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Volume 5 Issue 1 2015

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EDITORIAL BOARD Sooyoun Ahn

Hae-Yeong Kim

University of Florida, USA

Kyung Hee University, South Korea

Walid Q. Alali

Woo-Kyun Kim

University of Georgia, USA

Kenneth M. Bischoff

M.B. Kirkham

NCAUR, USDA-ARS, USA

Kansas State University, USA

Debabrata Biswas

Todd Kostman

University of Maryland, USA

University of Wisconsin, Oshkosh, USA

Claudia S. Dunkley

Y. M. Kwon

University of Georgia, USA

University of Arkansas, USA

Michael Flythe

Maria Luz Sanz

USDA, Agricultural Research Service

MuriasInstituto de Quimica Organic General, Spain

Lawrence Goodridge

Byeng R. Min

McGill University, Canada

Tuskegee University in Tuskegee, AL

Leluo Guan

Melanie R. Mormile

University of Alberta, Canada

Missouri University of Science and Tech., USA

Joshua Gurtler

Rama Nannapaneni

ERRC, USDA-ARS, USA

Mississippi State University, USA

Yong D. Hang

Jack A. Neal, Jr.

Cornell University, USA

University of Houston, USA

Armitra Jackson-Davis

Benedict Okeke

Alabama A&M University, USA

Auburn University at Montgomery, USA

Divya Jaroni

John Patterson

Oklahoma State University, USA

Purdue University, USA

Weihong Jiang

Toni Poole

Shanghai 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 Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 5, Issue 1 - 2015

EDITORIAL STAFF EDITOR-IN-CHIEF Steven C. Ricke University of Arkansas, USA

EDITORS Todd R. Callaway FFSRU, USADA-ARS, USA Philip G. Crandall University of Arkansas, USA Janet Donaldson Mississippi State University, USA

MANAGING and LAYOUT EDITOR Ellen J. Van Loo Ghent, Belgium

TECHNICAL EDITOR Jessica C. Shabatura Fayetteville, USA

ONLINE EDITION EDITOR C.S. Shabatura Fayetteville, USA

Ok-Kyung Koo Korea Food Research Institute, South Korea

ABOUT THIS PUBLICATION Mailing Address: 2138 Revere Place . Fayetteville, AR . 72701 Agriculture, Food & Analytical Bacteriology (ISSN 2159-8967) is published quarterly. Instructions for Authors may be obtained at the back of this issue, or online via our website at www.afabjournal.com Manuscripts: All correspondence regarding pending manuscripts should be addressed Ellen Van Loo, Managing Editor, Agriculture, Food & Analytical Bacteriology: ellen@afabjournal.com Information for Potential Editors: If you are interested in becoming a part of our editorial board, please contact Editor-in-Chief, Steven Ricke, Agriculture, Food & Analytical Bacteriology: editor@afabjournal.com

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Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 5, Issue 1 - 2015

TABLE OF CONTENTS ARTICLES 6

Salmonella Transfer to the Lymph Nodes and Synovial Fluid of Experimentally Orally Inoculated Swin P.R. Broadway, J.A. Carroll, J.C. Brooks, J.R. Donaldson, N.C. Burdick Sanchez, T.B. Schmidt, T.R. Brown, and T.R. Callaway

Efficacy of Elimination of Listeria spp., Salmonella spp. and Pseudomonas spp. in Single and Mixed Species Biofilms by Combination of Hydrogen Peroxide Pre-treatment and Cleaning Process B. T. Q. Hoa, T. Sajjaanantakul, V. Kitpreechavanich, W. Mahakarnchanakul

Hops (Humulus lupulus) ß-Acid as an Inhibitor of Caprine Rumen Hyper-Ammonia-Producing Bacteria In Vitro M. D. Flythe, G. E. Aiken1, G. L. Gellin, J. L. Klotz, B. M. Goff, K. M. Andries

Introduction to Authors 41

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. 5, Issue 1 - 2015

Salmonella prevalence of lymph nodes and synovial fluid of orally inoculated swine P.R. Broadway1, J.A. Carroll2, J.C. Brooks1, J.R. Donaldson3, N.C. Burdick Sanchez2, T.B. Schmidt4, T.R. Brown1, and T.R. Callaway5 Department of Animal and Food Sciences, Texas Tech University, Lubbock, TX Livestock Issues Research Unit, Agricultural Research Service, USDA, Lubbock, TX 3 Department of Biological Sciences, Mississippi State University, Mississippi State, MS 4 Department of Animal Science, University of Nebraska Lincoln, Lincoln, NE 5 Food and Feed Safety Research Unit, Southern Plains Agricultural Research Center, Agricultural Research Service, USDA, College Station, TX 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 no approval of the product, or exclusion of others that may be suitable.” USDA is an equal opportunity provider and employer

ABSTRACT Salmonella is a foodborne pathogen that may be associated with the consumption of meat products. Failure of current interventions to control Salmonella in the food supply of the U.S. has led researchers to believe that atypical carcass reservoirs may be partially responsible for harboring this pathogen. In this two phase study, pigs (n = 36/Phase 1; n = 38/Phase 2) were experimentally infected orally with Salmonella Typhimurium to monitor the spread of the organism within the animal body. Fecal samples were collected 24, 48, and 72 h post-infection and tested for the presence of the Salmonella. After the pigs were euthanized, Ileocecal, subiliac, popliteal, and mandibular lymph nodes were collected, and synovial fluid was collected from the coxofemoral, shoulder, and stifle joints at the same post-infection timepoints to test for the experimentally inoculated bacteria. Fecal prevalence tended to be greater in Phase 1 (P = 0.06; 52.8 versus 31.6%). Ileocecal lymph node prevalence was 41.67% for Phase 1 and 37.00% for Phase 2. Both mandibular and subiliac lymph node prevalence was determined to be 2.78% in Phase 1; however, no Salmonella were detected in Phase 2. Examination of synovial fluid yielded a prevalence of 2.63% in all locations (from a single pig) in Phase 2 but was not different from Phase 1 (P = 0.30) in which no samples were positive for Salmonella. These results suggest that it is possible for orally contracted Salmonella to migrate to musculoskeletal lymph nodes. Contamination in these areas may lead to cross-contamination of meat products. Further research is needed to determine routes and migration patterns of Salmonella from the gastrointestinal tract to peripheral tissues to further elucidate how these infections impact food safety. Keywords: swine, Salmonella, lymph node, synovial Agric. Food Anal. Bacteriol. 5: 6-14, 2015

Correspondence: Jeff Carroll, Jeff.Carroll@ars.usda.gov Tel: +1 -806-746-5353 Ext. 120 Fax: +1-806-746-5028

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INTRODUCTION Salmonella was the most commonly reported foodborne bacterial infection in 2011 (16.42 cases per 100,000 people), thus failing to meet the objectives to reduce the incidence of foodborne Salmonella illness set forward by the 2010 national health objective (6.8 cases per 100,000 people; Centers for Disease Control and Prevention; CDC, 2012). The “Healthy People 2010” report, a publication of the United States Department of Health and Human Services, indicated a failure to mitigate Salmonella illness (Johnston, 2012). A broader understanding of etiological and ecological characteristics and strat-

In addition to lymph nodes, synovial fluid may also harbor Salmonella, thus serving as another potential source of contamination within food products. While investigations into Salmonella and other foodborne pathogens in pork synovial fluid are limited, an increased prevalence of Salmonella in human joints after trauma or invasive procedures has been reported, and supports the hypothesis that joint synovial fluids can harbor Salmonella (Fihman et al., 2007). Nairn (1973) associated Salmonella and other pathogenic bacteria with osteomyelitis and synovitis in commercial turkeys. Additionally, 55 to 93% of commercial swine suffer from hind foot lesions, abrasions, and/or infections (Gentry et al., 2002; Mouttotou et al., 1999).

egies to reduce and control the prevalence of this pathogen remains a priority in all meat producing species. Recent estimates suggest that among foodborne illness in the U.S., 9 to 15% of all Salmonella infections, and 7.5% of Salmonella enterica serotypes Enteritis and Typhimurium infections, are associated with the consumption of pork or pork products (Hald et al., 2004; Pires et al., 2010). The risk of Salmonella infection from pork consumption is often considered minimal when compared to Salmonella infections stemming from the consumption of other food products (especially poultry). However, the commonality in serotypes isolated from pigs and human infection (USDA, 2013) combined with Salmonella’s ubiquitous nature in swine production settings makes the pathogen an area of focus for the pork industry. With regard to Salmonella in the food supply, lymph nodes have recently become a harbor of interest. Lymph nodes in musculoskeletal tissues of beef carcasses are often included in retail cuts and/ or ground beef, and have been targeted as an atypical reservoir of Salmonella (Arthur et al., 2008; Gragg et al., 2013). Lymph nodes collected from the gastrointestinal tract and head of pigs at harvest have also been reported to harbor Salmonella (Vieira-

Thus, the prevalence of hind foot lesions/infections combined with the ubiquitous nature of Salmonella supports the theory of Salmonella manifestation in bone joints of infected and/or sick animals, making synovial fluid a possible contamination vector. There is limited information within the literature to elucidate the mechanisms of translocation of Salmonella from the gastrointestinal tract into the circulating lymph system and ultimate migration to musculoskeletal lymph nodes in apparently healthy animals. Thus, the overall objective of this study was to determine if pigs that were orally inoculated with Salmonella would harbor the pathogen in mesenteric and musculoskeletal lymph nodes, as well as synovial fluid.

Pinto et al., 2005); however, there has been minimal research conducted to evaluate the occurrence of Salmonella in peripheral lymph nodes of seemingly healthy swine that are more likely to be introduced into the food supply.

Pigs, diet, and experimental design This experiment was conducted in two phases in which Yorkshire/Duroc crossbred pigs (n = 36 for Phase 1; n = 38 for Phase 2; average 10 ± 1.4 kg BW) were purchased from a commercial swine producer

MATERIALS AND METHODS All procedures in this study were reviewed and approved by the USDA-ARS, Livestock Issues Research Unit’s Institutional Animal Care and Use Committee (IACUC protocol 2010-10-JAC8).

Animals

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and transported to the USDA Livestock Issues Research Unit’s Swine Facility in Lubbock, TX. Pigs were fed a non-medicated commercial diet composed of (dry matter basis): ground corn 56.2%, soybean meal 23.25%, rice bran 9%, fish meal 4%, soyhulls 3.3%, Pork Flex 110 2.75%, tallow 1%, L-threonine 0.2%, L-lysine 0.15%, and methionine 0.15%. The diet was formulated according to Nutrient Requirements of Swine recommendations and pigs were allowed ad libitum access to water. Pigs were individually penned and housed in an environmentally controlled facility with an average air temperature of 28.2 ± 0.4°C. Fecal samples were collected from each pig upon arrival and each subsequent day

growth in the anaerobic intestinal tract. This strain was made resistant to novobiocin and nalidixic acid (20 and 25 µg/mL, respectively) by repeated transfer and selection in the presence of sub-lethal concentrations of each antibiotic. This resistant phenotype was stable through multiple unselected transfers in batch culture and through repeated culture vessel turnovers in continuous culture (data not shown). Overnight cultures (1 L) contained populations of Salmonella Typhimurium that were determined to be 4 x 109 CFU/ml by serial dilution and plating.

(for 5 d) during the dietary/facility adaptation period to verify that no growth occurred on novobiocin (20 µg/ml) and nalidixic acid (25 µg/ml) supplemented Brilliant Green agar (BGANN). Prior to inoculation, no colonies grew on any of the BGANN plates. Fecal samples were also analyzed during the adaptation period by enrichment for the presence of bacteriophages that could lyse the Salmonella Typhimurium (Callaway et al., 2010), strain used in the present inoculation study. In phase one, pigs were supplemented via the diet with phosphate buffered saline (PBS) or PBS with Enterobacter cloacae. In phase 2, pigs were fed with and without the inclusion of yeast cell wall products. For both phases, at the end of the 5 d adaptation period, each pig was inoculated with Salmonella Typhimurium (2 x 1010 CFU/pig) via oral gavage (10 mL total volume per pig) at 0 h. The concentration of Salmonella Typhimurium was utilized to ensure relatively elevated concentrations of Salmonella in the gastrointestinal tract to further enhance the possible transfer of the pathogen into systemic lymph.

Pigs (n = 12/d) in Phase 1 of the study were humanely euthanized at 24, 48, and 72 h after inoculation with Salmonella. For Phase 2, all pigs were euthanized 72 h post-inoculation in an effort to increase the possibility of finding Salmonella in peripheral lymph nodes and joints based on results from Phase 1. Ileocecal lymph nodes were aseptically collected and enriched following maceration (Figure 1). Digesta and epithelial tissues from the terminal rectum were also aseptically collected upon necropsy.

Gastrointestinal sample collection

Lymph node collection Mandibular, subiliac, and popliteal lymph nodes were collected from each animal following collection of gastrointestinal contents. Each lymph node was collected aseptically from the right side of the carcass. Samples were subjected to a surface disinfectant dip with 70% ethanol and were subsequently macerated and placed in tetrathionate broth for enrichment. Isolation of the inoculated Salmonella strain was conducted as described below.

Bacterial cultures

Synovial Fluid Collection

Salmonella enterica serotype Typhimurium (ATCC

Synovial fluid was collected from each pig (n = 74)

BAA-186) from the USDA Food and Feed Safety Research Unit culture collection was repeatedly grown (4 passages) by 10% (vol/vol) transfer in anoxic (85% N2, 10% CO2, 5% H2 atmosphere) Tryptic soy broth (TSB) medium at 37ºC to adapt the culture for

at three anatomical locations (i.e., shoulder, coxofemoral, and stifle) on the right side of the carcass. These joints were selected because each of these anatomical locations represent an area of the carcass that may be included in a retail cut or a joint that may

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Figure 1. Graphical representation of lymph node and synovial sampling locations. a. Mandibular lymph node, b. Shoulder joint, c. Subiliac lymph node, d. Stifle Joint, e. Popliteal lymph node, f. Hip joint

be exposed during fabrication that could lead to possible cross contamination. Joints were exposed using leverage on one side of the joint along with a sterile scalpel cutting the skin on the opposite side of the joint. Both the skin of the animal and the instruments used were disinfected with 70% ethanol prior to incision. Expressed synovial fluid was collected with environmental sponges (EZ 10BPW; World Bioproducts, Woodinville, WA) that were placed in 10 mL buffered peptone water (BPW). Sponges were stomached at 230 RPM for 2 min. (Stomacher 400 Circulator, Seward, Davie, FL), and the enrichment was incubated at 37ºC prior to detection.

Salmonella Detection To qualitatively confirm the presence of inoculated Salmonella Typhimurium in lymph nodes, synovial fluid, rectal contents, and epithelial samples, macerated samples and sponges were incubated overnight in tetrathionate broth at 39ºC and a 200 µL aliquot was transferred to Rappaport-Vassiliadis R10 Broth which was subsequently incubated at 42ºC for 24 h. Following this secondary enrichment, samples

were streaked on BGANN plates. Plates that exhibited colonies after 24 h incubation were classified as positive for experimentally innoculated Salmonella Typhimurium. Unless otherwise noted, all media and agar were from Difco Laboratories (Sparks, MD). Reagents and antibiotics were obtained from Sigma Chemical Co., St. Louis, MO. Post-enrichment synovial samples were subjected to real-time PCR analysis (BAX ®; Dupont, Wilimington, DE; AOAC 100201) to confirm the presence of Salmonella.

Statistical Analysis The experimental unit in both phases was the individual pig. Pigs were randomly assigned to a harvest day for Phase 1. Data from pigs positive for Salmonella were analyzed using Pearson Exact Chi Square analysis of SAS (v. 9.3 SAS Inst. Inc., Cary, NC). Interactions between fecal and illeocecal lymph node prevalence were analyzed using binomial logistic regression in PROC GLIMMIX of SAS. For Phase 1, day was considered a random variable. Significance was determined at P < 0.05 for all data.

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Table 1. The prevalence of Salmonella detected in four different lymph nodes from pigs experimentally inoculated with Salmonella. % Positive for Salmonella

Feces

Illeocecal

Popliteal

Mandibular

Subiliac

Phase 11

52.7

41.6

0.0

2.7

Phase 22

31.5

37.0

0.0

P-value3

0.0

0.6

1.0

0.3

n = 36 pigs n = 38 pigs 3 SEM = 2.34 1 2

RESULTS AND DISCUSSION Animals Pigs did not demonstrate any visual signs or symptoms of disease during this short term infection study. However, quantified immunological markers were consistent with an infection. Additionally, a small, yet significant, febrile response was observed after the Salmonella challenge (data not shown).

Fecal Fecal prevalence of the experimentally inoculated Salmonella in Phase 1 was 52.8% (85%, 46%, and 30% for 24, 48, and 72 h post-inoculation, respectively) and 31.6% in Phase 2 of the study (Table 1). There was a tendency (P = 0.06) for Phase 1 fecal prevalence to be greater than Phase 2 across all collection timepoints. This tendency is understandable due to Phase 2 sample collection only occurring at 72 h post-infection. Also, there was no fecal x day interaction (P = 0.18). The presence of Salmonella in the feces of pigs plays a major role in the crosscontamination of pork carcasses and ultimately the food supply. While the pigs in this study were in a controlled environment, many factors can influence infection and fecal shedding of Salmonella such as transportation (Hurd et al., 2002), lairage (Hurd et al., 10

2001), and co-mingling with other animals and new environments (Hurd et al., 2001). Fecal prevalence in pigs raised in commercial swine production operations has been reported to be between 1 and 33% (Davies et al., 1998; Rodriguez et al., 2006; Barber, 2002; Foley et al., 2008). Gebreyes et al., (2004) reported that swine herds with a greater prevalence of fecal Salmonella had the greatest incidence of carcass contamination at harvest, further solidifying the correlation between fecal and carcass contamination. Furthermore, Ojha and Kostrzynska (2007) stated that pigs may shed 10 million cells/g in feces during a Salmonella infection. Salmonella shedding in feces may transfer to other pigs or lead to crosscontamination via lairage, feed, or water.

Salmonella in Lymph Nodes In Phase 1 of the study, a total of 41.8% of pigs (n = 15) experimentally inoculated were positive for Salmonella in ileocecal lymph nodes at necropsy. The pigs were euthanized at three timepoints (24, 48, and 72 h post-inoculation; n = 13, 13, 10, respectively) in Phase 1, and ileocecal lymph node prevalence of Salmonella at these time points was 46.15%, 46.15%, and 30.00%, respectively (Table 1). Salmonella prevalence in ileocecal lymph nodes was less than expected considering the concentration and dosage of innocula introduced into the gastrointestinal tract of the animal. Callaway and colleagues

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(2011) reported that in a trial in which pigs were inoculated with a similar dose of Salmonella, 77 to 83% of ileocecal lymph nodes were positive for Salmonella. In Phase 1 of the present study, all popliteal lymph nodes collected tested negative for Salmonella, but subiliac and mandibular lymph nodes were positive for the experimentally infected Salmonella strain at a prevalence of 2.78% (Table 1). Interestingly, all of these positive peripheral lymph nodes were isolated from pigs necropsied only at 48 h after inoculation. A total of 36.80% of ileocecal lymph nodes collected from pigs in Phase 2 (at 72 h post-inoculation) of the study were positive for the inoculated Salmonella strain; however, no peripheral lymph nodes col-

and 50% less than the prevalence of Salmonella in feces from the same animals. In the present study, there was no interaction between fecal and ileocecal lymph node prevalence (P = 0.15). These previously reported data as well as data from the current study help elucidate a possible disconnect between fecal shedding of Salmonella from pigs and the incidence of Salmonella harbored in the lymphatic system (Foley et al., 2008). Berends et al. (1996) stated that the gastrointestinal tract and lymph nodes may be major sources of Salmonella carcass contamination and subsequent transfer to human consumers. Positive correlations have been reported between Salmonella in feces and

lected from Phase 2 pigs harbored Salmonella (Table 1). These results are consistent with data reported by Gray et al., (1996) which indicated that lymph nodes near the ileocolic junction were the lymph nodes with the greatest Salmonella prevalence at harvest. While there were numerical differences, there were no statistical differences in lymph node prevalence of Salmonella between the two phases of the present study for ileocecal (P = 0.67), popliteal (P = 1.00), mandibular (P = 0.30), and subiliac (P = 0.30) lymph nodes. Our results associated with the presence of Salmonella in subiliac and mandibular lymph nodes are of great importance to food safety as these lymph nodes are anatomically located in areas of the pork carcass that are commonly included in trim and retail cuts. These lymph nodes may also be exposed and evaluated during post-mortem USDA inspection and ultimate fabrication of the carcass. Exposure of infected lymph tissues during fabrication could potentially lead to cross contamination of equipment and/ or other carcasses. The presence of Salmonella in the feces of live pigs has been previously examined in conjunction with subiliac lymph nodes; however, little association was present to suggest feces as a predictor of lymph node contamination (Wang et al., 2010). Studies conducted by Hurd et al. (2001, 2002)

on carcasses (Berends et al., 1997) and between Salmonella in the intestinal tract and carcasses (Swanburg et al., 1999). A study conducted by Vieira-Pinto et al, (2005) sampled carcasses that swabbed positive for Salmonella, and the researchers reported that 18.8% of ileocecal lymph nodes from those carcasses contained Salmonella. Additionally, pig mandibular lymph nodes and tonsils have also been reported to harbor Salmonella (12.9 and 9.9%, respectively; Vieira-Pinto et al. 2005). Based on data from the current study, we can infer that there is a relatively miniscule possibility that Salmonella (contracted orally) will be harbored in musculoskeletal lymph nodes at 72 h post-inoculation. While the Salmonella prevalence of these lymph nodes was very limited at 72 h postinfection, further investigation should be conducted to elucidate the timeframe of migration of oral Salmonella infections as well as other factors that may impact pathogen migration in the lymphatic system. While musculoskeletal lymph node prevalence was not great, a single contaminated lymph node could potentially contaminate thousands of pounds of product when comminuted together with other trim. Given that Salmonella can be internalized within the lymph nodes of pigs, the bacterial cells are not as susceptible to topical in-plant pathogen reduction

reported less incidence of Salmonella in feces from swine at the farm when compared to ileocecal lymph nodes positive for Salmonella at harvest. Wood et al. (1989) reported the prevalence of Salmonella infected ileocecal lymph nodes to be between 30

systems implemented by most packers and processors. For these reasons, some packers excise lymph nodes as a routine part of the fabrication process in an effort to reduce potential contamination of pork trim.

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Table 2. The prevalence of Salmonella detected in synovial fluid from three different joints from pigs experimentally inoculated with Salmonella. % Positive for Salmonella

Shoulder

Hip

Stifle

Phase 1

0.0

Phase 2

2.6

P-value

0.3

n = 36 pigs n = 38 pigs 3 SEM = 1.32 1

Synovial Synovial fluid prevalence was 0 and 2.63% for Phase 1 and 2 respectively. There was no statistical difference (P = 0.30; Table 2) between Salmonella positive synovial samples in Phase 1 and Phase 2 of the study. Of the synovial swabs collected (n = 222; 3/pig), only three swabs were positive for Salmonella, all of which were from the same pig. This particular pig was noted to have a large abscess on the abdomen at the time of harvest. While this phenomenon was only observed in one animal, we hypothesize that the immunocompromised state of the pig during the experimental infection may have played a role in the transmission of Salmonella into the synovia. This hypothesis is supported by the fact that no visible lesions were noted on this particular animal at any of the joint sampling locations. While little is known about Salmonella in the joints of swine, Varley and Wiseman (2001) suggest that immunosuppression due to porcine reproductive and respiratory syndrome (PRRS) may predispose the swine to synovial infections by Haemophilus parasuis. More research is needed to determine if immunocompromised pigs translocate infections from the gastrointestinal tract to other peripheral tissues. Similar to lymph nodes, Salmonella in synovial fluid is not susceptible to post-harvest topical pathogen reduction interventions applied to the carcass. The original hypothesis of this study stated that synovial fluid may harbor Salmonella and be a possible vec12

tor for potential cross-contamination when exposed during fabrication. However, based on the data from the current study, we can infer that the possibility of Salmonella cross-contamination via synovial fluid from pigs that orally acquire this pathogen is relatively low.

CONCLUSIONS Overall, this study determined that experimental oral inoculation of pigs with Salmonella may result in Salmonella penetrating the lymphatic system and reaching peripheral lymph nodes. While most of the infection was localized to the illeocecal lymph nodes and feces, the infection was also shown to reach peripheral lymph in some animals. While only being observed in one animal, we hypothesize that oral infection with Salmonella may be able to influence Salmonella prevalence in the joints of immunocompromised animals. Elucidating the transmission routes of Salmonella to peripheral tissues in the carcasses of pigs that enter the food chain is vital to the establishment of interventions and control points to prevent foodborne illness and cross contamination in the pork production process. More research needs to be conducted to determine how Salmonella infections with different routes of entry migrate through the lymphatic system and how these infections impact animal health and food safety.

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Miller. 2002. Alternative housing systems for pigs: influences on growth, composition, and pork quality. J. Anim. Sci. 80:1781-1790. Gragg, S. E., G.H. Loneragan, M.M. Brashears, T.M. Arthur, J.M. Bosilevac, N. Kalchayanand, R. Wang,

types using a Bayesian microbial subtyping approach for source attribution. Foodborne Pathog. Dis. 7:143-151. Rodriguez, A., P. Panlgoli, H.A. Richards, J.R. Mount, and F.A. Draughon. 2006. Prevalence of Salmo-

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nella in diverse environmental farm samples. J. Food Prot. 69:2576-2580. Swanburg, M., H.A.P. Urlings, D.A. Keuzenkamp, and J.M.A. Snijders. 1999. Tonsils of slaughtered pigs as a marker sample for Salmonella positive pork. In: Bahnson, P.B. (ed.). Proceedings of the 3rd International Symposium on the Epidemiology and Control of Salmonella in Pork, pp. 264-265. Washington DC, USA. Varley, M.A. and J. Wiseman. 2001. The weaner pig: nutrition and management. P. 243. CABI Publishing, New York, NY. Vieira-Pinto, M., P. Temudo, and C. Martins. 2005. Occurrence of Salmonella in the ileum, ileocolic lymph nodes, tonsils, mandibular lymph nodes and carcasses of pigs slaughtered for consumption. J. Vet. Med. 52:476-481. Vieira-Pinto, M., P. Temudo, and C. Martins. 2005. Occurrence of Salmonella in the ileum, ileocolic lymph nodes, tonsils, mandibular lymph nodes and carcasses of pigs slaughtered for consumption. J. Vet. Med. 52:476-481. Wang, B., I. V. Wesley, J. D. McKean, and A. M. O’connor. 2010. Sub-iliac lymph nodes at slaughter lack ability to predict Salmonella enterica for swine farms. Foodborne Path. Dis. 7:795-800. Wood, R.L., A. Pospischil, and R. Rose. 1989. Distribution of persistent Salmonella typhimurium infection in internal organs of swine. Am. J. Vet. Res. 50:1015-1021.

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Efficacy of Sanitizers on Listeria, Salmonella, and Pseudomonas Single and Mixed Biofilms in a Seafood Processing Environment B. T. Q. Hoa1, W. Mahakarnchanakul1, T. Sajjaanantakul1, V. Kitpreechavanich2 Department of Food Science and Technology, Faculty of Agro-Industry, Kasetsart University, Bangkok 10900, Thailand 2 Department of Microbiology, Faculty of Science, Kasetsart University, Bangkok 10900, Thailand

ABSTRACT Control and elimination of biofilm formation in the food processing environment is vital for food safety. This study was designed to investigate the efficacy of hydrogen peroxide (H2O2) pre-treatment combined with the regular daily cleaning procedure used in a shrimp plant to control biofilm formation. Single and mixed species biofilms of Listeria, Salmonella and Pseudomonas were used as the test model. Single biofilms on stainless steel (SS) coupons were formed under nutrient stress and harvested at 3 and 7 days to assess four cleaning procedures. Using 2% alkaline detergent for 10 minutes followed by two types of quaternary ammonium compounds (QACs) - based sanitizers completely eliminated single biofilms of Listeria and Salmonella. When alkaline was replaced to acidic type, microbial reduction achieved 5 log colony forming units (CFU)/cm2 (or more). For the mixed species biofilm study, biofilms were formed under the simulated seafood processing plant conditions for 7 days, on SS, Teflon and rubber coupons. After pre-treated mixed species biofilms with H2O2 at 1% and 2% for 5 and 10 minutes followed by the regular cleaning procedure, 2% of H2O2 for 10 minutes reduced microorganisms by 6 log CFU/cm2. Mixed biofilm on SS was easier to remove compared to the other surfaces. Overall these results suggest that the application of H2O2 prior to the regular cleaning process in food processing facilities may help to reduce and control biofilm formation, particularly biofilms composed of mixed species. Keywords: Biofilm, Listeria, Salmonella, Pseudomonas, hydrogen peroxide, cleaning, sanitizer, stainless steel, Teflon, rubber Agric. Food Anal. Bacteriol. 5: 15-28, 2015

INTRODUCTION Microorganism contamination on food products has been increasing in the food processing environment even though routine cleaning and sanitizing using various detergents and chemicals is employed Correspondence: Warapa Mahakarnchanakul,fagiwpm@ku.ac.th Tel: +66-2562-5036 Fax: +66 2 562 5021

(Bridier et al., 2015; Food & Water-Watch, 2007; Norhana et al., 2010). Contaminated food causes harm to consumer’s heath and economic issues as well as losses related to the brand name of the respective food producer (Bohme et al., 2013). Biofilm formation is a serious concern due to inappropriate cleaning process (Brooks and Flint, 2008; Chmielewski and Frank, 2003; Giaouris et al., 2014; Gibson et al., 1999). Moreover, not only spoilage bacteria

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(e.g., Pseudomonas, Klebsiella) but also most of the foodborne pathogens (e.g., Listeria monocytogenes, Salmonella enterica serovar Typhimurium, E. coli H7: O157) are able to adhere to biofilms on most materials and under almost all of the environmental conditions in food production plants (Beauchamp et al., 2012; Bridier et al., 2015; Giaouris et al., 2012; Joseph et al., 2001; Marchand et al., 2012; Ryder et al., 2007). In addition, viable microorganisms in biofilm are tolerant to sanitizers because some microorganisms have specific mechanisms to resist sanitizing agents. A few examples include Pseudomonas spp., Listeria spp., Salmonella spp., Escherichia coli, etc. (Bridier et al., 2015; Chmielewski and Frank, 2003; Dourou et

et al., 2005). Therefore, the objectives of this study were focused on the assessment of cleaning process at food processing plants through single mature biofilm formation under nutrient stress followed by a proposed modified cleaning method that combines EPS degradation by H2O2 and a cleaning process to assess the effectiveness of removing a mixed species biofilms under simulated food processing ecosystem conditions.

MATERIALS AND METHODS Sanitizers and detergents

al., 2011; Duong, 2012; Ibusquiza et al, 2011; Myszka and Czaczyk, 2011). Furthermore, extracellular polymeric substances (EPS), which are produced by microorganisms to protect themselves against other species or under stress growth conditions, act as glue which help microorganisms to effectively adhere on the surfaces of equipment, leading to the failure of the removal of the biofilm during the cleaning process (Bridier et al., 2015; Giaouris et al., 2012; 2013). Therefore, finding the appropriate method for removing biofilm is the interest of both food scientists and food manufacturers. Numerous studies on evaluation of the effectiveness of sanitizer on single biofilms of both pathogenic and spoilage bacteria have been reported (Beauchamp et al., 2012; Belessi et al., 2011; Choi et al., 2012; Elmali et al., 2012; Ölmez and Temur, 2010; Oz et al., 2012). However, research on the efficacy of sanitizers to remove the mixed species biofilms at a mature stage remain limited, especially mixed species biofilm formation under simulated food processing ecosystem conditions. In addition, most research only focuses on the effect of sanitizers to eliminate organisms in biofilms (Aase et al., 2000; Fatemi and Frank, 1999; Joseph et al., 2001; Norwood and Gilmour, 2000). There is still a lack of studies that fully applies the simulation of likely cleaning process

Three types of bacteria (Listeria monocytogenes 101; Salmonella enterica serovar Aberdeen and Pseudomonas aeruginosa) were obtained from the Department of Food Science and Technology, Agro-Industry Faculty, Kasetsart University, Thailand. These bacteria (Listeria monocytogenes 101, Salmonella enterica serovar Aberdeen and Pseudomonas aeruginosa) were commonly present on foodborne pathogenic and spoilage bacteria, especially in seafood processing (Gram and Huss, 1996; Gram et al., 1987; Koonse et al., 2005; Norhana et al., 2010). All

at the seafood processing facility, using both cleaning with detergent and disinfectant with sanitizers. Furthermore, employing a method to degrade the EPS in a biofilm before cleaning procedure has not been fully investigated (Giaouris et al., 2014; Xavier

bacterial species were cultured (37°C, 24 h) and subcultured (37°C, 18 h) individually in 10 mL tryptic soy broth (TSB; Difco). Cells of the individual cultures were then harvested by centrifugation (10000 ×g at 4 °C for 10 min) followed by washing twice in 1 mL

Sanitizers and detergents were supplied by a local seafood processing facility in Thailand where these chemicals are regularly used in cleaning process. These included an alkaline foam cleaner (Superp foam), acidic foam cleaner (Dilac Z – descaler), QACs (Spectrum- broad spectrum liquid disinfectant), PAA, oxidizing disinfectant- peracetic acid (Zal Perax II) and QACs- based (Quatdet clear -broad disinfectant, fogging). These chemicals were products of Diversey Hygiene (Thailand) Co., Ltd. Hydrogen peroxide (grade AR) was a product of QReC, New Zealand.

Bacteria cultures and stock preparation

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phosphate buffered saline (PBS, pH 7.4) (Bae et al., 2012; Stewart and Costerton, 2001). Washed cell pellets of each species were resuspended in 1 mL TSB. To prepare the single species stock, the resuspension of each species was mixed with sterilized glycerol 36% by a ratio of 1: 1 (v/v). Finally, stock cultures were stored in a refrigerator at -20°C. Similar with the method to apply for preparing single species stock, the mixed species stock, which was used for the mixed species experiment, was prepared with 80% of volume cell suspension of Pseudomonas spp., 10% of volume of Listeria spp., and 10% of volume of Salmonella spp. Following this, the mixed species were added to glycerol and kept in a

Forty eight coupons were set up under the same conditions. First, coupons were soaked overnight in commercial detergent solution, degreased with 70% ethanol, thoroughly rinsed with tap water and distilled water (Hoa et al., 2015), and coupons were subsequently vertically placed sequentially at coupon gaps on a frame of the biofilm reactor (bioreactor). These CDC bioreactors were autoclaved at 121° C for 15 min prior to use (Pan, 2005; Stewart and Costerton, 2001). Next, four liters of broth (1% TSB, 16°C) and 1 mL of mixed species stock were added into the bioreactor. The bacterial population in the bioreactor was approximately 4.4 × 106 CFU/mL. Following this, the bioreactor was placed in an incubator for 7 days,

refrigerator at -20°C.

at 16°C under simulated food processing ecosystem with daily released and refreshed 4 liters of broth. Bacteria in the biofilm were subjected to broth for 8 hours/day, followed by starvation for nearly 16 hours /day. Finally, mixed species biofilms were harvested for conducting the experiment.

Biofilm formation Single species biofilm formation Stocks of three species Listeria spp., Salmonella spp., and Pseudomonas spp. were individually cultured (37°C, 24 h) and sub-cultured (37°C, 18 h) again to prepare single species biofilms for this experiment. Sterilized SS coupons (304, finish # 2B) with dimensions of 5 × 2 × 0.08 cm were transferred to 50 mL conical centrifuge tubes containing 40 mL of TSB (1% or 10%; Difco); which were inoculated with a 0.1mL suspension of sub-cultured bacterial cells; the bacterial population in each sample was approximately 2 × 107 CFU/mL. Following this, samples were incubated at 16°C for 3 days or 7 days. At sampling time (3 days or 7 days) single species biofilms were collected to conduct the experiments.

Cleaning process and modified cleaning process by pretreatment with hydrogen peroxide

Mixed species biofilm formation under a simulated food processing ecosystem (SFP) The SS coupons (304, finish # 2B); teflon® coupons and rubber coupons representing common materials that have been employed in food processing systems were used in this study. A modified CDC biofilm

Cleaning process The cleaning process used at the seafood company was simulated for this experiment. The regime for the cleaning process consisted of rinsing, cleaning with detergent, rinsing, disinfecting, rinsing, redisinfecting and rinsing. The four cleaning processes were respectively, (Fig. 1), D1S1- regular daily cleaning; D1S2- weekly cleaning; D2S1 and D2S2 - bimonthly cleaning. To perform the cleaning process, biofilm coupons were placed in a cleaning holding coupon frame which had been designed for assessing cleaning performance. The cleaning procedure was conducted as indicated by the flow chart shown in Fig. 1. The spraying method was applied for all steps of each treatment. Distilled water, detergents,

reactor (Centers for Disease Control, CDC; BioSurface Technologies Corp., Bozeman, MT, USA) (modified from Hadi et al., 2010) was used for conducting this experiment. It included two main components, namely, a bioreactor and a frame holding coupons.

sanitizers and hydrogen peroxide were contained in the same type of bottles (distilled water bottle 16 oz # 500 mL). Spraying distance was approximately 10 cm from the top of bottle to coupons. Spraying time was approximately 30 seconds per treatment (4 cou-

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Figure 1. Flow chart of four cleaning processes (D1S1) daily; (D1S2) weekly; (D2S1 & D2S2) bimonthly

pons and both sides of coupons). Spraying volume was approximate 60 mL per treatment. At the end of each step, samples were covered and placed in a laminar flow biological safety cabinet for specified exposure times. The concentration of detergents, concentration of sanitizers and exposure time were determined based on the flow chart of each cleaning process. Detergents and sanitizers were diluted and placed in the refrigerator for an hour prior to the cleaning process. After the cleaning process, biofilm coupons were collected in duplicate and placed into sterilized petri dishes for evaluating the efficacy of each cleaning process.

was performed with two levels of concentrations (1% and 2%) and two levels of exposure time (5 and 10 minutes). Following this, biofilm coupon continuous performances were evaluated over a regular daily cleaning process (D1S1). For D1S1 samples (no pretreatment with H2O2), coupons were rinsed with 10 mL distilled water, followed by continuous performance evaluations over a regular daily cleaning process (D1S1).

Determination of number of surviving bacteria Surviving bacteria on inoculated coupons with

Modified cleaning process by pre-treatment with H2O2 Similarly, a modified cleaning process was applied the pre-treatment with H2O2 before cleaning with a detergent-based stage. The pre-treatment with H2O2 18

single species or mixed species bacteria were detached carefully using two cotton swabs. For control samples (Biofilm, no cleaning process), coupons were rinsed twice with 10 mL distilled water in order to remove the loosely attached cells. Following this,

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all samples were swabbed thoroughly and the swab heads were broken off into a glass tube containing 10mL sterilized saline peptone water (SPW) (0.85% of salt and 0.1% of bacteriological peptone, Difco; Bagge-Ravn et al., 2003; Gibson et al., 1999). Next, suspensions were left for 30 minutes at 16°C to recover cells after treatment with the sanitizing agent. The bacteria on the swabs were re-suspended by vortexing for 1 min at high speed (Vortex Genie 2 G – 560E, speed 8). The re-suspension fluid was serially diluted in SPW and spread in duplicate on Tryptic soy agar (TSA) (Difco). The TSA plates were incubated at 37°C for 1 day and bacterial viability was quantified (Bae et al., 2012; Nguyen and Yuk, 2013).

siderable ability to form single and mixed species biofilms (Bridier et al., 2015; Dourou et al., 2011; Giaouris et al., 2013; Slama et al., 2012). However, the ability to eliminate bacteria in biofilms depends on the characteristics of the biofilms and the cleaning process. In this experiment, biofilms which were formed on SS with two conditions of nutrient stress (TSB 10% or TSB 1%) and two different ages of biofilms (3 days or 7 days) and three species bacteria at 16 °C. These results are demonstrated in Fig. 2. Four cleaning procedures which have been used in the seafood plant were applied to examine the efficacy of each cleaning process. Populations of Listeria, Pseudomonas and Salmo-

Effect of growth conditions on biofilm population Listeria spp., Salmonella spp., and Pseudomonas spp. have been demonstrated to be significant hazards in food production environments, especially in

nella biofilms are presented as log CFU/cm2 values. Figure 2 presents the population of single species biofilms of L. monocytogenes, Salmonella spp., and Pseudomonas spp. ranging from 3.5 log CFU/cm2 to 7.7 log CFU/cm2. In general, it was observed that the population of cells in biofilm depended on the type of cultures and nutrient levels (TSB 10% and TSB 1%). There was no significant difference in the population of cells for 3 days biofilms or 7 days biofilms. Similar results were obtained from a study of Giaouris et al. (2013) for Pseudomonas putida at 18°C. Their study showed that during a 10 day sampling interval, Pseudomonas putida generated two biofilm-formed cycles, one reached on day 4 and another at day 8. The cell population was approximately 7 log CFU/cm2. The results of this experiment indicated that the population of Pseudomonas cells was significantly higher than the population of Salmonella cells, but there was no significant difference with Listeria population in TSB 10%. This result may be explained due to the temperature of this experiment. In this experiment, the temperature of biofilm formation was set up at 16°C to simulate the working temperature of the seafood processing plant. Therefore, this temperature may not have influenced the growth of Listeria or Pseudomonas since they are

seafood processing ecosystems (Ababouch et al., 2005; Bridier et al., 2015; Giaouris et al., 2014; Van Houdt and Michiels, 2010). In addition, L. monocytogenes, Salmonella spp. and Pseudomonas spp. have been widely studied and shown to have con-

psychrotrophic microorganisms, whereas Salmonella is a mesophilic bacteria. Moreover, due to the presence of a complex enzymatic system, Pseudomonas spp. can metabolize various materials around them to serve as their nutrient sources (Franzetti and Scar-

Data analysis Results of experiments were converted into log values and averages were reported. Mean values and standard deviations were determined for results of two biological independent tests in duplicates. To calculate means and standard deviations, a value of 0.69 was assigned to sample outcomes that were below the lower limit of detection (<10 CFU/ mL) of the spread method. This value was equal to 100 CFU/1 coupon (# 20 cm2), or 5 CFU/cm2. SPSS version 18 was used for statistical analyses. Statistical significance was set at P-value less than 0.05.

RESULTS AND DISCUSSION Assessment of efficacy of four cleaning processes by removing the differences of mature single species biofilms

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Figure 2. Population of viable cells in single biofilms of Listeria, Pseudomonas and Salmonella for 3 days or 7 days in TSB 1% and 10%, at 16°C. Results were expressed as mean values ± standard deviation from two independent tests in duplicate. Among different nutrient conditions, incubation times and type of cultures (a, b, c); the mean values with the same letters are not significantly different (p>0.05). 9.0

8.0

7.0

abc bc

Log (CFU/cm2)

6.0

bc cd

bcd

5.0 4.0

bcd

abc

3.0 2.0 1.0 0.0

Listeria TSB 1% - 3 days

Pseudomonas TSB 10% - 3 days

pellini, 2007). Hence, under nutrient stress in TSB 1%, the population of Pseudomonas biofilm reached approximately 5.5 log CFU/cm2 whereas the Listeria biofilms reached only 3.5 and 4.5 log CFU/cm2 for 3 days and 7 days biofilms, respectively.

TSB 1% - 7 days

Salmonella TSB 10% - 7 days

Efficacy of four cleaning processes on different single biofilms We applied cleaning procedures (Fig. 1), that were used in the seafood factory in daily, weekly and bimonthly. This procedure included: 1. rinsing with fresh water, 2. cleaning with 2% alkaline or acidic detergent for 10 min, 3. rinsing with fresh water to remove detergent, 4. sanitizing with QACs 1 or PAA

Among the four cleaning processes, treatment D1S1 was the least effective and treatment D2S2 was the most effectiveness for eliminating bacteria in biofilms (Table 1 and Table 2). In terms of efficacy to remove biofilm of various single species, the results show that the Pseudomonas biofilms had the highest ability to survive after the cleaning process followed by Salmonella and Listeria. All the cleaning processes were effective at removing biofilms of Salmonella and Listeria. According to Somers and Wong (2004) a combination of cleaning and sanitizing was more effective than sanitizer alone in terms of eliminating Listeria monocytogenes biofilms. Moreover, QACs, which was used in four cleaning processes as sani-

0.2% for 10 min, 5. repeating the rinsing process with fresh water to remove the sanitizers, 6. re-sanitizing by fogging with Quatdet clear - a QACs 2 based 1% for 10 min. Finally, equipment was rinsed with fresh water again and allowed to dry before production.

tizer and re-sanitizer, was more effective against Salmonella spp. (Gram–negative) (Sinde and Carballo, 2000) and Listeria spp. (Gram-positive) (Buffet-Bataillon et al., 2012); even though they differ in their cell wall characterstics. Therefore the Pseudomonas

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Table 1. Efficacy of 4 cleaning processes on different single biofilms of Pseudomonas, Listeria, Salmonella for 3 and 7 days in TSB 1% and TSB 10%

Organism

Time TSB (days) (%)

log (CFU/cm2) Treatment Biofilm

D1S1

D1S2

D2S1

D2S2

Pseudomonas

5.50 ± 0.11

2.63 ± 0.01ab

0.79 ± 1.11ce

Listeria

3.51 ± 0.04

0.70 ± 0.00c

Salmonella

5.49 ± 0.06

NDd

Pseudomonas

7.65 ± 0.54

4.13 ± 1.19a

2.87± 0.66ab

1.55 ± 1.20bc

Listeria

6.96 ± 0.38

0.70 ± 0.00c

Salmonella

4.33 ± 1.38

Pseudomonas

5.36 ± 0.19

1.74 ± 0.13bc

0.59 ± 0.83ce

0.35 ± 0.49ce

Listeria

4.55 ± 0.23

Salmonella

4.75 ± 1.90

Pseudomonas

5.84 ± 0.73

2.71 ± 0.75ab

0.50 ± 0.71ce

1.61 ± 0.81bc

0.70 ± 0.00c

Listeria

5.88 ± 0.98

Salmonella

5.45 ± 0.64

a,b,c

significant decrease within 4 cleaning processes (D1S1, D1S2, D2S1 and D2S2) in both rows and columns

ND, not detectable, less than 0.69 log CFU/cm2

one sample detectable

Results were expressed as mean values ± standard deviation from two independent tests in duplicate. The mean values with the same letters are not significantly different (p>0.05).

biofilm would be considered a main concern. The results in Table 1 and Table 2 showed that treatment D1S1 and D1S2 removed only 3 to 4 log CFU/cm2 Pseudomonas; whereas treatment D2S1 or D2S2 removed 5 to 6 log CFU/cm2 Pseudomonas. The main difference between the four treatments was the types of detergents; treatment D1S1 and D1S2 using alkaline detergent, (D1- superp foam, 2%) and treatment D2S1 and D2S2 using acidic detergent

rent study. Our results were similar to the results obtained by Gibson et al. (1999). The investigators showed that Pseudomonas aeruginosa was more resistant to the detergent products, greater than 3 log CFU/cm2 still remained on SS coupons after cleaning; their research also reported that the maximum removal cells was a 4 log CFU/cm2 reduction (Gibson et al., 1999). The reason Pseudomonas biofilms were more resistant than Listeria and Salmonella

(D2 - Dilac Z, 2%). The characteristics of different bacteria, which contribute to the formation of their corresponding biofilms, affects their survival under the cleaning and disinfecting process was examined in the cur-

biofilms may be because of the difference in colonization mechanisms (Gibson et al., 1999). Pseudomonas aeruginosa attached to surfaces produces a variety of extracellular polymeric substance (EPS) such as cellulose, alginate, Pel and PsI exopolysac-

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Table 2. Percentage reduction of 4 cleaning processes on different single biofilms of Pseudomonas, Listeria, Salmonella for 3 and 7 days in TSB 1% and TSB 10% Percentage reduction* Organism

Time (days)

TSB (%)

Treatment D1S1

D1S2

D2S1

D2S2

Pseudomonas

99.8685

99.9943

100.0000

Listeria

99.8435

100.0000

99.8435

100.0000

Salmonella

100.0000

Pseudomonas

99.9235

99.9980

99.9998

100.0000

Listeria

99.9999

100.0000

99.9999

100.0000

Salmonella

100.0000

Pseudomonas

99.9764

99.9968

99.9989

Listeria

100.0000

Salmonella

100.0000

Pseudomonas

99.9232

99.9996

99.9935

99.9996

Listeria

100.0000

Salmonella

100.0000

*Each value is a mean of duplicate replication of two independent tests.

charides which help them to strongly adhere and form strong biofilms on surfaces (Giaouris et al., 2013; Gibson et al., 1999). Moreover, EPS also forms a matrix around the cells, which can protect the cells from adverse conditions (Gibson et al., 1999). In addition, the larger population of Pseudomonas meant thicker biofilm compared to those of Listeria or Salmonella biofilms, causing less diffusion of sanitizers in Pseudomonas biofilms. Therefore, Pseudomonas

CFU/cm2), there was significant differences in terms of viable cells of Pseudomonas and Salmonella biofilms; Pseudomonas spp. remained at approximately 2.7 log CFU/cm2 while no Salmonella spp. colonies were recovered. Similar results were demonstrated by Giaouris et al. (2013). Gram-negative P. putida showed higher tolerance to benzalkonium chloride (BC- QACs -based) compared with the Gram positive L. monocytogenes.

spp. may survive better than the others under the same treatment (Giaouris et al., 2013). The results revealed that under simultaneous growth conditions (3 days - TSB 1% or 7 days - TSB 10%) and having a similar cellular populations (approximately 5.5 log

When comparing the effectiveness of biofilm removal based on detergent pH, an acidic detergent (Dilac Z, pH 1.6; treatment D2) was more effective than an alkaline product (Superp foam, pH 12.2; treatment D1) in terms of effect on cell viability. Sim-

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Figure 3. Efficacy of combination of H2O2 pre-treatment and regular daily cleaning process (D1S1) on mixed species biofilms on SS, Teflon and rubber. Results were expressed as mean values ± standard deviation from two independent tests in duplicate. Among different biofilm and cleaning processes (A, B, C, D, E) and among different materials at the same cleaning process (a, b, c); the mean values with the same letters are not significantly different (p>0.05).

8.0

a a

Stainless steell Teflon Rubber

7.0

Log (CFU/cm2)

6.0

5.0

b c

4.0

a a

3.0

a a a

b ab

a a

2.0

b b

1.0 0.0

A Biofilm Biofilm

B D1S1

1%, 5 min

1%, 10 min

D 2%, 5 min

E 2%, 10 min

Pre-treatment H2O2 + D1S1

(Before cleaning)

Assessment cleaning and sanitizing procedure based on the efficacy to remove mixed species biofilms

ilarly, Gibson et al. (1999) found that the acidic detergent reduced the viable population of attached Staphylococcus aureus but not P. aeruginosa. Our results may be explained by source of Pseudomonas, Pseudomonas genus which can be found as an bundance species in fish and seafood ecosystem grew at pH 6 to 9; they did not growth at pH 4 (Shivaji et al., 1989) thus leading to acidic detergent being more effective in cleaning than the alkaline detergent. Indeed, PAA may have a stronger effect to antimicrobial activity compared with QACs, because PAA had both oxidizing and low pH functions in killing

Although combining detergent and sanitizer in the cleaning process showed more effectiveness to eliminate biofilms (Somers and Wong, 2004), Pseudomonas biofilms were still present on SS coupons after the cleaning process, particularly the D1S1 method. Hence, it can be concluded that the regular daily cleaning process D1S1 is an improper cleaning procedure to remove Pseudomonas biofilms.

bacteria whereas QACs had only one antimicrobial mechanism, namely, interaction with cell membranes, disruption of membranes integrity and leakage of cellular content (Buffet-Bataillon et al., 2012; Giaouris et al., 2013).

However, the cleaning process D1S1 was still applied daily because alkaline detergents have a higher potential to remove organic material and prevent the corosion of equipment in the food procesing environment. In addition, the ability to eliminate micro-

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organisms in biofilms depends on many factors such as the microbial population of the biofilm, quorum sensing, and material where biofilms form and inhabit. From previous experiments, it was demonstrated that mixed species biofilms may be formed by numerous different species. Moreover, mixed species biofilms are usually more stable than single species biofilm (Giaouris et al., 2014). Consequently, there was a need to evaluate the efficacy of cleaning process on mixed species biofilms of Listeria, Salmonella and Pseudomonas under simulated food processing conditions in the current experiment. According to the result from Gram et al. (1987), Guðbjörnsdóttir et al.(2005) and our previous experiment

Moreover, limiting the diffusion of sanitizers within the biofilm occurred because these strong biofilms formed under nutrient limitation (they were subjected to TSB 1%, 8 h per day) and 7 days. Indeed, Giaouris et al. (2014) reported that one of the four mechanisms leading to increase resistance of organisms with sanitizers is a physical barrier formed by the EPS matrix. Therefore, disruption EPS matrix will increase the effectiveness of the cleaning procedure by increasing the penetration of sanitizer into biofilms (Simões et al., 2010; Xavier et al., 2005). There are several methods which could be used to destabilize the EPS matrix of biofilms including biological methods, physical methods and chemi-

(data unpublished), the microflora in seafood plants was shown to be 80% Gram negative microorganisms; Pseudomonas spp. was shown to be the predominant species. Therefore, the mixed species stock which included 80% of volume cell suspension of Pseudomonas spp., 10% of volume of Listeria spp., and 10% of volume of Salmonella spp. was used for this experiment. The efficacy of the cleaning process was determined by the surviving bacteria attached on the surface of coupons after performing the cleaning process. The results were reported as a mean with duplicate replication of two independent experiments. The modified cleaning process was applied to remove mixed species of 7 day biofilms on different substratum such as SS, Teflon and rubber. Biofilm formation on these materials ranged from 7.2 to 7.7 log CFU/cm2. There were no significant difference in the population of cells adhering on SS, Teflon and rubber for 7 days biofilms. D1S1 sample consisted of no treatment with H2O2 using only the cleaning process of D1S1. Overall, the surviving bacterial cells of all samples decreased when H2O2 concentration and exposure time were increased (Fig. 3). Without pretreatment with H2O2, viable cells on SS, Teflon and rubber were 3.3, 4.3 and 5.2 log CFU/cm2, respec-

cal methods; among them H2O2 (chemical method) was selected because of its advantages, namely, low-cost, ease of use, minimal impact on the environment, broad spectrum capabilities and high potential to degrade the EPS matrix (Back et al., 2014; Gao et al., 2014; Imamura et al., 2010). According to Imamura et al. (2010) H2O2 molecules degenerate and produce °OH radicals by accepting an electron from the metal surface. Due to an extremely high oxidation potential, °OH radicals are generated on the metal surface at high concentrations (Imamura et al., 2002). As a consequence, organic substances, which include in EPS, are instantaneously oxidized. The oxidized substances become soluble fragments which can easily be removed from surfaces by a regular cleaning procedure (Imamura et al., 2002, 2010). The effectiveness of pretreatment with H2O2 is associated with H2O2 concentration and exposure time. Average survival of microbial cells in different substratum ranged from 3.1 to 1.3 log CFU/cm2 followed by H2O2 1% for 5 minutes and H2O2 2% for 10 minutes, respectively. The results showed that there was more than a 6 log CFU/cm2 reduction in terms of pre-treatment with H2O2 2% for 10 minutes. A similar result was obtained by DeQueiroz and Day (2007). In their study, when sodium hypochlorite was

tively. The low efficacy for removing mixed species biofilms may be attributed to the large population of microbial cells in the biofilms, and interactions between cells and surfaces (Giaouris et al., 2013, 2014; Lagha et al., 2014; Norwood and Gilmour, 1999).

combined with H2O2, cell numbers were reduced by 5 log to 6 log of P. aeruginosa biofilms after 1 min exposure while sodium hypochlorite reduced viable numbers by 3 log to 4 log under an equivalent concentration. In addition, P. aeruginosa biofilm

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formation on SS and aluminum surfaces were also removed (DeQueiroz and Day, 2007). The results of this experiment was supported by Choi et al. (2012) where pathogen biofilms on the SS surfaces were reduced when treated with aerosolized hydrogen peroxide-based sanitizer. However, there was no significant difference between pre-treatment with H2O2 1% for 10 minutes and H2O2 2% for 5 minutes. It was suggested that the treatment of 1% of H2O2 for 10 minutes or 2% of H2O2 for 5 minutes should result in a low concentration or short exposure time for °OH radicals to decompose the EPS matrix of the highly mixed microbial species generated in these biofilms. Attachment or detachment of microorganisms on

nas biofilms demonstrated higher adaptability for all growth conditions; as indicated by the higher number of cells in biofilms. Four existing cleaning processes were effective against Listeria and Salmonella biofilms, with Pseudomonas biofilms being the exception. For mixed-species biofilm, when combined with pretreatment of H2O2, the cleaning process was more effective when compared to using the existing cleaning process alone; there was a 4 log CFU/cm2 reduction for the control method compared to a 6 log CFU/cm2 reduction for the method that combined pre-treatment H2O2 2% for 10 minutes and a cleaning process for cleaning mixed species biofilms on SS coupons. SS is ideally suited for food indus-

surfaces depends on both characteristics of bacteria and material surfaces (Sinde and Carballo, 2000). Among the three materials SS, Teflon and rubber, there were no significant difference in the number of adhesive cells but there was a significant decrease in the number of viable cells during the cleaning process. The number of viable cells on rubber remained high in both the control and the pre-treatment with H2O2. In fact, when applying the pre-treatment H2O2 2% for 10 minutes to the biofilm formation on the rubber coupon, there was greater than a 5 log CFU/ cm2 reduction, however there remained 2 log CFU/ cm2 of viable cells on the surfaces of rubber coupons. While applying the same treatment condition, biofilms on SS and Teflon were removed, reaching a 6 log CFU/cm2 reduction. Higher efficacy in cleaning process to SS and Teflon could be explained because a high concentration of °OH radicals was produced on the metal surface (Imamura et al., 2010) and the more hydrophobic characteristics of the Teflon surface.

try, especially in terms of eliminating and preventing biofilm formation. Therefore, the combination of H2O2 pre-treatment and cleaning process can be used as an alternative method to remove mixed species biofilms in food processing equipment.

ACKNOWLEDGEMENTS The authors are grateful for financial support from: the Faculty of Agro-Industry, Kasetsart University, Thailand (through the Scholarships for International Graduate Students 2012) and Agri-Biotech and Fisheries project, Vietnamese Government grant for this study. The authors would like to thank Professor Dr. Steven C. Ricke, Editor-in-Chief and Professor Dr. Md. Latiful Bari, University of Dhaka, Bangladesh for their valuable comments and suggestions for this manuscript.

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Hops (Humulus lupulus) ß-Acid as an Inhibitor of Caprine Rumen Hyper-Ammonia-Producing Bacteria In Vitro M. D. Flythe1, 2, G. E. Aiken1, 3, G. L. Gellin1, J. , L. Klotz1, 2, B. M. Goff3, K. M. Andries4 1

Forage-Animal Production Research Unit, Agricultural Research Service, United States Department of Agriculture 2 Department of Animal & Food Sciences, University of Kentucky 3 Department of Plant & Soil Sciences, University of Kentucky 4 College of Agriculture, Food Science & Sustainable Systems, Kentucky State University “Proprietary or brand names are necessary to report factually on available data; however, the USDA neither guaran-

tees nor warrants the standard of the product, and the use of the name by the USDA implies no approval of the product, nor exclusion of others that may be suitable.”

ABSTRACT Antimicrobial plant secondary metabolites increase rumen efficiency and decrease waste products (i.e. ammonia, methane) in some cases. A promising source of bioactive secondary metabolites is the hops plant (Humulus lupulus L.), which produces ß-acid, a suite of structurally similar, potent antibacterial compounds. The efficacy of hops has been shown in bovines. Additionally, the ß-acid mechanism of antimicrobial action was determined on rumen bacteria of bovine origin. The objective of the current study was to determine the effect of hops ß-acid on amino acid degradation and ammonia production by goat rumen bacteria. The growth of two rumen hyper-ammonia-producing bacteria of caprine origin (Peptostreptococcus spp. BG1 and BG2) was inhibited by ß-acid (≥ 45 ppm and ≥ 4.5 ppm, respectively) when either amino acids or peptides were the growth substrate. Uncultivated, mixed rumen microorganisms harvested from goats produced approximately 35 mM ammonia from amino acids and 50 mM ammonia from peptides during 24 h incubation. The addition of ß-acid reduced the final ammonia concentration, and there was a dose-response relationship between the ß-acid concentration and the ammonia concentration. Peptide catabolism was more sensitive to ß-acid inhibition than free amino acid catabolism to ß-acid inhibition because as little as 3 ppm resulted in less ammonia production than the control (P < 0.05). These results demonstrate that hops ß-acid is effective against caprine HAB and suggest that hops could be useful in controlling wasteful metabolic processes in the caprine rumen. Keywords: antibiotic alternative, ammonia, beta-acid, bypass protein, feed efficiency, hops, goat, hyper ammonia producing bacteria, inhibition, ionophore, lupulone, phytochemical, phytoprotectant, plant secondary metabolite, rumen Agric. Food Anal. Bacteriol. 5: 29-36, 2015

Correspondence: Michael D. Flythe, michael.flythe@ars.usda.gov Tel: +1 -859-421-5699 Fax: +1-859-257-3334

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INTRODUCTION A symbiotic relationship with the rumen microflora is the adaptation that gives ruminants metabolic access to fibrous plant tissues (Russell and Rychlik, 2001). This dense community of microorganisms includes bacteria, archaea, fungi and protists, which collectively degrade fiber and convert the resulting sugars to fermentation acids and gases. The fermentation acids can be transported across the rumen epithelium and utilized by the ruminant host, and the gases, which are essential for reducing equivalent cycling in the microorganisms, are eructated (Russell and Rychlik, 2001).

Shazly, 1964) and to a guild of bacteria termed Hyper Ammonia-producing Bacteria, HAP or HAB (Russell et al., 1988). The contribution of each type of microorganism to rumen ammonia depends on host, diet and other factors. However, ammonia production in defaunated ruminants supports the hypothesis that bacteria are sufficient for amino acid degradation (Abou Akkada and El-Shazly, 1964). Fortunately, both HAB and ciliates are susceptible to ionophores, which are included in many ruminant diets (Chen and Russell, 1989; Dennis et al., 1986). Ionophores (e.g. monensin, lasalocid) inhibit ammonia production, increase feed efficiency, promote growth and decrease amino-nitrogen release into the environment

In addition to carbohydrates, plant proteins, peptides and amino acids are also catabolized. Rumen proteolysis and deamination (diagrammed in Figure 1) are not inherently harmful to ruminants. In fact, some cellulolytic bacteria, such as Ruminococcus albus, require the aromatic- and branched-acids produced by deamination of particular amino acids (Caldwell and Bryant, 1966). Bacteria can assimilate some of the ammonia allowing the ruminant host to utilize the resulting microbial protein (Satter and Slyter, 1974). The ruminant converts excess ammonia to urea, some of which can be recycled into the rumen, and goats are exceptionally good at this process (Kohn et al., 2005). Ruminants have lower nitrogen clearance rates than non-ruminants and goats tend to be lower than other ruminants (Kohn et al., 2005). The urea transport rate through the rumen epithelum and kidney increases as nitrogen in the diet decreases (Muscher et al., 2010; Starke et al., 2014). When urea is not recycled it is lost in the urine. The loss of feed amino-nitrogen in the urine is an economic loss to the rancher or dairyman, and a source of environmental pollution (Tedeschi et al., 2003). Additionally, amino acid fermentation results in the production of indole and related compounds, which can cause flavor notes in milk that are undesir-

(Tedeschi et al., 2003; Callaway et al., 2008). In spite of the production and environmental advantages and limited clinical utility of ionophores, the fact that they are antibiotics has led to some resistance to their use in industrialized nations (Russell and Houlihan, 2003). For more than a decade, researchers have looked for phytochemical alternatives to control ammonia production and other aspects of rumen fermentation (Wallace, 2004). Some phytochemicals, like those derived from the hops plant (Humulus lupulus) even have an ionophore-like mechanism of action on HAB (Flythe 2009). Antimicrobial phytochemicals have the added advantage of production from local botanical sources. For example, hops might be an appropriate choice in northern Europe or northern North America where this plant is cultivated. Spearmint might be a better local source of phytochemicals in the Middle East, and spearmint essential oil has been shown to have antimicrobial activity on HAB from Mehraban sheep (Taghavi-Nezhad et al., 2013). The use of regionally available antimicrobial phytochemicals in ruminant production raises the question of which animals and microorganisms should be used to test these compounds. Most rumen microbiology research has been conducted on dairy cows

able to some consumers (Attwood et al., 2006). Proteolysis is carried out by a variety of rumen microorganisms, but most of the ammonia production from the resulting peptides and amino acids is attributable to ciliate protozoa (Abou Akkada and El-

in North America and Western Europe, but goats are the predominant domestic ruminants in many parts of the world (FAO, 2011). Goat production increased more than other species, except poultry, in both developed and developing countries (FAO,

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Figure 1. Simplified schematic of amino-nitrogen cycling the rumen. Processes are labeled: 1- Proteolysis by proteolytic microorganisms, 2- “By-pass” protein that is not deconstructed in the rumen, 3- Deamination by HAB and other microorganisms, 4- Assimilation of ammonia and amino acids by microorganisms for anabolic purposes.

2011). The goat industry in the US increased by 23% between 1997 and 2007, this increase was 116% for the 12 Southeastern states (USDA 2004 and 2009; USDA-APHIS 2005), which is where the current work was conducted. The size of the industry steadied between 2002 and 2012 goat numbers increased by just over 1% in these same states (USDA 2004 and 2014). The purpose of the study was to test the effect of a model antimicrobial phytochemical on: 1) the growth of caprine HAB, and 2) ammonia production by uncultivated rumen microorganisms from goats. Hops ß-acid is a suite of structurally similar compounds (lupulone, colupulone, adlupulone) that are potently inhibitory to Gram-positive bacteria. It was selected

MATERIALS AND METHODS

as the model antimicrobial phytochemical because inhibition and the mechanism of action have already been determined on bovine HAB (Flythe 2009).

and 0.25 kg head-1d-1 soya-based supplement (16% crude protein; Southern States Cooperative, Richmond, Virginia, USA). Water and a mineral mixture (Southern States Cooperative, Richmond, Virginia, USA) were provided free choice.

Animals and diet All animal husbandry and procedures were approved by the University of Kentucky care and use committee. Rumen fistulated Kiko goat wethers (n=4; 2 y, 40-50 kg) were used as rumen digesta donors. They were maintained on pasture in a herd of Kiko goats at the University of Kentucky’s Research Farm, Lexington, Kentucky, USA. The botanical composition of the pasture is shown in Table 1. The herd was supplemented with 1.0 kg head-1 d-1 orchard grass hay (Dactylis glomerata; 18% crude protein),

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Table 1. The botanical composition of the pasture Pasture Component

Percent

Forage Tall Fescue

Lolium arundinaceum (Schreb.) Darbysh.

44.4

Orchardgrass

Dactylis glomerata L.

14.9

White Clover

Trifolium repens L.

14.5

Kentucky Bluegrass

Poa pratensis L.

8.9

Red Clover

Trifolium pratense L.

3.3

Bermudagrass

Cynodon dactylon (L.) Pers.

1.4

Weeds*

12.6

* Weed species include: Rumex crispus L., Taraxacum officinale F.H. Wigg., Plantago coronopus L., Solanum carolinense L., Convolvulus arvensis L., Digitaria sp., Ambrosia artemisiifolia L.

Bacterial strains and media composition The isolation and characterization of the HAB cultures (Peptostreptococcus spp. BG1 and BG2) used in these experiments were previously reported (Flythe and Andries, 2009). The HAB medium contained (per liter): 240 mg K2HPO4, 240 mg KH2PO4, 480 mg NaCl, 480 mg Na2SO4, 64 mg CaCl2·2H2O, 100 mg MgSO4·7H2O, 600 mg cysteine hydrochloride, mineral and vitamin solutions as previously described (Russell et al., 1988). The initial pH was adjusted to 6.5 with 20% NaOH. The liquid was autoclaved to remove O2 and cooled under O2 -free CO2. The buffer (4.0 g Na2CO3) was added before dispensing anaerobically and autoclaving again for sterility. Casamino acids and Trypticase (Fisher BioReagents, Fair Lawn, New Jersey, USA) were prepared separately using anaerobic technique, as described above. They were aseptically added when indicated (15 mg ml-1 final concentration).

Pure culture growth experiments The HAB medium was amended with Casamino acids or Trypticase (15 mg ml-1). Overnight Pepto32

streptococcus spp. BG1 and BG2 cultures were used as the inocula (10% v/v). “Beta-Bio” hops extract was added to the media, vigorously mixed and serially diluted with to achieve the concentration indicated. The propylene glycol-based extract contained 45% ß-acid and no detectable ß-acid, as reported by the manufacturer (S.S. Steiner, Inc., Yakima, Washington, USA). The concentrations were verified by HPLC (Dionex; Sunny Vale, California). The Phenomineex 250×4.6 mm Luna 5u C18 column (Torrance, California) was 30˚C. The mobile phase was 80% methanol, 20% water, pH 2.5. Eluting compounds were detected by UV absorbance (314 nm). The standard curve was generated using hops standards obtained from the American Society of Brewing Chemists (St. Paul, Minnesota). The limits of detection were 3.3 ppm and 4.1 ppm for colupulone and adlupulone, respectively. The extract was not autoclaved, but was added to uninoculated media and incubated as a control. Controls for the effect of the propylene glycol carrier were also performed. The incubations were conducted in a shaking incubator (150 rpm, 39˚C). Growth was determined by optical density at 24 and 48 h (600 nm).

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Figure 2. Effect of hops extract on ammonia production by uncultivated microorganisms from the goat rumen. Ammonia concentrations from peptides (squares) or amino acids (circles) after 24 h incubation (39 ° C) are shown. Error bars indicate standard error of the mean. Asterisks indicate treatments that are different than the 0 ppm control (P < 0.05).

Ammonia production by uncultivated rumen microorganisms The rumen digesta (approx. 0.5 kg) was collected from each goat individually (n=4) and transported back to the laboratory in an insulated, airtight container within 30 min. Rumen contents were removed and filtered through four layers of muslin cheesecloth into centrifuge bottles. The fluid was subjected to low-speed centrifugation (100 x g, 10 min) to remove feed particles. The supernatants were transferred to new bottles and subjected to high-speed centrifugation (25,600 x g, 10 min) to harvest the microorganisms. The pellets were resuspended in HAB medium without a growth substrate, and low-speed centrifugation was repeated to wash the cells. The pellets were resuspended in HAB medium, pooled into a glass vessel (500 ml), and sparged with O2-free CO2. Microscopic examination revealed that the cell suspension (approximately 15.0 OD, pH 6.7) contained no visible plant fiber and few protozoa. The

cell suspension was dispensed into serum bottles that contained CO2. Growth substrates (Trypticase or casamino acids, 15 mg ml-1) and hops extract were added as indicted. The bottles were incubated in a shaking water bath (39˚C 150 rpm). Samples were clarified by centrifugation and frozen (-20°C) until ammonia analysis. Ammonia concentrations were determined with a colorimetric method (Chaney and Marbach, 1962).

Replication and statistics The pure culture growth experiments were repeated three times with identical results. The in vitro ammonia production experiments were repeated four times using cell suspensions from a different goat each time. The data were subjected to an analysis of variance with Tukey’s test post hoc. Results were considered significant when P < 0.05.

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RESULTS Two caprine HAB pure cultures (Peptostreptococcus spp. BG1 and BG2) grew when either free amino acids (Casamino acids) or peptides (Trypticase) were the sole growth substrate, and produced ammonia at rates as great as 600 nmol mg cell protein-1min-1 (data not shown). Hops extract inhibited growth of the two HAB cultures when the ß-acid concentrations were ≥ 45 ppm and ≥ 4.5 ppm, respectively. The inhibitory concentration was not changed if amino acids or peptides were used as the growth substrate. Cell suspensions of uncultivated rumen microorganisms from goats produced approximately 35 mM ammonia from free amino acids (Casamino acids) and 50 mM ammonia from peptides (Trypticase) in 24 h (Fig. 2). The addition of hops extract decreased the total ammonia produced during the incubations. Ammonia from amino acids was not significantly less than the control unless the ß-acid concentration was 30 ppm or greater. When the ß-acid concentration was 30 ppm, ammonia production from amino acids was 30% less than the control (P < 0.05). When peptides were the growth substrate as little as 3 ppm ß-acid caused a decrease in ammonia production (P < 0.05). Ammonia production from peptides was decreased by 50% relative to the control when the ßacid concentration was 12 ppm or greater (P < 0.05).

croorganisms. Wang and colleagues (2010) showed the in vivo efficacy of hops in a feeding trial. In that experiment, the average daily gain of steers was improved by the inclusion of hops cones in the diet. The results of the current study indicate that hops, specifically the ß-acid component, can decrease ammonia production by microorganisms from the goat rumen. Inhibition of caprine HAB in pure culture demonstrated that the decrease in ammonia could be accomplished by antimicrobial of the ß-acid on HAB. These results are consistent with previous work that showed antimicrobial action on three bovine HAB species (Flythe, 2009).

ACKNOWLEDGEMENTS This work was funded by the United States Department of Agriculture, Agricultural Research Service. S.S. Steiner, Inc. (Yakima, WA) donated the hops extract used in this study. The authors acknowledge the technical assistance of Adam Barnes and Tracy Hamilton.

REFERENCES

Several other experiments have been performed to assess the usefulness of hops as a feed additive, and all of these employed cows or microorganisms of bovine origin. Previous in vitro experiments indicated that hops ß-acid did not inhibit bovine ruminal protozoa (Schmidt et al., 2006). However, hops ß-acid did inhibit Streptococcus bovis and decreased the acetate: propionate ratio during in vitro experiments

Abou Akkada, A. R., and K. El-Shazly. 1964. Effect of absence of ciliate protozoa from the rumen on microbial activity and growth of lambs. Appl. Microbiol. 12: 384-390. Attwood, G., D. Li, D. Pacheco, and M. Tavendale. 2006. Production of indolic compounds by rumen bacteria isolated from grazing ruminants. J. Appl. Microbiol. 100: 1261-1271. Caldwell, D. R., and M. P. Bryant. 1966. Medium without rumen fluid for non-selective enumeration and isolation of rumen bacteria. Appl. Microbiol. 14: 794-801. Callaway, T. R., T. S. Edrington, J. L. Rychlik, K. J.

(Flythe and Aiken, 2010). S. bovis is proteolytic (Russell et al., 1981) thus, the results of the latter study is consistent with recent work by Lavrenčič and colleagues (2013), who determined that two varieties of hops decreased proteolysis by bovine rumen mi-

Genovese, T. L. Poole, Y. S. Jung, K. M. Bischoff, R. C. Anderson, and D. J. Nisbet. 2003. Ionophores: Their use as ruminant growth promotants and impact on food safety. Curr. Issues Intest. Microbiol. 4: 43-51.

DISCUSSION

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Chaney, A. L., and E. P. Marbach. 1962. Modified reagents for determination of urea and ammonia. Clin. Chem. 8: 130-132. Chen, G. J., and J. B. Russell. 1989. More monensinsensitive, ammonia-producing bacteria from the rumen. Appl. Environ. Microbiol. 55: 1052-1057. Dennis, S. M., T. G. Nagaraja, and A. D. Dayton. 1986. Effect of lasalocid, monensin and thiopeptin on rumen protozoa. Res. Veter. Sci. 41: 251-256. Flythe, M. 2009. The antimicrobial effects of hops (Humulus lupulus l.) on ruminal hyper ammoniaproducing bacteria. Letters Appl. Microbiol. 118: 242-248. Flythe, M., and K. Andries. 2009. The effects of mo-

Enrichment and isolation of a ruminal bacterium with a very high specific activity of ammonia production. Appl. Environ. Microbiol. 54: 872-877. Russell, J., and J. Rychlik. 2001. Factors that alter rumen microbial ecology. Sci. 11: 1119-1122. Schmidt, M. A., M. L. Nelson, J. J. Michal, and H. H. Westberg. 2006. Effects of hop acids. II. Beta acids on ruminal methane emission, protozoal population, fermentation, and coM concentration in cannulated finishing steers. J. Anim. Sci. Suppl. 84: 240. Satter, L. D., and L. L. Slyter. 1974. Effect of ammonia concentration on rumen microbial protein production in vitro. Brit. J. Nutr. 32: 199-208.

nensin on amino acid catabolizing bacteria isolated from_the Boer goat rumen. Small Ruminant Res. 81:178–181. Flythe, M. D., and G. E. Aiken. 2010. Effects of hops (Humulus lupulus l.) extract on volatile fatty acid production by rumen bacteria. J. Appl. Microbiol. 109: 1169-1176. Russell, J.B., Bottje, W.G., and M.A Cotta. 1981. Degradation of protein by mixed cultures of rumen bacteria: identification of Streptococcus bovis as an actively proteolytic rumen bacterium. J. Anim. Sci. 53: 242-252. Russell, J. B., and A. J. Houlihan. 2003. Ionophore resistance of ruminal bacteria and its potential impact on human health. FEMS Microbiol. Rev. 27: 65-74. Kohn, R. A., M. M. Dinneen; and E. Russek-Cohen. 2005. Using blood urea nitriogen to predict nitrogen excretion and efficiency of nitrogen utilization in cattle, sheep, goats, horses, pigs, and rats. J. Anim. Sci. 83:879-889. Lavrenčič, A., A. Levart, I. J. Košir, and A. Čerenak. 2013. Influence of two hop (Humulus lupulus L.) varieties on in vitro dry matter and crude protein degradability and digestibility in ruminants. J. Sci. Food Agricult. 94: 1248-1252.

Starke, S., A. S. Muscher, N. Hirschhausen, E. Pfeffer, G. Breves, and K. Huber. 2012. Expression of urea transporters is affected by dietary nitrogen restriction in goat kidney. J. Anim. Sci. 90:38893897. Taghavi-Nezhad, M., D. Alipour, M. D. Flythe, P. Zamani, and G. Khodakaramian. 2013. The effect of essential oils of Zataria multiflora and Mentha spicata on the in vitro rumen fermentation, and growth and deaminative activity of amino acidfermenting bacteria isolated from Mehraban sheep. Anim. Prod. Sci. 54: 299-307. Tedeschi, L. O., D. G. Fox, and T. P. Tylutki. 2003. Potential environmental benefits of ionophores in ruminant diets. J. Environ. Qual. 32: 1591-1602. USDA-APHIS. 2005. The goat industry structure, concentration, demand and growth. Electronic Report. http://www.aphis.usda.gov/animal_health/ emergingissues/downloads/goatreport090805. pdf USDA. 2004. 2002 Census of Agriculture. AC02-A-51. http://www.agcensus.usda.gov/Publications/2002/USVolume104.pdf USDA. 2009. 2007 Census of Agriculture . AC07-A-51. http://www.agcensus.usda.gov/Publications/2007/Full_Report/Volume_1,_Chapter_1_

Muscher, A. S., B. Schröder, G. Breves, and K. Huber. 2010. Dietary nitrogen reduction enhances urea transport across goat rumen epithetium. J. Animal Sci. 88:3390-3398. Russell, J. B., H. J. Strobel, and G. J. Chen. 1988.

US/usv1.pdf USDA. 2014. 2012 Census of Agriculture. AC12-A-51. http://www.agcensus.usda.gov/Publications/2012/Full_Report/Volume_1,_Chapter_1_ US/usv1.pdf

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Wallace, R. 2004. Antimicrobial properties of plant secondary metabolites. Proceedings of the Nutrition Society 63: 621-629. Wang, Y., A. V. Chaves, F. L. Rigby, M. L. He, and T. A. McAllister. 2010. Effects of hops on ruminal fermentation, growth, carcass traits and shedding of Escherichia coli of feedlot cattle. Livestock Sci. 129: 135-140.

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VOLUME 4 ISSUE 1 Case Studies 8

Introduction Special Issue P. G. Crandall

A Personal Hygiene Behavioral Change Study at a Midwestern Cheese Production Plant J. A. Neal, C. A. O’Bryan and P. G. Crandall

Preventing Post-Processing Contamination in a Food Nugget Processing Line When Language Barriers Exist J. A. Neal, C. A. O’Bryan and P. G. Crandall

Behavioral Change Study at a Western Soup Production Plant C. A. O’Bryan, J. A. Neal, and P. G. Crandall

Salmonella in Cantaloupes: You Make Me Sick! B. A. Almanza

The Hurricane Sandy Dilemma B. A. Almanza

Intellect-u-ale: A Smart Approach to Quality Assurance in a Micro-Brewery A. J. Corsi, M. Goodman, and J. A. Neal

Introduction to Authors 61

Instructions for Authors

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VOLUME 4 ISSUE 2 ARTICLES 96

Contribution of Chemical and Physical Factors to Zoonotic Pathogen Inactivation during Chicken Manure Composting M.C. Erickson, J. Liao, X. Jiang, and M.P. Doyle

REVIEW 76

Antibiotic Use in Livestock Production Broadway, P. R., J. A. Carroll, and T. R. Callaway

Effects of Co-nutrients in Foods and Bioremediation in the Environment on Methylmercury P. G. Crandall, C. A. O’Bryan

109 Alternative antimicrobial supplements that positively impact animal health and food safety Broadway, P. R., J. A. Carroll, and T. R. Callaway

122 Human Health Benefits of Isoflavones from Soybeans k. Kushwaha, C. A. O’Bryan, D. Babu, P. G. Crandall, P. Chen, and S.-O. Lee

Introduction to Authors 147 Instructions for Authors

The publishers do not warrant the accuracy of the articles in this journal, nor any views or opinions by their authors. 38

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VOLUME 4 ISSUE 3 ARTICLES 164 The Prevalence of E. coli O157:H7 in the Production of Organic Herbs and a Case Study of Organic Lemongrass Intended for Use in Blended Tea

S. Zaman, Md. K. Alam, S. S. Ahmed, Md. N. Uddin, and Md. L. Bari

177

Batch Culture Characterization of Acetogenesis in Ruminal Contents: Influence of Acetogen Inocula Concentration and Addition of 2-Bromoethanesulfonic Acid P. Boccazzi and J. A. Patterson

195 The Effect of Phytochemical Tannins-Containing Diet on Rumen Fermentation Characteristics and Microbial Diversity Dynamics in Goats Using 16S rDNA Amplicon Pyrosequencing B. R. Min, C. Wright, P. Ho, J.-S. Eun, N. Gurung, and R. Shange

212 Characterization of the Novel Enterobacter cloacae Strain JD6301 and a Genetically Modified Variant Capable of Producing Extracellular Lipids

J. R. Donaldson, S. Shields-Menard, J. M. Barnard, E. Revellame, J. I. Hall, A. Lawrence, J. G. Wilson, A. Lipzen, J. Martin, W. Schackwitz, T. Woyke, N. Shapiro, K. S. Biddle, W. E. Holmes, R. Hernandez, and W. T. French

224 Survival of Salmonella enterica and Listeria monocytogenes in manure-based compost mixtures at sublethal temperatures

M.C. Erickson, C. Smith, X. Jiang, I.D. Flitcroft, and M.P. Doyle

Introduction to Authors 243 Instructions for Authors

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Aerobic microbiology Aerobiology Anaerobic microbiology Analytical microbiology Animal microbiology Antibiotics Antimicrobials Bacteriophage Bioremediation Biotechnology Detection Environmental microbiology Feed microbiology Fermentation Food bacteriology Food control

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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.

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

Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 5, Issue 1 - 2015

“(James et al., 2010a,b)” in text, refers to “James, Smith, and Elliot. 2010a” and “James, West, and Adams. 2010b” in the reference section.

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.

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.

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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