ISSN: 2159-8967 www.AFABjournal.com
Volume 3, Issue 1 2013
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EDITORIAL BOARD Sooyoun Ahn
W.K. Kim
University of Florida, USA
University of Manitoba, Canada
Walid Q. Alali
M.B. Kirkham
University of Georgia, USA
Kansas State University, USA
Kenneth M. Bischoff
Todd Kostman
NCAUR, USDA-ARS, USA
University of Wisconsin, Oshkosh, USA
Debabrata Biswas
Y.M. Kwon
University of Maryland, USA
University of Arkansas, USA
Claudia S. Dunkley
Maria Luz Sanz
University of Georgia, USA
MuriasInstituto de Quimica Organic General, Spain
Lawrence Goodridge
Melanie R. Mormile
Colorado State University, USA
Missouri University of Science and Tech., USA
Leluo Guan
Rama Nannapaneni
University of Alberta, Canada
Mississippi State University, USA
Joshua Gurtler
Jack A. Neal, Jr.
ERRC, USDA-ARS, USA
University of Houston, USA
Yong D. Hang
Benedict Okeke
Cornell University, USA
Auburn University at Montgomery, USA
Divya Jaroni
John Patterson
Oklahoma State University, USA
Purdue University, USA
Weihong Jiang Shanghai
Toni Poole
Institute for Biol. Sciences, P.R. China
FFSRU, USDA-ARS, USA
Michael Johnson
Marcos Rostagno
University of Arkansas, USA
LBRU, USDA-ARS, USA
Timothy Kelly
Roni Shapira
East Carolina University, USA
Hebrew University of Jerusalem, Israel
William R. Kenealy
Kalidas Shetty
Mascoma Corporation, USA
North Dakota State University, USA
Hae-Yeong Kim Kyung Hee University, South Korea Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 3, Issue 1 - 2013
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EDITORIAL STAFF EDITOR-IN-CHIEF Steven C. Ricke University of Arkansas, USA
EDITORS Todd R. Callaway FFSRU, USADA-ARS, USA Cesar Compadre University of Arkansas for Medical Sciences, USA
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TECHNICAL EDITOR Jessica C. Shabatura Fayetteville, USA
ONLINE EDITION EDITOR C.S. Shabatura Fayetteville, USA
Philip G. Crandall University of Arkansas, USA
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Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 3, Issue 1 - 2013
TABLE OF CONTENTS REVIEW 17
Greenhouse Gas Emissions from Livestock and Poultry C. S. Dunkley and K. D. Dunkley
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Can Salmonella Reside in the Human Oral Cavity? S. A. Sirsat
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Shiga Toxin-Producing Escherichia coli (STEC) Ecology in Cattle and Management Based Options for Reducing Fecal Shedding T. R. Callaway, T. S. Edrington, G. H. Loneragan, M. A. Carr, D. J. Nisbet
ARTICLES 6
Growth of Acetogenic Bacteria In Response to Varying pH, Acetate Or Carbohydrate Concentration R. S. Pinder, and J. A. Patterson
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Independent Poultry Processing in Georgia: Survey of Producers’ Perspective E. J. Van Loo, W. Q. Alali, S. Welander, C. A. O’Bryan, P. G. Crandall, S. C. Ricke
Introduction to Authors 79
Instructions for Authors
The publishers do not warrant the accuracy of the articles in this journal, nor any views or opinions by their authors. Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 3, Issue 1 - 2013
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www.afabjournal.com Copyright © 2013 Agriculture, Food and Analytical Bacteriology
Growth of Acetogenic Bacteria In Response to Varying pH, Acetate Or Carbohydrate Concentration R. S.Pinder1,2 and J. A. Patterson1
Animal Science Dept, Purdue University, West Lafayette, IN 47907-1026 2 Current address; 7855 South 600 East, Brownsburg, IN 46112
1
ABSTRACT Acetogens have only been isolated in low numbers from ruminal contents, even though the majority of acetogens isolated from ruminal contents are capable of utilizing both H2 and soluble carbohydrates present in ruminal fluid (e.g. glucose and cellobiose). The much higher methanogenic affinity for hydrogen has been suggested to determine the prevalence of methanogens over acetogens in many ecosystems, suggesting that other environmental factors determine the number of acetogens present in ruminal fluid. We report the effects of carbohydrate concentration, pH and acetate concentration on the growth of two ruminal acetogenic isolates (A10 and H3HH). The minimum amount of glucose necessary for growth (threshold) of A10 (111 μM) and H3HH (56 μM) was greater than the glucose concentration previously detected in bovine ruminal fluid (8-17 μM). However, the threshold of H3HH on cellobiose (14 μM) was much lower than the actual concentration previously detected in ruminal fluid (110-175 μM), suggesting that this organism could survive in the rumen using cellobiose as an energy source. Isolate A10 had a sufficiently high threshold for cellobiose (139 μM) to suggest that, at least for certain periods, the concentration of cellobiose in ruminal contents could be too low to support growth of this isolate. The growth rate of isolate A10 was decreased by 50 % when the pH of the growth medium was lowered from 6.6 to 5.5. A similar decrease in growth rate was observed with isolate H3HH. Increasing the acetate concentration of the growth medium decreased the growth of both isolates as well. Moreover, the effect of high acetate concentration was more discernible at lower pH. The present results suggest that pH and volatile fatty acid concentration may be key factors limiting the growth of acetogens isolated from ruminal contents. Keywords: acetogens, rumen, hydrogen, pH, carbohydrate concentration Agric. Food Anal. Bacteriol. 3: 6-16, 2013
Correspondence: J. A. Patterson, jpatters@purdue.edu Tel: +1 -765-494-4826 Fax: +1-765-494-9347
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INTRODUCTION
MATERIALS AND METHODS
Hydrogen produced during microbial degradation of cellulose in ruminal contents must be eliminated in order to maintain the appropriate conditions for efficient conversion of cellulose to volatile fatty acids (Zinder, 1993). Normally, methanogens remove H2 from the rumen through methane production from H2/CO2. However, substantial interest exists to reduce methane emissions from ruminants to minimize this contribution to global warming. One potential approach is to use acetogenic bacteria to produce acetate as an alternate H2 sink. Acetogens are a diverse group of organisms that are capable
Organisms Isolates A10 and H3HH, previously described ruminal isolates (Boccazzi and Patterson, 2011; Pinder and Patterson, 2011, 2012) were used in all the experiments described herein. These organisms were maintained in acetogenic medium.
Growth medium The acetogen medium of Boccazzi and Patterson (2011) was used except where indicated otherwise.
of utilizing H2/CO2 to form acetate, a product useful to the host animal. However, acetogens are typically isolated in low numbers from ruminal contents (Morvan et al., 1994; LeVan et al., 1998) despite being capable of utilizing some of the carbohydrates glucose, cellobiose) present in ruminal fluid (Boccazzi and Patterson, 2011; Genthner et al., 1981). This apparent contradiction suggests that other ruminal environmental factors limit the growth of acetogens in ruminal contents. Several possibilities exist, including: 1) chemical factors (pH and volatile fatty acid concentration) which interfere with the growth of acetogens; 2) acetogens are not capable of utilizing carbohydrates at the concentration present in ruminal fluid; 3) acetogens are not capable of growing at a pace fast enough to avoid being washed out of rumen contents; 4) acetogens cannot compete with methanogens for H2/CO2. The goal of the research presented herein was to determine 1) the minimum concentration of glucose, maltose and cellobiose needed for growth of acetogens, 2 ) the growth rates of acetogens, and 3) the influence of volatile fatty acid concentrations and pH on growth of acetogens. We report that the growth of isolates A10 and H3HH, two acetogenic bacteria isolated and characterized from ruminal contents
In experiments where the initial concentration of carbohydrates was the treatment tested, ruminal fluid was omitted to reduce the amount of carbohydrates present in basal medium. In experiments where the initial pH of the medium was one of the treatments imposed, the composition of the medium was modified by eliminating the Na2CO3 which buffered the medium too strongly for reliable pH adjustment below 6.5. In addition, the medium was gassed with oxygen-free 100 % N2 rather than 100 % CO2, during preparation and dispensing. Preliminary experiments showed that the buffering capacity of this modified medium was sufficient for short-term (less than 12 h) experiments with low (0.05 g/L) amounts of carbohydrate substrate.
(Boccazzi and Patterson, 2011; Pinder and Patterson, 2011, 2012) may be affected by the carbohydrate concentrations found in ruminal fluid but that growth of these organisms is severely influenced by the combination of low pH and high acetate concentration.
where the initial carbohydrate concentration was the treatment regimen imposed, the growth rate (at each carbohydrate concentration) was determined and the threshold of carbohydrate utilization was estimated as the point where growth rate was greater than
Batch cultures Glucose, maltose or cellobiose (from 0.2 μ-filter sterilized stock solutions) were added as indicated to tubes containing 10 mL of acetogen medium. Tubes were inoculated with 0.1 mL of an overnight culture, measured for the initial OD (absorbance at 660 nm) and placed in a 37°C water bath. The contents of the tubes were mixed and the OD of the tubes measured and recorded at intervals as shown. For experiments
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baseline (without added carbohydrates). For experiments where the pH and acetate concentration of the media were the treatments imposed, growth rates of the isolates was estimated by plotting the OD values for each culture and determining the apparent maximum growth rate from the steepest portion of the growth curve.
kit (Sigma Chemical Co., St. Louis, MO). The initial glucose concentration was determined on uninoculated control bottles. After sampling, the media flow to the growth vessels was adjusted to obtain the next dilution rate.
RESULTS Continuous cultures An anaerobic continuous culture system as described by Boccazzi and Patterson, (2011) was used. The growth medium consisted of acetogen medium supplemented with 1.4 mM glucose. The medium
Effect of dilution rate on growth of isolate A10 and H3HH Previously, we determined the maximum growth rate of isolate A10 to be approximately 0.47 h-1 (Pin-
reservoir and growth vessels were gassed with 100 % CO2 at a flow of approximately 50 mL min-1. After the system was reduced, the growth vessels (operating capacity; approximately 215 mL) were inoculated with 20 mL of an overnight culture. The bacterial culture was allowed to grow as a batch culture for 6 h after which the medium flow was initiated. The cultures were allowed at least ten turnovers to reach steady state conditions before sampling. Turnover rate was determined by collecting the outflow from the growth vessels. Sample collection was accomplished by diverting the outflow from the growth vessels for 10 min into ice-cooled collecting vessels. Aliquots of the outflow were analyzed for volatile fatty acids (VFA), glucose and bacterial DM as described below. Concentrations of volatile fatty acids in the culture supernatant were determined using a Hewlett Packard 5890 gas chromatograph (Hewlett Packard Co., Palo Alto, CA) fitted with a glass column packed with GP 60/80 carbopack C / 0.3% Carbowax M / 0.1 % H3PO4 (Supelco, Inc.; Bellefonte, PA). The injector and detector temperatures were set at 200°C while the column temperature was set at 135°C. Formate concentrations in the culture supernatant were determined using a formate dehydrogenase assay (Schaller and Triebig, 1983). The pH of the culture
der and Patterson, 2012). Both isolate A10 and isolate H3HH attached to the surfaces of the growth vessels, perhaps due to extracellular polysaccharide. The net result was that apparent maximum growth rates in continuous culture approached 1.2 h-1 (Figure 1), This is much higher than the maximum growth rate estimated in batch culture and is probably a result of the wall growth continuously inculating the medium. Similar problems have been reported for Eubacterium limosum, an acetogen also isolated from ruminal contents, which produces copious amounts of extracellular polysaccharide (Loubiere et al., 1992). Nevertheless, the batch culture experiments provided sufficient information to obtain an estimate of the maximum growth rate of isolate H3HH. The highest growth rate (approximately 0.6 h-1) for isolate H3HH was obtained when this organism was grown on glucose (42 mM). The maximum growth rate observed for isolate A10 was approximately 0.5 h-1 in agreement with previous experiments.
was measured immediately after sampling for volatile fatty acids with an Ag combination electrode connected to a Fisher Accumet pH meter (Fisher Scientific, Pittsburg, PA). Glucose concentration in the culture medium was assayed with a glucose oxidase
The purpose of this series of experiments was to assess the minimum concentration of carbohydrate necessary for growth of isolates A10 and H3HH. The approach used in the present experiments was to determine the growth rate of isolates A10 and H3HH
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Minimum carbohydrate concentration necessary for growth Boccazzi and Patterson (2011) described the array of carbohydrates utilized by isolates A10 and H3HH.
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Figure 1. Continuous culture of isolate H3HH. A continuous culture system was established as previously described The system was inoculated (approx. 10 % v/v) with an overnight culture and allowed at least ten turnovers to reach steady state conditions. An aliquot of the outflow was collected to determine bacterial mass (●) and acetate content (□). After sampling the media flow rate was adjusted and the cultures were allowed to reach steady state conditions again.
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Figure 2. Influence of carbohydrate concentration on the growth rate of isolate H3HH in batch culture. Acetogen media (10 mL) containing various amounts of either glucose (♦), cellobiose (■) or maltose (●) was inoculated (0.1 mL) with an overnight culture of isolate H3HH. The OD (absorbance at 660 nm) was measured at appropriate time intervals. The growth curve of each culture was plotted and the highest growth rate at each carbohydrate concentration was determined. The growth rate not attributable to the carbohydrates added (i.e., growth rate of cultures without added carbohydrates) was subtracted from the other growth rates. The vertical lines drawn on the graph represent the range of glucose and cellobiose concentrations detected in ruminal fluid. Each point is the mean of at least two (but usually three) tubes.
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in acetogen medium containing concentrations of glucose, maltose and cellobiose that ranged from very low (10 μM, as found in the rumen) to very high (> 1 mM, as used in substrate utilization assays). The apparent maximum growth rates obtained at each carbohydrate concentration were plotted as a function of the carbohydrate concentration present at inoculation. The minimum carbohydrate concentration necessary for growth was estimated as the carbohydrate concentration below which the growth rate of the cultures was similar to that of cultures without added carbohydrate. Using this definition, H3HH had a lower minimum threshold for maltose (14 μM) and cellobiose (14 μM) than for glucose
Factors other than growth rate, minimum concentration of carbohydrates needed to start growth may also be involved in maintaining relatively low numbers of acetogens in rumen contents. Physical parameters (pH and VFA) have been implicated in inhibiting growth of a number of bacterial species, thus, the effects of pH and VFA concentration on the growth rates of isolates A10 and H3HH were determined. Because of the growth on the wall of the growth vessel, continuous culture data was not reliable, and only batch culture data is presented. Isolate A10 and isolate H3HH were grown in acetogen medium without ruminal fluid, but with sodium acetate (0, 50 or 100 mM) and initial pH adjusted to 6.6
(111 μM) (Figure 2). In contrast, isolate A10 had a lower minimum threshold for maltose (28 μM) than for glucose (56 μM) or cellobiose (139 μM) (Figure 3). Furthermore, the concentration of carbohydrates needed for maximal growth rate of isolate A10 was one magnitude less than the concentrations needed by isolate H3HH (1 versus 10 mM).
or 5.5. Ruminal fluid was omitted from the medium to reduce the basal acetate concentration from approximately 2 mM to less than 0.1 mM. Both isolate A10 and isolate H3HH were affected by the initial pH and acetate concentration of the medium (Table 1). At neutral pH (6.6), there was a small decrease in growth rate with increasing acetate concentrations. However, the growth rate of both organisms at pH 5.5 was approximately half of that obtained at pH 6.6 when no acetate was added to the medium. An
Effect of pH and acetate concentration on growth
Table 1. Growth rate of isolates A10 and H3HH as affected by pH and acetate concentrations1 H3HH
A10
Acetate
pH 6.6
pH 5.5
pH 6.6
pH 5.5
0 mM
0.309
0.119
0.279
0.126
50 mM
0.260
0.014
0.270
0.023
100 mM
0.278
0.006
0.259
0.005
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Figure 3. Influence of initial carbohydrate concentration on the growth rate of isolate A10 in batch culture. Please refer to the legend of Figure 2 for details of this experiment.
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initial acetate concentration of 50 mM was sufficient to inhibit the growth of both organisms when the pH of the media was 5.5, and no growth occurred at pH 5.5 when initial acetate concentration was 100 mM.
The rumen evolved to provide the appropriate conditions for microbial degradation of cellulose and other plant structural carbohydrates (Weimer et al., 2009). Accordingly, a number of anaerobic, carbohydrate-fermenting microbial species grow well and can be isolated in high numbers from ruminal
available for ruminant utilization. The present results show that maximum growth rate of isolate A10 or H3HH would not limit establishment of high numbers in ruminal contents. Although we were unable to obtain reliable data from continuous culture experiments, data obtained from batch culture experiments indicated that the maximum growth rate of these organisms was approximately 0.6 h-1 for isolate A10 and 0.5 h-1 for isolate H3HH. These generation times are greater than the liquid turnover rate of ruminal contents which ranges between 0.05 and 0.1 turnovers/h (Hungate, 1966; Weimer, 1992, Zinder, 1993). The concentration of soluble carbohydrates in
fluid (Hungate, 1966; Oshio et al., 1987; Ricke et al., 1996). However, the rumen is not a hospitable environment for all microbes. The anaerobiosis, liquid turnover rate, concentration of soluble carbohydrates and the chemical (i.e., pH and VFA concentration) characteristics of the rumen prevent the growth of a number of bacteria that otherwise should be able to grow on the substrates in rumen fluid. The rumen is a highly reduced (Eh = -150 to -350 mV) anaerobic environment (Clarke, 1977) and oxygen introduced into ruminal contents through feeding and drinking is rapidly depleted by facultatively anaerobic organisms (Ellis et al., 1989). Many bacterial species are unable to tolerate the reduced and anaerobic character of ruminal contents. This anaerobic ecosystem provides the host animal with VFA, which it can utilize as an energy source (Lin and Luchi, 1991; Lindley et al., 1990). Although the acetogens used in these experiments are somewhat tolerant to aerobic conditions (Boccazzi and Patterson, 2011), as a group, acetogens isolated from ruminal contents are obligate anaerobes. The turnover rate of rumen contents also can affect which organisms colonize and persist in the rumen. For example, fatty acid-degrading bacteria (e.g., Anerovibrio lipoliticus) have an ample supply
ruminal fluid may also limit growth of certain bacteria. As an example, Streptococcus bovis, although routinely isolated from ruminal contents, is not found in high numbers unless relatively high concentrations of soluble carbohydrates (i.e. maltose) are present (Russell and Baldwin, 1979). This situation exists due to the relatively low affinity (> 1 mM) of S. bovis for carbohydrates. In previous research (Pinder et al., 2012), we determined that the concentration of soluble carbohydrates in rumen fluid of cattle consuming typical production diets ranges between 8 and 17 μM for glucose, and between 110 and 175 μM for cellobiose. The present results are important because they indicate that the soluble carbohydrate concentration of ruminal fluid may be sufficient for growth by acetogens and other factors may limit growth. The low numbers of acetogens in ruminal contents cannot be explained by the anaerobic nature of the isolates, growth rate, or minimum carbohydrate concentration necessarry for growth. Thus, other factors, such as pH or VFA concentration may be involved in restricting colonization. The rumen is an environment with a relatively high concentration of volatile fatty acids and frequently acidic conditions (Hungate, 1966; Oshio et al., 1987; Rus-
of substrates in ruminal contents. However, these organisms are unable to propagate in high numbers in ruminal contents because their maximum growth rate is less than the liquid turnover rate of ruminal contents (Hungate, 1966). The result is that VFA are
sell, 1992). The concentration of total fatty acids in ruminal fluid fluctuates considerably but is usually within the range of 60-140 mM (Owens and Goetsch, 1988). Many factors (e.g., diet of the animal and time in relation to feeding) affect ruminal fluid pH and
DISCUSSION
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VFA concentrations. For example, high concentrate diets cause an increase in the total concentration of VFA in ruminal fluid as compared to diets high in forages (Oshio et al., 1987). Moreover, ruminal VFA concentrations fluctuate diurnally as a result of feed ingestion by the animal. Total VFA concentrations typically start to rise soon after feeding and peak approximately 6 h post-feeding. Similar to VFA concentrations, ruminal fluid pH fluctuates (generally between pH 5 and 7) in response to diet and time post-feeding. The pH of ruminal fluid usually is lower in animals consuming high-concentrate diets compared to animals consuming high forage diets. Ruminal fluid typically decreases soon after feeding
to pH’s ranging from 7 to 5, the internal pH of the cells is maintained above 7 until the extracellular pH dropped below 5.5 (Baronofsky et al., 1984). These characteristics identify C. thermoaceticum as a neutrophile (pH optimum for growth is near 7). One explanation of the sensitivity of neutrophiles to low pH conditions, based on the chemiosmotic theory, proposes that volatile fatty acids act as uncouplers destroying the H+ gradient across the cell membrane and are thus capable of dissipating the proton motive force of cell membranes (Freese et al., 1973; Sheu et al., 1975; Baronofsky et al., 1984; Salmond et al., 1984; Wallace et al., 1989). The ability of acidophiles and acid-tolerant species to resist low pH and
and remains low for several h after feeding before returning to near neutral levels. Many bacteria (e.g., S. bovis) are capable of tolerating the low pH and high VFA concentrations similar to those often detected in ruminal contents after feeding (Russell, 1991). However, many other bacterial species (e.g., Fibrobacter succinogenes, R. flavefaciens, Salmonella, Veillonella, and Escherichia coli) are sensitive to decreasing pH, especially as pH is decreased to values of pH 6.0 and less (Dunkley et al., 2008; Hollowell and Wolin, 1965; Kwon et al., 1997; Ricke, 2003; Russell and Dombrowsli, 1980). Wolin (1969) determined that the growth of E. coli is markedly inhibited by the high volatile fatty acid concentrations of ruminal fluid and that this inhibition is pH dependent. Although disagreement exists as to the mechanism by which the presence of volatile fatty acids in acidic conditions inhibits the growth of certain bacteria (Russell, 1992), the fact remains that these low pH and high VFA sensitive bacteria are unable to grow in ruminal contents even when specific enrichment is attempted (Wallace et al., 1989). The non-ruminal Clostridium thermoaceticum, is the only acetogen that has been examined regarding its ability to tolerate acidic conditions and/ or high VFA concentrations. Clostridium thermo-
high VFA concentrations is conveyed by the ability to lower intracellular pH in the face of declining extracellular pH thus maintaining a constant proton motive force across the cell membrane (Russell, 1992). The data presented herein show that isolate A10 and isolate H3HH, are sensitive to the pH of the culture medium as well as the presence of volatile fatty acids in an acidic medium. These results suggest that acetogens isolated from ruminal contents are similar to C. thermoaceticum, E. coli and other neutrophilic bacteria which are sensitive to low pH and high VFA concentrations. It remains to be determined exactly how these environmental factors (i.e., low pH and high VFA concentrations) prevent the growth of acetogens isolated from ruminal contents. Although the carbohydrate concentrations in ruminal fluid may be marginally sufficient for growth of isolates A10 and H3HH, the inability to grow under low pH and high VFA conditions would suggest that establishment of a high number of these isolates in ruminal contents (especially with a single inoculation) would be unlikely. However, we did not determine if either isolate remains metabolically active under low pH and high VFA conditions. If acetogens are metabolically active under low pH and high VFA conditions, then the possibility of using repeat-
aceticum grows faster and produces more acetic acid when grown at neutral pH than when grown in acid medium (Schwartz and Keller, 1982; Wang and Wang, 1984). Furthermore ,when exponentiallygrowing cells of C. thermoaceticum were exposed
ed (monthly, weekly or daily) inoculations to maintain high numbers of acetogens in ruminal contents may be possible. Another, though more difficult, possibility is to isolate or generate mutants capable of tolerating low pH and high VFA conditions.
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Russell, J.B. and R. L. Baldwin. 1979. Substrate preferences in rumen bacteria: Evidence of catabolite regulatory mechanisms. App. Environ. Microbiol. 36:319-329. Russell, J.B. and D. B. Dombrowski. 1980. Effect of pH on the efficiency of growth by pure cultures of rumen bacteria in continuous culture. Appl. Environ. Microbiol. 39:604-610. Russell, J.B. 1992. Another explanation for the toxicity of fermentation acids at low pH: anion accumulation versus uncoupling. J. Appl. Microbiol. 73:363-370. Salmond, C.V., R. H. Kroll, and I. R. Booth. 1984. The effect of food preservatives on pH. J. Gen. Microbiol. 130:2845-2850. Schwartz, R.D. and F.A. Keller, Jr. 1982. Isolation of a strain of Clostridium thermoaceticum capable of growth and acetic acid production at pH 4.5. Appl. Environ. Microbiol. 43:117-123. Sheu, C.W., D. Salomon, T. Sheelvalsan, and E. Freese. 1975. Inhibitory effects of lipophilic acids and related compounds on bacteria and mammalian cells. Antimicrob. Agents and Chemother. 7:349-363. Wallace, R.J., M.L. Falconer, and P. K. Bhargava. 1989. Toxicity of volatile fatty acids at rumen pH prevents enrichment of Escherichia coli by sorbitol in rumen contents. Curr. Microbiol. 19:277-281. Wang, G. and D.I.C. Wang. 1984. Elucidation of growth inhibition and acetic acid production by Clostridium thermoaceticum. Appl. Environ. Microbiol. 47:294-298. Weimer, P. J. 1992. Cellulose degradation by ruminal microorganisms. Crit. Rev. Biotech. 12:189-223. Weimer, P.J. J.B. Russell, and R.E. Muck. 2009. Lessons from the cow: What the ruminant animal can teach us about consolidated bioprocessing of cellulosic biomass. Bioresour. Technol. 100:5323-5331. Wolin, M. J. 1969. Volatile fatty acids and the inhibition of Escherichia coli growth by ruminal fluid. Appl. Microbiol. 17:83-87. Zinder, S. H. 1993. Physiological ecology of methanogens. In: Methanogenesis. Ecology, physiology, biochemistry and genetics. J. G. Ferry (ed.), Chapman and Hall. New York, NY. 16
Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 3, Issue 1 - 2013
www.afabjournal.com Copyright © 2013 Agriculture, Food and Analytical Bacteriology
REVIEW Greenhouse Gas Emissions from Livestock and Poultry C. S. Dunkley1 and K. D. Dunkley2 Department of Poultry Science, University of Georgia, Tifton, GA School of Science and Math, Abraham Baldwin Agricultural College, Tifton, GA 1
2
ABSTRACT In 2008 the Environmental Protection Agency (EPA) estimated that only 6.4% of U.S. greenhouse gas (GHG) emissions originated from agriculture. Of this amount, 53.5% comes from animal agriculture. Agricultural activities are the largest source of N2O emissions in the U.S. accounting for 69% of the total N2O emissions for 2009. In animal agriculture, the greatest contributor to methane emissions is enteric fermentation and manure management. Enteric fermentation is the most important source of methane in beef and dairy production, while most of the methane from poultry and swine production originates from manure. The main cause of agricultural nitrous oxide emissions is from the application of nitrogen fertilizers and animal manures. Application of nitrogenous fertilizers and cropping practices are estimated to cause 78% of total nitrous oxide emissions. Based on the life cycle assessment of beef cattle, 86.15% of the GHGs are emitted during the production stage, while 68.51% of emissions take place during the production of pork and 47.82% of GHG emissions occur during the production stage of broiler chickens. The majority of the emissions from the beef cattle production comes from enteric fermentation while manure management is the major source during swine production and propane use during broiler poultry production. Keywords: greenhouse gas, LCA, poultry emissions, beef emissions
Agric. Food Anal. Bacteriol. 3: 17-29, 2013
Correspondence: C. S. Dunkley, cdunkley@uga.edu Tel: +1 -229-386-3363 Fax: +1-229-386-3239
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INTRODUCTION The primary greenhouse gases emitted by agricultural activities are carbon dioxide (CO2) methane (CH4) and nitrous oxide (N2O) (Johnson et al., 2007). Livestock production contributes GHGs to the atmosphere both directly and indirectly (IPCC, 2006). The emissions can be classified based on the source of the emission; 1) Mechanical, and 2) Non-mechanical. The majority of direct CO2 emissions from animal agriculture are usually from fossil use, for example; the use of propane or natural gas in furnaces or incinerators and the use of diesel gas to operate farm equipment and generators results mostly in CO2
son et al., (2007). In 2011 the US Environmental Protection Agency (EPA) reported that the Agricultural Sector was responsible for a total of 410.6 Tera gram CO2 equivalents (Tg CO2e in 2005). Enteric fermentation and manure management contributed a total of 200.4 Tg CO2e which represented about 48% of the total emissions from the agricultural Sector. During this period (Figure 1.) enteric fermentation was responsible for 136.5 Tg CO2e and managed manure was responsible for 63.9 Tg CO2e. In 2007, the emissions from the Agricultural sector were 425.8 Tg CO2e a 3.7% increase. The emissions from enteric fermentation during this period were 141 Tg CO2e a 3.3% increase
emissions (Dunkley unpublished data), this type of emission can be described as “mechanical emissions.” The use of electricity on animal production farms results in indirect emissions since the emissions do not occur on site. For non-mechanical emissions, direct emissions can be a by-product of digestion through enteric fermentation (CH4 emissions). Direct emissions also occur from the decomposition and nitrification/denitrification of livestock waste (manure and urine) where CH4 and N2O are emitted. Managed waste that is collected and stored also emits CH4 and N2O. Indirect emission of N2O occurs when nitrogen is lost from the system through volatization as NH3 and Nx. Also, indirect emissions can result from nitrogen that is runoff or leached from manure management systems in a form other than N2O and is later converted to N2O offsite (IPCC, 2006). Methane from enteric fermentation and manure management are the main sources of CH4 emissions from agricultural activities and of all domestic livestock, dairy and beef cattle are the largest emitters of CH4. Agricultural activities are the largest source of N2O emissions in the US accounting for 69% of the total N2O emissions for 2009 (EPA, 2011). The majority of the N2O emission from animal agriculture is from manure management
over the 2005 period, while manure management emissions increased to 68.8 Tg CO2e a 7.7% increase. The GHG emissions from agriculture showed a 1.5% reduction to 419.3 Tg CO2e in 2009 when compared to 2007. This reduction was reflected slightly in enteric fermentation which was down by 0.8% to 139.8 Tg CO2e and a 2% reduction in manure management emission to 49.5 Tg CO2e (IPCC, 2010).
which is the second largest (a far second to cropping practices) N2O emitter in the agricultural sector (IPCC, 2010). Application of nitrogenous fertilizers and cropping practices are estimated to cause 78% of total nitrous oxide emissions according to John-
emissions are dependent on how the manure is managed. Beef cattle raised on pasture/range exhibit relatively high N2O emissions. In this system the manure and urine from the cattle are deposited directly on the soil reducing the likelihood of much methane
18
EMISSIONS BASED OF MANURE MANAGEMENT SYSTEMS The type of manure management system that is used in livestock production can affect the amount of emissions and the type of gases that are emitted. A variety of livestock production systems operates in the U.S. and different manure management systems are utilized depending on the type of livestock or poultry produced (Del Grosso et al., 2008). Among the manure management systems practiced in the US are; pit storage, poultry with/without litter (that is, poultry raised on a bedding material or poultry raised in cages), dry-lot, anaerobic lagoon, pasture, etc. (Table 1). Beef cattle can be raised using different manure management systems and the amount of
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Figure 1. The distribution of livestock GHG emissions by source in 2005, 2007 and 2009
160 141
136.5
140
139.8
Tg CO2 equivalent
120 100 80
68.8
63.9
67.4
60 40 20 0 2005
2007
Manure Management
emission. When cattle are raised under conditions where the manure is collected and spread daily and there is no storage before it is spread onto the soil there is low CH4 emissions and no N2O emissions. Dairy cattle and swine reared in liquid/slurry manure management systems have moderate to high CH4 emissions, while emissions from swine and dairy cattle reared in anaerobic lagoon management systems have variable CH4 emissions as it is mostly dependent on the duration of time the manure and slurry are stored in the lagoons. In this system, the waste can be stored between 30 to 200 days; the longer the storage time, the more likely the CH4 emissions will be high. Both the liquid/slurry and anaerobic lagoon manure systems have low N2O emissions. Poultry reared in management systems with litter and using solid storage have relatively high N2O emissions but low CH4 emissions. This is because the manure is stock piled under aerobic conditions which limits the production of CH4 (USAFGGI, 2008). Broiler, pullets,
2009
Enteric Fermentation
and to an extent breeders, are reared using these manure management systems. Commercial layers are typically reared in high-rise cages or scrape-out/ belt systems. Here the manure is excreted onto the floor below with no bedding to absorb moisture. The ventilation system dries the manure as it is stored. In some broiler breeder houses a part of the manure is collected under the slats in the houses making it similar to the commercial layers. In this type of manure management system both CH4 and N2O emissions are relatively low (IPCC, 2000). The amount of CH4 or N2O that is emitted from livestock also depends on environmental conditions (Del Grosso et al., 2008). Methane is emitted under anaerobic conditions where oxygen is not available (Palmer and Reeve, 1993). Storage in tanks, ponds or pits, such as those used with liquid/slurry flushing systems encourages anaerobic conditions, therefore more CH4 is produced (USAF 2008). Conversely, solid waste storage in stacks or shallow pits promotes
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Table 1. Description of livestock waste deposition and storage pathways Relative Emissions Manure Management System
Description
CH4
N2O
Pasture/range/paddock
Manure and urine from pasture and grazing animals is deposited directly onto soil.
Low
High
Daily Spread
Manure and urine are collected and spread on fields (little or no storage prior to application).
Low
Minimal
Solid storage
Manure and urine with or without litter are collected and stored long term in bulk.
Low
High
Manure and urine are deposited directly on unpaved feedlots where it is allowed to dry. It is periodically removed.
Low
High
Manure and urine are collected and transported in liquid form to tanks for storage. The liquid/ slurry may be stored for long periods.
Moderate to high
Low
Ex. beef cattle
Ex. poultry Dry lot Ex. Beef cattle Liquid/slurry Ex. Swine/dairy cattle Anaerobic Lagoon Ex. Swine/dairy cattle Pit Storage Ex. Swine/poultry layers Poultry with litter Ex. Broiler/pullet/breeders Poultry without litter Ex. Poultry layers/broiler breeders
Manure and slurry are collected using a flush Variable system and transported to lagoons for storage. It remains in lagoons for 30-200 days.
Low
Combined storage of manure and urine in pits below livestock confinements.
Low
Moderate to high
Enclosed poultry houses utilize bedding material Low (ex. Wood shavings, peanut hull, rice hulls etc.). The bedding absorbs moisture and dilutes manure. Litter is cleaned out typically once per year.
High
In high-rise cages or scrape-out/belt systems, manure is excreted onto the floor below with no bedding to absorb moisture. The ventilation system dries the manure as it is stored.
Low
Low
Adapted from IPCC (2000) Chapter 4.
aerobic conditions which are more favorable for N2O emissions. High temperatures and increased storage time can also increase CH4 emissions (Del Grosso et al., 2008). Feed characteristics also play a role in CH4 emissions. Feed, diet, and growth rate have an
manure is composted, which will increase aeration limiting anaerobic production of CH4. Higher energy feeds result in manure with more volatile solids, which increases the substrates from which CH4 is produced (Del Grosso et al., 2008). Depending on
effect on the amount and quality of manure an animal produces (Monteny, 2006). Harper (2000) stated that there was a large effect on CH4 emissions that is contingent on the production and use of farmyard manure. Typically, in an organic system, stock piled
the species, this impact is somewhat offset because some higher energy feeds such as that fed to poultry are more digestible than lower quality forages fed to ruminant animals and therefore less waste is excreted. The energy content and quality of feed affects
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the amount of methane produced in enteric fermentation where lower quality feed and higher quantities of feed causes greater emissions (USAFGG, 2008). It was reported by the EPA (2011) that an animals feed quality and feed intake affects emission rates. In general, lower feed quality and / or higher feed intakes lead to higher emissions.The composition of the waste, the type of bacteria involved, and the conditions following excretion, all have an effect on the production of N2O from waste management systems (EPA, 2010). In order for N2O to be emitted, the waste must be handled aerobically where NH3 and organic nitrogen is converted to nitrates and nitrites (Del Grosso et al., 2008).
stock waste had increased by 4.69% from emission levels in 2005 to 209.8 Tg CO2e. This was as a result of increases in enteric fermentation from dairy cattle (101.6 Tg CO2e), beef cattle (32.4 Tg CO2e) swine (2.1 Tg CO2e) and horses. There were also increases in emissions from managed livestock waste in all the major livestock categories. Overall, during the two year period from 2005 to 2007, dairy cattle had a 2.5% increase in emissions (112.4 Tg CO2e), beef cattle had the highest percentage increase of 8.7% up to 62.4 Tg CO2e. Swine had an increase of 6.6% up to 24.3 Tg CO2e, while poultry had a 4.5% increase (4.6 Tg CO2e) during the 2005 to 2007 period (EPA, 2011). In 2009 (Table 4), a reduction in emissions of 1.28%
Ninety-one % of emissions from enteric fermentation and managed livestock manure are in the form of CH4 (EPA, 2011). When Monteny et al. (2001) compared the distribution of methane emissions from enteric fermentation among animal types; poultry had the lowest amount with 0.57 lbs methane/ animal/ year when compared to dairy cattle with 185 to 271 lbs methane/ animal/ year and swine with 10.5lbs methane/ animal/ year. In 2005, livestock emissions from enteric fermentation and manure management were 200.4 Tg CO2e (Table 2). Of this total, dairy cattle and beef cattle contributed 99.3 and 30.4 Tg CO2e respectively from enteric fermentation. Swine contributed 1.9 Tg CO2e from enteric fermentation while poultry contributed no emissions from enteric fermentation. For this same period, dairy cattle were responsible for 109.6 Tg CO2e from enteric fermentation and managed livestock waste combined; beef cattle contributed 57.4 Tg CO2e, swine contributed 22.7 Tg CO2e, while poultry contributed 4.4 Tg
from the 2007 levels was observed even though these emissions were not as low as in 2005. The emissions from enteric fermentation from the major livestock categories showed a reduction in enteric fermentation from dairy cattle (99.6 Tg CO2e), while beef cattle showed an increase (33.2 Tg CO2e). Enteric fermentation emissions from swine remained the same as in 2007. For the major livestock categories overall reductions in emissions from enteric fermentation and managed livestock waste combined were observed in all with the exception of beef cattle. Dairy cattle had a 2% reduction down to 110.1 Tg CO2e from the 2007 levels of 112.4 Tg CO2e. Beef cattle had a 1.7% increase up to 63.5 Tg CO2e, swine had a reduction of 4.9% (23.1 Tg CO2e) while poultry had a 6.5% reduction in the emissions from 2007. Of all the major livestock categories (dairy, beef cattle, swine and poultry) only poultry had an overall reduction (2.2%) in emissions from 2005 to 2009 (EPA, 2011). The emission estimates reported here were adapted from the EPA’s 2011 report. Several modifications to the estimates relative to the previous estimates had an effect on the emission estimates. The modifications included; the average weight assumed for mature dairy cows from 1550 pounds used in previous inventories to 1500 pounds. There were also
CO2e. The remaining emissions (5.66 Tg CO2e) were from other livestock animals which were not reared in large amounts. By 2007 (Table 3), the total amount of GHG emissions from enteric fermentation and managed live-
slight modifications from the 2008 numbers in the populations of calves, beef replacement and feedlot cattle. Swine populations were also modified so that the categories “<60 pounds” and “60- 119pounds” changed to “<50 pounds” and “50-119 pounds”.
EMISSIONS FROM ENTERIC FERMENTATION AND MANAGED MANURE FROM 2005 TO 2009
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Table 2. Greenhouse gas emissions by livestock category and source in 2005 Enteric Fermentation CH4 Animal Type
Managed Livestock Waste CH4
N2O
Total
Tg CO2 equivalent
Dairy Cattle
99.3
2.8
7.5
109.6
Beef Cattle
30.4
21.4
5.6
57.4
Swine
1.9
19.0
1.8
22.7
Horses
3.5
0.06
0.3
3.86
Poultry
0.00
2.7
1.7
4.4
Sheep
1.0
0.1
0.4
1.5
Goats
0.30
0.00
0.0
0.3
Total
136.5
46.6
17.3
200.4
Table 3. Greenhouse gas emissions by livestock category and source in 2007
Enteric Fermentation CH4 Animal Type
Managed Livestock Waste CH4
N2O
Total
Tg CO2 equivalent
Dairy Cattle
101.6
2.9
7.9
112.4
Beef Cattle
32.4
24.2
5.8
62.4
Swine
2.1
20.3
1.9
24.3
Horses
3.6
0.6
0.6
4.8
Poultry
0.00
2.8
1.8
4.6
Sheep
1.0
0.1
0.1
1.2
Goats
0.3
0.0
0.0
0.3
Total
141.0
50.7
18.1
209.8
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Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 3, Issue 1 - 2013
Table 4. Greenhouse gas emissions by livestock category and source in 2009 Enteric Fermentation CH4
Managed Livestock Waste CH4
Animal Type
N2O
Total
Tg CO2 equivalent
Dairy Cattle
99.6
2.7
7.8
110.1
Beef Cattle
33.2
24.5
5.8
63.5
Swine
2.1
19.0
2.0
23.1
Horses
3.6
0.5
0.3
4.4
Poultry
0.0
2.7
1.6
4.3
Sheep
1.0
0.1
0.3
1.4
Goats
0.3
0.0
0.0
0.3
Total
139.8
49.5
17.9
207.2
These changes attributed to an average reduction in emissions from dairy cattle of 11.5 Gg or 0.8% per year and beef cattle emissions decreased an average of 0.13 Gg or less that 0.01% per year over the entire time series relative to the previous inventory (EPA, 2011). Of course, in order to discuss emissions from enteric fermentation one must consider the size (weight) of the livestock and the number of each type of livestock grown each year. Larger animals will produce more methane than smaller animals and the amount of methane emitted is increased with increasing number of animals grown (Del Grosso et al., 2011). The type of digestive system will also determine the amount of methane produced. Cattle are ruminant animals with a four compartment stomach. Their digestive tract is designed for microbial fermentation of fibrous, high cellulose materials. One of the by-products of microbial fermentation is meth-
amount of CH4 that is emitted, poorer quality highfiber diets will likely result in greater CH4 emissions than higher quality diets that contains more protein (Del Grosso et al., 2011). Typically, CH4 is usually produced following the degradation of carbon components during digestion of feed and manure (Monteny et al., 2006). Husted (1994) stated that the rumen was the most important site of methane production in ruminants (breath), while in monogastric animals such as swine and poultry, methane is usually produced in the large intestines. The manner in which animal manure are stored whether indoors in sub-floor pits or outdoors are also relevant sources of CH4 production (Husted, 1994). Enteric fermentation is the most important source of methane in the dairy industry, while, the majority of CH4 emissions from the pig and poultry industries originates from manures (Monteny et al., 2006). There is also a range in the total emissions in dairy cows that is
ane (Stevens and Hume, 1998). Poultry and swine are mono-gastric animals with a simple stomach and little microbial fermentation taking place; therefore they have less enteric methane production (Frédéric et al., 2007). The feed quality also plays a role in the
caused by differences in diet and housing systems. For example; there are lower emission rates for tying stalls and higher rates for cubicle houses (Groot Koerkamp and Uenk, 1997).
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FARM-GATE AND LIFE CYCLE ASSESSMENT EMISSIONS Greenhouse gas emission from the different livestock categories can also be evaluated based on “Life Cycle Assessment” (LCA). This involves not only the farm-gate emissions but also an inventory of the material and energy inputs and the emissions associated with each stage of production. The LCA looks at the “cradle to grave” energy use (Guinee et al., 2001). This assessment could include; fertilizer production and transportation, crop production and transportation, feed additive manufacturing and transportation, animal production facilities, trans-
majority of the emissions from the production of edible eggs occurs at the farmgate (Figure 3) and as with chicken meat production, these emissions came from feed production, on-farm energy use, N2O from poultry litter and fuel combustion (EWG, 2011). The Environmental Working Group (2011) also reported LCAs from dairy production, reporting yogurt, cheese and 2% milk LCAs. The production of whole milk at the farm-gate resulted in 1.02 CO2e per Kg of edible whole milk, while only 0.67 kg CO2e was emitted per kg of edible 2% milk. Domestic cheese production at the farm-gate resulted in the emission of 9.09 kg per kg of edible cheese (Figure 3). For yogurt production, the majority of emissions occurred
portation to processing plants, processing, distribution to retail markets, consumer use of the product and disposal of packaging (Guinee et al., 2001). This can be a very complex process and researchers have used different boundaries when approaching the LCA for different livestock. The Environmental Working Group (2011) examined GHG emissions from beef cattle and poultry, based on “farm-gate” emissions and showed that each of the livestock category assessed displayed differences in various areas of production (Figure 2). Farm-gate emissions here are based on the emissions that occur within the bounds of the farm plus the feed production and did not include processing of the meat. The EWG reported that the majority (7.51 kg CO2e) of GHGs was emitted to produce 1 kg beef at the farm-gate was as a result of enteric fermentation. In poultry production the majority (1.26 kg CO2e) of GHGs emissions was from feed production and no GHGs emissions from enteric fermentation. To produce 1 kg of edible beef at the farm-gate resulted in the emissions of 1.75 kg CO2e of N2O from manure, while 0.28 kg CO2e N2O was emitted from manure to produce 1 kg edible chicken meat. Emissions of GHGs from energy use at the farm-gate can also be compared for different livestock categories. On-farm energy use to produce
post-farm gate (1.03 kg CO2e per kg yogurt). Methane emissions from enteric fermentation were the primary source of pre-farm-gate GHGs for cheese, milk and yogurt production (EWG, 2011). A number of different GHG emission values from LCA have been published for different livestock categories (Table 4). Based on these publications the emissions from beef production at the farm-gate ranged from 14.8 to 20 kg CO2e/kg of product at the farm-gate with an average of 16.25 kg CO2e/kg of product at the farm-gate. The figures for swine ranged from 3.4 to 6.4 kg CO2e/kg of product at the farm-gate with an average of 4.82 kg CO2e/kg of product at the farm-gate, while the emissions for poultry ranged from 2.33 to 4.6 kg CO2e/kg of product at the farm-gate with an average of 3.09 kg CO2e/kg of product at the farm-gate. According to reports by EWG (2011), beef cattle LCA emissions in kg CO2e/kg of consumed food was 27 kg. They also reported that the LCA for pork was 12.1 kg CO2e/kg of consumed food, while chicken had an LCA of 6.9 kg CO2e/kg of consumed food (Figure 3). The LCA emissions that were calculated by the EWG included the production emissions. This included the emissions before the product left the farm plus all avoidable and unavoidable waste. Cal-
1 kg of beef at the farm-gate resulted in the emission of 0.23 kg CO2e, while to produce 1 kg chicken meat at the farm-gate resulted in the emission of 0.26 kg CO2e GHGs. It was also reported that 4.8 kg CO2e was generated to produce 1 kg of edible eggs. The
culations were also done to include post-production emissions which included processing, transport, retail, cooking and waste disposal (EWG, 2011). Of the 27 kg CO2e emitted to produce 1 kg of beef (consumed) only 3.73 kg CO2e was post farm-gate emis-
24
Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 3, Issue 1 - 2013
Figure 2. Sources of beef and poultry production emissions
Farmgate Emissions per kg consumed meat 8 7 7.51
6 5 kg CO2e/kg 4 edible meat at farm-gate 3
4.67
2 1.26 1.75
1
0.28
0
0
0.59
Enteric Fermentation CH4
Feed Production
Manure N20
Beef Cattle
0
Manure CH4
0.23 0.26 On-farm Energy Use
Poultry
Figure 3. LCA production and post-production emissions of beef and dairy cattle, swine and poultry
LCA Emissions for Beef Cattle, Dairy, Swine and Poultry 30
kg CO2e/kg consumed meat
3.73
25 20 15 10
1.7
3.81
23.27 3.6 11.77
5
1.17 8.29
0 Beef Cattle
Cheese
Swine
Production Emissions
1.03 3.3
3.63
Poultry
Eggs
0.98
1.17
0.92
Yogurt
Milk (2%)
Post-production Emissions
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25
Figure 4. Percent production and post-production emissions for beef and dairy cattle, swine and poultry LCA
100% 90% 80% 70% 60% 50% 40% 30% 20% 10% 0% Beef Cattle
Cheese
Swine
Production Emissions
Poultry
Eggs
Yogart
Milk (2%)
Post-production Emissions
sions (Figure 3). A total of 3.81 kg CO2e was emitted post farm-gate to produce 1 kg of consumed pork, while 3.3 kg CO2e from the total 6.9 kg emitted to produce 1 kg chicken (consumed) was post farmgate. From the LCA emissions it is clear that the majority (86%) of the emissions from beef cattle production occur during the production stage while only 14% of the LCA emissions occur post-production (Figure 4). This is similar to swine where the majority (69%) of emissions was also observed during the production stage. A different scenario was observed
Of the major livestock animals reared, emissions from poultry production systems generate the lowest levels of emissions to produce one kg CO2e/kg meat at farm-gate while dairy cattle produce the lowest levels of emissions to produce one kg CO2e/ kg product at farm-gate. Dairy cattle emit the highest levels of GHG per animal followed by beef cattle and swine. The majority of the emissions from beef production come from enteric fermentation and feed production with the cow to calf and the steer calf stages generating more than 65% of the total GHG emissions from this livestock category. In all the
for the poultry LCA where 48% of the emissions were observed during the production stage.
stages of beef production, high levels of CH4 from enteric fermentation are generated. For dairy cattle, the majority of emissions are from enteric fermentation, similar to beef cattle production. Methane emission from manure storage and feed production
CONCLUSIONS 26
Agric. Food Anal. Bacteriol. â&#x20AC;˘ AFABjournal.com â&#x20AC;˘ Vol. 3, Issue 1 - 2013
Table 5. Greenhouse gas emissions by livestock category and source in 2007
Livestock Category
Average GHG emissions kg CO2e/kg Product at farm-gate from all references
GHG Emissions
Peer reviewed, independent,
kg CO2e/kg of
Government Sources
product at farm-gate 15.9 20
Beef
Swine
Poultry
16.25
4.82
3.09
DERFA, 2008 Phetteplace et al. 2001(US)
14.8
Pelletier et al., 2010
15.32
Subak, 1999
15.23
EWG, 2011
6.4
DERFA, 2008
3.4-4.2
Pelletier, 2010
5.5
Wiltshire, 2006
4.62
EWG, 2011
4.6
DERFA, 2008
2.36
Pelletier, 2008
3.1
Wiltshire, 2006
2.33
EWG, 2011
in dairy cattle production also contributes to high levels of GHGs. Swine production emits GHGs primarily from manure management and fuel combustion. Only a small amount of CH4 is emitted during digestion when compared to ruminants. At least one third of GHG emitted from swine production is from post farm-gate activities. The largest contributor to GHG emissions from poultry production is feed production. The highest emissions from poultry on-
quarter of the total emissions.
farm activities are from fuel combustion from energy use and manure management. In broiler production post farm-gate emission makes up more than half of all the emission, while post farm-gate emissions from egg farm operations accounts for less than one
Del Grosso, S.J., J. Duffield, M.D. Eve, L. Heath, S. Ogle, J. Smith, and T. Wirth. 2011 U.S. Agriculture and Forestry Greenhouse Gas Inventory: 19902008. Climate Change Program Office, Office of the Chief Economist, U.S. Department of Agricul-
REFERENCES ADAS. 2007. The Environmental Impact of Livestock Production. Report for Defra FFG. www.archive. derfa.gov.uk/foodanimal/document/envimpactslivestock.pdf
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ture. Technical Bulletin No. 1930. 159 pp. June, 2011. Del Grosso, S.J., Ogle, S., Wirth, J., Skiles, S. 2008. U.S. Agriculture and Forestry Greenhouse Gas Inventory: 1990-2005. United States Department of Agriculture Technical Bulletin 1921. Environmental Working Group. 2011. Hamerschlag, K. and K. Venkat. Meat eaters guide: Methodology. Frédéric, P., Stéphane, G., Stéphane, P. L., Robert, D. v. B., Sylvain, P., & Jean-Yves, D. 2007. Evaluation of Greenhouse Gas Emissions from Five Swine Production Systems Based on Life Cycle Assessment. Paper presented at the 2007 ASAE Annual Meeting. Retrieved from http://asae.frymulti.com/
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abstract.asp?adid=23065&t=5 Groot Koerkamp, P.W.G, and G.H. Uenk. 1997. Climatic conditions and aerial pollutants in and emissions from commercial production systems in the Netherlands. In: Voermans, J.A.M., Monteny, G.J. (Eds.) Proceedings of the International Symposium on Ammonia and Odor control from Animal Production Facilities. Research Station for Pig Husbandry (PV), Rosmalen, 139-144. Guinee, J., M. Gorree, R. Heijungs, G. Huppes, R. Kleijn, A. de Koning, L. van Oers, A. Weneger, S. Suh, H. Udo de Haes, H. de Bruin, R. Duin, and M. Huijbregts. 2001. Life Cycle Assessment: An operational guide to the ISO Standards Part 2. Ministry of Housing, Spatial Planning and Environment, The Hague, Netherlands. <http://www.leidenuniv. nl/cml/ssp/projects/lca2/part1.pdf>(accessed 01/2006). Harper, L.A., R.R. Sharpe, and T.B. Parkin. 2000. Gaseous nitrogen emissions from anaerobic swine lagoons: ammonia, nitrous oxide, and dinitrogen gas. J. Environ.Qual. 29:1356-1365. Husted, S. 1994. Seasonal variation in methane emission from stirred slurry and solid manures. J. Environ. Qual. 23:585-592. Intergovernmental Panel on Climate Change (IPCC).
Monteny, G.J., and G.W. Erisman. 1998. Ammonia emissions from dairy cow buildings: A review of measurement techniques, influencing factors and possibilities for reduction. Neth. J. Agric. Sci. 46:225-247. Netherlands. <http://www.leidenuniv. nl/cml/ssp/projects/lca2/part1.pdf> Palmer, J.R. and J. N. Reeve. 1993. In: Genetics and Molecular Biology of Anaerobic Bacteria” Ed. Sebald, M., Springer-Verlag, New York, Berlin, Heidelberg, London, Paris, Tokyo, Hong Kong, Barcelona, Budapest. p 13-35. Pelletier, N. 2008. Environmental performance in the US broiler poultry sector: Life cycle energy use and greenhouse gas, ozone depleting, acidifying and eutrophying emissions. Agri. Systems 98:67-73. Phetteplace, H., D. Johnson, and A. Seidl. 2001. Greenhouse gas emissions from simulated beef and dairy livestock systems in the United States. Nutrient Cycling in Agroecosystems. 60:99-102. Reicosky, D.C., J.L. Hatfield, and R.L. Sass. 2000. Agricultural contribution togreenhouse gas emissions. In: Reddy, R., Hodges, H. (Eds.), Climate Change and Global Crop Productivity. CABI Publishing, Wallingford, Oxon, UK, pp. 37-55. Stevens, C.E., and J. D. Hume. 1998. Contribution of microbes in vetebrate gastrointestinal tract to
2000. Penman, J., D. Kruger, I. Galbally, T. Hiraishi, B. Nyenzi, S. Emmanuel, L. Buendia, R. Hoppaus, T. Martinsen, J. Meijer, K. Miwa, and K. Tanabe. (Eds.). Good practice guidance and uncertainty management in national greenhouse gas invento-
production and conservation of nutrients. Physiol. Rev. 78:393-427. Subak, S. 1999. Global environmental costs of beef production. Ecol.Econ. 30:79-91. U.S. Agriculture and Forestry Greenhouse Gas In-
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ventory: 1990-2005. 2008. Global Change Program Office, Office of the Chief Economist, U.S. Department of Agriculture. Technical Bulletin No. 1921. 161 pp. August, 2008. http://www.usda.gov/ oce/global_change/AFGGInventory1990_2005. htm.U.S. Environment Protection Agency (EPA). 2010. Chapter 6: Agriculture. Inventory of U.S. greenhouse gas emissions and sinks. 1998-2008. http://www.epa.gov/climatechange/emissions/ downloads10/US-GHG-Inventory-2010_Chapter6Agriculture.pdf U.S. Agriculture and Forestry Greenhouse Gas Inventory: 1990-2005. Global Change Program Office, Office of the Chief Economist, U.S. Department of Agriculture. Technical Bulletin No. 1921. 161 pp. August, 2008. http://www.usda.gov/oce/ global_change/AFGGInventory1990_2005.htm.
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www.afabjournal.com Copyright © 2013 Agriculture, Food and Analytical Bacteriology
REVIEW Can Salmonella Reside in the Human Oral Cavity? Sujata A. Sirsat1 1
Conrad N. Hilton College of Hotel and Restaurant Management, University of Houston, Houston, TX 77204-3028
ABSTRACT The oral cavity is a dynamic environment with several niches for attachment of a variety of flora. The dominant flora in the mouth are comprised of anaerobic Gram-positive bacteria. Salmonella is a Gramnegative bacterium which, according to the literature, is found very rarely in the parotid gland of the mouth of humans. One of the most important characteristics of bacteria in the oral cavity is their ability to attach to the mucosal cells. Salmonella has displayed ability to attach to the epithelial cells of the intestine and have a variety of fimbiral lectins, type I fimbriae, and flagella which aid attachment to a variety of cells. This review details the ability of Salmonella to cause disease and the potential of Salmonella as a pathogen in the oral cavity of humans. Keywords: Salmonella, dental, oral cavity Agric. Food Anal. Bacteriol. 3: 30-38, 2013
INTRODUCTION Salmonella is a facultative intracellular Gram-negative pathogen transmitted by the ingestion of contaminated food and water. Depending on the species, Salmonella may cause either typhoid fever or enteritis. S. Typhi causes typhoid fever which is a serious systemic infection. S. Typhimurium causes gastroenteritis which is localized infection in the intesCorrespondence:Sujata A. Sirsat, sasirsat@central.uh.edu Tel: +1 -713-743-2624 Fax: +1-713-743-3696
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tines of humans (Takaya et al., 2005). The incidence of illness due to salmonellosis has risen in the past three decades. Foods such as eggs, poultry, peanut butter, raw sprouts and most commonly implicated in foodborne Salmonella outbreaks (Maki, 2009). One of the largest outbreak of Salmonella Typhimurium occurred in Chicago in 1984 when 200,000 people acquired the pathogen via pasteurized milk contaminated with non-pasteurized milk (Tafazoli et al., 2003). Salmonella outbreaks in foods can be prevented by proper food handling practices and thorough cooking of poultry and eggs. Improv-
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ing standards of quality testing and animal rearing practices will also be advantageous (Tafazoli et al., 2003). However, contaminated ready-to-eat (RTE) foods pose a challenge since these are not processed before consumption (Koo et al., 2012). More recently, Salmonella outbreaks have occurred in ground beef, mangoes, cantaloupe, and pet food (CDC, 2012a; CDC, 2012b; CDC, 2012c; CDC, 2013). The evidence of Salmonella in the oral cavity dates back to 430 BC (Sulonen et al., 2007). There have been a few cases reported of Salmonella infection in the parotid gland of the mouth (Sulavik et al., 1997; Suárez and Rüssmann, 1998; Stern et al., 2001; Sulakvelidze et al., 2001; Stone 2002; Sturny et al.,
been proven by testing the effects of these factors on Salmonella strains containing lacZ fusions in the invasion genes and performing ß–galactosidase assays and improved invasion of Salmonella in tissue culture cell lines (Cheung et al., 1999; Davidson and Harrison, 2002). In addition, (Sirsat et al., 2011) demonstrated that sublethal heat stress may increase the attachment ability of Salmonella in Caco-2 cells. SPI1 encodes a type III secretion system (TTSS) which is essential for the process of cell invasion. The protein forms a secretory “needle complex” which spans the inner and outer bacterial membrane (Jones, 2005). Control of invasion genes leads to the formation of the type III secretion apparatus at the point of infec-
2003; Strindelius et al., 2004; Stern et al., 2005; Stern et al., 2006). The parotid gland is the salivary gland in humans that facilitates the early digestion of starch and swallowing (Thesleff et al., 1988).These cases are most often caused by Salmonella due to secondary infections (Stern et al., 2001). As will be discussed in the following sections, Salmonella has a tremendous ability of attach to epithelial cells of the intestine of the host. The focus of this review is to discuss the potential ability of Salmonella to survive and attach to the oral mucosa and propose possible methods to study this interaction.
It is believed that Salmonella virulence has evolved as a result of horizontal gene transfer (Johnston et al., 1996). This concept is supported by the fact that large numbers of virulence genes are clustered within its chromosome. The five Salmonella Pathogenicity Islands (SPI) identified in Salmonella are located in various regions on the chromosomes and contain sets of virulence genes (Foster and Spector, 1995). Invasion is believed to be controlled
tion (Bowe et al., 1998). This secretion system is used by microorganisms to translocate virulence-associated effector proteins into the cytoplasm of the host cells resulting in a cross-talk which leads to down stream responses such as membrane ruffling on the surface of the host cell followed by the formation of phagosomes which may internalize the bacteria (Lucas and Lee, 2000; Liu et al.,2008). SPI-1 specifically codes for transcriptional regulators such as hilA (Davidson and Harrison, 2002), hilC (Chenoweth et al., 2007), and hilD (Chatfield et al., 1992). The hilA gene has been shown to be required for the expression of three other invasion genes: invF, sspC, and orgA (Davidson and Harrison, 2002). The gene invF is a transcriptional regulator (Enoch, 2007), sspC codes for an invasion protein (Libby et al., 1994), and orgA product is a component of an export machinery system (Leverentz et al., 2001). PhoPQ is a regulatory two-component system, which is not contained within SPI-1 but is crucial for invasion phenotype of Salmonella (Liljemark and Gibbons, 1972; Lichtensteiger and Vimr, 2003). This two-component system responds to extracellular cation levels (Lillard, 1980). In conditions of low cation concentration, sensor kinase PhoQ phospohorylates the regulator PhoP which activates pag transcription. Induction and ex-
by genetic and environmental regulators. Environmental signals such as high pH (Cheung et al., 1999), low osmolarity (Darwin and Miller, 1999) and low oxygen (Cirillo et al., 1998) are believed to increase invasion of Salmonella in the host. This has
pression of pag genes are required for the survival of the bacteria in the macrophage (Lewis et al., 2009). Other pathogenicity islands, SPI-2 and SPI-5, encode genes which are responsible for pathogenesis of Salmonella after invasion of the host (Shumway, 1990).
GENETICS OF SALMONELLA PATHOGENESIS
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Salmonella attachment to epithelial cells is dependent on lectin-like adhesions which recognize specific glycoconjugate receptors on the host cells. Salmonella has several types of fimbrial lectins including SEF14. SEF17, and SEF21 (Teplitski et al., 2003). Other fimbriae such as plasmid-encoded fimbriae (PEF) and long polar fimbriae (LPF) of Salmonella are also believed to be involved in the adhesion of the bacteria to villus of the small intestine and the Peyers patches (Terai et al., 2005; Mihaljevic et al., 2007). Studies done by (Teplitski et al., 2003) have shown that flagella also play an important role in the attachment of Salmonella to epithelial cells of the host by functioning to move to appropriate sites.
leads to downstream activation of both the T helper (Th) and T cytotoxic (Tc) cells (Thomson et al., 1977; Testerman et al., 2002). Tc cells either kill the intracellular bacteria or release live bacteria when they lyse. Th cells activate B cells which may produce antibodies which could aid in opsonizing the bacteria (Thomson et al.,1977; Testerman et al., 2002). Mice studies have shown that the depletion of Th cells had a more pronounced effect than the depletion of Tc cells. Transfer of Th cells to naïve recipients induced more effective vaccination compared to transfer of Tc cells. The following section discusses the microbial ecology of the mouth (Mittrucker and Kaufmann, 2000).
The next section discusses host immune reaction to combat virulent Salmonella.
ORAL MICROBIAL ECOLOGY IMMUNE REACTION TO COMBAT SALMONELLA Salmonella enters the host via contaminated food or water (Threlfall, 2006). The pathogen exhibits increased acid-tolerance response when exposed to the low pH of the stomach (Leistner and Gorris, 1995). Following this, Salmonella enters the small intestine and moves towards and adheres to the intestinal epithelium due to expression of several fimbriae genes (Leistner, 2000). Cytoskeletal rearrangements occurs once the bacteria adhere to the epithelial surface causing host membrane ruffling which leads to enclosing adherent bacteria in vesicles (Li et al., 2002). Salmonella invasion into epithelial cells leads to recruitment of neutrophils into the intestinal lumen. One of the significant factors that lead to neutrophil recruitment is the secretion of IL-8 by the epithelial cells (Takaya et al., 2005). Following this, Salmonella enters the macrophages leading to the activation of its virulence mechanisms which further leads to its survival and replication in the host. In some cases
Bacteria have been found in the mouths of infants shortly after birth. Organisms such as Escherichia coli are transient microorganisms. Others such as S. salivarius and lactobacilli are more characteristic of adult flora and are found within a few days (Takaya et al., 2002). Studies have shown that maternal saliva may act as a source of some Gram-negative anaerobes in the oral microflora of infants even before tooth eruption (Könönen et al., 2007). Oral flora rapidly changes once teeth erupt in the mouth since teeth provide an additional unique surface for bacterial colonization. Saliva is another common environmental influence in the mouth since it washes away non-adherent bacteria and has inherent buffering properties (Russell and Melville, 1978). Additionally, other factors such as diet, oral hygiene, and general health also influence the nature of oral flora (Tatusov et al., 2003). The mouth contains several different types of bacteria. This could be because of the availability of several different niches available for bacterial attachment. These surfaces are the teeth, mucosal epi-
Salmonella may enter the blood and cause secondary infections colonizing in various organs of the body (Lewis et al., 2009). Salmonella also has the potential to activate the adaptive immune system. Activation of cytokines
thelial cells, and the bacterial layers that constitute the plaque on the teeth (Whittaker et al.,1996). This topic has been extensively reviewed by (Tafazoli et al., 2003; Takaya et al., 2004). The majority of bacteria belong to streptococci genus which is a Gram-posi-
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tive filamentous organism. In particular, Streptococcus mutans are especially important as they cause dental caries (Gibbons and Houte, 1975; Gibbons, 1984). S. mitis and S. sanguis are other streptococci that are present in lower numbers (Liljemark and Gibbons, 1972). Other Gram-positive rods such as those belonging to the Actinomyces species are also present. These are involved in some periodontal diseases. Gram-negative organisms such as Neisseria and Veillonella may be also present on the oral cavity. Most of these bacteria are anaerobic and include members of Bacteroides, Fusobacterium, and Vibrio (Gibbons, 1984). The indigenous oral flora exerts competition to-
polymerase chain reaction (PCR) and screened six different pathogens using specific DNA primers. They concluded that the pathogen in the dental pulp of the skeletons was Salmonella or a pathogen very closely related to Salmonella. The parotid gland is the salivary gland that is often affected due to inflammation (Takaya et al., 2004; Takaya et al., 2005). Parotitis usually manifests in infants, elderly persons, and the immunocompromised (Takaya et al., 2005). The most common pathogen that is associated with bacterial parotitis is Staphylococcus aureus (Takaya et al., 2005). Other organisms such as Haemophilus influenzae, Klebsiella pneumonia, and Salmonella spp. are not as frequently
ward bacteria found elsewhere in the human body and hence it is rare to isolate Salmonella, E. coli or Staphylococcus aureus from the mouth. This is because all the attachment sites are taken up by the indigenous flora and their numbers are a lot higher as compared to the potential pathogenic bacteria (Tavazoie et al., 1999). There are several forces that regulate the colonization of bacteria in the mouth. The most important factor however is bacterial adherence to surfaces in the mouth. Attachment properties are especially crucial for microorganisms since there is a risk of being washed out of the oral cavity entirely (Tafazoli et al., 2003). The mouth has proven to be a particularly interesting model for studying adhesion between host and pathogens since it has several different microcosms which can be easily sampled. Additionally, there is increased interest to study these interactions because microorganisms cause periodontal diseases. It has been demonstrated that dietary sucrose is required for accumulation of microorganisms on teeth surfaces (Taitt et al., 2004).
isolated (Takaya et al., 2004). Nine cases of parotid abscess due to Salmonella has been reported in the literature to date (Sulavik et al., 1997; Suárez and Rüssmann, 1998; Stern et al., 2001; Sulakvelidze et al., 2001; Stone 2002; Sturny et al., 2003; Strindelius et al., 2004; Stern et al., 2005; Stern et al., 2006). As described in the previous sections, Salmonella causes these types of infections after it enters the gastrointestinal tract and is disseminated into the bloodstream. The secondary areas of colonization are usually diseased tissue and trauma sites (Stern et al., 2001). (Stern et al., 2001) presented a case study of a 50-year old patient who had parotid abscess. This individual may have been infected orally or it could have been a result of a secondary infection due to Salmonella. However, the patient did not show symptoms of gastroenteritis and hence may have been a carrier of the pathogen eventually resulting in Salmonella spreading to the bloodstream and the manifestation of parotid abscess. Saliva contains the enzymes lysozyme and lactoferrin (Tafazoli et al., 2003) and Gram-negative pathogens such as Salmonella and E. coli have been shown to be sensitive toward lactoferrin as it acts as an antimicrobial (Takaya et al., 2002). In addition, these antimicrobial enzymes interact with each other
SALMONELLA IN THE ORAL CAVITY Evidence of Salmonella in the oral cavity has been seen as far back as 430 BC when (Sulonen et al., 2007) set out to determine the probable cause of the “Plague of Athens” using skeleton dental pulp from the mass burial site. The researchers performed
in a synergistic or additive manner (Tenovuo, 1998). Lysozyme acts on Gram-negative bacteria by hydrolyzing 1,4-beta-linkages in the bacterial cell walls (Proctor et al., 1988). Lactoferrin interacts with the lipolysacharide of the Gram negative cell membrane
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Figure 1. Flow chart of an experimental method to test the ability of bacterial attachment to human epithelial cells from the oral cavity
Collect human epithelial cells by scraping oral cavity with wooden stick
Wash cells to remove unattached bacteria and diluted to 105 cells/ml
Mix cell suspensions with bacteria and incubate at 35°C for 30 min
Wash cells to removed unattached bacteria
Stain bacteria and enumerate under the microscope
(Appelmelk et al., 1994). This may explain why such few cases of Salmonella persistence are evident in the oral cavity. Independent studies have found that specific Salmonella mutations may lead to resistance against lysozyme (Sanderson et al., 1974).
As discussed in previous sections, the key to bac-
be effective against Gram-negative organisms such as Salmonella and E. coli (Takaya et al., 2002). One of the initial studies to examine Salmonella in the oral cavity would be to test the effect of saliva on the viability of Salmonella. This would give investigators an indication if the concentration of lysozyme in saliva is inhibitory to Salmonella. This can be done by collecting saliva from humans and adding cultures of Salmonella grown overnight and testing viability by the plate count method.
terial presence and survival in the mouth is attachment to various sites in the oral cavity. This is especially challenging in the mouth due to the constant washing of non-adherent bacteria by saliva. Also, saliva contains lysozyme which has been shown to
Tenor et al. (2004) prepared epithelial cell suspensions to test the adherence of bacteria to the cells of oral cavity and similar studies could be performed for Salmonella to test its capability of adherence to these types of cells. A flow-chart of the method is
POTENTIAL STUDIES
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demonstrated in Figure 1. The method can be described briefly as follows. The researchers collected human epithelial cells by scraping oral mucosal cells with a wooden stick and twirled the applicator in saline to dislodge the cells. The cells were subsequently washed free of unattached bacteria and adjusted with saline to 105 cells/mL. Following this, the authors tested the ability of various bacteria to adhere to the epithelial cells. This was done by mixing cell suspensions of the bacteria with cell suspensions of the epithelial cells and incubating the mixture at 35°C for 30 min. The epithelial cells were washed to remove non-adherent bacteria. The bacteria were stained in order to observe and enumerate the cells
monella; however, some strains may be resistant to these enzymes. Additional studies are needed to understand the implications of Salmonella in the oral cavity and complications that may results downstream. The proposed experiments may be able to identify whether Salmonella possesses gene(s) that encodes for the necessary fimbrial lectins to attach to surfaces of the oral cavity.
under the microscope. A more quantitative method would involve conducting viable bacteria studies by recovering cells as petri plate counts and assessing the number of Salmonella that adhere to the human epithelial cells. This study would offer the means for assessing the capability of Salmonella to compete with the normal flora of the human oral cavity. Also, the results may indicate whether Salmonella has the necessary mechanisms to attach to surfaces of the mouth. Tenor et al. (2004) performed electron microscopy studies after mixing bacterial cultures and epithelial cells to observe the adherence property of bacteria. Similar studies could be performed with Salmonella and human epithelial cells. The electron microscopy analyses would be particularly useful in demonstrating visually, the ability of Salmonella to adhere to these cells in the presence of other bacterial competition.
The ability to adhere to a variety of surfaces available in the oral cavity of the mouth is of utmost important to the flora that primarily resides in the
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CONCLUSIONS
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REVIEW Shiga Toxin-Producing Escherichia coli (STEC) Ecology in Cattle and Management Based Options for Reducing Fecal Shedding T. R. Callaway1, T. S. Edrington1, G. H. Loneragan2, M. A. Carr3, D. J. Nisbet1 Food and Feed Safety Research Unit, USDA/ARS, 2881 F&B Rd., College Station, TX 77845 2 Department of Animal and Food Sciences, Texas Tech University, Lubbock, TX 79409 3 Research and Technical Services, National Cattlemen’s Beef Association, Centennial, CO 80112 1
Proprietary or brand names are necessary to report factually on available data; however, the USDA neither guarantees nor warrants the standard of the product, and the use of the name by the USDA implies neither approval of the product, nor exclusion of others that may be suitable.
ABSTRACT Cattle can be naturally colonized with foodborne pathogenic bacteria such as Shiga Toxin-producing E. coli (STEC) in their gastrointestinal tract. While these foodborne pathogens are a threat to food safety, they also cause human illnesses via cross contamination of other foods and the water supply, as well as via direct animal contact. In order to further curtail these human illnesses and ensure a safe and wholesome food supply, research into preharvest pathogen reduction controls and interventions has grown in recent years. This review addresses the ecology of STEC in cattle and potential controls and interventions that have been proposed or implemented to reduce STEC in cattle. We focus in this review on the use of management practices and the effects of diet and water management. Implementation of preharvest strategies will not eliminate the need for good sanitation procedures in the processing plant and during food preparation and consumer handling. Instead, live-animal management interventions must be implemented as part of a multiple-hurdle approach that complements the in-plant interventions, so that the reduction in pathogen entry to the food supply can be maximized.
Keywords: E. coli O157:H7, EHEC, cattle, management
Agric. Food Anal. Bacteriol. 3: 39-69, 2013
of Shiga Toxin-producing Escherichia coli (STEC) bacteria, which are part of the natural reservoir in ruminant animals such as cattle (Karmali et al., 2010).
INTRODUCTION One of the largest food safety (and economic) impacts on the cattle industry has been the emergence Correspondence: Todd Callaway, todd.callaway@ars.usda.gov Tel: +1-979-260-9374 Fax: +1-979-260-9332.
STEC-caused illnesses cost the American economy more than $1 billion each year in direct and indirect costs from more than 175,000 human illnesses (Scallan et al., 2011; Scharff, 2010). Furthermore, since the emergence of the “poster child” of STEC, E. coli
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O157:H7, more than $2 billion dollars have been spent by the cattle industry to combat STEC in processing plants (Kay, 2003). While post-harvest pathogen-reduction strategies have been largely successful at reducing direct foodborne illness, these processing interventions have not been perfect (Arthur et al., 2007; BarkocyGallagher et al., 2003), in large part because avenues of human exposure include indirect routes (LeJeune and Kersting, 2010; Nastasijevic, 2011). In order to further curtail human illnesses and ensure a safe and wholesome food supply, research into preharvest pathogen reduction controls and interventions has grown in recent years (Callaway et al., 2004; LeJeune
EHEC, STEC, VTEC AND NON-O157:H7’S: A PRIMER
and Wetzel, 2007; Oliver et al., 2008; Sargeant et al., 2007). The impact of using pathogen reduction strategies focused on the environmental contamination and exposure routes at the live animal stage are likely to have large impacts on resulting human illnesses (Rotariu et al., 2012; Smith et al., 2012). Reduction of STEC can also yield public health improvements in rural communities (LeJeune and Kersting, 2010) and amongst attendees of agricultural fairs, rodeos and open farms (Keen et al., 2006; Lanier et al., 2011). Thus, the logic underlying focusing on reducing foodborne pathogenic bacteria in live cattle is straightforward: 1) reducing the amount of pathogens entering processing plants will reduce the burden on the plants and render the in-plant interventions more effective; 2) reducing horizontal pathogen spread from infected animals (especially in “supershedders”) in transport and lairage; 3) will reduce the pathogenic bacterial burden in the environment and wastewater streams; and 4) will reduce the direct risk to those in direct contact with animals via petting zoos, open farms, rodeos and to animal workers. This review addresses the microbial ecology of STEC colonization of cattle and controls and interventions that have been proposed or imple-
ers refer to these pathogens often interchangeably as Enterohemorrhagic E. coli (EHEC), Shiga toxinproducing E. coli (STEC), or Verotoxin-producing E. coli (VTEC). While E. coli O157:H7 was the first of the STEC’s to be recognized as a major food safety threat, recently the other “non O157:H7 STEC” have been increasingly implicated in human illness outbreaks (Bettelheim, 2007; Fremaux et al., 2007). Because of this linkage, the “gang of six” non-O157 serogroups (O26, O45, O103, O111, O121, and O145) have joined O157:H7 as being classified as adulterants in beef (USDA/FSIS, 2012). Since this declaration, focus has shifted on understanding the generalized STEC ecology, rather than simply focusing on E. coli O157:H7 (Gill and Gill, 2010). For years, researchers (including the present authors) assumed that in general, all of the non-O157 STEC would behave similarly to O157:H7 in a physiological and ecological sense. However, recent research has found that in addition to the genetic divergence seen in O157:H7 lineages (Zhang et al., 2007), there appear to be significant physiological differences between and within non-O157 STEC which may play a role in the ecological niche occupied in the ruminant gastrointestinal tract by these non-O157 serotypes (Bergholz and Whittam, 2007;
mented to reduce STEC in live cattle in the areas of: 1) Management practices and transport, and 2) Cattle water and feed management.
Free et al., 2012; Fremaux et al., 2007). While these and other physiological differences need to be investigated further, and their roles in the gastrointestinal microbial population must be determined, it appears that the O157:H7 serotype is well adapted to survive
40
Although the relatively recent (1982) emergence of E. coli O157:H7 into public view makes it seem that this organism is a new arrival in the food chain, data indicates that this organism is far more ancient, having arisen between 400 and 70,000 years ago (Law, 2000; Riley et al., 1983; Wick et al., 2005; Zhou et al., 2010). Although a variety of acronyms have been applied to the “hamburger bug”, they belong to a single group that acquired toxin genes from Shigella via a gene transfer event (Kaper et al., 2004; Karmali et al., 2010; Wick et al., 2005). Research-
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in cattle (García et al., 2010; O’Reilly et al., 2010) and that other STEC serotypes can also live in the gastrointestinal tract of cattle (Arthur et al., 2002; ChaseTopping et al., 2012; Joris et al., 2011; Monaghan et al., 2011; Polifroni et al., 2012; Thomas et al., 2012), and be transferred into ground beef (Bosilevac and Koohmaraie, 2011; Fratamico et al., 2011). While we understand much of how E. coli O157:H7 behaves in the gastrointestinal tract and farm environment, we know very little about the ecology of other STEC in those environments (Monaghan et al., 2011; Polifroni et al., 2012). Thus this review focuses on pre-harvest pathogen interventions based upon E. coli O157:H7 data. We hypothesize that non-
rens and Hovde, 2011). This transmission most frequently occurs during summer months, and is linked to a summer increase in the prevalence of E. coli O157:H7 in cattle (Edrington et al., 2006a; Lal et al., 2012; Naumova et al., 2007; Ogden et al., 2004; Wells et al., 2009), not just an increase in consumption or a change of cooking habits by consumers (Money et al., 2010; Williams et al., 2010a). It has been suggested that neuroendocrine factors may play a role in E. coli O157:H7 (Edrington et al., 2006a; Green et al., 2004), as may signaling between host and intestinal microbial populations or within STEC populations via quorum-sensing (Edrington et al., 2009b; Sperandio, 2010; Sperandio et al., 2001). Further
O157:H7 STEC will largely behave in a broadly similar fashion to E. coli O157:H7 in the gastrointestinal and farm environment; however this is an educated assumption and that imposed limitation must be understood. Thus readers must be aware that most of the research referred to in this review is based upon E. coli O157:H7 specifically, and may or may not apply to all STEC.
possible interactions within the microbial ecosystem of the rumen are demonstrated in the preferential consumption of E. coli O157:H7 by ruminal protozoa (Epidinium), and increased populations in the presence of Dasytricha (Stanford et al., 2010). Because of the nature of STEC survival in the ruminant gut, it is no surprise that it persists in fecal deposits (Dargatz et al., 1997; Jiang et al., 2002; Maule, 2000; Yang et al., 2010) and in soils (Bolton et al., 2011; Semenov et al., 2009; Van Overbeek et al., 2010). This allows E. coli O157:H7 to cycle within pens and farms in a fecal-oral route (Russell and Jarvis, 2001), recirculating within groups or individual animals (Arthur et al., 2010). The presence of supershedding cattle (Chase-Topping et al., 2008) in the population can further enhance this horizontal transmission within a herd or a pen of cattle (Arthur et al., 2010; Arthur et al., 2009; Cobbold, 2007; LeJeune and Kauffman, 2006). However, the host/dietary/microbial factors underlying the “supershedder” status of cattle remains unknown, as do factors that allow simple gut colonization by E. coli O157:H7. Thus it is apparent that the farm/pen/facility environment plays an important role in STEC colonization and recirculation, as well as via direct and indirect transmission to human farm workers/visitors and consumers
ECOLOGY OF STEC AND GASTROINTESTINAL COLONIZATION Because E. coli O157:H7 (and to some degree non-O157 STEC) co-evolved along with its host it is uniquely well-fitted to survive in the gastrointestinal tract of cattle as a commensal type organism (Law, 2000; Wick et al., 2005). While E. coli O157:H7 can live in the rumen of cattle (Rasmussen et al., 1999), the site of primary colonization is the terminal rectum (Naylor et al., 2003; Smith et al., 2009a). This organism produces a potent cytotoxin (Shiga toxin) that does not seriously impact its preferred host (cattle) because they lack toxin receptors (PruimboomBrees et al., 2000), but this same toxin causes serious illness in humans colonized by E. coli O157:H7 (Karmali et al., 2010; O’Brien et al., 1992). Unfortunately, this means that the natural commensal-type relationship between STEC (including O157 and non-O157) and cattle ensures that this organism can be passed on to meat products and consumers of beef (Fe-
(Ihekweazu et al., 2012; LeJeune and Kersting, 2010; Smith et al., 2012; Stacey et al., 2007; Strachan et al., 2006).
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MANAGEMENT PRACTICES TRANSPORTATION
AND
Good management of cattle is critical for efficient animal production, but to date no typical “management” procedures have been shown to affect colonization or shedding of foodborne pathogens (EllisIversen et al., 2008; Ellis-Iversen and Van Winden, 2008; LeJeune and Wetzel, 2007), some practices may reduce horizontal transmission and recirculation of STEC within a herd of cattle (Ellis-Iversen and Watson, 2008). However, the use of management tools like the squeeze chute (crush) to process cattle has been shown to increase the odds of hide contamination with E. coli O157(Mather et al., 2007). Yet, in spite of this lack of evidence regarding impacts on food safety, good management practices are critical to ensuring animal health and welfare (MorrowTesch, 2001).
Bedding and pen surfaces E. coli O157:H7 can live for a long period of time in manure, soil and other organic materials (Jiang et al., 2002; Maule, 2000; Winfield and Groisman, 2003) and can be transmitted successively through their environment (Semenov et al., 2010; Semenov et al., 2009). Cattle, especially dairy cows, are bedded on materials that are largely chosen on animal health and welfare grounds. Unfortunately, bedding material can harbor bacteria that are responsible for mastitis, as well as foodborne pathogenic bacteria that can be spread between cattle (Davis et al., 2005; Oliver et al., 2005; Richards et al., 2006; Wetzel and LeJeune, 2006). Researchers have shown that urine increases growth of E. coli O157:H7 on bedding, potentially by providing substrate for growth (Davis et al., 2005). Modeling research has shown that an increase in bedding cleaning frequency would increase the death rate of E. coli O157:H7 (Vosough Ahmadi et al., 2007). Further studies have demonstrated that the use of very dry bedding reduced E. coli O157:H7 prevalence on farms (Ellis-Iversen et al., 2008; EllisIversen and Van Winden, 2008). Researchers have shown that sand bedding reduced transmission of E. 42
coli O157:H7 between dairy cows, resulting in lower populations of E. coli O157:H7 in cattle bedded on sand compared to sawdust (LeJeune and Kauffman, 2005; Westphal et al., 2011). It is suspected that this difference was due to desiccation or reduced nutrient availability. Feedlot surfaces were thought to contain manure-like bacterial populations, but recent molecular studies have indicated that the bacterial communities of feedlot surfaces are complex, yet utterly distinct from fecal bacterial populations (Durso et al., 2011). This suggests that traits that favor survival in the gastrointestinal tract (anaerobic, warm, dark) do not favor survival on the feedlot surface (aerobic, cooler, sunlit). Surfaces such as pond ash do not impact survival of E. coli O157 (Berry et al., 2010), however studies and anecdotal evidence indicates that a greater number of cattle shed E. coli O157:H7 when housed in muddy pen conditions than cattle from pens in normal condition and that the condition of the pen floor may influence the prevalence of cattle shedding the organism and the ability of E. coli O157:H7 to survive dry conditions (Berry and Miller, 2005; Smith et al., 2001; Smith et al., 2009b). Studies have recently demonstrated that sunlight can reduce E. coli O157:H7 populations on pen surfaces (Berry and Wells, 2012) and in water systems (Jenkins et al., 2011). Overall, bedding or pen cleaning will not eliminate E. coli O157:H7 from any farm or feedlot environment, but it may slow spread within a herd or between penmates.
Manure Management E. coli O157:H7 and other STEC survive in manure and can persist for a lengthy period of time (up to 21 months) (Bolton et al., 2011; Fremaux et al., 2007; Hutchison et al., 2005; Kudva et al., 1998; Varel et al., 2008). Although there are differences amongst STEC strains in their ability to persist in manure, these appear to be related to the oxidative capacity of each strain (Franz et al., 2011). The presence of a native bacterial population in manure reduces E. coli O157:H7 survival in soils (Van Overbeek et al., 2010). The amendation of manure in soil can result in
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STEC uptake directly by plants, including food crops (Franz and Van Bruggen, 2008; Jiang et al., 2002; Semenov et al., 2010; Semenov et al., 2009). Rainfall events can also wash STEC from cattle feces (stored or in fields) into drinking or irrigation water supplies (Anonymous, 2000; Cook et al., 2011; Ferguson et al., 2007; Oliveira et al., 2012; Pachepsky et al., 2011). As the mean temperature of manure rises during storage the survival of E. coli O157:H7 is reduced (Semenov et al., 2007), indicating that composting can enhance manure safety, thus reducing human illnesses (Graham and Nachman, 2010; Kelley et al., 1999). There have been few studies that have isolated STEC consistently from cattle waste lagoons (Purdy
ding (Gunn et al., 2007; Synge et al., 2003). Ruminant animals other than cattle do carry E. coli O157:H7 (French et al., 2010; Hussein et al., 2000; Sargeant et al., 1999), and this includes sheep and deer that often share the same pasture, feed bunks and water supplies (Bolton et al., 2012; Branham et al., 2005). Other researchers have found that flies and other insects on farms can carry STEC from one location to another (Ahmad et al., 2007; Hancock et al., 1998; Keen et al., 2006; Talley et al., 2009). Furthermore, wild migratory birds such as starlings (Carlson et al., 2011a; Carlson et al., 2011b; Cernicchiaro et al., 2012; Wallace et al., 1997; Wetzel and LeJeune, 2006), cowbirds and egrets (Callaway, unpublished
et al., 2010). This is potentially due to the oxidized nature of the lagoon, or the presence of a native microbial population. In waste water lagoons, there are protozoa that preferentially consume E. coli O157:H7 (Ravva et al., 2010), possibly explaining at least some of the difference between E. coli O157:H7 survival in manure with that of limited survival in dairy lagoons (Ravva et al., 2006). Research has demonstrated that the addition of chemical oxidants to wastewater lagoons can reduce pathogen populations (LusterTeasley et al., 2011).
data) can carry STEC (and other foodborne pathogen) between pens, and even between farms long distances apart. While these effects are probably minimal in their direct impact on food safety within a farm, they represent vectors for pathogens to move between “clean” groups of cattle or farms.
Cattle grouping
Farm biosecurity is critical for animal health and welfare, especially in regard to animal diseases, but to date there has been little direct impact demonstrated on foodborne pathogenic bacteria such as E. coli O157:H7 (Ellis-Iversen and Van Winden, 2008). Research has shown that other animal species, rodents, insects and birds and boars can carry STEC at least transiently (Branham et al., 2005; Cernicchiaro et al., 2012; French et al., 2010; Rice et al., 2003; Sánchez et al., 2010; Wetzel and LeJeune, 2006). Mixing of sheep with cattle has been shown to increase the risk of cattle shedding STEC (Stacey et al., 2007),
Many farms are closed to entry by animals from other farms to prevent animal disease transmission. Closed herds prevent spread of E. coli O157:H7 (and other pathogens) from one farm to another (EllisIversen et al., 2008; Ellis-Iversen and Van Winden, 2008; Ellis-Iversen and Watson, 2008). However some studies have shown that a closed farm does not impact E. coli O157:H7 incidence on farms (Cobbaut et al., 2009). The results of this study suggest that E. coli O157:H7 should be considered common to groups of feedlot cattle housed together in pens (Smith et al., 2001), thus keeping groups together throughout their time on a farm, or in a feedlot, without introducing new members to groups appears to reduce horizontal transmission between animals. A further benefit of grouping cattle involves the use of age as a segregating factor. Young cattle (es-
and a positive correlation between cattle and sheep density was found, at least in the UK (Strachan et al., 2001). Other diverse factors such as the presence of dogs, pigs, or wild geese on the farm have been linked to an increased risk of E. coli O157:H7 shed-
pecially heifers) shed more E. coli O157:H7 than do older cattle (Cobbaut et al., 2009; Cray and Moon, 1995; Smith et al., 2001). While it is not possible to segregate calves from cows in the beef industry, there is potential benefit to keeping same-age
Biosecurity
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groups of calves together as they are transported and enter backgrounding or feedlot operations to prevent horizontal transmission between groups. Off-site rearing of dairy heifers may be an important solution to reducing foodborne pathogens, as has been shown in regard to Salmonella (Hegde et al., 2005), and the risk of transmission back to the farm by heifers returning from an off-site facility was found to be low (Edrington et al., 2008). Animal density may also play a role in the horizontal spread of E. coli O157:H7 and other foodborne pathogens (Vidovic and Korber, 2006) as well as the vertical spread to humans (Friesema et al., 2011b). Densely packed animals have a great chance of con-
Handling and transport to processing plants or feedlots or other farms causes stress (see below) and may spread E. coli O157:H7 due to physical contact or fecal contamination, and trailers used may spread pathogens between lots or loads of cattle (Mather et al., 2007). Studies have indicated that transport did not affect STEC populations in cattle, however in these studies E. coli O157:H7 populations were very low initially (Barham et al., 2002; Minihan et al., 2003; Reicks et al., 2007; Schuehle Pfeiffer et al., 2009). However, other studies have found that transport caused an increase in fecal shedding of E. coli O157:H7 (Bach et al., 2004). Researchers found that transporting cattle more than 100 miles doubled the
tamination with fecal spread. However increased animal density reduces the physical footprint and may allow for more efficient and effective waste handling. It has been shown that higher animal density can be linked to an increased risk of carriage of some STEC, including O157:H7 (Frank et al., 2008; Vidovic and Korber, 2006). Other European studies have also found an effect of animal density on human STEC illnesses (Friesema et al., 2011a; Haus-Cheymol et al., 2006), yet Canadian researchers found a variable impact (Pearl et al., 2009). Further studies found that increased stocking density increased shedding of STEC, independent of group size (Stacey et al., 2007; Strachan et al., 2006). The issue of “supershedders” complicates research into effects of animal density and pathogen shedding (Arthur et al., 2009; Cernicchiaro et al., 2010; Chen et al., 2012; LeJeune and Kauffman, 2006; Stanford et al., 2005). If supershedders do exist long term, rather than simply being a transient phase of infection, then there are interactive effects of animal density and pathogen density in the animal that must be accounted for (Matthews et al., 2006; Matthews et al., 2009). The role of super-shedding animals (even if a transient phenomenon) cannot be discounted in the contamination of hides during transport and
risk of having positive hides at slaughter compared to cattle shipped a short distance, though the question of time in close-confinement versus being in transit was not examined (Dewell et al., 2008). In another study, longer transport times were correlated with increased levels of fecal shedding of E. coli O157:H7 (Bach et al., 2004). It was also demonstrated that a combination of transport and lairage did not lead to an increase in the number or prevalence of E. coli O157:H7 from cattle (Fegan et al., 2009). The presence of a high shedding animal in a trailer has been shown to increase the odds of other animals within the load being hide-positive for E. coli O157:H7 (Arthur et al., 2010; Arthur et al., 2009; Fox et al., 2008). However, it should be noted that both low- and high-shedding cattle can be responsible for the spread within and between truckloads (Dodd et al., 2010). Cattle trailers can be important fomites of E. coli O157:H7 to uninfected cattle and are frequently positive for E. coli O157:H7 when sampled (Barham et al., 2002; Cuesta Alonso et al., 2007; Reicks et al., 2007). It has been shown that the incidence of E. coli O157:H7 in transport trailers increases the risk of transmission to farms and feedlots from cattle on these trailers (Cuesta Alonso et al., 2007). To date however, the washing of trailers has
lairage, especially in dense conditions (Arthur et al., 2010; Arthur et al., 2009).
only been shown to be effective against Salmonella contamination in swine (Rajkowski et al., 1998), yet it is an intuitive, feasible solution to prevent some degree of cross-contamination of cattle during a stressful period.
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Lairage and holding facilities are further locations that can impact the prevalence and concentration of E. coli O157:H7 on hides of cattle, which is an important route of entry to the food supply (Arthur et al., 2007). Studies have shown that the transfer of E. coli O157:H7 to hides that occurs in lairage at processing plants accounted for more of the hide and carcass contamination than did the population of cattle leaving the feedlot (Arthur et al., 2008). Furthermore, the presence of supershedding cattle in these pens can increase the spread of E. coli O157:H7 between animals from different farms or feedlots (Arthur et al., 2010; Cernicchiaro et al., 2010). The exact role of lairage and transport/trailers in the spread of E. coli
of E. coli O157:H7 in dairy calves (Edrington et al., 2011). Other stresses such as movement have been identified as playing a role in E. coli O157:H7 shedding in Scottish cattle (Chase-Topping et al., 2007), but this has not been clearly defined in U.S. cattle systems. When calves were preconditioned to transport stress, they were found to be less susceptible to infection from the environment than were calves not preconditioned to this stressor (Bach et al., 2004). Cattle with excitable temperaments were less likely to shed E. coli O157:H7 than were “calm” cattle (Brown-Brandl et al., 2009; Schuehle Pfeiffer et al., 2009). In studies with pigs, it was found that the so-
O157:H7 (and other pathogens) in cattle is unclear, and is likely time- and animal density-dependent, and may also be affected by stress.
cial stress/excitement of mixing penmates increased fecal shedding of Salmonella (Callaway et al., 2006), but this has not been shown to date in cattle, however this implies a potential role of social stresses in cattle during lairage. Heat stress (and methods to alleviate it) can have effects on animal health and productivity (BrownBrandl et al., 2003), as well as shedding of E. coli O157:H7 and Salmonella (Callaway et al., 2006). Water sprinkling to alleviate heat stress in cattle increased measures of animal well-being and decreased E. coli O157:H7 populations on the hides of cattle, but did not affect fecal populations (Morrow et al., 2005). In another study with dairy cattle, researchers found that heat stress had no impact on STEC shedding, but Salmonella shedding was increased (Edrington et al., 2004). Other researchers have also found that heat stress did not impact E. coli O157:H7 shedding in cattle (Brown-Brandl et al., 2009).
Stress While we understand stress intuitively, any discussion of “stress” in animals is fraught with anthropomorphism and complexity (Rostagno, 2009; Verbrugghe et al., 2012). Long-term stress may depress immune function in cattle (Carroll and Forsberg, 2007; Kelley, 1980; Salak-Johnson and McGlone, 2007), making them more susceptible to colonization, but the short term effects of stress from weaning, handling or transport on immune status are unknown. Catecholamines rise when animals are under stress, and catecholamines (along with other hormones) have been demonstrated to have an effect on the microbial population, including pathogens (Freestone and Lyte, 2010; Lyte, 2010; Walker and Drouillard, 2012). To date the effect of stress on colonization or shedding of E. coli O157:H7 is unclear. Weaning is stressful to calves, and was shown to increase colonization with STEC (Herriott et al., 1998) and E. coli O157:H7 (Chase-Topping et al., 2007). In other studies however, these researchers demonstrated that weaning does not affect the likelihood of shedding (Synge et al., 2003). Interestingly, calving was seen to reduce the likelihood of E. coli O157:H7 shedding (Synge et al., 2003). Further studies found that weaning stresses alone did not impact shedding
CATTLE WATER AND FEED MANAGEMENT Diet and water supplies can be used to reduce horizontal transmission of STEC between animals on the same farm or in the same feedlot pen. The underlying biology behind these effects has not been elucidated to this point, but it has been suggested that difference could be due to increased fecal pH or
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intermediate endproducts of the yeast fermentation (e.g., vitamins, organic acids, L-lactic acid), however these suggestions remain hypothetical (Wells et al., 2009). While the magnitude of the dietary impacts effects is relatively small, it underlines the point that dietary composition can potentially significantly impact E. coli O157:H7 populations in the gut of cattle.
Cattle water troughs can harbor E. coli O157:H7 for long periods of time (Hancock et al., 1998; LeJeune et al., 2001a; LeJeune et al., 2001b; Murinda et al., 2004; Polifroni et al., 2012; Wetzel and
Cattle can be fasted for up to 48 h before and during their transport to slaughter, which can affect the prevalence of E. coli O157:H7 (Pointon et al., 2012). Ruminal and intestinal VFA concentrations limit the proliferation of E. coli because of toxicity of the VFA to the bacteria (Hollowell and Wolin, 1965; Russell and Diez-Gonzalez, 1998; Wolin, 1969). This has created the demand for the use of organic acids/VFA as methods to alter the ruminal fermentation and to reduce pathogen populations in the gut (Ohya et al., 2000; Prohaszka and Baron, 1983; Van Immerseel et al., 2006). However, fasting causes levels of VFA to decline rapidly (Harmon et al., 1999). Fasting increased E. coli, Enterobacter and total
LeJeune, 2006), and as many as 25% of cattle farm water samples have been shown to contain E. coli O157:H7 (Sanderson et al., 2006). These results suggest that these common-use troughs can be vectors for horizontal transmission of E. coli O157:H7 within a group of animals. The organic material in the water troughs tends to harbor and protect the STEC, and modeling research has shown that an increase in water trough cleaning frequency would increase the death rate of E. coli O157:H7 (Vosough Ahmadi et al., 2007) as well as exposure to sunlight (Jenkins et al., 2011). Chlorination of water supplies has long been used to reduce bacterial populations in municipal water supplies, and this also can be used in cattle water troughs to reduce E. coli O157:H7 populations. However, sunlight and organic material in the water reduces the effectiveness of chlorination over time, as has been seen in real world chlorination studies with cattle water troughs (LeJeune et al., 2004). Electrolyzed oxidizing (EO) water has been shown to reduce STEC populations as high as 104 CFU/mL (Stevenson et al., 2004), and can be used as an in-plant hide cleaning strategy (Bosilevac et al., 2005). Other treatments such as cinnamaldehyde and sodium caprylate addition to water supplies have been shown to reduce STEC populations, but
anaerobic bacterial populations throughout the intestinal tract of cattle (Buchko et al., 2000b; Gregory et al., 2000), and increased Salmonella and E. coli populations in the rumen (Brownlie and Grau, 1967; Grau et al., 1969). More recent research has demonstrated that fasting can cause “apparently E. coli (O157:H7) negative animals to become positive” (Kudva et al., 1995). Fasting made calves more susceptible to colonization by inoculated E. coli O157:H7 (Cray et al., 1998). Cattle fasted for 48 h prior to slaughter contained significantly greater E. coli populations throughout the gut than cattle fed forage (Gregory et al., 2000). In contrast, it was demonstrated that a fasting period had no effect on E. coli O157:H7 shedding (Harmon et al., 1999). When culled dairy cows were reconditioned through feeding high energy diets for 28 d before harvest, the prevalence of E. coli O157:H7 declined from 14% to 6% (Maier et al., 2011). In general, studies examining the intestinal environment have repeatedly indicated that low pH and high concentrations of short chain VFA result in lower STEC populations (Bach et al., 2002a; Bach et al., 2005b; Cobbold and Desmarchelier, 2004; Pointon et al., 2012; Shin et al., 2002). Thus the bulk of research supports the concept that fasting increases shedding or population
the effects on palatability are not currently known (Amalaradjou et al., 2006; Charles et al., 2008).
concentrations, or makes cattle more susceptible to colonization due to decreased short chain VFA and increased pH in the gastrointestinal tract. Because feed withdrawal and/or starvation results in decreased VFA concentrations in the gut, it has been
Drinking Water treatments
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suggested that this shift plays a role in the effects of transport and/or lairage on the shedding of STEC.
The first dietary practice shown in early studies to significantly increase the risk of STEC shedding among heifers was feeding corn silage (Herriott et al., 1998). In adult cows, the inclusion of animal byproducts in the diet (currently discontinued) was shown to increase STEC shedding (Herriott et al., 1998). Other studies linked feeding whole cottonseed with reduced E. coli O157 shedding (Garber et al., 1995; Hancock et al., 1994). Fecal samples from
The industrial fermentation of corn to produce ethanol has increased more than 4-fold between 2001 and 2007, and its use doubled by 2010 (Richman, 2007). Thus, an economic incentive to increase the utilization of distillers grains (DG) by-product feeds in the cattle industry has increased dramatically in recent years, especially given DG’s role as cost-effective feed supplements for finishing and lactating cattle (Firkins et al., 1985). The inclusion DG into cattle rations has been shown to be an effective replacement for common feedstuffs and has demonstrated an increased daily gain in beef cattle (Al-Suwaiegh et al., 2002) and milk yield and feed efficiency in dairy cows (Kleinschmit et al., 2006). This
cattle fed dry rolled corn, high-moisture corn and wet corn gluten feed did not contain different populations of generic E. coli, or extreme acid-resistant E. coli during a limit-feeding period (Scott et al., 2000). However, feces from cattle fed wet corn gluten ad libitum contained significantly higher concentrations of extreme acid resistant E. coli (resistant to an acid shock simulating passage through the human stomach) than did feces of cattle fed dry-rolled or high moisture corn (Scott et al., 2000). Barley is often fed to cattle and is ruminally fermented more rapidly than corn by the commensal microbial population. More starch is fermented in the lower gut of corn-fed cattle than in barley-fed cattle, resulting in barley-fed cattle having higher fecal pH and lower VFA concentrations compared with corn-fed animals (Bach et al., 2005a; Berg et al., 2004; Buchko et al., 2000a). Barley feeding was linked (albeit at a low correlation) to increased E. coli O157:H7 shedding (Dargatz et al., 1997); and in experimental infection studies barley feeding was again associated with increased shedding of E. coli O157:H7 by feedlot cattle (Buchko et al., 2000a). Survival of E. coli O157:H7 in manure from corn-and barley fed cattle is approximately equal, therefore simple survival in the feces is not responsible for the increased
improvement is likely due to the fact that DG alters the population structure and function of the microbial ecosystem of the rumen and throughout the gastrointestinal tract (Callaway et al., 2010a; Durso et al., 2012; Williams et al., 2010b). Cattle fed 40% corn wet distiller’s grains (WDG) were very different than the fecal populations in cattle fed a non DGcontaining diet, and populations of generic E. coli were higher in their feces (Durso et al., 2012), and in previous studies the survival of E. coli O157:H7 in feces was increased by increasing levels of DG supplementation (Varel et al., 2008). Unfortunately, research has suggested a potential association between DG feeding and an increased prevalence and fecal shedding of the foodborne pathogen E. coli O157:H7 in cattle (Jacob et al., 2008a; Jacob et al., 2008b; Yang et al., 2010). Distillers grains were shown to increase the shedding of E. coli O157:H7 in cow-calf operations in Scotland (Synge et al., 2003). Other researchers found that feeding a related product (brewers grain) to cattle was also associated with increased E. coli O157 shedding, and increased the odds of shedding by more than 6-fold (Dewell et al., 2005). The individual animal prevalence of feedlot cattle shedding E. coli O157 on d 122 (but not d 136) was higher in cattle
prevalence of E. coli O157:H7 in barley-fed cattle (Bach et al., 2005b).
fed 25% wet distiller’s grain compared to control diets lacking WDG (Jacob et al., 2008b), but the penlevel shedding was unaffected by WDG feeding. Pen floor fecal sample prevalence of E. coli O157 was significantly higher across a 12 week finishing period
Feed types
Distiller’s grains
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in cattle fed 25% DDG and either 15% or 5% corn silage compared with cattle fed 0% DDG and 15% corn silage (Jacob et al., 2008a). However, follow-up studies found no differences in E. coli O157:H7 fecal shedding in cattle fed DG (Edrington et al., 2010; Jacob et al., 2009), with no indications of why this difference in results was observed. In a further study utilizing both dry and wet-distillers grains, researchers found that higher levels (40% of the ration) of DG inclusion did increase fecal E. coli O157:H7 shedding (Jacob et al., 2010). When cattle were fed 40% wet DG, they had higher populations of E. coli O157:H7 as well as higher pH values and lower concentrations of L-lactate (Wells et al., 2009). Further studies
or by increasing fecal starch concentrations (Depenbusch et al., 2008).
found that the DG-associated increase in fecal E. coli O157:H7 populations could be mitigated by reducing WDG concentrations to 15% or less for 56 d prior to slaughter (Wells et al., 2011). When corn or wheat DDG were supplemented into cattle on a primarily barley-based diet, there was no difference in impact of DDG supplementation, likely because barley inclusion had already increased the E. coli O157:H7 populations through some complementary mechanism (Hallewell et al., 2012). Interestingly, researchers found that the numbers of E. coli O157:H7 were greater in fecal in vitro incubations that contained corn DG than with wheat DG (Yang et al., 2010).
Other scientists have examined the form of corn included in cattle rations can impact E. coli O157:H7. In feedlot cattle, steam-flaked grains increased E. coli O157 shedding in feces compared to diets composed of dry-rolled grains (Fox et al., 2007). This difference was theorized to be due to dry rolling allowing the passage of more starch to the hindgut where it was fermented to produce VFA thereby killing E. coli O157 (Fox et al., 2007). This theory is supported by the fact that post-ruminal starch infusion
these studies the calves that consistently shed the highest concentrations of E. coli O157:H7 were fed a high concentrate (grain) diet (Tkalcic et al., 2000). Ruminal fluid collected from steers fed a high-forage diet allowed E. coli O157:H7 to proliferate to higher populations in vitro than did ruminal fluid from highgrain fed steers (Tkalcic et al., 2000). This was possibly due to differences in VFA concentrations between the ruminal fluids. Other researchers found that feeding forage actually increased the shedding of E. coli O157:H7 in cattle (Van Baale et al., 2004). When cattle were fed forage E. coli O157:H7 was shed for 60 d compared to 16 d for cattle on a grain-based diet (Van Baale et al., 2004). Studies examining the effects of forage on survival of E. coli O157:H7 in manure found that low quality forages caused a faster rate of death of E. coli O157 populations (Franz et al., 2005), indicating a possible role of forage chemical or secondary plant components (such as tannins, see below) in fecal shedding (Min et al., 2007). Feces from cattle fed grain had higher VFA concentrations and lower pH which allowed E. coli O157:H7 populations to survive longer than feces from grass-fed cattle (Lowe et al., 2010). Other studies have found that feeding forage rich secondary compounds such as sainfoin, might
increased generic E. coli populations in the lower gut numerically (Van Kessel et al., 2002). However, to date studies have shown no effect on E. coli O157:H7 populations of increasing starch concentrations in the diet (Nagaraja, T. G., personal communication)
be a method to manipulate fecal populations of E. coli O157:H7 to a limited extent (Aboaba et al., 2006; Berard et al., 2009). Although E. coli O157:H7 populations are generally lower in cattle fed forage diets, it must be
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Forage feeding Escherichia coli can and does thrive in the lower gut of animals fed forage diets (Hussein et al., 2003a; Hussein et al., 2003b; Jacobson et al., 2002). Comparing grain-fed to forage-fed cattle indicates that more E. coli (including O157:H7) are present in the feces of cattle fed grain diets. The effects of high grain or high forage diets on the duration or shedding of fecal E. coli O157:H7 populations in experimentally inoculated calves have been examined. In
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emphasized that STEC are still isolated from cattle solely fed forage, so forage feeding should not be viewed as a magic bullet (Hussein et al., 2003b; Thran et al., 2001). Many outlets have claimed that grassfed cattle contain fewer pathogens than do cattle fed grain; however this has not been demonstrated scientifically. Researchers have found no difference in food safety parameters of beef from grass-fed cattle versus grain fed cattle (Zhang et al., 2010). Furthermore, research into organic versus conventional rearing systems have demonstrated no difference in the incidence of E. coli O157:H7 shedding (Jacob et al., 2008c; Reinstein et al., 2009).
Dietary shifts
not impact carcass characteristics; however, when cattle were fed hay during the final portion of the finishing period, they had lower dry matter intake and lost 2.2 lb/head/d (Stanton and Schutz, 2000). Hay feeding did not significantly impact carcass weight, dressing percentage, carcass grades, or quality parameters, but significantly reduced total coliform counts and (generic) E. coli counts (Stanton and Schutz, 2000), but the impact was not as large as that reported by Diez-Gonzalez et al. (1998). Cattle fed hay for a 48 h period immediately prior to transport to slaughter did not lose more weight during transport than fasted or pasture fed animals (Gregory et al., 2000). Cattle with a natural E. coli O157:H7 in-
A sudden shift from grain to hay appears to cause a severe, widespread disruption in the gut microbial flora population, much like an earthquake in a macrobiological environment (Fernando et al., 2010). Thus the effects of rapid dietary shifts on the microbial population in regards to E. coli O157:H7 populations have been examined. Early studies investigating (generic) E. coli and dietary effects indicated that a sudden decrease in hay intake by cattle increased fecal E. coli populations (Brownlie and Grau, 1967). Other studies using experimentally infected sheep found a sudden switch from an alfalfa pellet diet or a corn/alfalfa ration to a poor-quality forage diet increased E. coli O157:H7 shedding (Kudva et al., 1995; 1997). Cattle fed feedlot-type ration contained (generic) E. coli populations that were 1000-fold higher than cattle fed a 100% good-quality hay diet (Diez-Gonzalez et al., 1998). When these cattle were abruptly switched from a 90% grain finishing ration to a 100% hay diet, fecal E. coli populations declined 1000-fold within 5 d (Diez-Gonzalez et al., 1998). However, it is important to note that in this study no E. coli O157:H7 were detected. Based on these results
fection (53%) were divided into two groups and one was fed grain and the other abruptly switched to hay, 52% of the grain-fed controls remained E. coli O157:H7 positive, but only 18% of the hay-fed cattle continued to shed E. coli O157:H7; but this switch resulted in a BW decrease of 1.25 lb/hd/d compared to controls (Keen et al., 1999). Other researchers found that cattle fed a high-concentrate diet and switched to a diet containing 50/50% corn silage/alfalfa hay diet had lower E. coli counts (0.3 log10) after just 4 days (Jordan and McEwen, 1998). Cattle that were fed an 80% barley ration, fasted for 48 h and then subsequently switched to 100% alfalfa silage did not exhibit any change in E. coli O157:H7 shedding (Buchko et al., 2000b). However, when these same animals were again fasted for 48 h and re-fed alfalfa silage, the prevalence of E. coli O157:H7 shedding increased significantly (Buchko et al., 2000b). Researchers found that experimentally-infected cattle fed hay shed E. coli O157:H7 significantly longer than did grain-fed cattle (42 d vs. 4 d), but E. coli O157:H7 populations shed were similar between diets (Hovde et al., 1999). Cattle abruptly switched from a finishing diet that contained wet corn gluten feed to alfalfa hay for 5 d showed an increase in colonic pH and total E. coli populations decreased ap-
the authors suggested that feedlot cattle could be switched from high grain diets to hay for 5 days prior to slaughter to reduce E. coli contamination entering the abattoir (Diez-Gonzalez et al., 1998). Research indicated that a brief (5 d) period of hay-feeding did
proximately 10-fold (Scott et al., 2000). Conversely, it was found that when cattle were switched from forage-type diets to a high grain finishing ration, fecal and ruminal generic E. coli concentrations increased (Berry et al., 2006). In another
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study slightly outside of the “normal” dietary switch structure, switching cattle from pasture to hay for 48 h prior to slaughter significantly reduced the E. coli population throughout the gut (Gregory et al., 2000). Gregory et al., found that hay feeding increased intestinal Enterococci populations that are capable of inhibiting E. coli populations in a fashion similar to that of a competitive exclusion culture. However, the effects of high grain versus forage diets were not examined in this New Zealand-based study (Gregory et al., 2000). Based on their data, the authors concluded, “the most effective way of manipulating gastro-intestinal counts of E. coli was to feed hay” (Gregory et al., 2000).
demonstrated to significantly inhibit the growth of E. coli O157:H7 in vitro and generic E. coli populations in cattle (Berard et al., 2009; Cueva et al., 2010; Min et al., 2007; Wang et al., 2009). Other researchers have found that the phenolic acids that comprise lignin also demonstrated antimicrobial activity against E. coli O157:H7 in fecal slurries, and highly lignified forages showed a reduced period of E. coli O157:H7 shedding compared with cattle fed only corn silage (Wells et al., 2005). Phenolic compounds in cranberry extract and sorrel are also effective against E. coli O157:H7 growth in vitro (Caillet et al., 2012; Fullerton et al., 2011), also the anthocyanins/proanthocyanidins from lowbush blueberries demonstrated in vitro
Collectively, these results emphasize that while dietary manipulations such as shifting cattle from a high grain to forage ration could be a powerful method to reduce E. coli/STEC populations in cattle prior to harvest, the mechanism remains unknown and the effect is very inconsistent. It appears that a factor in this inconsistency involves forage quality and type, but this remains a hypothesis. It does appear that the presence of endproducts of fermentation (e.g., VFA) and some secondary plant compounds in forages play some role in pathogen population levels. While a dietary switch to forage in feedlots is not advocated due to feasibility, weight loss and other logistical issues, other high fiber feedstuffs (e.g., soy hulls, cottonseed meal) or feedstuffs rich in phenolics or essential oils (see below), may be a more feasible alternative strategy to decrease in E. coli O157:H7 populations.
potential to inhibit E. coli O157:H7 growth (Lacombe et al., 2012). Essential oils are most often associated with aromatic compounds in various plants used as spices or extracts (Barbosa et al., 2009). Many of these essential oils exhibit antimicrobial acitivity (Dusan et al., 2006; Fisher and Phillips, 2006; Kim et al., 1995; Pattnaik et al., 1996; Reichling et al., 2009; Turgis et al., 2009), often through the mode of action of dissolving bacterial membranes (Di Pasqua et al., 2007; Turgis et al., 2009). As a result, many plant products have been used for centuries for the preservation and extension of the shelf life of foods (Dabbah et al., 1970). Essential oils have been proposed as potential modifiers of the ruminal fermentation (Benchaar et al., 2008; Benchaar et al., 2007; Boadi et al., 2004; Patra and Saxena, 2009) and to reduce E. coli O157:H7 in the live animal via in vitro studies (Benchaar et al., 2008; Jacob et al., 2009). Some essential oils have been shown to penetrate biofilms and kill E. coli O157:H7 (Pérez-Conesa et al., 2011), which could potentially play a role in reducing colonization in the rumen and/or terminal rectum.
Tannins, phenolics, and essential oils Plants contain phenolic and polyphenolic compounds, such lignin and tannins, that can affect the microbial ecosystem of the gastrointestinal tract through antimicrobial action (Berard et al., 2009; Cowan, 1999; Hristov et al., 2001; Jacob et al., 2009; Patra and Saxena, 2009). It is theorized that some of these compounds may penetrate biofilms and have an anti-quoroum-sensing effect, which may play a role in STEC colonization (Edrington et al., 2009b; Kociolek, 2009; Sperandio, 2010). Tannins have been 50
Seaweed (Tasco) Brown seaweed (Tasco-14) is a feed additive that has been included in cattle diets to improve carcass quality characteristics and shelf life, increase antioxidants and to improve ruminal fermentation efficiency (Anderson et al., 2006; Braden et al., 2007;
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Leupp et al., 2005). In vitro studies have indicated that Tasco-14 can reduce populations of E. coli and Salmonella (Callaway, unpublished data), and more recent results have linked this antipathogen activity to presence of phlorotannins in the brown seaweed (Wang et al., 2009). The phlorotannin anti- E. coli activity was greater than that found in other studies with terrestrial tannin sources (Min et al., 2007; Wang et al., 2009). Studies in vivo found that Tasco-14 feeding reduced fecal and hide prevalence of E. coli O157 in cattle (Braden et al., 2004). Because Tasco-14 is currently available in the market place, this is a product that can be included in cattle rations; however the extent of anti-pathogen activity in vivo is still not clear, therefore the cost of addition must be weighed carefully by the producer.
Citrus products Orange peel and citrus pulp have excellent nutritional characteristics for cattle and have been included as low-cost ration ingredients in dairy and beef cattle rations for many years (Arthington et al., 2002). Citrus fruits contain a variety of compounds, including essential oils and phytophenols that exhibit antimicrobial activity against foodborne pathogens (Friedly et al., 2009; Mkaddem et al., 2009; Nannapaneni et al., 2008; Viuda-Martos et al., 2008). Other studies have found that limonids from grapefruit may play a role in inhibiting secretion and intercellular communication by E. coli O157:H7 (Vikram et al., 2010). Research has demonstrated that the addition of > 1% orange peel and pulp reduced populations of E. coli O157:H7 and Salmonella Typhimurium in mixed ruminal fluid fermentations in the laboratory (Callaway et al., 2008; Nannapaneni et al., 2008). Further studies have demonstrated that feeding orange peel and pulp reduced intestinal populations of diarrheagenic E. coli in weaned swine (Collier et al., 2010). In ruminants, researchers demonstrated that feeding of orange peel and citrus pellets (a 50/50 mixture) at levels up to 10% DM reduced artificially inoculated populations of E. coli O157:H7 and Salmonella Typhimurium in sheep (Callaway et al., 2011a; b).
When studies were performed using only dried pelleted orange peel, the reduction in pathogen populations disappeared (Farrow et al., 2012), likely due to the inactivation of essential oils (limonene and terpeneless fraction) during the pelleting process. Continuing studies have demonstrated that orange oils offer a potential method for reducing both STEC and Salmonella on beef carcasses as well (Pendleton et al., 2012; Pittman et al., 2011). To date, orange peel feeding has not been examined in large-scale feeding studies, but retains promise as a potential on farm strategy to reduce the burden of pathogens on the farm, reducing environmental contamination and re-infection.
Organic acids Organic acids have been used in animal nutrition to modify the ruminal fermentation by providing some members of the microbial ecosystem a competitive advantage, and by inhibiting other species (Grilli et al., 2010; Martin and Streeter, 1995; Nisbet and Martin, 1993; Piva et al., 2007). Some organic acids (such as lactate, acetate, propionate, malate) have been shown to have antimicrobial activity against E. coli O157:H7 (Harris et al., 2006; Sagong et al., 2011; Vandeplas et al., 2010; Wolin, 1969). These acids have been used on hide and carcass washes to reduce pathogen populations, but only recently has interest in using organic acids to reduce pathogens in live animals received interest (Callaway et al., 2010b; Nisbet et al., 2009). Preliminary results do show some success in inhibiting pathogens in the lower intestinal tract of animals (unpublished data), however, further research needs to be performed to be able to release the appropriate organic acid and concentration in the appropriate intestinal location to reduce populations of E. coli O157:H7 in cattle.
Ractopamine Ă&#x;-agonists, such as ractopamine, are used in cattle to improve animal performance and carcass leanness. In vitro, ractopamine showed no effect on growth parameters of E. coli O157:H7 (Edrington et
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al., 2006c); but when used in sheep, the fecal shedding and cecal populations of E. coli O157:H7 were increased (Edrington et al., 2006c). When feedlot cattle were fed ractopamine, the numbers of cattle shedding E. coli O157:H7 were decreased (Edrington et al., 2006b). In a follow-up study, researchers demonstrated a negligible effect of ß-agonist (ractopamine and zilpaterol) treatment on fecal shedding of E. coli O157:H7 in cattle (Edrington et al., 2009a; Paddock et al., 2011). Taken as a whole, these results indicate that the effects of ß-agonist feeding are minimal or non-existent. Interestingly however, in an in vitro swine model norepinephrine was shown to increase E. coli O157:H7 adherence (Green et al.,
a forage ration that included monensin shed E. coli O157:H7 for a shorter period of time than forage-fed cattle not supplemented with monensin, but monensin had no effect on shedding when cattle were fed a corn-based ration (Van Baale et al., 2004). In an in vitro study, it was found that monensin and the coapproved antibiotic tylosin (tylan) treatment reduced E. coli O157:H7 populations up to 2 log10 CFU/mL in ruminal fermentations from cows fed forage, but this did not extend to E. coli O157:H7 populations in ruminal fluid from cows fed corn (McAllister et al., 2006). These researchers later found that the inclusion of monensin and tylosin did not alter fecal shedding of experimentally-inoculated E. coli O157:H7
2004), though further research is obviously needed to determine if this applies to cattle colonization.
when included in barley (grain)-based diet fed to cattle (McAllister et al., 2006). These results suggest there may a potential interaction between diet and ionophore inclusion in the effects on E. coli O157:H7 populations. Further studies found that monensin decreased E. coli O157:H7 prevalence when fed at 44 mg/kg of feed, compared to the typical 33 mg/kg dosing (Paddock et al., 2011).
Ionophores Ionophores, such as monensin and lasalocid, are antimicrobial compounds included in most feedlot and dairy rations to inhibit gram-positive bacteria, thereby improving feed:gain ratios and production efficiency (Callaway et al., 2003). Because these feed additives affect the gram-positive portion of the microbial population, possibly giving gram-negative bacteria (such as E. coli) a competitive advantage, they have been investigated as to their role in the spread of E. coli O157:H7 in cattle. Because E. coli O157:H7 has a true gram-negative membrane physiology ionophores did not affect the growth of this pathogen in vitro when added at concentrations up to 3 fold higher than those normally found in the rumen (Bach et al., 2002b; Van Baale et al., 2004). Early studies demonstrated a marginal increase of STEC shedding by heifers fed ionophores (Herriott et al., 1998), but other studies found no effect (Dargatz et al., 1997). Further studies examining the effect of ionophoric feed additives (monensin, lasalocid, laidlomycin and bambermycin) on E. coli O157:H7 demonstrated no effect of these additives in vitro (Edrington et al., 2003b), or on fecal shedding or intestinal populations in experimentally-inoculated lambs in a short-term (12 d) trial (Edrington et al., 2003a). In an in vivo study using cattle, it was found that cattle fed 52
CONCLUSIONS While STEC of many serotypes can be viewed as a commensal organism in the gastrointestinal tract cattle, they represent a significant threat to human consumers and public health. Pre-harvest controls in cattle hold great potential to reduce STEC dissemination on farms, in the environment, and entering the food chain. However, none of the on farm management-based controls discussed herein will completely eliminate STEC from cattle and will certainly not eliminate the need for proper procedures in the processing plant. Instead the live-animal management controls must be installed in a complementary fashion to reduce pathogens in a multiple-hurdle approach (Nastasijevic, 2011) that complements the in-plant interventions as well, so that the reduction in pathogen entry to the food supply can be maximized.
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Wick, L. M., W. Qi, D. W. Lacher and T. S. Whittam. 2005. Evolution of genomic content in the stepwise emergence of Escherichia coli O157:H7. J. Bacteriol. 187:1783-1791. Williams, M. S., J. L. Withee, E. D. Ebel, N. E. Bauer,
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W. D. Schlosser, W. T. Disney, D. R. Smith and R. A. Moxley. 2010a. Determining relationships between the seasonal occurrence of Escherichia coli O157:H7 in live cattle, ground beef, and humans. Foodborne Path. Dis. 7:1247-1254. Williams, W. L., L. O. Tedeschi, P. J. Kononoff, T. R. Callaway, S. E. Dowd, K. Karges and M. L. Gibson. 2010b. Evaluation of in vitro gas production and rumen bacterial populations fermenting corn milling (co)products. J. Dairy Sci. 93:4735-4743. Winfield, M. D. and E. A. Groisman. 2003. Role of nonhost environments in the lifestyles of Salmonella and Escherichia coli. Appl. Environ. Microbiol. 69:3687-3694. Wolin, M. J. 1969. Volatile fatty acids and the inhibition of Escherichia coli growth by rumen fluid. Appl. Microbiol. 17:83-87. Yang, H. E., W. Z. Yang, J. J. McKinnon, T. W. Alexander, Y. L. Li and T. A. McAllister. 2010. Survival of Escherichia coli O157:H7 in ruminal or fecal contents incubated with corn or wheat dried distillers’ grains with solubles. Can. J. Microbiol. 56:890895. Zhang, J., S. K. Wall, L. Xu and P. D. Ebner. 2010. Contamination rates and antimicrobial resistance in bacteria isolated from “grass-fed” labeled beef products. Foodborne Path. Dis. 7:1331-1336. Zhang, Y., C. Laing, M. Steele, K. Ziebell, R. Johnson, A. K. Benson, E. Taboada and V. P. J. Gannon. 2007. Genome evolution in major Escherichia coli O157:H7 lineages. BMC Genomics. 8:121-137. Zhou, Z., X. Li, B. Liu, L. Beutin, J. Xu, Y. Ren, L. Feng, R. Lan, P. R. Reeves and L. Wang. 2010. Derivation of Escherichia coli O157:H7 from its O55:H7 precursor. PLoS ONE. 5:e8700-8714.
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Independent Poultry Processing in Georgia: Survey of Producers’ Perspective E. J. Van Loo1,2, W. Q. Alali3, S. Welander4, C. A. O’Bryan1, P. G. Crandall1, S. C. Ricke1 Department of Food Science and Center for Food Safety, University of Arkansas, Fayetteville, AR 2 Present address: Department of Agricultural Economics, Faculty of Bioscience Engineering, Ghent University, Ghent, Belgium 3 Center for Food Safety and Department of Food Science & Technology, University of Georgia, Griffin, GA 4 Georgia Organics, 200-A Ottley Drive, Atlanta, GA
1
ABSTRACT A survey was presented to Georgia independent poultry farmers to evaluate current processing options as well as desired future changes. A total of 82 Georgia farmers participated in the survey, 31 of whom were raising broilers at the time of the survey. Most of the farmers surveyed who were growing broilers at the time (81%) processed on-farm, but these were also the farmers who processed less than 1000 birds per year. The larger independent Georgia farmers processed off-farm in South Carolina and Kentucky, where there were processors that served small-scale farmers and provided USDA inspection. These out of state processing trips took place between 4 and 30 times per year for an average of 391 miles round trip. For farmers’ future needs, similar numbers of farmers wanted only on-farm or only off-farm drop off processing (22% and 25% respectively), but 40% of the farmers surveyed were open to more than one processing option. The farmers were also asked to evaluate the importance of several attributes of processing facilities, and they chose quality of service to be the most important processing facility attribute, followed by cost of processing, distance from the farm, and USDA inspection. Keywords: Poultry processing, independent growers, pastured poultry, mobile processing units, on-farm processing Agric. Food Anal. Bacteriol. 3: 70-77, 2013
INTRODUCTION The U.S. broiler industry produced more than 8.6 billion broilers in 2009, estimated at a retail equivalent of $45 billion (ERS, 2011). Georgia is the number one broiler production state producing 1.3 billion Correspondence: Steven C. Ricke, sricke@uark.edu Tel: +1-479-575-4678 Fax: +1-479-575-6936
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broilers in 2009, accounting for 15% of the total U.S. broiler production (ERS, 2011). Federal laws exempt farms processing fewer than 20,000 birds per calendar year from USDA bird-by-bird inspection (USDA, 2006). However, some states have laws that are stricter and prohibit this exemption, which is the case in Georgia. The small-scale poultry industry in Georgia is limited in growth due to challenges it faces imposed by the strict state rules. Without the exemp-
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tion, small farmers who process more than 1,000 birds/year have to meet the federal inspection requirements for commercial processors. Procedures and practices need to be well-documented and include documents such as Standard Operating Procedures and Hazard Analysis Critical Control Point (HACCP) plans. Complying with these regulations to meet federal inspection standards is costly but is manageable for large commercial processors. However, it is too costly for small farmers and the farmers may also need help in developing all the required documentation (Geering, 2011a,b). The poultry industry in the U.S. is strongly vertically integrated, leaving limited options for off-farm
MATERIALS AND METHODS
processing for small-scale poultry farms. At the time of conducting the survey in Georgia, the only available processing facilities were owned by the large poultry companies, and there were no independent federally inspected processing facilities. Thus, the Georgia farmers who processed more than 1,000 birds/year had to travel to other states such as South Carolina and Kentucky to process their birds. This results in additional travel and accommodation expenses, requires additional time and results in a tremendous reduction in profitability for the grower. This lengthy transportation also increases the environmental impact of the locally grown poultry products and may have negative effects on the animal welfare which are both important to many of the local poultry product consumers. The local Georgia poultry farmers’ community is in need of safe and legal processing options, both on- and off-farm, that provide both easy access to poultry processing for farms throughout the state and ensure a safe product for the Georgia consumers. The purpose of this survey was to evaluate processing options for the independent local poultry farms in Georgia for small-scale poultry processing that would (1) ensure safe products for Georgia consumers, (2) meet farmer needs for various safe pro-
the farm. Farmers were asked to rate importance of attributes on a Likert scale from 1 (not important) to 7 (very important). Table 1 contains the questions and possible answers. Frequency tables, mean values and standard deviations were determined using JMP (release 9.0.0: SAS Institute, Inc.).
cessing options, both on- and off-farm, (3) provide easily accessible poultry processing options, (4) lay the foundation for future growth of pastured poultry production in Georgia.
ducers. With the vertical integration of the industry, poultry processing is highly consolidated and not available to independent producers (Heffernan and Hendrickson, 2002). This lack of processing is a real barrier and greatly restricts the ability of pastured
Notice of the survey was posted in the Georgia Organics print newsletter, and in an electronic newsletter, as well as by a targeted email to a list of interested parties amassed based on connections Georgia Organics made at conferences and meetings. The link to the survey was also posted on Georgia Organics’ website. A total of 82 Georgia farmers took the survey between September of 2008 and July of 2010. The survey consisted of questions about (i) current processing methods; (ii) interest in other processing options; (iii) importance of different processing facilities attributes; (iv) demographics of
RESULTS AND DISCUSSION Current Processing Methods A total of 82 Georgia farmers participated in the survey, of which 31 farmers were raising broilers at the time of the survey totaling 37,642 birds/year. Most of the farmers who were currently growing broilers (28 of the 31) processed on-farm but grew only 43% of the total pastured birds produced by the respondents in Georgia each year (Figure 1, Figure 2). The majority (60%) of the 28 farmers that processed onfarm produced less than 500 birds/year. Pastured poultry is actually not a new concept; until the 1950’s, all poultry was raised outdoors (Sustainable Agriculture Network, 2006). The renewed interest in pastured poultry has served to highlight the lack of suitable processing available to small pro-
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Table 1. Farmers survey questions and possible answers
1. Indicate the number of broilers you are currently processing per year at the different processing sites listed. a) On-farm b) USDA-inspected facility in South Carolina c) USDA-inspected facility in Kentucky d) Other off-farm processing facility (please describe) e) Not currently processing (enter 0) 2. If you currently process birds off-farm, please answer the following questions: (Enter your response in numbers only, with no dollar signs. Decimal points are ok.) a) Roundtrip driving miles to the facility b) Number of trips made per year c) Average # of birds processed per visit c) Approximate processing cost per bird charged by facility 3. Please indicate the annual number of broilers that you would like to raise under the following processing scenarios. Please enter your response in numbers only. a) Processing on-farm b) Shared-use facility with USDA inspection c) Drop Off/Pick Up facility with USDA inspection d) Contract grower 4. Do you want to sell your poultry direct to consumers, under your own farm’s label? a) Yes, I want to direct market my own processed poultry b) No, I want to raise pastured poultry but have someone else sell it to the consumer c) I want to pursue both options d) Not sure e) I am not interested in raising pastured poultry at this time. 5. What is the maximum distance you would be willing to drive for processing? 6. Rate the importance of the following items in considering a Georgia based processing facility where 7 = Very important, and 1 = Not important at all. Distance from farm, Processing price, Quality of service, Certified organic, USDA Inspection, Marketing services 7. If you were paying a facility to process your chickens for you (Drop off/pick up arrangement), at what price per bird processed would this service be a) Absolutely too expensive to ever use? b) Getting expensive? c) Inexpensive? d) A great value for the money? e) So cheap I would not trust what was happening in there. 8. Are you interested in participating in a co-op of producers to explore: a) processing options? b) Other infrastructure issues? c) What other issues/comments do you have? 9. Where is your farm located? Please provide answers to at least two of the following options. State: County: City/Town:
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Figure 1. Percentages of current place of processing for 31 Georgia (GA) pastured poultry farmers
16%
3%
On-farm in GA Off-farm SC Off-farm KY
81%
Figure 2. Percentage of current poultry processing of 31 Georgia farmers by total broiler processed per year (total 37,642 broilers/year)
7% 43%
On-farm in GA Off-farm SC Off-farm KY 50%
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Figure 3. Type of processing preferred by Georgia pastured poultry farmers (n=82) when presented with multiple choice options: 1) on-farm, 2) shared-use/rental facility, 3) drop off/pick up, 4) contract grower facility. Farmers had the choice to select/be interested in some of the 4 or all 4 processing options.
60 All 4 methods Shared-use + drop-off + contract
50
On-farm + shared-use + drop-off On-farm + shared-use 40
Contract + drop-off
# Farmers
Shared + drop-off On-farm + drop-off
30
Only this method
20
10
0 On-farm Shared-use Drop-off
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Contract
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Table 2. Importance of processing facility attributes to Georgia pastured poultry farmers (average Likert scale value with 1 = Not important and 7 = Very important)
Mean Likert scale value
St. dev.
Quality of service
6.42
0.96
Processing price
6.09
1.18
Distance from farm
5.95
1.20
USDA inspection
4.76
2.06
Certified organic
3.64
2.04
Marketing services
2.92
1.80
Processing facilities’ attributes
poultry producers to sell and market their products; if a farmer wants to slaughter more than 1,000 birds a year, they must meet stringent inspection standards established by the U.S. Department of Agriculture for large-scale poultry processing operations. Farmers raising up to 1,000 birds per year may slaughter on their own farms and sell the whole birds on site directly to consumers, or sell frozen birds directly to consumers at farmers markets, provided they have a mobile vehicle license. These on farm processors generally consist of family members, are able to process about 50 to 100 birds per day, and the work is very labor intensive (Fanatico, 2003). Some of the Georgia farmers (19% of the 31 growing broilers at the time of the survey) processed offfarm in South Carolina and Kentucky, where there were processors that serve small-scale farmers and provide USDA inspection (Figure 1). These were the larger farms that processed more than 1,000 birds/ year each, and raised 57% of the total amount of birds raised each year (Figure 2). These farmers traveled between 4 and 30 times per year for an average of 391 miles round trip to processing facilities out of state. Not only do the long travel times and distances cost farmers’ time and money, it may also
of transport and found higher mortality for broilers transported for 9 h as compared to 4 h (0.3% v. 0.2%). These last findings, when taken together with those of Bayliss and Hinton (1990), indicate that it is the time of transport and stationary waiting time that affects mortality, rather than distance driven. More bruising has been seen on carcasses with increasing transport duration (Scholtyssek and Ehinger, 1977). Ehinger and Gschwindt (1981) found that meat pH measured post mortem decreased with duration of transport, and live weight has been found to decrease as much as 3% with transport duration of 4.5 h (Scholtyssek et al., 1977). Blood corticosterone is an indication of stress in broilers and has been found to be higher in broilers following transport for 4 h compared to 2 h duration (Freeman, 1984).
adversely affect the survival or quality of the birds themselves. Vecerek et al. (2006) found that mortality was higher in broilers transported up to 185 mi compared those transported up to 30 mi (0.9% v. 0.2%). Warriss et al. (1992) examined the effects of time
included: (1) shared-use facility where the farmers do the processing, and sell their birds under their own farm label, (2) drop off/pick up facility where the facility processes as a service to the farmer, and the farmer sells the birds under their own label and (3)
Interest in Potential Processing Options and Importance of Different Processing Facilities Attributes Interest in on-farm and three different off-farm processing options was evaluated with a multiple choice question which was answered by all 82 respondents (Figure 3). Off-farm processing options
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contract grower facility where the farmer sells the live birds to the facility, and the facility processes and sells the birds under the facilityâ&#x20AC;&#x2122;s label. A similar number of farmers wanted only on-farm processing (22%) or only off-farm drop off processing (24%). However, 40% of the farmers were more flexible and were open to more than one processing option. The off-farm drop-off scored the best with 49 farmers (60%) being interested. This was followed by on-farm processing (43 farmers or 52%), off-farm rental (27 farmers or 33%), and contract growers (11 farmers or 13%). Overall, off-farm processing (including drop-off, shared use and contract growers) was more popular than on-farm processing. Comparing
there a need to find a solution in order to keep small poultry production as a profitable business. Off-farm processing, in particular drop-off facilities might offer an efficient solution; however, determination of the location might be an issue as distance to the processing facility is an important factor for many farmers. Of course, these services need to be offered at an affordable processing price and the quality of the service is very important to the farmers. The second preferred processing option, on-farm processing, also offers some opportunities. One innovative solution that is already being used in several other states is mobile processing. These consist essentially of small trailers equipped with everything
the three off-farm processing options, drop-off was the preferred off-farm processing option. Availability of off-farm processing has potential to significantly increase the volume of pastured birds raised in Georgia. Based on the reported volumes that the farmers would like to produce, an off-farm drop-off facility has the potential to increase the total smallscale poultry production from 37,642 birds/year to 232,205 birds/year, a potential 6-fold increase. However, it may be difficult to find a location for one single processing facility to be conveniently located for all interested farmers. The farmers evaluated the importance of several attributes of processing facilities. The farmers reported quality of service to be the most important processing facility attribute (Table 2), followed by the processing price, the distance from the farm and USDA inspection. It was less important if the processing plant was certified organic and if the marketing services were provided by the processing plant. Small-scale farmers in Georgia were having difficulties catching up with demand for local or pasture raised poultry due to the processing regulations in Georgia. Georgia is not alone in the need to address these issues; many other states are innovating processing solutions to address the chronic need
needed to process poultry, and travel to the farm for processing, providing necessary infrastructure and less environmental impact than a fixed facility. Several mobile processing units have shown to be successful in other states (eXtension, 2011), such as Kentucky (Skelton, 2011). These mobile processing units have the advantages compared to a fixed processing unit including lower processing costs, reduced stress on animals, lower capital investment, and less resistance from municipalities and neighbors (Simon, 2008). With a rising consumer interest and awareness in local and sustainable foods, it will be increasingly important to make local food products more widely available in the U.S. food market. Different solutions exist to help the small local farmers with limited resources to gain profits and make the local poultry production as a viable industry.
for small-scale slaughter facilities in rural communities while protecting and preserving the health and safety of the food supply. The production of smallscale poultry producers is insufficient to justify long trips to processors and additional fees, and thus is 76
ACKNOWLEDGEMENTS This research was funded by SARE grant LS11-245 (WQA, PGC and SCR) and USDA-NIFSI grant #200851110-04339 (SCR and PGC) and a farm aid grant (SW and Georgia Organics, Atlanta, GA).
REFERENCES Bayliss, P.A., and M.H. Hinton. 1990. Transportation
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of broilers with special reference to mortality rates. Appl. Animal Behaviour Sci. 28:93–118. Economic Research Service. 2011. U.S. broiler industry: background statistics and information. Available at : http://www.ers.usda.gov/News/broilercoverage.htm Accessed 12 January 2012. Ehinger, F., and B. Gschwindt. 1981. The influence of different transport duration on meat quality and physiological traits in broilers of different origins. Archiv. Geflugelk. 45: 260–265. eXtension. 2011. Mobile slaughter/processing units currently in operation. Available at http://www. extension.org/pages/19781/mobile-slaughterprocessing-units-currently-in-operation Accessed 12
ing to Connecticut? Cutting-edge process could benefit state farmers and consumers. Available at http://www.workingtheland.com/feature-mobileslaughterhouse.htm Accessed 12 January 2012. Sustainable Agriculture Network. 2006. Profitable poultry: raising birds on pasture. Available at: http://www.sare.org/Learning-Center/Bulletins/ National-SARE-Bulletins/Profitable-Poultry Accessed 11 January 2011. USDA. 2006. Guidance for determining whether a poultry slaughter or processing operation is exempt from inspection requirements of the poultry products inspection act. Available at: http:// www.fsis.usda.gov/oppde/rdad/fsisnotices/poul-
January 2012. Freeman, B.M. 1984. Transportation of poultry. World’s Poult. Sci. J. 40: 19–30. Geering, D. 2011a. In Georgia, local birds may take scenic route to your plate. Atlanta Magazine. Available at: http://www.atlantamagazine.com/covereddish/localfoods/blogentry. aspx?BlogEntryID=10241894 Accessed 12 January 2012 Geering, D. 2011b. Small farmers go to great lengths to process birds. Atlanta Magazine. Available at: http://www.atlantamagazine.com/covereddish/ localfoods/blogentry.aspx?BlogEntryID=10244161 Accessed 12 January 2012. Heffernan, W.D., and M. K. Hendrickson. 2002. Multi-national concentrated food processing and marketing systems and the farm crisis. Presented at conference: The farm crisis: how the heck did we get here? American Association for the Advancement of Science Symposium: Science and Sustainability, Boston, Massachusetts, February 2002. Available at: http://www.foodcircles.missouri.edu/paper.pdf Accessed 11 January 2011. Scholtyssek, S., F. Ehinger, and F. Lohman. 1977. Influence of transport and fasting on the slaughter quality of broilers. Archiv. Geflugelk. 41:27–30.
try_slaughter_exemption_0406.pdf Accessed 12 January 2012. Vecerek, V., S. Grbalova, E. Voslarova, B. Janackova, M. Malena. 2006. Effects of travel distance and the season of the year on death rates of broilers transported to poultry processing plants. Poult. Sci. 85:1881–1884. Warriss, P.D., E.A. Bevis, S.N. Brown, and J.E. Edwards. 1992. Longer journeys to processing plants are associated with higher mortality in broilerchickens. Br. Poult. Sci. 33:201–206.
Skelton, S. 2011. Kentucky mobile poultry processing unit. Available at http://www.extension.org/ pages/Kentucky_Mobile_Poultry_Processing_Unit Accessed 18 January 2012. Simon, K. 2008. Is a mobile slaughterhouse comAgric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 3, Issue 1 - 2013
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PEER REVIEW PROCESS Authors will be requested to provide the names and complete addresses including emails of five (5) potential reviewers who have expertise in the research area and no conflict of interest with any of the authors. Except for manuscripts designated as Rapid Communication each reviewer has two (2) weeks to review the manuscript, and submit comments electronically to the editorial office. Authors have three (3) weeks to complete the revision, which shall be returned to the editorial office within six (6) weeks after which the authors risk having their manuscript removed from AFAB files if they fail to ask the editorial office for an extension by email. Deleted manuscripts will be reconsidered, but they will have to be processed as new manuscripts with an additional processing fee assessed upon submission. Once reviewed, the author will be notified of the outcome and advised accordingly. Editors handle all initial correspondence with authors during the review process. The editor-in chief will notify the author of the final decision to accept or reject. Rejected manuscripts can be resubmitted only with an invitation from the editor or editor-in chief. Revised versions of previously rejected manuscripts are treated as new submissions.
PRODUCTION OF PROOFS Accepted manuscripts are forwarded to the editorial office for technical editing and layout. The manuscript is then formatted, figures are reproduced, and author proofs are prepared as PDFs. Author proofs of all manuscripts will be provided to the correspond82
ing author. Author proofs should be read carefully and checked against the typed manuscript, because the responsibility for proofreading is with the author(s). Corrections must be returned by e-mail. Changes sent by e-mail to the technical editor must indicate page, column, and line numbers for each correction to be made on the proof. Corrections can also be marked using “track changes” in Microsoft Word or using e-annotation tools for electronic proof correction in Adobe Acrobat to indicate necessary changes. Author alterations to proofs exceeding 5% of the original proof content will be charged to the author. All correspondence of proofs must be agreed to by the editorial office and the author within 48 hours or proof will be published as is and AFAB will assume no responsibility for errors that result in the final publication.
PUBLICATION CHARGES AFAB has two publication charge options: conventional page charges and rapid communication. The current charge for conventional publication is $25 per printed page in the journal. There is no additional charge for the publication of pages containing color images, micrographs or pictures. For authors who wish to have their papers processed as a rapid communication, authors will pay the rapid communication fee when proofs are returned to the editorial office in addition to twice the conventional page charges. Charges for rapid communications are $1000 per manuscript for guaranteed peer review within one week and $100 per journal page.
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MANUSCRIPT CONTENT REQUIREMENTS Preparing the Manuscript File Manuscripts must be written in grammatically correct English. AFAB offers a fee based language service upon request (language@afabjournal.com). Manuscripts should be typed double-spaced, with lines and pages numbered consecutively. All documents must be submitted in Microsoft Word (.doc or .docx, PC or Mac). All special characters (e.g., Greek, math, symbols) should be inserted using the symbols palette available in this font. Tables and figures should be placed in separate sections at the end of the manuscript (not placed in the text). Failure to follow these instructions will cause delays of the processing and review of the manuscript.
Title Page At the very top of the title page, include a title of not more than 100 characters. Format the title with the first letter of each word capitalized. No abbreviations should be used. Under the title, the authors names are listed. Use the author’s initials for both first and middle names with a period (full-stop) between initials (e.g., W. A. Afab). Underneath the authors, a list affiliations must be listed. Please use numerical superscripts after the author’s names to designate affiliation. If an authors address has changed since the research was completed, this new information must be designated as “Current address:”. The corresponding author should be indicated with an asterisk e.g., * Corresponding author. The title page shall include the name and full address of the corresponding author. Telephone and e-mail address must also be provided for the corresponding author, and emailaddresses must be provided for all authors.
at the beginning of the manuscript. In vivo, in vitro and bacterial names must be italicized (obligatory). Authors must avoid single sentence paragraphs and merge such paragraphs appropriately. Authors must not begin sentences with “Figure or Table shows…” as these are inanimate objects and cannot “show” anything. When number are reported in text or in tables, always put a zero in front of decimal numbers: “0.10” instead of “.10”.
MANUSCRIPT SECTIONS Abstract The abstract provides an abridged version of the manuscript. Please submit your abstract on a separate page after the title page. The abstract should provide a justification of your work, objectives, methods, results, discussion and implications of study or review findings . Your abstract must consist of complete sentences without references to other work or footnotes and must not exceed 250 words. On the same page as your abstract, please provide at least ten (10) keywords to be used for linking and indexing. Ideally, these keywords should include significant words from the title.
Introduction The introduction should clearly present the foundation of the manuscript topic and what makes the research or the review unique. The introduction should validate why this topic is important based on previously published literature, and the relevance of the current research. Overall goals and project objectives must be clearly stated in the final sentence of the last paragraphs of the introduction.
Materials and Methods Editing Author-derived abbreviations should be defined at first use in the abstract and again in the body of the manuscript. If abbreviations are extensive authors may need to provide a list of abbreviations
Information on equipment and chemicals used must include the full company name, city, and state (country if outside the United States or Province if in Canada) [i.e., (Model 123, ACME Inc., Afab, AR)].
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Variability, Replication, and Statistical Analysis To properly assess biological systems independent replication of experiments and quantification of variation among replicates is required by AFAB. Reviewers and/or editors may request additional statistical analysis depending on the nature of the data and it will be the responsibility of the authors to respond appropriately. Statistical methods commonly used in the bacteriology do not need to be described in detail, but an adequate description and/or appropriate references should be provided. The statistical model and experimental unit must be designated when appropriate. The experimental unit is the smallest unit to which an individual treatment is imposed. For bacterial growth studies, the average of replicate tubes per single study per treatment is the experimental unit; therefore, individual studies must be replicated. Repeated time analyses of the same sample usually do not constitute independent experimental units. Measurements on the same experimental unit over time are also not independent and must not be considered as independent experimental units. For analysis of time effects, assess as a rate of change over time. Standard deviation refers to the variability in the biological response being measured and is presented as standard deviation or standard error according to the definitions described in statistical references or textbooks.
Results Results represent the presentation of data in words and all data should be described in same fashion. No discussion of literature is included in the results section.
Discussion The discussion section involves comparing the current data outcomes with previously published work in this area without repeating the text in the results section. Critical and in-depth dialogue is encouraged.
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Results and Discussion Results and discussion can be under combined or separate headings.
Conclusions State conclusions (not a summary) briefly in one paragraph.
Acknowledgments Acknowledgments of individuals should include institution, city, and state; city and country if not U.S.; and City or Province if in Canada. Copies being reviewed shall have authors’ institutions omitted to retain anonymity.
References a) Citing References In Text Authors of cited papers in the text are to be presented as follows: Adams and Harry (1992) or Smith and Jones (1990, 1992). If more than two authors of one article, the first author’s name is followed by the abbreviation et al. in italics. If the sentence structure requires that the authors’ names be included in parentheses, the proper format is (Adams and Harry, 1982; Harry, 1988a,b; Harry et al., 1993). Citations to a group of references should be listed first alphabetically then chronologically. Work that has not been submitted or accepted for publication shall be listed in the text as: “G.C. Jay (institution, city, and state, personal communication).” The author’s own unpublished work should be listed in the text as “(J. Adams, unpublished data).” Personal communications and unsubmitted unpublished data must not be included in the References section. Two or more publications by the same authors in the same year must be made distinct with lowercase letters after the year (2010a,b). Likewise when multiple author citations designated by et al. in the text have the same first author, then even if the other authors are different these references in the text and the references section must be identified by a letter. For example
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“(James et al., 2010a,b)” in text, refers to “James, Smith, and Elliot. 2010a” and “James, West, and Adams. 2010b” in the reference section. b) Citing References In Reference Section In the References section, references are listed in alphabetical order by authors’ last names, and then chronologically. List only those references cited in the text. Manuscripts submitted for publication, accepted for publication or in press can be given in the reference section followed by the designation: “(submitted)”, “(accepted)’, or “(In Press), respectively. If the DOI number of unpublished references is available, you must give the number. The year of publication follows the authors’ names. All authors’ names must be included in the citation in the Reference section. Journals must be abbreviated. First and last page numbers must be provided. Sample references are given below. Consult recent issues of AFAB for examples not included in the following section. Journal manuscript: Author(s). Year. Article title. Journal title [abbreviated]. Volume number:inclusive pages.
Book Chapter: Author(s) of the chapter. Year. Title of the chapter. In: author(s) or editor(s). Title of the book. Edition or volume, if relevant. Publisher name, Place of publication. Inclusive pages of chapter.
Examples: O’Bryan, C. A., P. G. Crandall, and C. Bruhn. 2010. Assessing consumer concerns and perceptions of food safety risks and practices: Methodologies and outcomes. In: S. C. Ricke and F. T. Jones. Eds. Perspectives on Food Safety Issues of Food Animal Derived Foods. Univ. Arkansas Press, Fayetteville, AR. p 273-288. Dissertation and thesis: Author. Date of degree. Title. Type of publication, such as Ph.D. Diss or M.S. thesis. Institution, Place of institution. Total number of pages.
Maciorowski, K. G. 2000. Rapid detection of Salmonella spp. and indicators of fecal contamination in animal feed. Ph.D. Diss. Texas A&M University, College Station, TX.
Examples: Chase, G., and L. Erlandsen. 1976. Evidence for a complex life cycle and endospore formation in the attached, filamentous, segmented bacterium from murine ileum. J. Bacteriol. 127:572-583.
Donalson, L. M. 2005. The in vivo and in vitro effect of a fructooligosacharide prebiotic combined with alfalfa molt diets on egg production and Salmonella in laying hens. M.S. thesis. Texas A&M University, College Station, TX.
Jiang, B., A.-M. Henstra, L. Paulo, M. Balk, W. van Doesburg, and A. J. M. Stams. 2009. A typical one-carbon metabolism of an acetogenic and hydrogenogenic Moorella thermioacetica strain. Arch. Microbiol. 191:123-131.
Van Loo, E. 2009. Consumer perception of ready-toeat deli foods and organic meat. M.S. thesis. University of Arkansas, Fayetteville, AR. 202 p.
Book: Author(s) [or editor(s)]. Year. Title. Edition or volume (if relevant). Publisher name, Place of publication. Number of pages.
Examples: Hungate, R. E. 1966. The rumen and its microbes Academic Press, Inc., New York, NY. 533 p.
Web sites, patents: Examples: Davis, C. 2010. Salmonella. Medicinenet.com. http://www.medicinenet.com/salmonella /article. htm. Accessed July, 2010. Afab, F. 2010, Development of a novel process. U.S. Patent #_____
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Abstracts and Symposia Proceedings: Fischer, J. R. 2007. Building a prosperous future in which agriculture uses and produces energy efficiently and effectively. NABC report 19, Agricultural Biofuels: Tech., Sustainability, and Profitability. p.27 Musgrove, M. T., and M. E. Berrang. 2008. Presence of aerobic microorganisms, Enterobacteriaceae and Salmonella in the shell egg processing environment. IAFP 95th Annual Meeting. p. 47 (Abstr. #T6-10) Vianna, M. E., H. P. Horz, and G. Conrads. 2006. Options and risks by using diagnostic gene chips. Program and abstracts book , The 8th Biennieal Congress of the Anaerobe Society of the Americas. p. 86 (Abstr.)
Data Presentation in Tables and Figures Figures and tables to be published in AFAB must be constructed in such a fashion that they are able to “stand alone” in the published manuscript. This
means that the reader should be able to look at the figure or table independently of the rest of the manuscript and be able to comprehend the experimental approach sufficiently to interpret the data. Consequently, all statistical analyses should be very carefully presented along with variation estimates and what constitutes an independent replication and the number of replicates used to calculate the averages presented in the table or figure. Each table and figure must be on a separate page in the submitted paper. In addition, you will need to submit all data for charts, tables and figures in native format when possible (e.g., Microsoft Excel, Powerpoint). Photographs should be submitted as high-resolution (600 dpi) .jpg or tif. files. All figures should be clearly presented with well defined axis and units of measurement. Symbols, lines, and bars must be made distinct as “stand alone” black and white presentations. Stippling, dashed lines etc. are encouraged for multiple comparison but shades of gray are discouraged. Color images, micrographs, pictures are recommended and there is no additional fee for their submission.
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