Influence of Stressors on Normal Intestinal Microbiota, Intestinal Morphology, and Susceptibility to Salmonella Enteritidis Colonization in Broilers K. M. Burkholder, K. L. Thompson, M. E. Einstein, T. J. Applegate, and J. A. Patterson1 Department of Animal Science, Purdue University, West Lafayette, IN 47907-2009 ABSTRACT In modern poultry production systems, environmental stressors may influence bird performance and susceptibility to pathogens such as Salmonella Enteritidis. Experiments were conducted to determine the influence of 24-h feed withdrawal and 24-h exposure to high temperature (30°C) on intestinal characteristics of broilers. Attachment of Salmonella Enteritidis to ileal tissue was determined using an in vitro ileal loop assay. Changes in commensal intestinal microbial populations were determined using denaturing gradient gel electrophoresis, and alterations in ileal morphology were determined histologically. Ex vivo attachment of Salmonella Enteritidis to ileal tissues increased by 1.5 logs (9.05 log10 vs. 7.59 log 10 Salmonella Enteritidis/g of
ileal tissue; P = 0.0006) in broilers fasted for 24 h. Similarly, ileal tissues from birds subjected to 30°C for 24 h had increased ex vivo attachment of Salmonella Enteritidis (8.77 log10 vs. 8.50 log10 Salmonella Enteritidis/g of ileum; P = 0.01) compared with birds held at 23°C. Exposure to 30°C for 24 h also altered microbial community structure in the ileum and cecum. Subjecting birds to 30°C for 24 h reduced crypt depth (6.0 vs. 7.8 μm, respectively; P = 0.002), but had no effect on villus height or villus:crypt ratio. This research shows that acute stressors in poultry production systems can cause changes in the normal intestinal microbiota and epithelial structure, which may lead to increased attachment of Salmonella Enteritidis.
Key words: broiler, heat stress, Salmonella Enteritidis, intestinal microbiota, intestinal morphology 2008 Poultry Science 87:1734–1741 doi:10.3382/ps.2008-00107
INTRODUCTION Salmonella is the leading cause of bacterial foodborne infection in the United States and causes approximately 1.2 million cases of human Salmonellosis each year (Centers for Disease Control and Prevention, 2007). The most commonly implicated source of foodborne Salmonellosis is consumption and handling of undercooked poultry products (Smithermann et al., 1984; Bryan and Doyle, 1995; Antunes et al., 2003). The correlation between poultry and foodborne illness has focused efforts toward identification of factors contributing to Salmonella infection in poultry, with the goal of decreasing carcass contamination and reducing animal morbidity and mortality. Environmental stress has been shown to be a factor that may induce colonization of food animals by enteric pathogens, facilitate horizontal transmission of pathogens between animals, increase pathogen shedding, and contribute to carcass contamination during processing (Rigby and Pettit, 1980; Mulder, 1995; Isaacson et al., 1999; Poppe, 1999; Jones et al., 2001). ©2008 Poultry Science Association Inc. Received March 7, 2008. Accepted May 19, 2008. 1 Corresponding author:
[email protected]
Stress is an important consideration in poultry production systems, because birds are routinely subjected to stressors such as feed withdrawal, temperature fluctuations, and confinement during transportation (Abeyesinghe et al., 2001; St-Pierre et al., 2003; Humphrey, 2006). Broilers are subjected to fasting to reduce the volume of intestinal contents before slaughter, and thus minimize the risk of carcass contamination by rupture of the intestinal tract during processing (May and Deaton, 1989). However, feed withdrawal has been associated with increased Salmonella colonization of the crop (Humphrey et al., 1993; Ramirez et al., 1997) and intestine (Bierer and Eleazer, 1965; Ramirez et al., 1997). Exposure to extreme temperature is an additional stressor encountered in seasonal environments, particularly during the summer months, and is also associated with increased intestinal colonization and fecal shedding of pathogens in poultry (Bailey, 1988). Stress can have a profound effect on overall physiology, animal health, and productivity. The gastrointestinal tract is particularly responsive to stressors, which can cause a variety of changes including alteration of the normal, protective microbiota (Tannock and Savage, 1974; Bailey and Coe, 1999, 2004) and decreased integrity of the intestinal epithelium (Saunders et al., 1994; Meddings and Swain, 2000; Soderholm et al., 2002). The commensal intestinal bacterial popula-
1734
STRESSORS ALTER NORMAL INTESTINAL CHARACTERISTICS IN BROILERS
tions can protect the host from pathogen colonization by competing for epithelial binding sites and nutrients, strengthening the intestinal immune response, and by producing antimicrobial bacteriocins (Guarner and Malagelada, 2003; Sansonetti, 2004; MacDonald and Monteleone, 2005). In addition, the intestinal epithelium serves as a barrier between luminal pathogens and the underlying blood supply (Sansonetti, 2004). Therefore, stress-induced perturbation of the normal intestinal microbiota or the integrity of the gut epithelium reduces innate protective mechanisms and may increase the potential for pathogens such as Salmonella to bind to and colonize the intestinal epithelium. Such colonization in poultry will increase the risk of carcass contamination during processing, and will increase the potential for Salmonella to translocate to the reproductive tract, where it can contaminate eggs during formation. The objective of the following study was to determine the influence of feed withdrawal and heat stress on intestinal microbial populations, small intestinal morphology, and intestinal susceptibility to Salmonella Enteritidis colonization in broilers. It was expected that stressors alone would induce changes in microbial community structure of the intestine and decrease intestinal integrity, and would increase intestinal susceptibility to Salmonella Enteritidis attachment.
MATERIALS AND METHODS Birds For feed withdrawal and heat stress experiments, birds were obtained from a larger study (Thompson and Applegate, 2006) in which 960 male Ross 308 broilers were allocated to 24 floor pens with 40 birds/pen. Birds were raised to 42 or 44 d of age on a standard corn-soybean meal diet that met or exceeded National Research Council (1994) nutrient requirements and had ad libitum access to feed and water. Initial ambient temperature was held at 35°C for newly hatched chicks, and was gradually decreased to 23°C by 21 d and held at 23°C for the duration of the 6-wk experiment. Animal procedures were approved for these studies by the Purdue Animal Care and Use Committee.
Stress Conditions Feed Withdrawal Study. At d 42, 10 birds were randomly chosen, killed by carbon dioxide asphyxiation, and sampled. Feed was withdrawn (0 h) from the remaining flock, and birds were kept on litter and given access to water for 4 h before being placed in transport crates for 20 h. After 24 h of feed withdrawal, 10 additional birds were killed by CO2 asphyxiation and tissues were sampled. Heat Stress Study. At d 44, 20 birds were randomly chosen and 10 were immediately sacrificed and sam-
1735
pled, whereas the remaining 10 were subjected to 30°C ambient temperature in floor pens for 24 h, with full access to feed and water, before euthanasia with CO2 overdose and sampling.
Intestinal Sampling Feed Withdrawal Study. A 10-cm section of the ileum was taken, starting 3 cm proximal to the ileo-cecal junction, from each bird for an ex vivo ileal challenge assay and was gently flushed with 0.05 M PBS (pH 7.2). Tissue sections were immediately placed in icecold Dulbecco’s Modified Eagle Medium + l-glutamine (DMEM, Mediatech, Herndon, VA) and kept on ice until used for an in vitro Salmonella challenge assay as described below. Heat Stress Study. Intestinal tissue and contents were obtained immediately after euthanasia, starting 3 cm proximal to the ileo-cecal junction and were collected as follows: 1) a 10-cm section of the ileum was taken for an in vitro Salmonella challenge assay as described below; 2) a 4-cm ileal section, 13 cm from the ileo-cecal junction, was collected for denaturing gradient gel electrophoresis (DGGE) analysis of microbial community structure. Ileal tissues were opened and contents were gently removed, placed in microfuge tubes, and immediately frozen at −20°C. Ileal tissues were gently flushed with 0.05 M PBS (pH 7.2), and were frozen at −20°C; 3) a 2-cm section of ileal tissue, 17 cm from the ileo-cecal junction, was collected for analysis of intestinal morphology, flushed with PBS and fixed in 10% neutral buffered formalin for 48 h; and 4) a 4-cm tissue section of the center of the cecum and cecal contents were obtained for DGGE analysis and processed as described above.
Ileal Loop Assay for Attachment of Salmonella Enteritidis Challenge Microorganism. Salmonella enterica serovar Enteritidis (NVSL 7759) was a gift from Tom Stable (National Animal Disease Center, Ames, IA) and was transfected with a kanamycin-resistance plasmid to allow selection in the presence of kanamycin (courtesy of Bruce Applegate, Purdue University, Lafayette, IN). The stock culture was grown in Luria Bertani (LB) broth containing 50 µg of kanamycin/mL (LB-kan) and stored with 20% (vol/vol) added glycerol. Fresh cultures were grown statically overnight in LB-kan broth, transferred to fresh LB-kan broth, and grown overnight for the challenge study. Bacterial cells were harvested by centrifugation at 6,000 × g at 4°C for 15 min, and were washed 3 times in equal volumes of sterile PBS. Cells were resuspended in DMEM to an optical density (OD)600 of 0.4 (approximately 1 × 106 cells/mL as determined by a prior standard curve). The inoculum was serially diluted and plated on LB broth to obtain the actual number of cells in the inoculum.
1736
Burkholder et al.
Ileal Loop Assay. The organ culture procedure of Naughton et al. (2001) was used. Briefly, the ileal sections were removed from DMEM, sealed at one end with 35-mm dialysis clamps, and inoculated with approximately 6 mL of Salmonella Enteritidis culture suspended in DMEM. The open end of the ileal section was sealed with dialysis clamps, the exterior was rinsed with PBS, and the ileal loops were incubated in 100 mL of DMEM for 1 h at 37°C in a rotary water bath in a 10% CO2 atmosphere. After incubation, ileal contents were removed, the interior and exterior of each section was rinsed with PBS, tissues were homogenized (T-25 basic homogenizer, Ultra-Turrax, IKA-Werke, Wilmington, NC), serially diluted in buffered peptone broth, and plated on LB agar plates containing 50 μg/ mL kanamycin. Plates were incubated at 37°C for 24 h and were enumerated for Salmonella Enteritidis. Differences due to feed withdrawal (0 vs. 24 h) or heat stress were determined and analyzed using the GLM procedure in SAS (SAS Inst. Inc., Cary, NC).
Intestinal Morphology After fixation in 10% neutral buffered formalin, a single 0.5-cm sample was cut from each ileal section, dehydrated with increasing concentrations (70, 80, 95, and 100%) of ethanol, cleared with xylene (SubX, Surgipath Medical Industries, Richmond, IL), and placed into polyfin embedding wax (Polysciences, Warrington, PA). Tissue sections (5 µm) were cut, floated onto slides, stained with hematoxylin (Gill #2, Sigma, St. Louis, MO) and eosin (Sigma), and measured for villus height and crypt depth using light microscopy and a micrometer. Measurements for villi length were taken from the tip of the villus to the valley between individual villi and measurements for crypt depth were taken from the valley between individual villi to the basolateral membrane. Eight villi and villus-associated crypts were measured for each sample. Morphology data were analyzed using the GLM procedure in SAS.
DGGE Genomic DNA was isolated from intestinal digesta and tissue samples using the Ultraclean Fecal DNA kit (MoBio, Solana Beach, CA), according to the manufacturer’s protocol. Samples were diluted 1:1 with sterile distilled water, and 0.25 g of the diluted sample was added to a bead beating tube containing beads, bead solution, and lysis solution. Cells were lysed by a combination of detergent and mechanical action using a MoBio Vortex Adapter (MoBio) on a standard vortex. From the lysed cells, the released DNA was bound to a silica spin filter. The filter was washed and DNA was eluted using DNase-free Tris buffer. The DGGE was performed according to previously described methods with modification (Muyzer et al., 1993; Muyzer and Smalla, 1998), using bacteria-specific PCR primers to conserved regions flanking the
variable V3 region of 16S rDNA. Each PCR reaction mixture contained 0.02 nmol of reverse primer (534r): 5′-ATT ACC GCG GCT GCT GG-3′ and 0.02 nmol of forward primer with a GC clamp (341FGC): 5′CGC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GGG GCC TAC GGG AGG CAG CAG-3′ (Integrated DNA Technologies, Skokie, IL), 3.75 units of Taq DNA Polymerase (Promega, Madison, WI), 5 to 10 ng of template DNA, 10× DNA Polymerase Buffer (containing 10 mM Tris-HCl, 50 mM KCl, and 0.1% Triton X-100), and 25 mM MgCl2. Amplifications were performed using a Gene Amp PCR System 9700 (Applied Biosystems, Foster City, CA) using the following program: 1) denaturation at 94°C for 3 min; 2) subsequent denaturation at 94°C for 1 min; 3) annealing at 65°C for 1 min, −0.5°C per cycle; 4) extension at 72°C for 1 min; 5) steps 2 to 4 repeated for 30 cycles; 6) denaturation at 94°C for 1 min; 7) annealing at 55°C for 1 min; 8) extension at 72°C for 1 min; steps 6 to 8 repeated for 7 cycles; 9) extension at 72°C for 7 min; 10) 4.0°C final holding temperature. Polyacrylamide gels (8% acrylamide-bisacrylamide ratio 37:5:1) were cast with a 40-60% urea:deionized formamide gradient. The 100% denaturing acrylamide contained 7 M urea and 40% deionized formamide. Amplified DNA was mixed with a 20% volume of 5× loading buffer [0.025% (wt/vol) bromophenol blue, 0.025% (wt/ vol) xylene cyanol, 47% (vol/vol) 0.1 M EDTA and 47% (vol/vol) glycerol], and 20 µL was loaded into each sample well (20-well comb). Gels were placed in a DCode Universal Mutation Detection System (Bio-Rad, Hercules, CA) and electrophoresed in 0.5× Tris-acetateEDTA buffer at 60 for 10 min at 200 V, followed by 16 h at 70 V. Gels were silver stained as described by Sambrook et al. (1989), using the following procedure: fixation solution for 3 min (77% ethanol, 4% glacial acetic acid, 16% distilled water), silver stain for 10 min (0.2% AgNO3, 99.8% fixation solution), developer for 15 to 20 min (0.001% NaBH4, 25% 1.5 M NaOH, 75% formaldehyde), and preservation solution for 7 min (27% ethanol, 12% glycerol, 61% distilled water). Fragment pattern relatedness was determined using Bionumerics Software (version 2.5; Applied Maths, Austin, TX), which determined the number of bands per sample and similarity coefficients for banding patterns between pairs of samples. A distance matrix was calculated using the DICE function and dendrograms were constructed from this matrix using the unweighted pair group means average (UPGMA) function. The degree of similarity of banding patterns between pairs of samples was represented as a similarity coefficient. All DGGE data were analyzed using the Mixed Model of SAS (SAS Inc., Cary, NC). Similarity coefficients between pairs of samples were segregated by treatment and similarity coefficients across treatments were used as an estimate of similarity assuming no treatment effect. Tukey-Kramer means separation was used to identify treatment differences. Significance was determined using P-value < 0.05.
1737
STRESSORS ALTER NORMAL INTESTINAL CHARACTERISTICS IN BROILERS
Table 1. Denaturing gradient gel electrophoresis similarity coefficients within control (23°C) and acute heat-stress (30°C for 24 h) treatments, and across treatments (cross-products)1 Treatment Item Ileal contents Ileal tissue Cecal contents Cecal tissue Figure 1. Effect of 24-h feed withdrawal on in vitro Salmonella Enteritidis attachment to broiler ileal tissue; n = 10 birds/time point. Error bars represent SEM.
RESULTS Effect of 24-h Feed Withdrawal on Ileal Susceptibility to Salmonella Enteritidis Attachment Intestinal tissues from fasted birds were more susceptible to pathogen attachment than tissues from control birds, with a 1.5 log increase (P = 0.01) in Salmonella Enteritidis associated with the ileal tissue of fasted birds compared with nonfasted controls (Figure 1).
Influence of an Acute High Temperature on Intestinal Susceptibility to Salmonella Enteritidis Adhesion, Commensal Microbial Populations, and Morphology of the Small Intestine of Broilers Broilers placed in the 30°C room showed behavioral signs of heat stress such as panting and spreading of wings. In the ileal loop assay, numbers of Salmonella Enteritidis associated with the ileum were greater (P = 0.0006) in heat-stressed birds (8.77 log10 cfu/g) compared with nonstressed birds (8.50 log10 cfu/g of ileum; Figure 2). Amplicon profiles for bacteria in the intestinal contents and tissues revealed differences in banding patterns between heat-stressed and nonstressed birds. Birds held at 30°C exhibited lower (P = 0.0001) similarity coefficients for microbial communities in ileal contents (Table 1) than did birds at 23°C. In all intestinal samples (Table 1), the similarity coefficients calculated across the 30 and 23°C treatments (crossproduct) were lower (P = 0.0001) than the similarity coefficients within individual treatments. The ileal tissue of heat-stressed animals contained fewer (P = 0.0251) amplicon fragments (bands) than the ileal tissue of nonstressed birds (Table 2). Influence of 24-h heat stress on ileal morphology was evaluated. In birds subjected to heat stress, crypt depth was reduced (P = 0.002) compared with nonstressed birds (60 vs. 78 µm, respectively; Table 3). Villus height and the villus:crypt ratio were unchanged in response to 24-h heat stress.
23°C
30°C
a
b
68.0 73.7a 58.5a 53.4b
Cross-products2
55.4 72.5a 60.4a 64.7a
c
41.7 46.3b 52.4b 44.5c
SEM 2.4 2.3 1.5 2.8
a–c Means within rows with different superscript letters are significantly different (P < 0.05). 1 Results represent the mean similarity coefficients for 10 birds/ treatment. 2 Indicates similarity coefficients for across treatment comparison.
DISCUSSION Salmonella Enteritidis and Salmonella Typhimurium are widely recognized as causative agents in poultry-related foodborne illness outbreaks. Since 1996, the incidence of Salmonella Typhimurium-induced Salmonellosis has declined in the United States, but the rate of Salmonella Enteritidis foodborne infection has remained steady (Centers for Disease Control and Prevention, 2006). Therefore, identification of on-farm risk factors for Salmonella colonization in poultry is necessary to develop strategies for reducing Salmonella load in animals at time of slaughter and processing. Because stressors such as feed withdrawal and high environmental temperature have been linked to increased incidence of enteric disease in poultry and other foodproducing animals, our study aimed to identify ways in which these stressors affect normal intestinal conditions, such as the protective commensal microbiota and the integrity of the intestinal epithelium (Abeyesinghe et al., 2001; St-Pierre et al., 2003; Humphrey, 2006). We also examined how these stressors influenced the binding of Salmonella Enteritidis to intestinal tissue.
Adhesion of Salmonella Enteritidis to Intestinal Tissue Following Stress Exposure The ileal loop assays demonstrated that stress due to 24-h feed withdrawal and exposure to high temperatures is associated with increased susceptibility of intestinal tissues to Salmonella Enteritidis colonization. Intestinal tissues from fasted birds had significantly greater attachment of Salmonella than did tissues from control birds (Figure 1). These results are in agreement with others who have observed increases in enteric pathogen colonization and shedding in birds subjected to feed withdrawal (Bierer and Eleazer, 1965; Humphrey et al., 1993; Ramirez et al., 1997). Less is known about the effect of heat stress on pathogen colonization of the intestinal mucosa. In the study reported here, numbers of Salmonella Enteritidis associated with the ileum were greater in heat-stressed birds (8.77 log10 cfu/g) compared with nonstressed birds (8.50 log10 cfu/g
1738
Burkholder et al.
Table 2. Influence of heat stress on number of bands present in various intestinal ecosystems of broilers as determined by denaturing gradient gel electrophoresis analysis1
Table 3. Influence of heat stress on small intestinal morphological characteristics, including villus height, crypt depth, and the villus:crypt ratio1 Treatment
Treatment Item
23°C
30°C
SEM
Ileal contents Ileal tissue Cecal contents Cecal tissue
15.0 18.4a 18.5 10.7
14.6 13.7b 19.4 8.9
1.1 1.9 1.3 2.0
a,b
Means within rows with different superscript letters are significantly different (P < 0.05). 1 Results represent the mean similarity coefficients for 10 birds/ treatment.
ileum; Figure 2), indicating that stress may contribute to increased intestinal colonization by Salmonella. This is supported by previous findings that heat stress may have damaging effects on mucosal structure (Sengupta and Sharma, 1993). In addition, heat shock proteins, whose expression can be induced by high ambient temperature and other environmental stressors (Lindquist, 1986; Lindquist and Craig, 1988), may act as epithelial surface receptors for pathogen binding (Dziewanowska et al., 2000; Wampler et al., 2004).
Changes in Intestinal Microbial Populations Influenced by Stressors The normal gut microbiota is a complex ecosystem that can benefit the host by serving as a barrier to pathogen colonization (Van der Waaij, 1989). Alteration of this protective barrier may leave the host more susceptible to colonization by enteric pathogens (Durant et al., 1999). There is a linear reduction in the mucus lining the intestinal tract over a 24-h fast, as well as changes in intestinal morphology (Thompson and Applegate, 2006). Hinton et al. (2000) showed that there were increases in intestinal Enterobacteriaceae and cecal aerobes with a concurrent decrease in lactic acid bacteria in broilers subjected to a 24-h feed withdrawal. Neurohormones associated with stress can increase growth and virulence factor expression in microbes including Escherichia coli, Yersinia enteroco-
Figure 2. Influence of heat stress on ileal susceptibility to Salmonella Enteritidis attachment; n = 10 birds/time point. Error bars represent SEM.
Item
23°C
30°C
SEM
Villus height, µm Crypt depth, µm Villus:crypt
450.0 78.0a 5.7
390.0 60.0b 6.9
27.6 5.1 1.0
a,b Means within rows with different superscript letters are significantly different (P < 0.05). 1 Means represent 10 birds per treatment.
litica, and Pseudomonas aeruginosa in vitro (Lyte and Ernst, 1992; Lyte et al., 1997b; Rahman et al., 2000). Release of norepinephrine in the intestinal tract increased the number of gram-negative bacteria within the lumen (Lyte and Bailey, 1997a). Most studies to date used traditional microbiological culturing techniques to examine the influence of stressors on a few specific bacterial populations. In our study, we used DGGE to evaluate stress-induced changes in the overall intestinal microbial profile, rather than focusing on a few specific microorganisms. One possible limitation of analyzing similarity coefficients within treatments is that each treatment analysis is separate; thus, one may end up with numerically comparable similarity coefficients, but banding patterns within treatment may differ, as the data from ileal tissue (Table 1 and Figure 3b) demonstrate. The dendrogram shows that banding patterns were highly similar within each treatment, with similarity coefficients of 73.7 and 72.5 for birds at 23 and 30°C, respectively. However, the banding patterns are obviously different between treatments. Thus, we used across-treatment pairwise comparisons (cross-product) to determine differences between treatments when similarity coefficients were not significantly different. In essence, the cross-products represent the similarity coefficient of the population as a whole. If treatment similarity coefficients are significantly different than the cross-product, even if they are numerically similar to each other, then the microbial community structure between treatments is different. The DGGE data show changes in the commensal intestinal bacterial populations of birds subjected to heat-stress, as indicated by changes in similarity coefficients in intestinal tissue and contents and the decreased number of bands in the ileal mucosa of birds subjected to 24-h heat stress (Tables 1 and 2, Figure 3). Bacterial communities in the ileal contents from birds at 30°C had lower similarity coefficients than in the ileal contents from birds at 23°C. The decrease in similarity of bacterial communities in heat-stressed birds does not necessarily mean that there is an increase in the number of bacterial species (diversity) within individual birds, but that there is increased variation in the composition of intestinal bacteria between birds. In contrast, bacterial communities in the cecal tissue of birds at 30°C had greater similarity values than those
STRESSORS ALTER NORMAL INTESTINAL CHARACTERISTICS IN BROILERS
1739
Figure 3. Dendrogram representing similarity of microbial banding patterns in ileal contents (a) and ileal tissue (b) of heat stressed (30°C) and control (23°C) birds. Relative similarity of band patterns is indicated by their grouping on the dendrogram. Each line on the dendrogram represents an individual bird. The percentage similarity coefficient is indicated on the graduated bar at the top of each panel.
in birds at 23°C, suggesting that the bacterial communities between birds were more similar. Figure 3a shows a slightly more complex dendrogram. It is difficult with complex dendrograms to visually determine treatment differences; thus, the use of the cross-product helps differentiate treatment differences. In all intestinal samples, the similarity coefficients across treatments were lower than the similarity coefficients within individual treatments, indicating that the intestinal microbial community structure was significantly changed when birds were exposed to high temperatures for 24 h. The ileal tissue from birds held at 30°C contained fewer amplicon fragments than the ileal tissue from birds at 23°C (Table 2), indicating that exposing birds to high temperatures for 24 h caused a reduction in microbial species associated with the ileal wall. These data show that greater changes occur in the ileum than the cecum when birds are subjected to elevated temperatures for 24 h, indicating that microbial populations in the ileum may be more sensitive to short-term changes than those in the cecum.
Influence of Stress on Intestinal Morphology Maintenance of normal morphology and structural integrity of the small intestine are imperative for preventing bacterial translocation from the intestinal tract. Extensive research has been conducted on the influence of stress due to fasting or weaning on small
intestinal morphology. These nutritional stressors have deleterious effects on the absorptive epithelium of the intestine, resulting in reduction in villus height and crypt depth (Mayhew, 1987; Yamauchi et al., 1995; Yamauchi et al., 1996). Less is known about the effect of high environmental temperature on intestinal morphology. Birds subjected to 30°C for 24 h had reduced crypt depth compared with birds at 23°C (Table 3). Villus height and the villus:crypt ratio were unchanged in birds exposed to 30°C. There are several possible reasons why ileal villus height was unchanged in response to elevated temperature, including the short duration of the stressor and the resistance of the ileum to structural change compared with other regions of the small intestine (Yamauchi et al., 1995). It is likely that changes in cell proliferation would be observed first in the stem cells of the crypt rather than the villus because of the high proliferative activity of the crypt (Yamauchi et al., 1995). Morphological changes in response to fasting occur more rapidly in the proximal two-thirds of the small intestine than in the ileum. In feed withdrawal studies with chickens, the structure of the duodenum and jejunum mucosa changes rapidly, often within 36 h of the onset of stress, whereas the ileum maintains its normal morphology longer and requires extended periods of stress to influence its structure (Yamauchi et al., 1995, 1996). Data indicate that acute (24-h) stressors can invoke significant changes in the normal intestinal microbiota, intestinal morphology, and in vitro susceptibility
1740
Burkholder et al.
for Salmonella Enteritidis attachment to the ileum in broilers. The findings from the 24-h heat stress study suggest that stressors could act in several ways to increase intestinal susceptibility to in vitro Salmonella Enteritidis attachment. By disrupting the normal protective microbiota and altering intestinal morphology, stressors create an opportunity for pathogens to colonize the intestine. Exposure to stressors may also cause subtle changes in epithelial mucus production and mucus composition, which may occur before gross anatomical changes in the intestinal morphology. Alterations in the mucus layer could change the attachment capabilities of both commensal and pathogenic microorganisms (Deplancke and Gaskins, 2001), which could account for changes in DGGE community profiles and for changes in intestinal susceptibility to Salmonella Enteritidis attachment. These findings are important for the poultry industry, where broilers are routinely fasted for gastrointestinal emptying before transportation, and where birds are often exposed to high environmental temperatures during the summer months. Future work should focus on the use of prophylactic measures to prevent or reduce the stress-induced alteration of the intestinal microbiota and changes in gut integrity. Probiotic organisms such as lactobacilli, bifidobacteria, some strains of E. coli, and yeast offer promise for reducing pathogen colonization when fed orally (Line et al., 1997; Patterson and Burkholder, 2003; Wehkamp et al., 2004; Parvez et al., 2006). When administered consistently, probiotic organisms can colonize and form a niche in the intestine and may be a useful dietary treatment if administered before feed withdrawal or transportation and processing. In addition, nondigestible oligosaccharides, or prebiotics, have been shown to enhance intestinal growth of probiotic or beneficial commensal organisms (Bouhnik et al., 2004; Macfarlane et al., 2008). The fermentation of prebiotic complex carbohydrates by intestinal microbes produces volatile fatty acids, which promote epithelial cell proliferation and renewal (Macfarlane et al., 2008). Such activity may enhance integrity of the intestinal epithelium. Therefore, dietary supplementation of the flock with prebiotics before periods of anticipated stress may also limit the damage to the intestinal epithelium elicited by stressors. In conclusion, understanding the mechanism by which stress alters normal intestinal characteristics and induces susceptibility to enteric infection is an important first step in designing on-farm strategies aimed at reducing pathogen contamination in poultry.
ACKNOWLEDGMENTS These studies were conducted as part of Hatch project IND051084, administered by Purdue University. We thank Jessamyn Bacon (Pacific Veterinary Specialist and Emergency Services, Soquel, CA) for her technical assistance.
REFERENCES Abeyesinghe, S. M., C. M. Wathes, C. J. Nicol, and J. M. Randall. 2001. The aversion of broiler chickens to concurrent vibrational and thermal stressors. Appl. Anim. Behav. Sci. 73:199–215. Antunes, P., C. Réu, J. C. Sousa, L. Peixe, and N. Pestana. 2003. Incidence of Salmonella from poultry products and their susceptibility to antimicrobial agents. Int. J. Food Microbiol. 82:97–103. Bailey, J. S. 1988. Integrated colonization control of Salmonella in poultry. Poult. Sci. 67:928–932. Bailey, M. T., and C. L. Coe. 1999. Maternal separation disrupts the integrity of the intestinal microflora in infant rhesus monkeys. Dev. Psychobiol. 35:146–155. Bailey, M. T., and C. L. Coe. 2004. Prenatal stress alters bacterial colonization of the gut in infant monkeys. J. Pediatr. Gastroenterol. Nutr. 38:414–421. Bierer, B. W., and T. H. Eleazer. 1965. Clinical salmonellosis accidentally induced by feed and water deprivation of oneweek-old broiler chicks. Poult. Sci. 44:1606–1607. Bouhnik, Y., L. Raskine, G. Simoneau, E. Vicaut, C. Neut, B. Flourie, F. Brouns, and F. R. Bornet. 2004. The capacity of nondigestible carbohydrates to stimulate fecal bifidobacteria in healthy humans: A double-blind, randomized, placebo-controlled, parallel-group, dose-response relation study. Am. J. Clin. Nutr. 80:1658–1664. Bryan, F. L., and M. P. Doyle. 1995. Health risks and consequences of Salmonella and Campylobacter jejuni in raw poultry. J. Food Prot. 58:326–344. Centers for Disease Control and Prevention. 2006. FoodNet surveillance report for 2004. Final report. www.cdc.gov/ foodnet/annual/2004.pdf Accessed Oct. 22, 2007. Centers for Disease Control and Prevention. 2007. Foodborne infections: General information and technical information. Division of Bacterial and Mycotic Diseases http://www.cdc. gov/nczved/dfbmd/disease_listing/salmonellosis_gi.html Accessed October 2007. Deplancke, B., and H. R. Gaskins. 2001. Microbial modulation of innate defense: Goblet cells and the intestinal mucus layer. Am. J. Clin. Nutr. 73(Suppl.):1131(S)–1141(S). Durant, J. A., D. E. Corrier, J. A. Byrd, L. H. Stanker, and S. C. Ricke. 1999. Feed deprivation affects crop environment and modulates Salmonella enteritidis colonization and invasion of Leghorn hens. Appl. Environ. Microbiol. 65:1919–1923. Dziewanowska, K., A. R. Carson, J. M. Patti, C. F. Deobald, K. W. Bayles, and G. A. Bohach. 2000. Staphylococcal fibronectin binding protein interacts with heat shock proteins 60 and integrins: Role in internalization by epithelial cells. Infect. Immun. 68:6321–6328. Guarner, F., and J. R. Malagelada. 2003. Gut flora in health and disease. Lancet 361:512–519. Hinton, A. Jr., R. J. Buhr, and K. D. Ingram. 2000. Physical, chemical, and microbiological changes in the ceca of broiler chickens subjected to incremental feed withdrawal. Poult. Sci. 79:483–488. Humphrey, T. 2006. Are happy chickens safer chickens? Poultry welfare and disease susceptibility. Br. Poult. Sci. 47:379–391. Humphrey, T. J., A. Baskerville, A. Whitehead, B. Rowe, and A. Henley. 1993. Influence of feeding patterns on the artificial infection of laying hens with Salmonella Enteritidis phage type 4. Vet. Rec. 132:407–409. Isaacson, R. E., L. D. Firkins, R. M. Weigel, F. A. Zuckermann, and J. A. Di Petrio. 1999. Effect of transportation and feed withdrawal on shedding of Salmonella typhimurium among experimentally infected pigs. Am. J. Vet. Res. 60:1155–1158. Jones, P. H., J. M. Roe, and B. G. Miller. 2001. Effects of stressors on immune parameters and on the faecal shed-
STRESSORS ALTER NORMAL INTESTINAL CHARACTERISTICS IN BROILERS ding of enterotoxigenic Escherichia coli in piglets following experimental inoculation. Res. Vet. Sci. 70:9–17. Lindquist, S. 1986. The heat-shock response. Annu. Rev. Biochem. 55:1151–1191. Lindquist, S., and E. A. Craig. 1988. The heat-shock proteins. Annu. Rev. Genet. 22:631–677. Line, J. E., J. S. Bailey, N. A. Cox, and N. J. Stern. 1997. Yeast treatment to reduce Salmonella and Campylobacter populations associated with broiler chickens subjected to transport stress. Poult. Sci. 76:1227–1231. Lyte, M., and M. T. Bailey. 1997a. Neuroendocrine-bacterial interactions in a neurotoxin-induced model of trauma. J. Surg. Res. 70:195–201. Lyte, M., A. K. Erickson, B. P. Arulanandan, C. D. Frank, M. A. Crawford, and D. H. Francis. 1997b. Norepinephrineinduced expression of the K99 pilus adhesion of enterotoxigenic Escherichia coli. Biochem. Biophys. Res. Commun. 232:682–686. Lyte, M., and S. Ernst. 1992. Catecholamine-induced growth of gram-negative bacteria. Life Sci. 50:203–212. MacDonald, T. T., and G. Monteleone. 2005. Immunity, inflammation, and allergy in the gut. Science 307:1920– 1925. Macfarlane, G. T., H. Steed, and S. Macfarlane. 2008. Bacterial metabolism and health-related effects of galactooligosaccharides and other prebiotics. J. Appl. Microbiol. 104:305–344. May, J. D., and J. W. Deaton. 1989. Digestive tract clearance of broilers cooped and deprived of water. Poult. Sci. 68:627–630. Mayhew, T. M. 1987. Quantitative ultrastructural study on the responses of microvilli along the small bowel to fasting. J. Anat. 154:237–243. Meddings, J. B., and M. G. Swain. 2000. Environmental stress induced a generalized increase in gastrointestinal permeability mediated by endogenous glucocorticoids in the rat. Gastroenterology 119:1019–1028. Mulder, R. W. A. W. 1995. Impact of transport and related stresses on the incidence and extent of human pathogens in pig meat and poultry. J. Food Saf. 15:239–246. Muyzer, G., E. C. de Waal, and A. G. Uitterlinden. 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol. 59:695–700. Muyzer, G., and K. Smalla. 1998. Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Antonie Van Leeuwenhoek 73:127–141. National Research Council (NRC). 1994. Nutrient Requirements of Poultry. 9th rev. ed. National Academy Press, Washington, DC. Naughton, P. J., L. L. Mikkelsen, and B. B. Jensen. 2001. Effects of nondigestible oligosaccharides on Salmonella enterica serovar Typhimurium and nonpathogenic Escherichia coli in the pig small intestine in vitro. Appl. Environ. Microbiol. 67:3391–3395. Parvez, S., K. A. Malik, S. Ah Kang, and H. Y. Kim. 2006. Probiotics and their fermented food products are beneficial for health. J. Appl. Microbiol. 100:1171–1185. Patterson, J. A., and K. M. Burkholder. 2003. Application of prebiotics and probiotics in poultry production. Poult. Sci. 82:627–631. Poppe, C. 1999. Epidemiology of Salmonella enterica serovar Enteritidis. Pages 3–18 in Salmonella enterica in Humans and Animals. Iowa State University Press, Ames.
1741
Rahman, H., R. Reissbrodt, and H. Tschape. 2000. Effect of norepinephrine on growth of Salmonella and its enterotoxin production. Indian J. Exp. Biol. 38:285–286. Ramirez, G. A., L. L. Sarlin, D. J. Caldwell, C. R. Yezak Jr., M. E. Hume, D. E. Corrier, J. R. Deloach, and B. M. Hargis. 1997. Effect of feed withdrawal on the incidence of Salmonella in the crops and ceca of market age broiler chickens. Poult. Sci. 76:654–656. Rigby, C. E., and J. R. Pettit. 1980. Changes in the Salmonella status of broiler chickens subjected to simulated shipping conditions. Can. J. Comp. Med. 44:374–381. Sambrook, J., E. F. Fritsch, and T. Maniatis. (1989) Molecular Cloning: A Laboratory Manual. 2nd ed. Cold Spring Harbor Laboratory Press, Woodbury, NY. Sansonetti, P. 2004. War and peace at mucosal surfaces. Nat. Rev. Immunol. 4:954–964. Saunders, P. R., U. Kosecka, D. M. McKay, and M. H. Perdue. 1994. Acute stressors stimulate ion secretion and increase epithelial permeability in rat intestine. Am. J. Physiol. Gastrointest. Liver Physiol. 267:G794–G799. Sengupta, A., and R. K. Sharma. 1993. Acute heat stress in growing rats: Effect on small intestinal morphometry and in vivo absorption. J. Therm. Biol. 18:145–151. Smithermann, R. E., C. A. Genigeorgis, and T. B. Farver. 1984. Preliminary observations on the occurrence of Campylobacter jejuni at four California chicken ranches. J. Food Prot. 47:293–298. Soderholm, J. D., D. A. Yates, M. G. Gareau, P.-C. Yang, G. MacQueen, and M. H. Perdue. 2002. Neonatal maternal separation predisposes adult rats to colonic barrier dysfunction in response to mild stress. Am. J. Physiol. Gastrointest. Liver Physiol. 283:G1257–G1263. St-Pierre, N. R., B. Cobanov, and G. Schnitkey. 2003. Economic losses from heat stress by US livestock industries. J. Dairy Sci. 86(E Suppl.):E52–E77. Tannock, G. W., and D. C. Savage. 1974. Influences of dietary and environmental stress on microbial populations in the murine gastrointestinal tract. Infect. Immun. 9:591– 598. Thompson, K. L., and T. J. Applegate. 2006. Feed withdrawal alters small-intestinal morphology and mucus of broilers. Poult. Sci. 85:1535–1540. Van der Waaij, D. 1989. The ecology of the human intestine and its consequences for overgrowth by pathogens such as Clostridium difficile. Annu. Rev. Microbiol. 43:69–87. Wampler, J. L., K.-P. Kim, Z. Jaradat, and A. K. Bhunia. 2004. Heat shock protein 60 acts as a receptor for the Listeria adhesion protein in Caco-2 cells. Infect. Immun. 72:931–936. Wehkamp, J., J. Harder, K. Wehkamp, B. Wehkamp-von Meissner, M. Schlee, C. Enders, U. Sonnenborn, S. Nuding, S. Bengmark, K. Fellermann, J. M. Schröder, and E. F. Stange. 2004. NF-κB- and AP-1-mediated induction of human beta defensin-2 in intestinal epithelial cells by Escherichia coli Nissle 1917: A novel effect of a probiotic bacterium. Infect. Immun. 72:5750–5758. Yamauchi, K., H. Kamisoyama, and Y. Isshiki. 1996. Effects of fasting and refeeding on structures of the intestinal villi and epithelial cells in White Leghorn hens. Br. Poult. Sci. 37:909–921. Yamauchi, K., K. Yamamoto, and Y. Ishiki. 1995. Morphological alterations of the intestinal villi and absorptive epithelial cells in each intestinal part in fasted chickens. Jpn. Poult. Sci. 32:241–251.
