0 0 7 37 71 0 0 0 0 0 0 0 0
Kanamycin conventional - - 0 0 0 1 4 9 5 0 0 0 109 organic - - 0 0 0 0 10 50 5 1 0 0 49
Clidamycin conventional 0 0 0 5 21 9 4 7 4 72 6 0 0 organic 0 0 0 52 35 19 1 0 1 5 2 0 0
Erythromycin conventional 0 0 2 0 1 3 8 8 6 5 1 2 92 181 organic 0 1 13 17 10 35 24 7 0 0 0 0 8
Ciprofloxacin conventional 0 3 14 10 17 0 1 2 13 60 7 1 0 organic 0 4 38 29 39 4 0 0 0 0 1 0 0
Norfloxacin conventional 0 0 3 12 20 9 0 2 0 1 37 41 3 organic 0 0 4 70 35 4 1 0 0 0 0 0 1
Nalidixic acid conventional - - 0 0 0 0 5 24 17 0 1 22 59 organic - - 0 0 0 0 61 45 8 0 0 0 1 a Thin vertical lines indicate the breakpoint between susceptible and intermediate strains. Thick vertical lines indicate the breakpoint between intermediate and resistant strains (except for Nalidixic acid that indicates between susceptible and resistant strains). b C. coli isolates from conventional poultry farms (n = 128) c C. coli isolates from organic poultry farms (n = 115)
Table 2.3 MIC distributions of C. coli isolated from conventional and organic poultry farms
181
MIC /MIC (µg/ml) of Campylobacter strains Agar 50 90 MIC Antimicrobial dilution breakpoints C. jejuni C. coli agents test range (µg/ml) (µg/ml) Conventional Organic Conventional Organic (n=240) (n=211) (n=128) (n=115) Ampicillin 0.06 - 128 ≥ 32 8/32 4/16 16/128 8/16 Tetracycline 0.06 - 128 ≥ 16 64/128 16/>128 128/>128 32/>128 Gentamicin 0.06 - 128 ≥ 16 1/1 0.5/1 1/1 1/1 Kanamycin 0.25 - 128 ≥ 64 8/>128 8/>128 >128/>128 8/>128 Clindamycin 0.06 - 128 ≥ 4 1/32 1/2 32/32 1/2 Erythromycin 0.06 - 128 2/>128 1/4 >128/>128 2/8
182 ≥ 8 Ciprofloxacin 0.008 - 128 ≥ 4 8/32 0.25/1 32/32 0.5/1 Norfloxacin 0.06 - 128 ≥ 16 64/128 0.5/1 64/128 0.5/1 Nalidixic acid 0.25 - 128 ≥ 32 128/>128 4/8 128/>128 4/8
Table 2.4 MICs of nine antimicrobial agents against C. jejuni and C. coli isolated from conventional and organic poultry farms
182 Number (%) of resistant Campylobacter strains Antimicrobial C. jejuni C. coli agents Conventional Organic Conventional Organic (n=165) (n=108) (n=2)† (n=57) Ampicillin 0a* 4 (3.7)b 0 1 (1.75)a,b
Tetracycline 139 (84.24)a 69 (63.89)b 2 (100) 30 (52.63)b
Gentamicin 0 0 0 0
Kanamycin 17 (10.30)a 12 (11.11)a 2 (100) 16 (28.07)b
Clindamycin 1 (0.61)a 3 (2.78)a 1 (50) 6 (10.53)a,b
Erythromycin 0a 3 (2.78)a 0 12 (21.05)b
Ciprofloxacin 76 (46.06)a 0b 0 0b
Norfloxacin 77 (46.67)a 0b 0 0b
Nalidixic acid 77 (46.67)a 0b 0 0b
† C. coli isolates from conventional broiler farms are excluded from chi-square analysis because of low sample size. * Numbers in the same row with different superscripts are significantly different (P<0.05); numbers with the same superscript do not differ significantly (a chi-square test with Yates correction for continuity).
Table 2.5 Resistance rates of C. jejuni and C. coli isolated from conventional and organic broiler farms
183 Number (%) of resistant Campylobacter strains Antimicrobial C. jejuni C. coli agents Conventional Organic Conventional Organic (n=75) (n=103) (n=126) (n=58) Ampicillin 30 (40)a* 7 (6.80)b 33 (26.19)a 3 (5.17)b
Tetracycline 74 (98.67)a 41 (39.81)b 112 (88.89)c 40 (68.97)d
Gentamicin 0 0 0 0
Kanamycin 46 (61.33)a 17 (16.50)b 107 (84.92)c 33 (59.90)a
Clindamycin 36 (48)a 2 (1.94)b 93 (73.81)c 3 (5.17)b
Erythromycin 46 (61.33)a 2 (1.94)b 114 (90.48)c 3 (5.17)b
Ciprofloxacin 52 (69.33)a 2 (1.94)b 84 (66.67)a 1 (1.72)b
Norfloxacin 52 (69.33)a 2 (1.94)b 82 (65.08)a 1 (1.72)b
Nalidixic acid 53 (70.67)a 2 (1.94)b 82 (65.08)a 1 (1.72)b
* Numbers in the same row with different superscripts are significantly different (P<0.05); numbers with the same superscript do not differ significantly (a chi-square test with Yates correction for continuity).
Table 2.6 Resistance rates of C. jejuni and C. coli isolated from conventional and organic turkey farms
184 Number (%) of resistant Campylobacter strains Antimicrobial C. jejuni C. coli agents Conventional Organic Conventional Organic (n=240) (n=211) (n=128) (n=115) Ampicillin 30 (12.5)a* 11 (4.58)b 33 (25.78)c 4 (3.49)b
Tetracycline 213 (88.75)a 110 (52.13)b 114 (89.06)a 70 (60.87)b
Gentamicin 0 0 0 0
Kanamycin 63 (26.25)a 29 (13.74)b 109 (85.16)c 49 (42.61)d
Clindamycin 37 (15.42)a 5 (2.37)b 93 (72.66)c 9 (7.83)a
Erythromycin 46 (19.17)a 5 (2.37)b 114 (89.06)c 15 (13.04)a
Ciprofloxacin 128 (53.33)a 2 (0.95)b 84 (65.63)c 1 (0.87)b
Norfloxacin 129 (53.75)a 2 (0.95)b 82 (64.06)c 1 (0.87)b
Nalidixic acid 130 (54.17)a 2 (0.95)b 82 (64.06)c 1 (0.87)b
* Numbers in the same row with different superscripts are significantly different (P<0.05); numbers with the same superscript do not differ significantly (a chi-square test with Yates correction for continuity).
Table 2.7 Resistance rates of C. jejuni and C. coli isolated from conventional and organic poultry farms
185 Number (%) of resistant Campylobacter strains isolated from Antimicrobial Conventional Conventional Organic broiler Organic turkey agents broiler farms turkey farms farms (n=165) farms (n=161) (n=167) (n=201) Ampicillin 0a* 5 (3.03)a 63 (31.34)b 10 (6.21)a
Tetracycline 141 (84.43)a 99 (60)b 186 (92.54)c 81 (50.31)b
Gentamicin 0 0 0 0
Kanamycin 19 (11.38)a 28 (16.97)a 153 (76.12)b 50 (31.06)c
Clindamycin 2 (1.20)a 9 (5.45)a 129 (64.18)b 5 (3.11)a
Erythromycin 0a 15 (9.09)b 160 (79.60)c 5 (3.11)d
Ciprofloxacin 76 (45.51)a 0b 136 (67.66)c 3 (1.86)b
Norfloxacin 77 (46.11)a 0b 134 (66.67)c 3 (1.86)b
Nalidixic acid 77 (46.11)a 0b 135 (67.16)c 3 (1.86)b
* Numbers in the same row with different superscripts are significantly different (P<0.05); numbers with the same superscript do not differ significantly (a chi-square test with Yates correction for continuity).
Table 2.8 Resistance rates of Campylobacter strains isolated from different poultry production systems
186
Number (%) of Campylobacter strains resistant to different groups of Number (%) of antibiotics Operation types multidrug-resistant 0 1 2 3 4 5 6 Campylobacter strains
Conventional 9 94 49 15 broiler farms - - - 15 (8.98) (5.39) (56.29) (29.34) (8.98) (n=167)
Organic broiler 56 83 15 2 9 - - 11 (6.67) farms (n=165) (33.94) (50.30) (9.09) (1.21) (5.45)
Conventional 13 25 23 30 87 23 turkey farms - 163 (81.09) 187 (n=201) (6.47) (12.44) (11.44) (14.93) (43.28) (11.44)
Organic turkey 55 74 24 2 4 2 - 8 (4.97) farms (n=161) (34.16) (45.96) (14.91) (1.24) (2.48) (1.24)
Table 2.9 Multidrug resistance of Campylobacter strains isolated from conventional and organic poultry farms
187 Number (%) of resistant Operation types Major resistance patternsa Campylobacter strains C. jejuni C. coli Conventional broiler TET-KAN-CIP/NOR/NAL 15 (8.98) 0 farms (n=167)
Organic broiler farms TET-KAN-CLI-ERY 3 (1.82) 6 (3.64) (n=165) AMP-TET-KAN 1 (0.61) 1 (0.61)
TET-KAN-CLI-ERY- 19 (9.45) 58 (28.86) CIP/NOR/NAL
AMP-TET-KAN-CLI-ERY- 7 (3.48) 16 (7.96) CIP/NOR/NAL
AMP-TET-CLI-ERY 5 (2.49) 3 (1.49)
AMP-TET-KAN-ERY- 4 (1.99) 3 (1.49) CIP/NOR/NAL
Conventional turkey TET-KAN-CLI-ERY 0 7 (3.48) farms (n=201) TET-KAN-ERY- 4 (1.99) 3 (1.49) CIP/NOR/NAL
AMP-TET-CIP/NOR/NAL 5 (2.49) 1 (0.50)
TET-KAN-CIP/NOR/NAL 5 (2.49) 0
AMP-TET-KAN-CLI-ERY 0 3 (1.49)
TET-KAN-ERY 2 (0.99) 1 (0.50)
Organic turkey farms TET-KAN-CLI-ERY 1 (0.62) 2 (1.24) (n=161) aAMP, ampicillin; CIP, ciprofloxacin; CLI, clindamycin; ERY, erythromycin; KAN, kanamycin; NAL, nalidixic acid; NOR, norfloxacin; TET, tetracycline
Table 2.10 Major multidrug resistance patterns of C. jejuni and C. coli isolated from conventional and organic poultry farms
188
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Figure 2.1. PFGE patterns of Campylobacter strains isolated from conventional broiler and turkey farms and from organic broiler and turkey farms. Lane 1 and 15, molecular weight marker (Promega-makers® Lambda ladders); lane 2, C. coli isolate from conventional turkey farm A; lane 3, C. coli isolate from conventional turkey farm B; lane 4, C. jejuni isolate from organic turkey farm D; lane 5, C. jejuni isolate from conventional broiler farm H; lane 6-8, C. jejuni isolates from conventional broiler farm J; lane 9-10, C. coli isolates from organic broiler farm A; lane 11, C. jejuni isolate from organic broiler farm A; lane 12-13, C. coli isolates from organic broiler farm C; lane 14, C. jejuni isolate from organic broiler farm C.
189
CHAPTER 3
ANTIMICROBIAL SUSCEPTIBILITY TESTING OF CAMPYLOBACTER BY
THE AGAR DILUTION AND THE AGAR DISK DIFFUSION METHODS
3.1 ABSTRACT
The agreement between the agar disk diffusion method and the standardized agar
dilution method for antimicrobial susceptibility testing of Campylobacter was
investigated. Six hundred and sixty-eight Campylobacter strains (431 C. jejuni and 237
C. coli) isolated from intestinal tracts of healthy slaughter-age broilers and turkeys with
different histories of antibiotic exposure were included in this study. In general, if only
susceptible and resistant Campylobacter strains were included in the data analysis, the
percent agreement between the agar dilution method and the disk diffusion method in
identification of susceptible and resistant Campylobacter isolates would be very high
ranging from 87.79% for ampicillin to 99.85% for gentamicin with the average percent
agreement around 95.72% for other antibiotics. In addition, the kappa statistic also
showed almost perfect agreement (kappa > 0.8) between these two tests for almost every class of antibiotics except for ampicillin and tetracycline. Because of a high level of agreement between the agar dilution and the agar disk diffusion methods in determining fluoroquinolones (e.g. ciprofloxacin) and aminoglycosides (e.g. gentamicin) susceptible
190 or resistant Campylobacter isolates, the agar disk diffusion method is likely to be an
interesting alternative for antimicrobial susceptibility testing of thermophilic
Campylobacter to these classes of antimicrobial agent. However, until the standard
resistance breakpoints specific for Campylobacter is established and validated, the agar
dilution method should be used to determine antimicrobial resistance of Campylobacter
spp. to ampicillin, tetracycline, clindamycin, and erythromycin.
3.2 INTRODUCTION
Campylobacter species particularly C. jejuni has been recognized as an important
cause of food-borne bacterial diarrhea in human worldwide (Allos, 2001). This organism
is generally carried in the intestinal tracts of food animals especially poultry and it is
often present on food of animal origin through fecal contamination during processing
(Jacobs-Reitsma, 2000). Although most patients with Campylobacter infections do not
require treatment, antimicrobial therapy seems to be essential in patients with systemic,
severe or prolonged infections (Allos, 2001; Blaser, 1997). In this circumstance,
macrolides (e.g. erythromycin) and fluoroquinolones (e.g. ciprofloxacin) are considered
to be the drugs of choice (Allos, 2001; Blaser, 2000). However, other antibiotics such as
gentamicin, tetracycline, clindamycin, and ampicillin may be listed as alternative drugs
for treatment of systemic Campylobacter infections as well (Blaser, 2000). In general,
antimicrobial susceptibility testing prior to treatment of Campylobacter infections may be unnecessary; however, it may be useful especially in the area where the prevalence of antimicrobial resistance in Campylobacter isolates is high. Several antimicrobial
191 susceptibility testing methods including agar dilution, broth microdilution, epsilometer
test (E-test), and disk diffusion test have been used to test for antimicrobial resistance of
Campylobacter species (Alfredson et al., 2003; Engberg et al., 1999; Fernandez et al.,
2000a; Frediani-Wolf and Stephan, 2003; Gaudreau and Gilbert, 1997; Ge et al., 2002;
Huang et al., 1992; Huysmans and Turnidge, 1997; Luber et al., 2003a; McDermott et al.,
2004; Oncul et al., 2003). For the disk diffusion method, an interpretation criterion
indicating whether the isolate is susceptible or resistant to antibiotics is based on the diameter of the zone of inhibition around the antibiotic disk (Caprioli et al., 2000). In contrast, a criterion to interpret whether the isolate is susceptible or resistant to antibiotics by the agar dilution method relies on the growth of the organism on a medium containing a two-fold concentration series of antibiotics (Caprioli et al., 2000). The lowest concentration of antimicrobial agent that completely inhibits the visible growth is defined as the minimal inhibitory concentration (MIC) (Caprioli et al., 2000; NCCLS, 2002a).
Unfortunately, there are no antimicrobial resistance breakpoints specific for
Campylobacter available currently. Hence, the resistance breakpoints of enteric bacteria in the family Enterobacteriaceae have been used to determine antimicrobial resistance of
Campylobacter spp. (Ge et al., 2002; Luber et al., 2003a; Nachamkin et al., 2000b).
However, the resistance breakpoints for Enterobacteriaceae approved by different organizations, e.g., the National Committee for Clinical Laboratory Standard (NCCLS), the British Society for Antimicrobial Therapy (BSAC), the World Health Organization
(WHO), the Société Francaise de Microbiologie (SFM), the Deutsches Institute fur
Normung (DIN), the Swedish Reference Group for Antibiotics (SIR), and the Werkgroep
192 Richtlijnen Gevoeligheidsbepalingen (WRG) seem to be different (Acar and Goldstein,
1996; Andrews, 2004; NCCLS, 2002a; SFM Antibiogram Committee, 2003). Since there
are no internationally accepted standard resistance breakpoints specific for
Campylobacter species, different resistance breakpoints have been used making a comparison of antimicrobial resistance results from various studies more difficult; for example, the breakpoints for susceptible Campylobacter strains to erythromycin varied
from ≤ 0.5 to ≤ 8 µg/ml, while the breakpoints for erythromycin-resistant Campylobacter
strains ranged from ≥ 2 to ≥ 64 µg/ml (Acar and Goldstein, 1996; Engberg et al., 1999;
Huysmans and Turnidge, 1997; King, 2001; NCCLS, 2002a; SFM Antibiogram
Committee, 2003; Shanker and Sorrell, 1983). In order to compare the antimicrobial
susceptibility patterns of thermophilic Campylobacter isolated from food animals, food
of animal origin, and humans from different laboratories, the same standardized methods
and interpretation criteria should be applied (Engberg et al., 1999; Nachamkin et al.,
2000b). Recently, the agar dilution method is considered as a standard antimicrobial
susceptibility testing method for Campylobacter species and Campylobacter jejuni
ATCC 33560 is recommended to use as a quality control organism (McDermott et al.,
2004; NCCLS, 2002a). Although the agar dilution method is very reliable, highly reproducible, and provides quantitative MIC values, it is labor-intensive, time- consuming, and quite expensive (Caprioli et al., 2000). On the other hand, the agar diffusion test such as disk diffusion method seems to be relatively easy to perform, not
expensive, and can be reproducible if it is conducted carefully with a high level of
standardization and quality control (Caprioli et al., 2000; Potz et al., 2004). Over the last
193 ten years, several comparison studies of the agreement between different antimicrobial
susceptibility testing methods for Campylobacter species have been conducted
(Alfredson et al., 2003; Engberg et al., 1999; Fernandez et al., 2000a; Frediani-Wolf and
Stephan, 2003; Gaudreau and Gilbert, 1997; Ge et al., 2002; Huang et al., 1992;
Huysmans and Turnidge, 1997; Luber et al., 2003a; Oncul et al., 2003). However, only a
few studies have compared the results obtained from the agar dilution and the disk
diffusion methods. Especially, after the standardized agar dilution method for
Campylobacter has been proposed, no information on the agreement between the
standardized agar dilution method and the agar disk diffusion method has been reported
yet. Hence, the aim of this study was to determine whether the agar disk diffusion test
could be used as a reliable method for antimicrobial susceptibility testing of
Campylobacter by analyzing the agreement between antimicrobial resistance results obtained from the disk diffusion test and the results obtained from the standardized agar dilution test.
3.3 MATERIALS AND METHODS
Bacterial isolates. Six hundred and sixty-eight Campylobacter strains isolated from the intestinal tracts of healthy slaughter-age broilers and turkeys with different histories of antibiotic exposure were selected for antimicrobial susceptibility testing. These
Campylobacter strains were isolated during August 2000 to November 2002 by the standard microbiological method and were stored in skim milk and glycerol at -85 °C until the antimicrobial susceptibility test was performed. One isolate per intestinal tract
194 was included in this study. These Campylobacter strains consisted of 431 C. jejuni and
237 C. coli on the basis of the hippurate hydrolysis test. In addition, reference standard bacterial strains including C. jejuni ATCC 33560 and E. coli ATCC 25922 were also
tested concurrently as quality control organisms in this study.
Antimicrobial agents. Nine antimicrobial agents tested in this study included ampicillin,
tetracycline, gentamicin, kanamycin, clindamycin, erythromycin, ciprofloxacin,
norfloxacin, and nalidixic acid. For the agar dilution method, all antimicrobial agents
were obtained from Sigma Chemical Co., St. Louis, MO except ciprofloxacin
(Serologicals Proteins, Inc., Kankakee, IL). The concentrations of most antimicrobial
agents tested in this study ranged from 0.06 to 128 µg/ml except for ciprofloxacin (0.008
to128 µg/ml) and for kanamycin and nalidixic acid (0.25 to 128 µg/ml). For the disk
diffusion method, all antibiotics, e.g., ampicillin (10 µg), tetracycline (30 µg), gentamicin
(10 µg), kanamycin (30 µg), clindamycin (2 µg), erythromycin (15 µg), ciprofloxacin (5
µg), norfloxacin (10 µg), and nalidixic acid (30 µg) were obtained from Becton
Dickinson and Company, Sparks, MD.
Antimicrobial susceptibility testing. All Campylobacter isolates were thawed and resubcultured onto blood agar plates and then incubated at 42 °C for at least 24 hours under a microaerophilic environment (approximately 5% O2, 10% CO2, and 85% N2)
prior to antimicrobial susceptibility testing. A few Campylobacter colonies were
randomly selected and inoculated into Mueller-Hinton (MH) broth (BBL Becton
195 Dickinson Microbiology Systems, Cockeysville, MD) and then adjusted to a turbidity equivalent to 0.5 McFarland standard by a colorimeter (BioMérieux, Inc., Hazelwood,
MO).
For the agar dilution method, approximately 104 CFU of adjusted samples were
applied onto Mueller-Hinton agar (BBL Becton Dickinson Microbiology Systems,
Cockeysville, MD) containing a two-fold concentration series of antibiotics and
supplemented with 5% defibrinated sheep blood (Cleveland Scientific, Cleveland, OH)
by a multipoint inoculator (a cathra replicator system) with 1 mm pins (Oxoid, Inc.,
Ogdensburg, NY). The inoculated plates were incubated in a Forma Series II water-
jacketed CO2 incubator (Thermo Electron Corporation, Marietta, OH) at 42 °C for 24
hours under a microaerophilic condition. The MIC breakpoints of ampicillin, tetracycline,
gentamicin, kanamycin, clindamycin, erythromycin, ciprofloxacin, and nalidixic acid
were determined according to the breakpoints used by the National Antimicrobial
Resistance Monitoring System (NARMS) (CDC, 2003). For norfloxacin, the breakpoints
defined by the National Committee for Clinical Laboratory Standard (NCCLS) for
Enterobacteriaceae were used (NCCLS, 2002a; NCCLS, 2002b). The resistance
breakpoints for each antimicrobial agent were as the followings: ≥ 4 µg/ml for
ciprofloxacin and clindamycin; ≥ 8 µg/ml for erythromycin; ≥ 16 µg/ml for tetracycline,
gentamicin, and norfloxacin; ≥ 32 µg/ml for ampicillin and nalidixic acid; and ≥ 64 µg/ml
for kanamycin (Table 3.1).
For the disk diffusion method, the adjusted samples were evenly streaked in three
directions onto Mueller-Hinton II agar (BBL Becton Dickinson Microbiology Systems,
196 Cockeysville, MD) to produce confluent lawn of bacterial growth by sterile cotton swabs.
Once the inoculum on the plates dried, antibiotic disks were distributed over the inoculated plates using a BBL Sensi-disc dispenser (BBL Becton Dickinson
Microbiology Systems, Cockeysville, MD). These plates were then incubated at 42 °C under the same condition as the agar dilution method. Zone of inhibition around each antibiotic disk was measured in millimeters and recorded. Zone diameter breakpoints indicating susceptible, intermediate, or resistant were determined according to the
NCCLS established guidelines for Enterobacteriaceae and bacteria isolated from animals
(NCCLS, 2002a; NCCLS, 2002b). The resistance breakpoints for each antimicrobial agent were as the followings: ≤ 12 mm for gentamicin and norfloxacin; ≤ 13 mm for
ampicillin, kanamycin, erythromycin, and nalidixic acid; ≤ 14 mm for tetracycline and
clindamycin; and ≤ 15 mm for ciprofloxacin (Table 3.1).
Data analysis. To measure the level of agreement between the results obtained from the
agar dilution method and the disk diffusion method, the percent agreement and the kappa
statistic were calculated as previously described by Luber et al. (2003a) and Dohoo et al.
(2003).
3.4 RESULTS
The results of antimicrobial susceptibility testing of Campylobacter spp. using the
agar dilution and the disk diffusion methods for each antibiotic were summarized in
Table 3.2. According to the resistance breakpoints currently used by the NARMS and the
197 NCCLS, a majority of Campylobacter isolates were classified as either susceptible or
resistant to ciprofloxacin, norfloxacin, gentamicin, and kanamycin by the agar dilution
and by the disk diffusion methods (Table 3.2). For erythromycin, clindamycin, and
ampicillin, a large number of Campylobacter isolates were classified as intermediate to these antimicrobial agents by both methods; for example, 413, 366, and 154
Campylobacter isolates were identified as intermediate by the agar dilution method to erythromycin, clindamycin, and ampicillin, respectively (Table 3.2). Although 49 and 31
Campylobacter isolates were recognized as intermediate to tetracycline and nalidixic acid by the disk diffusion method, respectively, only four Campylobacter isolates were classified as intermediate to tetracycline and none of the isolates was classified as intermediate to nalidixic acid by the agar dilution method (Table 3.2).
The agreement between the agar dilution and the disk diffusion methods for antimicrobial susceptibility testing of thermophilic Campylobacter was summarized in
Table 3.3 – 3.5. In general, the percent identification of resistant isolates by both methods was quite similar except for clindamycin and erythromycin that the percent of resistant strains identified by the agar dilution method was much higher than that identified by the disk diffusion method (76.20% versus 51.20% for clindamycin and 88.62% versus
37.13% for erythromycin) when the intermediate strains reclassified as resistant strains were included in the calculation (Table 3.4). The percent agreement between the agar dilution method and the disk diffusion method in identification of susceptible and resistant isolates ranged from 87.79% for ampicillin to 99.85% for gentamicin with the average percent agreement around 95.72% for other antibiotics when no intermediate
198 strains were included in the data analysis (Table 3.3). However, when the intermediate
strains reclassified as resistant strains were included in the calculation, the percent
agreement between these two methods changed significantly from 47.90% for
erythromycin to 99.85% for gentamicin (Table 3.4). On the other hand, when the
intermediate strains reclassified as susceptible strains were included in the analysis, the
percent agreement between the agar dilution and the disk diffusion methods varied from
83.21% for tetracycline to 99.85% for gentamicin (Table 3.5). In terms of the kappa
value, if only susceptible and resistant Campylobacter strains or the intermediate strains that were reclassified as susceptible strains were included in the kappa calculation, the kappa revealed almost perfect agreement (kappa > 0.8) between the agar dilution and the disk diffusion methods for susceptibility testing of C. jejuni and C. coli to almost every
class of antibiotics tested in this study except for ampicillin and tetracycline (Table 3.3
and 3.5). However, when the intermediate strains reclassified as resistant strains were
included in the kappa analysis, the weak agreement between these two methods for
susceptibility testing of thermophilic Campylobacter to erythromycin, clindamycin, and
ampicillin was observed with kappa 0.13, 0.30, and 0.40, respectively (Table 3.4). For
gentamicin, the kappa value could not be calculated because none of the isolates in this
study was resistant to this antibiotic causing the lack of the essential value for kappa
statistic calculation.
199 3.5 DISCUSSION
When the results of antimicrobial susceptibility testing of Campylobacter isolates
obtained from the disk diffusion method were compared to the results obtained from the
standardized agar dilution method, this study shows that the disk diffusion test may be able to use as a reliable screening method for determining antimicrobial resistance of
Campylobacter isolates from poultry to fluoroquinolones (e.g. ciprofloxacin) and
aminoglycosides (e.g. gentamicin and kanamycin). However, the disk diffusion method
may not be a good alternative to use for susceptibility testing of thermophilic
Campylobacter to ampicillin and tetracycline because the percent agreement between the
agar dilution and the disk diffusion methods in identification of susceptible and resistant
Campylobacter isolates to these antimicrobials as well as the kappa value of ampicillin and tetracycline between these two tests were not very high when compared to those of other antibiotics. Since a large number of Campylobacter isolates were classified as intermediate to erythromycin and clindamycin by the standardized agar dilution method but they were classified as either intermediate or susceptible to these antimicrobial agents by the disk diffusion method, until the standard resistance breakpoints specific for
Campylobacter is established, it is going to be difficult to determine whether the disk diffusion method is a reliable alternative for susceptibility testing of Campylobacter to
erythromycin and clindamycin or not.
In this study, Campylobacter strains that were resistant to fluoroquinolones and
aminoglycosides by the standardized agar dilution method were also resistant to these
antimicrobial agents by the disk diffusion method. Interestingly, no zone of inhibition
200 around these antibiotic disks was observed among the resistant Campylobacter strains,
while a large clear zone of inhibition averaged more than 37 mm in diameter was
observed around the disks of the susceptible Campylobacter strains. The drastic difference in the zone of inhibition diameter between the susceptible and resistant
Campylobacter strains to fluoroquinolone and aminoglycoside antibiotics in combination with the high percent agreement in identification of susceptible and resistant isolates and a high kappa value between the agar dilution and the disk diffusion methods indicate that the disk diffusion test may be able to use as a reliable screening method for antimicrobial susceptibility testing of thermophilic Campylobacter to these classes of antimicrobial agents. This finding is correlated well with the previous study by Gaudreau and Gilbert
(1997), who reported a complete agreement between the agar dilution method and the disk diffusion method for susceptibility testing of C. jejuni and C. coli to ciprofloxacin.
In addition, the previous study by Vanhoof et al. (1984) also showed a good correlation between the disk diffusion and the agar dilution methods for antimicrobial susceptibility testing of C. jejuni to many antibiotics. Likewise, Frediani-Wolf and Stephan (2003) suggested that the disk diffusion method can be used as a reliable and easy tool for monitoring the prevalence of ciprofloxacin-resistant C. jejuni strains although they found a weak correlation between the MIC and zone diameter results for ciprofloxacin- susceptible strains.
The MICs of erythromycin and clindamycin for most Campylobacter strains tested in this study ranged between 1 and 4 µg/ml for erythromycin and between 1 and 2
µg/ml for clindamycin, which were in the intermediate category according to the current
201 resistance breakpoints. However, if the interpretation criteria for resistance change, the
numbers of Campylobacter isolates that are susceptible, intermediate, or resistant to these
antibiotics will change. For example, if the tentative resistance breakpoints of
erythromycin for thermophilic Campylobacter proposed by Huysmans and Turnidge
(1997) are used, the numbers of intermediate strains in this study will reduce from 413 to
19 isolates for the agar dilution method and from 62 to 15 isolates for the disk diffusion method, whereas the numbers of susceptible strains will increase to 489 isolates by both methods and the numbers of resistant isolates will change to 160 for the agar dilution method and to 164 for the disk diffusion method. In addition, the percent agreement in identification of susceptible and resistant isolates to erythromycin as well as the kappa value will also change to 97.96% and 0.95, respectively. Likewise, as shown in Table 3.4 and 3.5, if the intermediate strains were reclassified as resistant or susceptible strains, the
percent agreement between the agar dilution method and the disk diffusion method in identification of erythromycin-susceptible and erythromycin-resistant isolates as well as the kappa value would change significantly. Particularly, when the intermediate strains were reclassified as resistant strains, the percent agreement between these two methods changed dramatically from 96.14% to 47.90% and the kappa also reduced significantly from 0.91 to only 0.13 (Table 3.4). However, this drastic change was not really surprising because most of the isolates classified as intermediate to erythromycin by the agar dilution method were classified as susceptible or intermediate by the disk diffusion method with the zone of inhibition diameter around 32 mm. Similar change was also observed on clindamycin. If the MIC and zone diameter breakpoints of clindamycin
202 recommended by the Société Francaise de Microbiologie (SFM) are used as the interpretation criteria (Acar and Goldstein, 1996), the numbers of intermediate strains will not be present, while the numbers of susceptible and resistant strains to clindamycin will change to 525 and 143 isolates by the agar dilution method and to 506 and 162 isolates by the disk diffusion method, respectively. In addition, the percent agreement between the agar dilution and the disk diffusion methods and the kappa value will change to 92.96% and 0.80, respectively as well. Likewise, when the intermediate strains were reclassified as resistant strains, the percent agreement in identification of clindamycin- susceptible and clindamycin-resistant strains between these two tests reduced significantly from 97.34% to 65.42%, while the kappa value also extremely reduced from
0.95 to 0.30 when compared to the percent agreement and the kappa value calculated without including the intermediate strains (Table 3.3 and 3.4).
Since the MICs and the zone of inhibition diameter between susceptible and resistant Campylobacter isolates to erythromycin and especially clindamycin were not drastically different like the case of fluoroquinolones, the resistance breakpoints used to determine whether the isolates were susceptible, intermediate, or resistant to erythromycin and clindamycin were very crucial. Unfortunately, the standardized interpretation criteria specific for Campylobacter have not been established yet. In addition, as mentioned earlier, the numbers of susceptible and resistant Campylobacter strains to erythromycin and clindamycin changed significantly when different resistance breakpoints were used. Therefore, it is quite difficult at this stage to determine whether the disk diffusion method is a reliable screening test for monitoring antimicrobial
203 resistance of thermophilic Campylobacter to erythromycin and clindamycin or not.
However, if the intermediate strains are not included in the data analysis, the disk
diffusion method seems to correlate well with the agar dilution method (Table 3.3).
Similarly, if the interpretation criteria proposed by Huysmans and Turnidge (1997) are
used to determine erythromycin resistance of Campylobacter isolates in this study, the percent agreement between the agar dilution method and the disk diffusion method as well as the kappa value of these two testes are also very high. A good correlation between the agar dilution method and the disk diffusion method for screening erythromycin- resistant Campylobacter strains was also previously reported by Alfredson et al. (2003) and Frediani-Wolf and Stephan (2003). Since 95% of the quality control organism C. jejuni ATCC 33560 and the majority of Campylobacter isolates tested in this study as well as in other studies (Baker, 1992; Gaudreau and Gilbert, 1997) had the MICs of erythromycin in the intermediate category (between 1 and 4 µg/ml) when the current
NCCLS interpretation criteria were applied, it might be a better idea if the MIC breakpoints for susceptible Campylobacter isolates are raised to less than or equivalent to
4 µg/ml instead of less than or equivalent to 0.5 µg/ml as currently use.
For ampicillin and tetracycline, the disk diffusion method may not be a reliable method to use for susceptibility testing of Campylobacter to these antibiotics because the percent agreement in identification of susceptible and resistant isolates between the agar dilution and the disk diffusion methods and the kappa value between these two tests were not very high even though different resistance breakpoints or different interpretation criteria have been used. For example, if the interpretation criteria for ampicillin
204 recommended by the British Society for Antimicrobial Therapy (BSAC) (Andrews,
2004) are used to determine antimicrobial resistance of Campylobacter isolates in this study, the percent agreement in identification of ampicillin-susceptible and ampicillin- resistant Campylobacter isolates between the agar dilution and the disk diffusion methods will be 85.82%, whereas the kappa value between these tests will be only 0.41%. As shown in Table 3.3 – 3.5, it does not matter whether the intermediate strains were reclassified as either susceptible or resistant strains or whether no intermediate strains were included in the statistical calculation, the kappa value of ampicillin ranged only between 0.39 and 0.56 (Table 3.3 – 3.5). Although the kappa value of tetracycline was not as low as that of ampicillin, it was still lower than those of fluoroquinolones and aminoglycosides tested in this study with the kappa ranging from 0.61 to 0.75 (Table 3.3
– 3.5). Since a moderate number of tetracycline-resistant Campylobacter isolates determined by the agar dilution method were classified as intermediate strains by the disk diffusion method, it was not usual that the percent agreement and the kappa value between these two tests were not very high. The previous study by Alfredson et al. (2003) also showed a low level of agreement (77%) between the agar dilution method and the disk diffusion method when they were used to determine ampicillin-resistant
Campylobacter isolates. However, this research group found that these two methods correlated well when they were used for screening of tetracycline-resistant
Campylobacter strains.
Although a correlation coefficient and a linear regression analysis have been the most commonly used statistics for determining the agreement between different
205 antimicrobial susceptibility testing methods, these statistics may not be the best statistics
to use because they measure the degree of association to which the results obtained from
one test varies linearly with the results obtained from the other test not the agreement
between the tests and they also highly influenced by the variation between individuals
(Altman and Bland, 1983; Dohoo et al., 2003). In addition, McDermott et al. (2004) also
suggested that it was impossible to correlate the disk diffusion results with the agar
dilution results by the linear regression analysis.
In conclusion, this study reveals that the disk diffusion method can be used as a reliable alternative method for susceptibility testing of thermophilic Campylobacter to fluoroquinolone and aminoglycoside antibiotics especially for monitoring the prevalence of resistant Campylobacter strains to these classes of antibiotics. Since the percent agreement and the kappa value of ampicillin and tetracycline between the standardized agar dilution method and the disk diffusion method obtained from this present study were not very high, the disk diffusion method should not be used for susceptibility testing of
Campylobacter to these classes of antimicrobials. When the current NCCLS resistance breakpoints for erythromycin and clindamycin were used, a large number of
Campylobacter isolates in this study were classified as intermediate by the agar dilution method. Since the numbers of Campylobacter strains susceptible, intermediate, or resistant to erythromycin and clindamycin changed drastically when the different resistance breakpoints were used, it is difficult at this stage to determine whether the disk diffusion method is a reliable alternative for susceptibility testing of Campylobacter to erythromycin and clindamycin or not. Until the standard resistance breakpoints specific
206 for Campylobacter is established and validated, the agar dilution method seems to be a
better method for antimicrobial susceptibility testing of Campylobacter to erythromycin and clindamycin. Nevertheless, we feel that the disk diffusion method may be able to use as a reliable screening method for susceptibility testing of thermophilic Campylobacter to erythromycin. Although the disk diffusion method is not complicated to perform as the agar dilution method and provides reliable results for several classes of antimicrobials, only qualitative data can be obtained from this method. In addition, another drawback of the disk diffusion method that we noticed in this study was the poor growth of
Campylobacter isolates on the plates causing difficulty in interpreting the results that
sometimes led to the retest. However, this method seems to be very useful especially
when several antimicrobial agents need to be tested against a few isolates. If the
quantitative data is required, other methods such as the agar dilution method or the E-test
should be performed.
3.6 ACKNOWLEDGEMENT
The authors wish to thank Ms. Sonya M. Bodeis at the Center for Veterinary
Medicine, Food and Drug Administration for her technical assistance in this study. The
authors would also like to thank Dr. Benjamas Promsopone, Ms. Elisabeth J. Angrick,
Ms. Lori Martin, Ms. Nicole Fehribach, Mr. Andew M. Krieger, and fellow colleagues at
the Avian Disease Investigation Laboratory at The Ohio State University for their help,
advice, and technical support. In addition, the assistance of Dr. Audrey Torres on
statistical analysis is also gratefully acknowledged.
207
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210 Antimicrobial Agar dilution method Disk diffusion method a Disk b Agar dilution test MIC breakpoints (µg/ml) Zone diameter breakpoints (mm) agents concentration range (µg/ml) S I R (µg) S I R
Ampicillin 0.06-128 ≤ 8 16 ≥ 32 10 ≥ 17 14-16 ≤ 13
Tetracycline 0.06-128 ≤ 4 8 ≥ 16 30 ≥ 19 15-18 ≤ 14
Gentamicin 0.06-128 ≤ 4 8 ≥ 16 10 ≥ 15 13-14 ≤ 12 Kanamycin 0.25-128 ≤ 16 32 ≥ 64 30 ≥ 18 14-17 ≤ 13 Clindamycin 0.06-128 ≤ 0.5 1-2 ≥ 4 2 ≥ 21 15-20 ≤ 14
211 Erythromycin 0.06-128 ≤ 0.5 1-4 ≥ 8 15 ≥ 23 14-22 ≤ 13
Ciprofloxacin 0.008-128 ≤ 1 2 ≥ 4 5 ≥ 21 16-20 ≤ 15
Norfloxacin 0.06-128 ≤ 4 8 ≥ 16 10 ≥ 17 13-16 ≤ 12
Nalidixic acid 0.25-128 ≤ 16 ≥ 32 30 ≥ 19 14-18 ≤ 13
a MIC breakpoints for enteric bacteria used by the NARMS and the NCCLS; S, susceptible; I, intermediate; R, resistance b Zone diameter breakpoints for Enterobacteriaceae approved by the NCCLS; S, susceptible; I, intermediate; R, resistance
Table 3.1 Breakpoints of the agar dilution method and the disk diffusion method used to determine antimicrobial susceptibility of Campylobacter isolates
211 Number of susceptible-, intermediate-, resistant- Campylobacter isolates identified by
Antimicrobial agents Agar dilution methoda Disk diffusion methoda S I R S I R Ampicillin 434 154 75 416 138 109
Tetracycline 158 4 505 195 49 423
Gentamicin 668 0 0 667 0 1
Kanamycin 418 2 248 415 10 243
Clindamycin 159 366 143 329 178 161 212 Erythromycin 76 413 179 449 62 157
Ciprofloxacin 444 10 214 437 2 229
Norfloxacin 451 4 213 439 0 229
Nalidixic acid 454 0 214 434 31 203
a S, susceptible; I, intermediate; R, resistant
Table 3.2 Antimicrobial susceptibility patterns of Campylobacter spp. identified by the agar dilution method and the disk diffusion method
212 Number of Campylobacter isolatesa b Resistant by % identification % identification % agreement Susceptible Susceptible Antimicrobial AD but Resistant by of resistant of resistant between AD c by both AD by AD but Kappa agents susceptible both AD and isolates by AD isolates by DD and DD and DD resistant by by DD DD methods method method methods methods DD methods methods Ampicillin 336 39 14 45 13.60 19.35 87.79 0.56 Tetracycline 143 14 50 408 74.47 68.62 89.59 0.75 Gentamicin 667 1 0 0 0 0 99.85 n/ad Kanamycin 408 1 6 242 37.75 36.99 98.94 0.98 Clindamycin 127 0 7 129 51.71 49.05 97.34 0.95 Erythromycin 74 0 9 150 68.24 64.38 96.14 0.91 Ciprofloxacin 422 20 5 209 32.62 34.91 96.19 0.92 213 Norfloxacin 431 20 4 209 32.08 34.49 96.39 0.92 Nalidixic acid 425 20 9 183 30.14 31.87 95.48 0.90
a Number of susceptible or resistant Campylobacter isolates identified by the agar dilution (AD) and the disk diffusion (DD) methods. b % agreement in identification of susceptible and resistant Campylobacter isolates by the agar dilution (AD) and the disk diffusion (DD) methods. c The magnitude of kappa indicates the level of agreement between the two tests as the followings: < 0.2, slight agreement; 0.2 to 0.4, fair agreement; 0.4 to 0.6, moderate agreement; 0.6 to 0.8, substantial agreement; > 0.8, almost perfect agreement. d Kappa value of gentamicin could not be calculated because none of the isolates was resistant to this antibiotic causing the lack of representative value essential for statistical calculation.
Table 3.3 The agreement between the agar dilution method and the disk diffusion method for antimicrobial susceptibility testing of Campylobacter (no intermediate strains were included)
213 Number of Campylobacter isolatesa b Resistant by % identification % identification % agreement Susceptible Susceptible Antimicrobial AD but Resistant by of resistant of resistant between AD c by both AD by AD but Kappa agents susceptible both AD and isolates by AD isolates by DD and DD and DD resistant by by DD DD methods method method methods methods DD methods methods Ampicillin 337 100 83 143 34.09 36.65 72.40 0.40 Tetracycline 143 15 52 457 76.31 70.77 89.96 0.74 Gentamicin 667 1 0 0 0 0 99.85 n/ad Kanamycin 408 10 7 243 37.43 37.87 97.46 0.95 Clindamycin 127 32 199 310 76.20 51.20 65.42 0.30 Erythromycin 74 2 346 246 88.62 37.13 47.90 0.13 Ciprofloxacin 422 22 15 209 33.53 34.58 94.46 0.88 214 Norfloxacin 431 20 8 209 32.49 34.28 95.81 0.91 Nalidixic acid 425 29 9 205 32.04 35.03 94.31 0.87
a Number of susceptible or resistant Campylobacter isolates identified by the agar dilution (AD) and the disk diffusion (DD) methods. b % agreement in identification of susceptible and resistant Campylobacter isolates by the agar dilution (AD) and the disk diffusion (DD) methods. c The magnitude of kappa indicates the level of agreement between the two tests as the followings: < 0.2, slight agreement; 0.2 to 0.4, fair agreement; 0.4 to 0.6, moderate agreement; 0.6 to 0.8, substantial agreement; > 0.8, almost perfect agreement. d Kappa value of gentamicin could not be calculated because none of the isolates was resistant to this antibiotic causing the lack of representative value essential for statistical calculation.
Table 3.4 The agreement between the agar dilution method and the disk diffusion method for antimicrobial susceptibility testing of Campylobacter (intermediate strains were classified as resistant strains)
214 Number of Campylobacter isolatesa b Resistant by % identification % identification % agreement Susceptible Susceptible Antimicrobial AD but Resistant by of resistant of resistant between AD c by both AD by AD but Kappa agents susceptible both AD and isolates by AD isolates by DD and DD and DD resistant by by DD DD methods method method methods methods DD methods methods Ampicillin 524 64 32 43 11.31 16.14 85.52 0.39 Tetracycline 147 15 97 408 75.71 63.42 83.21 0.61 Gentamicin 667 1 0 0 0 0 99.85 n/ad Kanamycin 419 1 6 242 37.13 36.38 98.95 0.98 Clindamycin 492 33 14 129 21.41 24.25 92.96 0.80 Erythromycin 482 7 29 150 26.80 23.50 94.61 0.86 Ciprofloxacin 434 20 5 209 32.04 34.28 96.26 0.92
215 Norfloxacin 435 20 4 209 31.89 34.28 96.41 0.92 Nalidixic acid 434 20 31 183 32.04 30.39 92.37 0.82
a Number of susceptible or resistant Campylobacter isolates identified by the agar dilution (AD) and the disk diffusion (DD) methods. b % agreement in identification of susceptible and resistant Campylobacter isolates by the agar dilution (AD) and the disk diffusion (DD) methods. c The magnitude of kappa indicates the level of agreement between the two tests as the followings: < 0.2, slight agreement; 0.2 to 0.4, fair agreement; 0.4 to 0.6, moderate agreement; 0.6 to 0.8, substantial agreement; > 0.8, almost perfect agreement. d Kappa value of gentamicin could not be calculated because none of the isolates was resistant to this antibiotic causing the lack of representative value essential for statistical calculation.
Table 3.5 The agreement between the agar dilution method and the disk diffusion method for antimicrobial susceptibility testing of Campylobacter (intermediate strains were classified as susceptible strains)
215
CHAPTER 4
TETRACYCLINE RESISTANCE OF CAMPYLOBACTER ISOLATES ON
ORGANIC POULTRY FARMS
4.1 ABSTRACT
A total of 245 pooled samples from one organic broiler farm were collected
weekly from the first week until the end of the production cycle. Similarly, 585 pooled
samples originated from 5 organic broiler farms and 3 organic turkey farms were also
collected and cultured for Campylobacter. Tetracycline resistance of these
Campylobacter isolates were identified by the agar dilution method, whereas the presence
of tet(O) gene were determined by the PCR method. The changes of tetracycline resistance rates on the organic broiler farm were observed during the production cycle.
Campylobacter strains isolated from the first few weeks of the production cycle were
susceptible to tetracycline, while tetracycline resistance was first noticed among
Campylobacter isolates from week 5 of the production cycle. The resistance rates reached to 100% at week 6 and 7 then dropped to 33.33% at week 10 of the production cycle.
Interestingly, only 13.79% of Campylobacter isolates from intestinal tracts of this flock were resistant to tetracycline. In addition, tetracycline resistance rates were different among organic broiler and turkey farms. The presence of tet(O) gene was detected in
216 97.79% of tetracycline-resistant Campylobacter isolates, while a few DNA samples directly extracted from unexposed environmental samples were positive for tet(O) gene.
This study reveals the complex nature of tetracycline resistance among Campylobacter isolates on organic poultry farms.
4.2 INTRODUCTION
Campylobacter jejuni, a leading cause of bacterial gastroenterocolitis in humans, is responsible for over 2 million cases of food-borne bacterial diarrhea each year in the
United States (Altekruse et al., 1999). Most illnesses associated with Campylobacter infections are usually due to consumption of contaminated foods or foods that are cross- contaminated with raw or undercooked poultry (Altekruse and Tollefson, 2003). In general, Campylobacter is carried in intestinal tract of domestic poultry and other livestock as commensal organisms and it is often present on foods of animal origin through fecal contamination during processing (Altekruse et al., 1999; Altekruse and
Tollefson, 2003). Since poultry is considered to be a major source of human
Campylobacter infections (Newell and Wagenaar, 2000), a development of antimicrobial resistance in Campylobacter isolated from poultry is a matter of concern.
Tetracycline is a broad-spectrum antibiotic that has been widely used in human and veterinary medicine since the late 1940s and the early 1950s (Chopra et al., 1992;
Chopra and Roberts, 2001; Roberts, 1996). This antibiotic inhibits protein synthesis in gram-positive and gram-negative bacteria by preventing the binding of aminoacyl-tRNA to the ribosomal acceptor (A) site on the 30S ribosomal subunit (Chopra et al., 1992;
217 Chopra and Roberts, 2001; Roberts, 1996; Speer et al., 1992). Currently, four different mechanisms associated with tetracycline resistance in bacteria have been reported including (i) an energy-dependent efflux system, which helps reduce the intracellular concentration of tetracycline by an integral membrane protein; (ii) ribosomal protection protein, which protects the ribosomal binding site of tetracycline; (iii) inactivation of tetracycline by modifying enzymes, which occurs rarely; and (iv) mutation of the 16S rRNA, which found in propionibacteria (Aarestrup and Engberg, 2001; Chopra et al.,
1992; Chopra and Roberts, 2001; Roberts, 1996; Ross et al., 1998; Schnappinger and
Hillen, 1996; Speer et al., 1992; Taylor and Chau, 1996; Trieber and Taylor, 2000).
However, only a ribosomal protection protein is associated with tetracycline resistance in
Campylobacter species (Aarestrup and Engberg, 2001; Trieber and Taylor, 2000). In general, the mechanism of tetracycline resistance in Campylobacter is primarily associated with tet(O) gene, which encodes a ribosomal protection protein designated as
Tet(O) (Taylor and Courvalin, 1988; Trieber and Taylor, 2000). Tet(O) protein interacts directly with the ribosome, the target of tetracycline, and displaces tetracycline from its primary binding site on the ribosome; thus, protecting the ribosome from the inhibitory effect of tetracycline (Connell et al., 2002; Connell et al., 2003; Gibreel et al., 2004b;
Manavathu et al., 1990; Pratt and Korolik, 2005; Trieber and Taylor, 2000). Although tet(O) gene is usually carried on transmissible plasmids (Aarestrup and Engberg, 2001;
Gibreel et al., 2004b; Lee et al., 1994; Pratt and Korolik, 2005; Sagara et al., 1987;
Taylor
218 Taylor and Courvalin, 1988; Trieber and Taylor, 2000), it has been found to be
chromosomally-located as well (Gibreel et al., 2004b; Lee et al., 1994; Pratt and Korolik,
2005).
Over the last decade, the emergence of tetracycline resistance in C. jejuni and C.
coli has increased drastically in many countries worldwide. For example, in Canada, the
prevalence of tetracycline resistance in Campylobacter isolates increased significantly
from 19.1% during 1985 - 1986 to 43%-68% during 1998-2001 (Gaudreau and Gilbert,
1998; Gaudreau and Gilbert, 2003). Similarly, a rapid increase in tetracycline resistance
was also observed among clinical Campylobacter isolates in Spain, where the prevalence
of tetracycline resistance increased dramatically from 23% during 1985 - 1987 to 72%
during 1995 - 1998 for C. jejuni isolates and from 15.2% to 97% for C. coli isolates
during the same period of time (Mirelis et al., 1999; Prats et al., 2000; Reina et al., 1994).
The prevalence of tetracycline resistance was also reported from other countries located
in different geographical regions of the world. In the United States, the prevalence of
tetracycline resistance in human isolates was around 43% (Gupta et al., 2004), while the
prevalence of tetracycline-resistant C. jejuni isolates in Europe ranged from 11% in
Denmark (Aarestrup et al., 1997) to 46% in Finland (Hakanen et al., 2003). In other
European countries, about 30% - 42% of C. jejuni and C. coli isolates from humans were resistant to tetracycine (Luber et al., 2003b; Pezzotti et al., 2003). A high frequency of tetracycline resistance was mainly observed in Asia, where 55% of C. jejuni isolates in
Japan and 95% of C. jejuni and 85% of C. coli isolates in Taiwan were resistant to this antibiotic (Li et al., 1998; Sagara et al., 1987). In terms of poultry isolates, about 69% -
219 81% of C. jejuni and 50% - 77% of C. coli isolates from poultry meat in the United States were resistant to tetracycline (Ge et al., 2003; Gupta et al., 2004). Interestingly, a wide variation of tetracycline-resistant C. jejuni and C. coli isolates from broilers and turkeys as well as from poultry meat was observed in Europe ranging from 2% in Denmark to
76.3% in Italy (Aarestrup et al., 1997; Avrain et al., 2003; Luber et al., 2003b; Pezzotti et al., 2003; Van Looveren et al., 2001). As mentioned earlier, a large number of
Campylobacter isolates from Asia were resistant to tetracycline. So, it was not unusual that 83% of C. jejuni and 96% of C. coli isolates from poultry meat in Taiwan were resistant to tetracycline (Li et al., 1998).
The emergence of antimicrobial resistance in zoonotic food-borne bacterial pathogens including Campylobacter seems to be an undesired consequence of antimicrobial usage in animal agriculture (Khachatourians, 1998; McEwen and Fedorka-
Cray, 2002; Witte, 1998). Many antibiotics including tetracycline have been widely used in animal production systems to control and prevent diseases as well as to promote growth and improve feed efficiency (Chopra and Roberts, 2001; Khachatourians, 1998;
McEwen and Fedorka-Cray, 2002; Roberts, 1996). Unfortunately, the extensive use of antibiotics for non-therapeutic purposes such as feeding of sub-therapeutic level of chlortetracycline to food-producing animals may have led to the development of antimicrobial resistance not only in pathogenic bacteria but also in non-pathogenic bacteria in the intestinal tracts of these animals (Chopra and Roberts, 2001;
Khachatourians, 1998; McEwen and Fedorka-Cray, 2002; Van den Bogaard and
Stobberingh, 1999; Witte, 1998). In addition, this practice may also lead to the
220 establishment of antimicrobial resistance gene in the environment (McEwen and
Fedorka-Cray, 2002; Van den Bogaard and Stobberingh, 1999). Since no antibiotics have
been used in the organic production system, bacterial isolates from organically-raised
animals should be susceptible to almost every class of antimicrobial agents, to which they
are not intrinsically resistant. Unfortunately, from our previous study, we found that about 50% to 60% of C. jejuni and C. coli isolates from organically-raised broilers and turkeys were resistant to tetracycline (Luangtongkum et al., 2005). In addition, the study by Sato et al. (2004) also indicated that a large number of Campylobacter isolates from
organic dairy farms was resistant to tetracycline. Since tetracyclines have been used as
feed additives for livestock and poultry for both therapeutic and sub-therapeutic purposes for long period of time (Chopra and Roberts, 2001; McEwen and Fedorka-Cray, 2002), it is possible that bacteria including Campylobacter may evolutionally become resistant to this class of antibiotic regardless of the sources of the isolates. However, there is no scientific information to support this assumption. The purpose of this study was to investigate tetracycline resistance of thermophilic Campylobacter on organic poultry
farms in order to get some scientific information that may help elucidate a high
prevalence of tetracycline resistance found among Campylobacter isolates from organic
production system. In addition, this study may help identify the source of tetracycline
resistance on organic poultry operations as well.
221
4.3 MATERIALS AND METHODS
Sample collection. This study focused on tetracycline resistance of thermophilic
Campylobacter on organic poultry farms. Five organic broiler farms and three organic turkey farms were utilized in this study. For one organic broiler farm, the samples were collected every week from the first week until the end of the production cycle.
Environmental samples such as feed, water, litter, and grass as well as fresh fecal samples were randomly collected throughout the farm to ensure that samples represented the whole production system and the whole-flock exposure and placed into sterile Nasco
Whirl-Pak™ (Nasco Agriculture Sciences, Fort Atkinson, WI) and sterile specimen
containers (Fisher Scientific, Pittsburgh, PA) and brought back to the laboratory within three hours of collection to culture for Campylobacter. Generally, each type of environmental samples was collected before and after coming in contact with the birds.
Approximately 1,075 environmental and fecal samples obtained from the whole production cycle of one organic broiler farm and about 1,630 environmental and fecal samples obtained from 5 organic broiler farms and 3 organic turkey farms were collected from August 2004 to November 2004. In addition, at the end of the production cycle, at least 30 intestinal tracts from each organic poultry farm were also collected and brought back to the laboratory within three hours of collection to culture for Campylobacter.
Bacterial isolation and identification. A total of 215 pooled environmental and fecal samples obtained from the whole production cycle of one organic broiler farm and 326
222 pooled environmental and fecal samples obtained from 5 organic broiler farms and 3
organic turkey farms as well as 283 intestinal tracts of organically-raised broilers and turkeys were cultured for Campylobacter by the standard microbiological method.
Since Campylobacter is susceptible to a variety of environmental conditions and is unlikely to survive for long period of time outside the host’s intestinal tract, the presence of Campylobacter in environmental samples was determined by a selective enrichment method, whereas the presence of this organism in poultry intestines was determined by a direct plating method. In addition, for water samples, besides a selective enrichment method, a direct plating method and a filtration method were also performed to increase a chance of Campylobacter detection from water samples. Likewise, both selective enrichment and direct plating methods were also conducted for fecal samples.
For direct plating method, each sample was directly streaked onto Campy CVA agar (BBL Becton Dickinson Microbiology Systems, Cockeysville, MD). The inoculated plates were then incubated at 42 °C for 48 hours under a microaerophilic environment
(approximately 5% O2, 10% CO2, and 85% N2) in a CampyPak II anaerobic system jar
with CampyPak gas generating system envelopes (BBL Becton Dickinson Microbiology
Systems, Sparks, MD).
For selective enrichment method, the procedure was performed according to the protocol previously published by Denis et al. (2001) with some modifications. Briefly, 10 grams of samples were added to 90 ml of Preston broth containing Nutrient broth No. 2
(CM0067) (Oxoid LTD., Basingstoke, Hampshire, England), Campylobacter growth supplement (SR084E/SR0232E) (Oxoid LTD., Basingstoke, Hampshire, England),
223 Preston Campylobacter selective supplement (SR0117E) (Oxoid LTD., Basingstoke,
Hampshire, England), and 5% laked horse blood (SR0048C) (Oxoid LTD., Basingstoke,
Hampshire, England) and homogenized for 30 seconds using a Seward stomacher® lab blender (model 400) (Seward Medical Limited, London, England). These samples were then incubated at 42 °C overnight under a microaerophilic environment in a CampyPak II anaerobic system jar with CampyPak gas generating system envelopes (BBL Becton
Dickinson Microbiology Systems, Sparks, MD). After incubation, the enriched samples were subcultured onto Campy CVA agar (BBL Becton Dickinson Microbiology Systems,
Cockeysville, MD) and incubated at 42 °C for another 48 hours under a microaerophilic environment in a CampyPak II anaerobic system jar with CampyPak gas generating system envelopes (BBL Becton Dickinson Microbiology Systems, Sparks, MD).
For filtration method, the procedure was performed according to the previously published protocols by Blaser and Cody (1986) and Pearson et al. (1993) with modifications. Briefly, 100 ml of water sample was filtered through a Nalgene® analytical
test filter funnel with 0.45-µm pore size, 47-mm diameter cellulose nitrate membrane
filter (Nalge Nunc International, Rochester, NY). Each filter was placed facedown onto
Campy CVA agar (BBL Becton Dickinson Microbiology Systems, Cockeysville, MD)
and incubated overnight at 42 °C under a microaerophilic condition in a Forma Series II
water-jacketed CO2 incubator (Thermo Electron Corporation, Marietta, OH). After
incubation, the filter was removed and the Campy CVA agar plate was resubcultured and
then incubated at 42 °C for another 48 hours under a microaerophilic environment in a
CampyPak II anaerobic system jar with CampyPak gas generating system envelopes
224 (BBL Becton Dickinson Microbiology Systems, Sparks, MD). If there were no suspect
Campylobacter colonies on the plates, these Campy CVA agar plates would be re-
incubated for up to four days before being reported as negative.
Suspect Campylobacter colonies were identified by colony morphology
characteristics as well as biochemical characteristics to Gram-stain, oxidase test, catalase test, and Campylobacter culture-plate latex agglutination confirmation test (INDX-
Campy [jcl]; PanBio INDX, Inc., Baltimore, MD). In addition, the hippurate hydrolysis test was also performed to differentiate between C. jejuni and C. coli.
Tetracycline resistance determination. The agar dilution method was used to determine the minimal inhibitory concentration (MIC) of tetracycline for Campylobacter isolates in this study as recommended by the National Committee for Clinical Laboratory Standard
(NCCLS) (NCCLS, 2002). Briefly, Campylobacter isolates grown on blood agar plates for at least 24 hours were inoculated into Mueller-Hinton (MH) broth (BBL Becton
Dickinson Microbiology Systems, Cockeysville, MD) and then adjusted to a turbidity equivalent to 0.5 McFarland standard by a colorimeter (BioMérieux, Inc., Hazelwood,
MO). Approximately 104 CFU of adjusted Campylobacter suspensions were applied onto
Mueller-Hinton agar (BBL Becton Dickinson Microbiology Systems, Cockeysville, MD)
containing a two-fold concentration series (0.06-128 µg/ml) of tetracycline (Sigma
Chemical Co., St. Louis, MO) and supplemented with 5% defibrinated sheep blood
(Cleveland Scientific, Cleveland, OH) by a multipoint inoculator (a Cathra replicator system®) with 1 mm pins (Oxoid, Inc., Ogdensburg, NY). Campylobacter jejuni ATCC
225 33560 was also inoculated onto each plate to serve as a quality control organism. The
inoculated plates were incubated in a Forma Series II water-jacketed CO2 incubator
(Thermo Electron Corporation, Marietta, OH) at 42 °C under a microaerophilic condition for 24 hours. The MIC was defined as the lowest concentration of antimicrobial agent that completely inhibited the visible growth on the plates. The MIC breakpoint for tetracycline resistance, which is ≥ 16 µg/ml, was determined according to the NCCLS established guideline for bacteria isolated from animals (NCCLS, 2002a).
DNA extraction. In order to detect the presence of tet(O) gene in environmental and fecal samples as well as in bacterial isolates, DNA was extracted from these samples by the Wizard® Genomic DNA Purification Kit (Promega Corporation, Madison, WI)
according to the manufacturer’s protocol and the protocol previously published by Denis
et al. (2001). Briefly, after the enrichment step, 1.5 ml of Preston broth was centrifuged at
5500 rpm for 1 minute to precipitate a residue using a 5590 Microcentrifuge-HP (Thermo
Electron Corporation, Marietta, OH). Then, the supernatant or the suspension of bacterial
cells was centrifuged at 13,500 rpm for 10 minutes to pellet the cells. After the
supernatant was discarded, bacterial cells were suspended in 600 µl nuclei lysis solution
and then incubated at 80 °C in a water bath for 5 minutes. After that 3 µl of RNase
solution were added to the tube and then incubated at 37 °C in a water bath for another 40
minutes. Then, 200 µl of protein precipitation solution were added to the tube. The tube
was then vortexed and placed on ice for 5 minutes. After centrifugation at 13,500 rpm for
3 minutes, the supernatant was transferred to a new microcentrifuge tube containing 600
226 µl of isopropanol and then centrifuged at 13,500 rpm for another 2 minutes. The pellet was then washed with 70% ethanol and centrifuged again at 13,500 rpm for 2 minutes.
The ethanol was aspirated and the pellet was air-dried for 15-20 minutes or until it was
completely dried. Finally, the DNA pellet was rehydrated in 100 µl of DNA rehydration
solution overnight at 4 °C. The DNA samples were stored at -20 °C until the detection of
tet(O) gene was performed.
Polymerase chain reaction (PCR). The presence of tet(O) and major outer membrane
protein (MOMP) of Campylobacter genes in environmental and fecal samples as well as
in bacterial isolates was determined by the PCR method. The PCR conditions for tet(O)
gene detection were performed according to the protocol previously published by
Aminov et al. (2001), while the PCR conditions for MOMP gene detection were followed
the protocol previously published by Zhang et al. (2000). Briefly, PCR was performed in
a volume of 50 µl with a final concentration of each component as the followings: 1x
PCR buffer, 2.5 mM MgSO4, 0.2 mM each deoxynucleoside triphosphate (dNTPs), 0.2
µM primers, and 1.25 U of Taq DNA polymerase. All PCR components except primers
(IDT® Integrated DNA Technologies, Inc., Coralville, IA) (Table 4.1) were obtained
from Promega Corporation, Madison, WI. PCR amplification was performed with a
GeneAmp® 2400 PCR System (Perkin-Elmer Corporation, Norwalk, CT) as the
followings: initial denaturation at 94 °C for 5 minutes, followed by 25 cycles of 94 °C for
30 seconds, 30 seconds of annealing temperature at 60 °C, and 30 seconds of extension at
72 °C for tet(O) gene detection, or 30 cycles of 94 °C for 1 minute, 1 minute of annealing
227 temperature at 58 °C, and 1 minute of extension at 72 °C for MOMP gene detection, and
then a final extension step at 72 °C for 10 minutes. This PCR amplification generated 170
bp and 1,400 bp DNA fragment of tet(O) and MOMP genes, respectively. PCR products
were determined by electrophoresis on a 2% agarose gel containing 0.5 µg/ml of
ethidium bromide solution (Promega Corporation, Madison, WI) and visualized with a
UV transilluminator and photographed by a digital imaging system (Alpha Innotech
Corporation, San Leandro, CA).
Statistical analysis. A Chi-square (χ2) test at a significance level of P < 0.05 (two-tailed)
with Yates correction for continuity was used for statistical analysis of tetracycline
resistance rates of Campylobacter spp. between different sources, different farms and
different studies.
4.4 RESULTS
For the organic broiler farm that was followed for the whole production cycle,
Campylobacter was not recovered from any environmental or fecal samples collected during the first and second week of the rearing cycle (Table 4.2). At the third week,
Campylobacter was first isolated from environmental and fecal samples with 73.33% of these samples positive for Campylobacter (Table 4.2). During the fourth to the tenth week of the production cycle, the prevalence of Campylobacter ranged from 61.11% to
88% (Table 4.2). At the end of the production cycle, Campylobacter was isolated from all
228 30 intestinal tracts (100%) of these organic broilers (Table 4.2). Interestingly, all
Campylobacter isolates from this organic broiler farm were C. jejuni on the basis of hippurate hydrolysis test.
One of the most interesting findings of this study was the changes in tetracycline resistance rates among Campylobacter isolates from each week of the production cycle.
Neither Campylobacter isolates from the third nor the fourth week of the production cycle were resistant to tetracycline, while more than 65% of the isolates from the fifth week were resistant to this antibiotic (Table 4.2). Tetracycline resistance rates reached to
100% during the sixth and seventh week then dropped to 72.22% and 81.82% in the eighth and ninth week of the production cycle, respectively (Table 4.2). At week 10, a few days before the birds were sent to the processing plant only 33.33% of these
Campylobacter isolates were resistant to tetracycline, whereas only 13.79% of
Campylobacter isolates from the intestinal tracts of these organically-raised broilers were resistant to this antibiotic (Table 4.2). In addition, all tetracycline-resistant
Campylobacter isolates except one isolate from the fifth week of the production cycle were positive for tet(O) gene, while none of the randomly selected tetracycline- susceptible Campylobacter isolates was positive for tet(O) gene when determined by
PCR method (Figure 4.1).
In terms of the prevalence and tetracycline resistance rates of Campylobacter isolates from other organic broiler and turkey farms, we found that the prevalence of
Campylobacter isolated from environmental and fecal samples of organic broiler farms ranged from 20.31% to 47.37%, whereas the prevalence of Campylobacter isolated from
229 environmental and fecal samples of organic turkey farms ranged from 46.15% to 59.38%
(Table 4.3 and 4.4). In addition, more than 90% of intestinal tracts of both organically-
raised broilers and turkeys were positive for Campylobacter (Table 4.3 and 4.4). As mentioned earlier, although a majority of Campylobacter isolates from organic broiler farms were C. jejuni, a proportion of C. jejuni and C. coli in organic turkey farms seemed to be different from farm to farm (Table 4.3 and 4.4). For example, none of
Campylobacter isolates from organic turkey farm A was C. coli, while 33.33% and
63.33% of Campylobacter isolated from environmental and fecal samples and from intestinal tracts of organic turkey farm C were C. coli, respectively (Table 4.4).
Interestingly, although none of Campylobacter isolated from environmental and fecal samples of organic turkey farm B was classified as C. coli, 35.71% of Campylobacter isolated from intestinal samples of this organic turkey farm B were classified as C. coli on the basis of hippurate hydrolysis test (Table 4.4).
When tetracycline resistance rates between organic broiler farms were compared, the resistance rates varied drastically from 0% in organic broiler farm D to 100% in organic broiler farm A (Table 4.3). Interestingly, although 100% of Campylobacter isolated from environmental and fecal samples of organic broiler farm A were resistant to tetracycline, only 7.14% of Campylobacter isolated from intestinal samples of this organic broiler farm A were resistant to this antibiotic (Table 4.3). In contrast, only
15.39% of Campylobacter isolated from environmental and fecal samples of organic broiler farm C were resistant to tetracycline, while 90% of Campylobacter isolated from intestinal tracts of the birds in this farm were resistant to tetracycline (Table 4.3). For
230 organic broiler farm B and farm E, although tetracycline resistance rates between
Campylobacter isolates from environmental/fecal samples and intestinal samples were not extremely different as those of organic broiler farm A and farm C, the tetracycline resistance rates between Campylobacter isolates from these two sources were still
significantly different from each other (P < 0.05) with a higher resistance rates observed
among the isolates from environmental and fecal samples for organic broiler farm B and
among the isolates from intestinal samples for organic broiler farm E (Table 4.3). Like
tetracycline resistance rates noticed in organic broiler farms, the resistance rates in
organic turkey farms were also varied from farm to farm with 100% and 11.77% of
Campylobacter isolated from environmental and fecal samples of organic turkey farm B
and farm C resistant to tetracycline, respectively (Table 4.4). As mentioned earlier, more
than 97% of tetracycline-resistant Campylobacter isolates from both organic broiler and
organic turkey farms also harbored tet(O) gene (Table 4.3 and 4.4).
In order to investigate the source of tetracycline resistance on organic poultry
farms, environmental samples such as feed, grit, litter, grass, and water samples from
separate storage places before coming in contact with the birds were collected and tested
for the presence of Campylobacter as well as tet(O) gene. In addition, other potential
sources of tetracycline resistance such as fecal samples of other animals in the farm were
also collected and tested for the presence of Campylobacter and tet(O) gene as well.
From 105 DNA samples directly extracted from enrichment broth of environmental
samples prior to coming in contact with the birds, 5 DNA samples from organic broiler
farm A and 2 DNA samples from organic broiler farm B were positive for tet(O) gene;
231 however, none of these tet(O) gene positive DNA samples were positive for MOMP gene
(Table 4.5). In addition, neither tet(O) gene nor MOMP gene was detected in DNA samples extracted from other organic broiler and turkey farms (Table 4.5). Although
Campylobacter was isolated from some of these clean environmental samples, all of them except one isolate from organic broiler farm C were susceptible to tetracycline (Table
4.5). Thus, it was unlikely that these environmental samples were the sources of tetracycline resistance on organic poultry farms; however, for organic broiler farm C, it is possible that tetracycline-resistant Campylobacter isolated from grass on the pasture may be the source of tetracycline resistance on this particular organic broiler farm.
4.5 DISCUSSION
For one organic broiler farm that was followed for the whole production cycle, the changes in tetracycline resistance rate of Campylobacter isolates were observed. None of
Campylobacter isolates from the third and the fourth week of the production cycle was resistant to tetracycline (Table 4.2). Tetracycline resistance was first noticed in
Campylobacter isolates during the fifth week of the production cycle (Table 4.2). During the sixth and seventh week, all Campylobacter isolates became resistant to tetracycline then this resistance rate dropped to 33.33% during the last week of the production cycle
(Table 4.2). Two days after the last environmental and fecal samples were collected from the farm; the intestinal tracts of these organically-raised broilers were collected and cultured for thermophilic Campylobacter. Interestingly, only 13.79% of these
Campylobacter isolates were resistant to tetracycline (Table 4.2). The similar change of
232 tetracycline resistance rate was also observed in another flock of this organic broiler farm
A, where 100% of Campylobacter isolated from environmental and fecal samples of nine
weeks old birds were resistant to tetracycline, while only 7.14% of Campylobacter
isolated from the intestinal tracts of the ten and a half weeks old birds were resistant to
this antibiotic (Table 4.3 and 4.6). The association between age of the birds and
tetracycline resistance rates were also observed among Campylobacter isolates from organic broiler farm E. Most of Campylobacter isolates from environmental and fecal samples of three weeks old birds in organic broiler farm E were susceptible to tetracycline, while more than 90% of Campylobacter isolates from six and seven weeks
old birds in organic broiler farm E were resistant to this antibiotic (Table 4.3 and 4.6).
Likewise, less than 41% of Campylobacter isolates from intestinal tracts of 10 weeks or older birds (farm A and farm B) were resistant to tetracycline, while as high as 92% of
Campylobacter isolates from intestinal tracts of 7 to 9 weeks old birds (farm C and farm
E) were resistant to this antibiotic (Table 4.3 and 4.6). This observation suggests that age of the organic broilers is likely to affect the prevalence of tetracycline resistance of
Campylobacter spp. on organic broiler farms. Since the birds from both organic broiler farm A and farm E were obtained from the same hatchery, it is interesting to see that
Campylobacter isolates from both farms have similar changes of tetracycline resistance patterns. For other organic broiler farms where the birds were obtained from different hatcheries, different tetracycline resistance patterns were observed.
When tetracycline resistance rates observed in this recent study were compared to the resistance rates observed in the previous study (Luangtongkum et al., 2005), we
233 notice that tetracycline resistance rates in Campylobacter isolates on organic broiler
farms seemed to decrease, while the resistance rates in organic turkey farms seemed to
increase (Table 4.7). One of the most drastic changes in tetracycline resistance rates
between our previous and recent studies was observed in Campylobacter isolates from
organic broiler farm D where the resistance rates to tetracycline extremely decreased
from 94.12% to 0% (Table 4.7).
In terms of the level of tetracycline resistance, most tetracycline-resistant
Campylobacter isolates in this study had MICs ranging between 32 and 128 µg/ml except
for the isolates from organic broiler farm E that the majority of tetracycline-resistant
isolates had MICs higher than 128 µg/ml. Similar observation was also observed by
Taylor et al. (1986), who reported that the MICs of tetracycline for tetracycline-resistant
Campylobacter isolates in Canada varied between 32 and 128 µg/ml. Recently, Gibreel et
al. (2004b) found that 37% of clinical Campylobacter isolates in Canada had MICs
ranging between 256 and 512 µg/ml, while Pratt and Korolik (2005) revealed that tetracycline-resistant C. jejuni and C. coli isolates in Australia had MICs ranging from 32
to >256 µg/ml. In addition, these previous studies (Taylor et al., 1986; Gibreel et al.,
2004b) also demonstrated that plasmid that harbored tet(O) gene could confer high-level
of resistance to tetracycline with MICs of 128 to 512 µg/ml.
In general, tetracycline resistance in Campylobacter spp. has been reported to be plasmid-mediated. Around 70-100% of tetracycline-resistant Campylobacter isolates
carried plasmid ranging in size from 42 to 133 kb (Gibreel et al., 2004b; Lee et al., 1994;
Pratt and Korolik, 2005; Sagara et al., 1987; Taylor and Courvalin, 1988; Tenover et al.,
234 1985). However, the rates of tet(O)-harboring plasmid seemed to vary significantly between studies ranging from 32.3% to 96% (Lee et al., 1994; Pratt and Korolik, 2005).
In addition, plasmid-free tetracycline-resistant Campylobacter strains were also reported by previous studies (Lee et al., 1994; Pratt and Korolik, 2005; Sagara et al., 1987).
Although the plasmid-encoded tet(O) gene is the primary tetracycline resistance determinant in Campylobacter species, the chromosomally-encoded tet(O) gene may play an important role in tetracycline resistance in C. jejuni and C. coli isolates as well especially among the isolates that lacked plasmids (Gibreel et al., 2004b; Lee et al., 1994;
Pratt and Korolik, 2005). Since tet(O) gene detected by PCR in this study originated from total genomic DNA, the location of tet(O) gene whether it was plasmid-encoded tet(O) gene or chromosomally-encoded tet(O) gene could not be identified in the present study.
As previously reported (Gibreel et al., 2004b; Pratt and Korolik, 2005), almost
100% (265 of 271) of tetracycline-resistant Campylobacter isolates in this study were positive for tet(O) gene. Since the mechanism of tetracycline resistance in Campylobacter spp. is mainly dependent on tet(O) gene, it is not surprising that most tetracycline- resistant Campylobacter isolates in this study harbor tet(O) gene. However, when the presence of tet(O) gene in DNA directly extracted from enrichment broth was investigated, some of these samples especially the ones directly extracted from clean environmental samples prior to coming in contact with the birds were positive for tet(O) gene, but they were negative for MOMP gene (Table 4.5). This finding suggests that these environmental samples may be contaminated with other bacteria that can also harbor tet(O) gene. At present, neither species nor the ability of these bacteria in
235 transferring tet(O) gene to Campylobacter is known. Although previous studies (Chopra and Roberts, 2001; Roberts et al., 1993; Taylor and Courvalin, 1988; Widdowson et al.,
1996; Zilhao et al., 1988) reported that other bacteria particularly gram-positive bacteria including Enterococcus spp., Streptococcus spp., Staphylococcus, Lactobacillus,
Molbiluncus, Aerococcus, and Peptostreptococcus can also harbor tet(O) gene, no
information whether these bacteria can transfer tet(O) gene to Campylobacter was
available. However, it is believed that Campylobacter species acquired tet(O) gene most
likely from gram-positive coccus (Taylor and Courvalin, 1988).
In terms of the prevalence of Campylobacter in organic poultry farms, the present
study along with other previous studies (Heuer et al., 2001; Luangtongkum et al., 2005)
found that the prevalence of Campylobacter in organic poultry farms was very high. On
the other hand, El-Shibiny et al. (2005) reported that only 68.5% of Campylobacter were
isolated from organic chickens in the United Kingdom. In addition, Campylobacter spp.
in the present study were first isolated from organic broilers at 3 weeks of age, while El-
Shibiny et al. (2005) found that not until 31 days of age that Campylobacter from organic
chicken flock were first isolated. When compared to our previous study (Luangtongkum
et al., 2005), the prevalence of C. jejuni isolated from intestinal tracts of organically-
raised broilers and turkeys in this study is higher than that in the previous study except
for the organic turkey farm C where the prevalence of C. jejuni in the recent study is
lower than that in the previous study (Table 4.7). For C. coli, the prevalence of this
organism in the previous and recent studies is opposite to that of C. jejuni (Table 4.7). In
general, C. jejuni was the main organism isolated from both organic broiler and turkey
236 farms in this study except for organic turkey farm C where C. jejuni was predominant in environmental and fecal samples, while C. coli was predominant in intestinal samples
(Table 4.4). The prevalence of C. jejuni and C. coli in turkeys seems to vary significantly among the studies. Some studies (Lee et al., 2005; Smith et al., 2004) reported that C. coli was the main organism isolated from turkeys. In contrast, other studies (Wallace et al.,
1998; Van Looveren et al., 2001) found that almost 100% of Campylobacter isolated from turkeys were C. jejuni. In addition, one survey study in the Midwest showed that
40.5% of Campylobacter isolated from turkeys from one processing plant were C. coli, while only 14.6% of the isolates from another processing plant were C. coli (Logue et al.,
2003).
Another interesting observation from this study was the association between the prevalence of C. jejuni and C. coli and the species of turkey (white or bronze turkey).
Fourteen intestinal tracts of white turkeys and 30 intestinal tracts of bronze turkeys were collected from organic turkey farm B and cultured for Campylobacter. Among 14 intestinal tracts of white turkeys, 12 intestines were positive for Campylobacter, while
Campylobacter was recovered from all 30 intestinal tracts of bronze turkeys. When the species of Campylobacter isolated from intestinal tracts of white turkeys were determined by the hippurate hydrolysis test, 11 C. jejuni and 1 C. coli were identified. In contrast, the majority of Campylobacter isolates from bronze turkeys were identified as C. coli on the basis of hippurate hydrolysis test. Although the prevalence of Campylobacter spp. between white and bronze turkeys is not different, C. jejuni seems to be more prevalent in white turkeys, while C. coli seems to be more prevalent in bronze turkeys.
237 Since Campylobacter is unlikely to survive for long period of time outside
intestinal tract, the selective enrichment method was used to culture Campylobacter from
environmental and fecal samples in this study in order to increase Campylobacter
recovery rate. The results of this study showed that there was no significant difference (P
> 0.05) between the direct plating method and the selective enrichment method for
Campylobacter isolation from fecal samples; however, the selective enrichment method
seemed to be very useful especially for isolation of Campylobacter from samples that
seemed to have low numbers of Campylobacter contamination such as feed or water
samples. In addition, when the selective enrichment method was compared to the filtration method for recovering Campylobacter from water samples, we noticed that both methods were not significantly different (P > 0.05) from each other although the selective enrichment method might be a little bit better than the filtration method.
In summary, this study reveals the complex nature of tetracycline resistance of
Campylobacter spp. on organic poultry farms. The changes of tetracycline resistance
rates in Campylobacter species were observed during the production cycle of one organic broiler flock. This information indicates that tetracycline resistance phenotype among
Campylobacter isolates from organic broilers is not stable. Since other bacteria that
contaminate outside environment may harbor tet(O) gene, it might be useful to identify
the genus and species of these bacteria as well as to investigate the ability of these
bacteria in transferring tet(O) gene to Campylobacter species.
238
4.6 ACKNOWLEDGEMENT
The authors gratefully acknowledge the organic poultry farms and processing
plant for participating in the study. The authors would like to thank Mr. Jerrel C. Meitzler and Mr. Jeremiah J. Meeks for their technical assistance on PCR. The authors would also like to thank Dr. Amna B. El-Tayeb, Ms. Elisabeth J. Angrick, Ms. Marisa Ames, and fellow colleagues at the Avian Disease Investigation Laboratory at The Ohio State
University for their help, advice, and technical support.
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244
Primer Sequence PCR annealing temperature (°C) Amplicon size (bp)
TetO-FW 5’ACGGARAGTTTATTGTATACC3’ 60 171 TetO-RV 5’TGGCGTATCTATAATGTTGAC3’
MOMP-FW 5’ATGAAACTAGTTAAACTTAGTTTA3’ 58 1400 MOMP-RV 5’GAATTTGTAAAGAGCTTGAAG3’
245 Table 4.1 PCR primers and annealing temperature (°C)
245 No. (%) of samples No. of No. of samples positive for Campylobacter No. (%) of positive for resistant to tetracycline- Campylobacter/ Production tetracycline/ No. resistant No. of samples period of Campylobacter cultured for Campylobacter positive for Campylobacter C. jejuni C. coli isolates tested tet(O) gene (%) (%)
Wk 1 0/9 (0) 0 0 0 0
Wk 2 0/14 (0) 0 0 0 0
Wk 3 11/15 (73.33) 11 (100) 0 0/11 (0) 0
Wk 4 24/30 (80) 24 (100) 0 0/24 (0) 0
Wk 5 21/30 (70) 21 (100) 0 14/21 (66.67) 13 (92.86)
Wk 6 21/24 (87.50) 21 (100) 0 18/18 (100)a 18 (100)
Wk 7 24/28 (85.71) 24 (100) 0 24/24 (100) 24 (100)
Wk 8 18/22 (81.82) 18 (100) 0 13/18 (72.22) 13 (100)
Wk 9 11/18 (61.11) 11 (100) 0 9/11 (81.82) 9 (100)
Wk 10 22/25 (88) 22 (100) 0 7/21 (33.33)a 7 (100)
Intestine 30/30 (100) 30 (100) 0 4/29 (13.79)a 4 (100)
a Number of Campylobacter isolates tested for tetracycline resistance were less than the number of Campylobacter isolated from samples because some isolates could not be recovered after they were stored at -85 °C.
Table 4.2 Campylobacter and tetracycline resistance rates on an organic broiler farm from the first week until the end of the production cycle
246
No. (%) of samples No. of samples positive for No. of No. (%) of positive for Campylobacter tetracycline- Campylobacter/ resistant to resistant Farm Sample No. of samples tetracycline/ No. of Campylobacter cultured for Campylobacter positive for Campylobacter C. jejuni C. coli isolates tested (%) tet(O) gene (%)
5/24 5 A Environment 0 5/5 (100) 5 (100) (20.83) (100) 28/30 28 Intestine 0 2/28 (7.14) 2 (100) (93.33) (100) 25/64 21 B Environment 4 (16) 19/23 (82.61)a 19 (100) (39.06) (84) 27/30 24 3 Intestine 11/27 (40.74) 11 (100) (90) (88.89) (11.11) 13/41 9 4 C Environment 2/13 (15.39) 2 (100) (31.71) (69.23) (30.77) 30/30 30 Intestine 0 27/30 (90) 27 (100) (100) (100) 15/37 15 D Environment 0 0/15 (0) 0 (40.54) (100) 30/30 15 Intestine 0 0/30 (0) 0 (100) (100) 18/38 17 1 E Environment 13/18 (72.22) 11 (84.62) (47.37) (94.44) (5.56) 27/30 27 Intestine 0 25/27 (92.59) 25 (100) (90) (100) a Number of Campylobacter isolates tested for tetracycline resistance were less than the number of Campylobacter isolated from samples because some isolates could not be recovered after they were stored at -85 °C.
Table 4.3 Campylobacter and tetracycline resistance rates on five organic broiler farms
247
No. (%) of samples No. (%) of No. of samples positive positive for No. of Campylobacter tetracycline- for Campylobacter/ resistant to tetracycline/ resistant Farm Sample No. of samples cultured No. of Campylobacter Campylobacter for Campylobacter (%) C. jejuni C. coli isolates tested (%) positive for tet(O) gene
A Environment 19/32 (59.38) 19 (100) 0 5/19 (26.32) 5 (100)
Intestine 29/30 (96.67) 29 (100) 0 13/29 (44.83) 12 (92.31)
B a Environment 33/56 (58.93) 33 (100) 0 14/14 (100) 14 (100) 248 Intestine 42/44 (95.46) 27 (64.29) 15 (35.71) 31/42 (73.81) 29 (93.55)
C Environment 18/39 (46.15) 12 (66.67) 6 (33.33) 2/17 (11.77)a 2 (100)
Intestine 30/30 (100) 11 (36.67) 19 (63.33) 13/21 (61.91)a 13 (100)
a Number of Campylobacter isolates tested for tetracycline resistance were less than the number of Campylobacter isolated from samples because some isolates could not be recovered after they were stored at -85 °C.
Table 4.4 Campylobacter and tetracycline resistance rates on three organic turkey farms
248
No. of samples b No. of samples No. of No. of positive for positive for tetracycline- Operation pooled Farm Campylobacter resistant tet(O) type samples by standard Campylobacter tet(O) and collecteda culture method isolates gene MOMP gene
A* 18 1 0 0 0
A 12 0 0 5 0
Organic B 17 0 0 2 0 broiler C 10 1 1 0 0
D 8 1 0 0 0
E 11 0 0 0 0
A 7 1 0 0 0 Organic B 13 0 0 0 0 turkey C 9 2 0 0 0
a Number of environmental samples before coming in contact with the organically-raised broilers and turkeys were collected. b The presence of tet(O) and MOMP genes in DNA directly extracted from enrichment broth detected by PCR method. * Organic broiler farm A followed for the whole production cycle.
Table 4.5 The presence of Campylobacter and tet(O) gene in environmental samples before coming in contact with the organically-raised broilers and turkeys
249
Age (week) of the birds Number of samples positive when samples were collected for tetracycline-resistant Operation from Campylobacter Farm type Environmental Environmental Intestinal Intestinal and fecal and fecal samples samples samples samples A 9 10.5 5 2
Ba 2, 3, 4-12 10.5-12.5 19c 11 Organic Cb 5, 6, 7, 8 8.5 2d 27 broiler Db 3, 7, 8 8-9 0 0
Ea 3, 6 7.5 13e 25
A 19 19.5 5 13 Organic B 18 19 14 31 turkey C 20-24 21.5-25.5 2 13
a The birds in organic broiler farm B were raised in three separate areas: nursery barn for 2 weeks old birds, nursery barn for 3 weeks old birds, and pasture for 4 to 12 weeks old birds, while the birds in organic broiler farm E were raised in two separate areas: nursery barn for 3 weeks old birds and pasture for 6 weeks old birds. b The birds in organic broiler farm C and D were raised in the same area but separate pens. For organic broiler farm C, samples were collected from pen#1 and #2 (8 weeks old), pen#3 and #4 (7 weeks old), pen#5 (6 weeks old), and pen#6 (5 weeks old). For organic broiler farm D, samples were collected from pen#1 (8 weeks old), pen#2 (7 weeks old), and pen#3 (3 weeks old). c Tetracycline-resistant Campylobacter strains were isolated from all three areas that samples were collected. d Only samples from 8 weeks old birds were positive for tetracycline-resistant Campylobacter strains. e Tetracycline-resistant Campylobacter strains were isolated from 6 weeks old birds.
Table 4.6 Age of the birds when environmental and fecal samples and intestinal samples were collected from each organic poultry farm
250
Percent of Percent of intestinal samples positive for Operation Farm Study tetracycline Campylobacter type C. jejuni C. coli resistance spp.
Recent 40.74 90 88.89 11.11 B Previous 50 81.48 59.09 40.91
Organic Recent 90 100 100 0 broiler C Previous 100 88.17 74.39 25.61
Recent 0 100 100 0 D Previous 94.12 91.40 63.53 36.47
Recent 73.81 95.46 64.29 35.71 B Organic Previous 37.50 93.62 55.68 44.32 turkey Recent 61.91 100 36.67 63.33 C Previous 57.14 100 78.57 21.43
Table 4.7 Prevalence of Campylobacter and tetracycline resistance in organically-raised broilers and turkeys between our previous and recent studies
251
500 bp
200 bp 100 bp
1 2 3 4 5 6 7 8 9 10 11 12 13 14
Figure 4.1. Tetracycline resistance gene [tet(O) gene] of Campylobacter strains isolated from organically-raised broilers and turkeys. Lane 1, 2, 4, 5, 6, 7, 8, 11, and 12, tetracycline-resistant Campylobacter strains; lane 3, 9, and 10, tetracycline-susceptible Campylobacter strains; lane 13, positive control (C. jejuni ATCC 43503); lane 14, 100 bp DNA ladder (Promega Corporation, Madison, WI).
252
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