Afab volume 4 issue 3 by cielo shabatura

ISSN: 2159-8967 www.AFABjournal.com

Volume 4, Issue 3 2014

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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. 4, Issue 3 - 2014

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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. 4, Issue 3 - 2014

TABLE OF CONTENTS 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

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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

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. â&#x20AC;˘ AFABjournal.com â&#x20AC;˘ Vol. 4, Issue 3 - 2014

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NEW EDITORIAL STAFF Dr. Byeng R. Min appointed to AFAB editorial board Agriculture, Food and Analytical Bacteriology is pleased to welcome Dr. Byeng R. Min to the editorial board. Dr. Byeng R. Min is an Animal nutritionist and rumen microbiologist in the Animal Science Unit at Tuskegee University in Tuskegee, AL. He has been with Tuskegee University since 2009. Byeng earned degrees in Animal science from Kon-Kuk University (B.S.), South Korea, and Massey University (M.S. and Ph.D.), New Zealand, and gained experience with utilization of phytochemical tannin-containing forages by small ruminants during his studies at Massey University in New Zealand. Dr. Min conducts research on sheep, goats, and cattle, focusing on providing technology to enable small to mid-size farmers to maximize profits and sustainability. His primary interest in rumen and intestinal microbial diversity as well as alternative control of food borne pathogens and gastro-intestinal parasites. Dr. Min is author and/or co-author of over 52 refereed journal articles, numerous technical/report papers, proceedings papers, and abstracts, and 2 book chapters.

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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. Zaman1, Md. K. Alam2, S. S. Ahmed4, Md. N. Uddin3, and Md. L. Bari1 Center for advanced Research in Sciences, University of Dhaka, Dhaka-1000, Bangladesh Institute of Food and Radiation Biology, Bangladesh Atomic Energy Commission, Savar, Dhaka, Bangladesh 3 Bangladesh Agricultural Research Institute, Gazipur 1701, Bangladesh 4 Kazi & Kazi Tea Estate Ltd. University of Liberal Arts Bangladesh (ULAB), Dhanmondi, Dhaka-1209, Bangladesh 1

ABSTRACT Tea blended with different herbs bring a world of flavors, aromas and colors and is usually made with dried tea leaves, or blended with other dried herbs and involves pouring boiling water over the leaves, letting them steep for few minutes followed by consumption. This study was done to evaluate the insights of potential microbial contamination of organic herbs production at the farm, after harvest, washing, before or after drying and packaging of dried herbs sample. Organic compost, water quality, worker hygiene status and overall food safety management systems were also evaluated to identify additional factors affecting microbiological contamination. In addition, effect of pouring hot water over contaminated dried leaves in a cup of tea was observed. The study was designed in such a way that reflects the actual tea preparation at home. Presence of higher numbers of generic E. coli and pathogenic E. coli O157:H7 was observed in dried tea, herbs and /or lemongrass samples, and blended tea mix lemongrass samples. However, no Salmonella was detected in any of the samples tested. When hot water was added into dried lemongrass or blended tea mix lemongrass samples in a cup of tea and held for 30, 60, 90, 120 or 180 seconds with or without a lid, no generic E. coli and pathogenic E. coli O157:H7 was observed in the prepared cup of tea in 30 seconds or above the holding time in selective medium. The bacteria might be severely injured by hot water treatment and did not appear on the selective plates. To confirm whether the bacteria were inactivated or injured, an enrichment study was done. Neither generic E. coli nor any pathogenic E. coli O157:H7 were detected in the prepared tea in the cup. The hot water temperature was recorded as 82˚C when added in the cup and after 60 seconds the temperature decreased to 78˚ C; further reduced to 73˚C after 3 minutes of holding and at the end of 5 minutes the temperature reached 64˚ C. In addition, the natural microflora was reduced to less than 100 CFU/ml. This finding suggested that addition of hot water (80˚C) in tea leaves resulted in complete elimination of pathogens and thus the present tea making practice could provide safe tea for drinking even though the tea leaves were contaminated. However, for sanitary reasons E. coli should be eliminated from the organic products prior to consumption. Keywords: Organic herbs, E. coli O157:H7, organic lemongrass, case study and blended tea Agric. Food Anal. Bacteriol. 4: 164-176, 2014 Correspondence: Md. Latiful Bari, latiful@univdhaka.edu Tel: 8801971560560 Fax: 8802-8615583

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INTRODUCTION Lemongrass (Cymbopogan flexuosus, family: Poaceae) is an aromatic plant which grows in many parts of tropical and sub-tropical South East Asia and Africa. Most of the species of lemon grass are native to South Asia, South-East Asia and Australia (USDA, 2008). Lemongrass naturally grows in tropical areas and can resist the heat of the sun. Fresh cut lemongrass often lasts for several days and can be preserved in fresh water for several months without losing any flavor or nutritional properties. However, these lemongrasses are dried easily, readily available worldwide and can be used to make tea.

These fresh herbs and leafy greens are potential transmission sources of enteropathogens. In a recent report from WHO/FAO on microbiological hazards in fresh fruits and vegetables (FAO/WHO 2008) it was stated that leafy green vegetables (including fresh herbs) “currently presented the greatest concern in terms of microbiological hazards.” This is because these products are grown and exported in large volumes, and they have been associated with many foodborne disease outbreaks affecting considerable numbers of people. Additionally, the production chain for leafy greens is highly complex. The microflora on these vegetables at harvest reflects the environment in which they are grown, if the temperature

Lemongrass is usually ingested as an infusion made by pouring boiling water on fresh or dried leaves and is one of the most widely used traditional plants in South American folk medicine (Blumenthal, 1998). It is used as an antispasmodic, antiemetic, and analgesic, as well as for the management of nervous and gastrointestinal (GI) disorders and the treatment of fevers (Leung, 1980). In India it is commonly used as an antitussive, antirheumatic, and antiseptic. In Chinese medicine, lemongrass is used in the treatment of headaches, stomach aches, abdominal pain, and rheumatic pain (Girón et al., 1991). Lemongrass is an important part of Southeast Asian cuisine, especially as a flavoring in Thai food. Lemongrass is used in Cuban folk medicine for hypertension and as an antiinflammatory (Lewinsohn et al., 1998). It is also used in Brazilian folk medicine in a tea called abafado as a sedative, and for gastrointestinal problems and fever (Martínez-de la Puente et al., 2009). Lemongrass and closely related species are popularly used as insect repellents (Wong et al., 2005; Tawatsin et al., 2001). They may be found in sprays, candles, and other repellent products. Various experimental studies support its use as an insecticide or insect repellant. Lemongrass has been shown to have antifungal properties in laboratory studies particularly against Candida species (Can-

and humidity is relatively high then the occurrence of enteropathogenic bacteria in this environment might be considerable. During cultivation, use of contaminated water for irrigation, application of biocides, and refreshing or washing of harvested crops, are potential sources of contamination. Contamination from contact with fresh manure used as fertilizer cannot be excluded. Heavy rainfall may also lead to fecal contamination from the environment. Direct sunshine will most likely have a disinfection effect, but if the plants are irrigated until harvest and the production hygiene during harvest and post-harvest is inadequate, there is a relatively high likelihood that the fresh herbs and leafy greens may be fecally contaminated. These fresh herbs and leafy greens and their products have been found to be contaminated with pathogenic bacteria such as Staphylococcus aureus, Escherichia coli, Salmonella enterica serovar Typhi, Shigella spp, Bacillus spp. amongst others that represent serious public health hazards (Abadias et al., 2008; Esimone et al., 2003; Oyetayo, 2008; Abba et al., 2009; Adeleye et al., 2005). Some of these pathogenic bacteria originate from soil and adhere to parts of plants (Lau et al., 2003) while most of them are being introduced into leafy products through processes of harvesting, drying, storage and manufacturing because of the

dida albicans, Candida glabrata, Candida krusei, Candida parapsilosis, and Candida tropicalis) (Warnke et al., 2009). In a preliminary study, lemongrass infusion had beneficial effects for the treatment of oral candidiasis in patients with HIV/AIDS (Wright et al., 2009).

unhygienic practices of the product handlers (Lau et al., 2003; Espen et al., 2008). In 2005, the Norwegian Food Safety Authority (Mattilsynet) conducted an ad hoc survey of 162 fresh herbs and green or leafy vegetables products, from South East Asia, and found

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that 28% were contaminated with Salmonella, and 35% with E. coli at greater than 100 CFU/gram. This resulted in a general import prohibition of such products from South East Asia, and now the EU accordingly requires certificate of analysis for Salmonella and E. coli before export (Olaimat and Holley, 2012). The objectives of this present study were to evaluate microbial contamination of organic herb production at the farm level, and a case study of food safety management in organic lemongrass production intended for blended tea. Organic compost, water quality, worker hygiene status and overall food safety management systems were also evaluated to identify the potential factors affecting microbiologi-

30, 2013. All samples were transported to the Food Analysis and Research Laboratory, Center for Advanced Research in Sciences (CARS) at the University of Dhaka using a cool box at the earliest convenience for processing and further analysis. All the microbial analysis was carried out according to the standard methods described in United States Food and Drug Administration (US-FDA) Bacteriological Analytical Manual.

cal contamination. In addition, effect of pouring hot water over contaminated dried leaves in a cup of tea was observed. The study was designed in such a way that reflects the actual tea preparation at home.

The critical sampling locations were selected based on the production scheme presented in Figure 1a and other sources of microbiological contamination as identified in literature reviews (Ilic et al., 2012; Vidal et al., 2004) i.e., soil, water, manure, food contact surfaces, or food handlers. For dried organic lemongrass production, 12 CSLs were selected (Figure 1b) including the lemongrass crop.

MATERIALS AND METHODS

Selected critical sampling locations (CSLs)

Sample collection Herb samples include, lemongrass, mint, neem and jasmine were obtained from an organic farm in Northern Bangladesh between May 15 and July

Figure 1a. Schematic flow diagram of lemongrass production chain (Farm to table).

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Total aerobic count and total coliform count Twenty five (25) g of each sample were homogenized in 225 mL of saline water (0.85% NaCl). Decimal dilutions were prepared upto 10-6 mL and appropriate dilutions were spread plated on Tryptic soy agar (Oxoid Ltd., Hampshire, England) and incubated at 35˚C for 24 hr for total aerobic bacterial counts and on MacConkey agar (Oxoid Ltd., Hampshire, England) with incubation at 35˚C and 42˚C for 24 hours for total coliform count. Total aerobic count indicates the quality and shelf life of the products and total coliform count indicates the unhygienic condition of the food preparation surfaces.

Escherichia coli, fecal coliform bacteria Twenty five (25) g of each sample were homogenized in 225 mL Enterobacteria enrichment brothMossel pre-enrichment medium (Oxoid Ltd., Hampshire, England) and incubated at 35˚C for 20 hours. One mL of pre-enriched cultures were mixed with nine mL of 2x EC medium (Nissui Co., Ltd., Tokyo, Japan) and incubated at 35˚C for 20 hours. To confirm the presence of fecal coliforms, one loopful of the culture was inoculated into 10 mL 1x EC medium with Durham fermentation tubes and incubated at

42˚C for 20 hours. Gas production in the tube indicates the presence of fecal coliforms. To isolate E. coli, one loopful of gas producing 1x EC culture broth was streaked on EMB agar plates (Nissui Co., Ltd., Tokyo, Japan) and the developed typical colonies were then confirmed using biochemical characterization (IMViC) and API 20E kit (bioMérieux, Durham, NC, USA). Presence of E. coli or fecal coliform bacteria was used as an indicator that the food is potentially contaminated with fecal material.

Escherichia coli O157, O111, O26 Twenty five (25) g of each samples were homogenized in 225 mL mEC medium (Nissui Co., Ltd., Tokyo, Japan) and incubated at 42˚C for 20 hours. The enriched cultures were streaked on Sorbitol MacConkey agar (Oxoid Ltd., Hampshire, England) supplemented with Cefixime and potassium tellurite admendments (Fluka, Sigma-Aldrich, Banglore, India) and characteristic colonies were subjected to biochemical tests (IMViC). Biochemically confirmed isolates were screened using Rainbow agar (Biolog, France) and CHROM agar (Kanto Co. Ltd., Kyoto, Japan). The colonies which gave the characteristic color were serotyped using O157, O111 and O26 specific antisera. The isolates were subsequently tested for the presence of stx1 and stx2 by NH-Immuno-

Figure 1b. Identification of selected Critical Sampling Locations (CSLs) in the production chain of dried lemongrass. S0: At the field; S1: weeks before harvest; S2: harvest and washing; S3: drying/sorting/grinding/ packaging;

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chromato VT1/2 and by polymerase chain reaction (PCR) assay using primer 5’-CAGTTAATGTGGTGGCGAAGG-3’ and 5’-CACCAGACAAATGTAACCGCTC-3’ for stx1 and 5’-ATCCTATTCCCGGGAGTTTACG-3’ and 5’-GCGTCATCGTATACACAGGAGC-3’ for stx2, respectively (Vidal et al., 2004).

Salmonella spp. Twenty five (25) g of each sample were homogenized in 225 mL of buffered peptone water (Merck, Darmstadt, Germany) and incubated at 35˚C for 20 hours. One mL pre-enrichment cultures was mixed with nine mL of Hanja Tetrathionate Broth (Eiken Chemical Co. Ltd., Tokyo, Japan) and incubated at 35˚C for 20 hrs and nine mL of Rappaport-Vassiliadis Broth (Eiken Chemical Co. Ltd., Tokyo, Japan) and incubated at 42˚C for 20 hrs. The broth the culture broths were subsequently streaked onto DHL and MLCB and characteristic colony were characterized with biochemical tests (TSI and LIM). Biochemically confirmed isolates were re-confirmed using Salmonella LA latex agglutination test and API 20E kits.

Hot water treatment in tea-cup & Enrichment study One gram of dried or blended herbs samples were added in a cup and 50 mL of hot water was poured over the dried leaves. The cup was kept with and without lid up to 5 minutes. In each 30 second interval microbiological parameters were done as described in the previous section on microbiological medium and conditions. For the enrichment study, one mL of hot water treated sample was added into 9 mL of Tryptic soy broth (TSB; Oxoid Ltd., Hampshire, England) medium and incubated at 37˚C for 6 hrs and then spread on to the selective medium of interest. If any bacteria survived or injured nonselective TSB medium was used to help resuscitate these cells and enable them to grow in selective microbiological medium.

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Statistical Analysis Three samples of each category were taken from the same farm. Reported plate count data represented in tables are the log10 mean values ± standard deviation of three individual trials, and each of these values were obtained from duplicate samples. Data were subjected to analysis of variance using the Microsoft Excel program (Redmond, Washington DC, USA). Significant differences in plate count data were established by the least-significant difference (P < 0.05) at the 5% level of significance.

RESULTS AND DISCUSSION The search for healthy, safe, and sustainable food production has increased the consumption of organic fresh produce. These products should be free of pesticide residues and other synthetic substances commonly used in conventional agriculture, such as soluble fertilizers (Oliveira et al, 2012). At the same time organic products have lower risks related to chemical contamination; however, several investigations have raised concerns related to the microbiological quality of these foods (Delaquis et al., 2007; Itohan et al., 2011). Among organic fresh produce, fresh and dried herbs stand out due to their flavors, aromas, colors and continual availability in the market as well as acceptability regardless of age or economic group of the human population worldwide (Esimone et al., 2003). Thirteen categories of herbs and tea including black tea, blend tea, neem blend herbs, neem tea, mint (fresh and dry), jasmine (fresh & dry), lemongrass and lemongrass blend tea were analyzed for total aerobic population (TAB), total coliform population (TCC) and presence of E. coli, E. coli O157:H7 and Salmonella spp. Table 1 presents the results of the distribution of natural aerobic population, coliform population and presence of E. coli, E. coli O157:H7 and Salmonella spp in different fresh and dry herbs; water and manure soil. Higher aerobic bacterial counts were recorded as 6.9 log CFU/g in liker base tea samples and the lowest aerobic

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Table 1. Distribution of natural aerobic population, coliform population and presence of E. coli, E. coli O157:H7 and Salmonella spp in different fresh and dry herbs; water and manure soila

Total Viable count (log CFU/g)

Total Coliform count (log CFU/g) )

Total E.coli Count (log CFU/g)

Black tea (Normal)

5.9 ± 0.08

6.9 ± 0.11

4.7± 0.14

3.8 ± 0.06

4.84

Blend tea

6.4 ± 0.11

5.9± 0.11

4.5 ±0.11

4.5 ± 0.12

5.01

Black tea (Original)

6.0± 0.14

6.0± 0.34

5.1±0.11

4.6± 0.11

5.40

Neem blend herbs

4.1 ± 0.15

ND*

4.98

Neem Tea

3.9 ± 0.12

5.04

Black tea (Premium)

5.4 ± 0.14

4.1 ± 0.21

4.0 ± 0.12

3.2 ± 0.11

5.10

liker base

6.9 ± 0.22

6.2 ± 0.22

5.7 ± 0.13

5.2 ± 0.15

7.13

Lemongrass

5.9 ± 0.11

5.8 ± 0.15

5.8 ± 0.23

4.8 ± 0.13

4.60

Lemongrass blended tea

5.5 ± 0.24

5.0 ± 0.09

4.7 ± 0.19

4.4 ± 0.11

3.7 ± 0.07

6.00

Jasmine fresh

5.7 ± 0.11

5.3 ± 0.12

5.1 ± 0.11

4.0 ± 0.12

1.0 ± 0.09

5.94

Jasmine dried

5.4 ± 0.13

5.2 ± 0.17

5.0 ± 0.11

3.9 ± 0.15

1.0 ± 0.12

5.94

Mint fresh

4.5 ± 0.12

4.4 ± 0.19

4.2 ± 0.12

3.2 ± 0.11

1.3 ± 0.14

5.99

Mint dried

4.4 ± 0.14

3.5 ± 0.23

3.4 ± 0.11

2.4 ± 0.12

5.94

Tap water

3.5 ±0.13

2.0 ± 0.14

1.8 ± 0.11

1.7± 0.13

ND*

6.60

Tank water

6.0 ±0.13

4.7 ± 0.09

3.9 ± 0.11

3.1 ± 0.11

7.50

Ground water

3.8 ± 0.13

3.4 ± 0.07

3.0 ± 0.11

2.1 ± 0.12

6.50

Manure soil

6.0 ± 0.14

5.8 ± 0.09

5.0 ± 0.11

4.7 ± 0.11

7.80

Herbs and tea Sample

E.coli O157:H7 Presence of counts Salmonella (log CFU/g) Spp.

*ND=Not detected; aResults are expressed in mean± standard deviation of three replicate samples, which are being calculated from duplicate plates.

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Table 2. Distribution of natural aerobic population, coliform population and presence of E. coli, E. coli O157:H7 and Salmonella spp at different steps of dried lemongrass productiona.

Lemongrass production & processing steps

Total aerobic counts (log CFU/g)

Total Coliform counts (log CFU/g)

Total E. coli counts (log CFU/g)

E. coli O157:H7 counts (log CFU/g)

Presence of Salmonella Spp.

At Harvest

5.9 ± 0.08

5.8± 0.18

4.8 ± 0.16

4.6

Cleaning & No washing

5.7 ± 0.12

5.4 ± 0.08

4.7 ± 0.08

3.7 ± 0.10

5.2

Cleaning & fresh water wash

5.0 ± 0.11

4.3± 0.16

4.3 ± 0.10

3.6± 0.20

5.1

Cleaning with fresh hot water

5.2 ± 0.14

4.5 ± 0.08

3.8± 0.19

3.5 ± 0.17

5.0

After dry heat at 900C for 20 min

4.4 ± 0.12

2.9 ± 0.12

2.8 ± 0.15

2.7 ± 0.12

5.2

After grinding at room temperature

4.2 ± 0.13

4.0 ± 0.09

3.7 ± 0.18

3.7 ± 0.16

5.4

After sorting at room temperature

5.6 ± 0.11

3.5± 0.20

3.3 ± 0.11

3.1 ± 0.11

5.3

After final streaming

4.4 ± 0.10

4.3 ± 0.08

2.3 ± 0.10

2.3 ± 0.12

5.4

*ND=Not detected; aResults are expressed in mean ± standard deviation of three replicate samples, which are being calculated from duplicate plates

counts were observed as 3.9 log CFU/g in neem tea samples (Table 1). Among the herb and tea sample tested, neem tea and neem blended herbs were determined to be microbiologically safe, because no coliform, fecal coliforms, E. coli, or Salmonella were recovered throughout the study. In contrast, jasmine, mint and tea blend with lemongrass were determined to be contaminated as the presence of E. coli O157:H7 and Salmonella was observed. The supply water used for irrigation, wash/rinse purposes, and compost used as fertilizer of soil were also analyzed.

Salmonella spp. was not detected in the manure sample tested (Table 1). In this study, water for irrigation and washing/rinse purpose was contaminated with E. coli O157H7, therefore it was concluded that there was a risk of contamination of final products. Foodborne outbreaks involving green vegetables contaminated by water have been reported in several studies around the world (Beuchat, 1996; Moyne, et al., 2011). Pathogenic bacteria such as E. coli O157:H7 are most often associated with outbreaks of waterborne diseases, resulting from inadequate

The water and composted manure was found to be heavily contaminated with enteric bacterial pathogens (Table 1). The total coliforms, E. coli and E. coli O157:H7 populations were enumerated as 5.0 log CFU/ml, 4.7 CFU/ml and 4.2 CFU/ml, respectively.

treatment of water used for irrigation and washing of fresh produce (Levantesi et al., 2012; Beraldo and Filho, 2011; Fischer-Arndt et al., 2010). Therefore, specific control measures should be developed in order to prevent final product contamination.

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Table 3. Recovery of natural aerobic population, coliform population and presence of E. coli, E. coli O157:H7 and Salmonella spp after corrective measures in processing and production of dried lemongrassa. Total aerobic count (log CFU/g)

Total Coliform count (log CFU/g)

Total E. coli Count (log CFU/g)

E.coli O157:H7 counts (log CFU/g)

Presence of Salmonella Spp.

Control

5.9 ± 0.08

5.8 ± 0.18

4.8 ± 0.16

4.6

Lemongrass (After corrective measures 1)

2.7 ± 0.18

ND*

5.7

0/3

Lemongrass (After corrective measures 2)

2.5 ± 0.12

5.8

0/3

Mint (After corrective measures 1)

2.7 ± 0.14

6.0

0/3

Mint (After corrective measures 2)

2.9 ± 0.11

1.3 ± 0.09

1.0 ± 0.11

5.6

1/3

Lemongrass production & processing steps

Positive / No of sample tested

*ND=Not detected; aResults are expressed in mean± standard deviation of three replicate samples, which are being calculated from duplicate plates. However when composted manure was analyzed, the presence of higher numbers of coliforms (5.8 log CFU/g), E. coli (5.0 log CFU/g), and E. coli O157:H7 (4.7 log CFU/g), were observed. Salmonella spp. was not detected in the compost samples (Table 1). Numerous published reports have indicated that the composting time and temperature of manure could effectively reduce microorganisms like E. coli, E. coli O157:H7, and Salmonella, which were routinely detected in fresh compost (James, 2006; MAFF, 2000; Millner, 2003; Johannessen, 2005). However, the organic fertilizer samples analyzed in the present study were above the detection limit (3.0 log MPN/g), indicating that the control of manure was not adequate. From the same farm, lemongrass production prac-

log CFU /g, 5.8 log CFU/g and 4.8 log CFU /g, respectively after harvest. However, Salmonella spp. was not detected in the lemongrass sample (Table 2). After harvest, the lemongrass sample was washed with water, in the hope of being able to remove the debris and to reduce the microbial load. Washing with tap water reduced the microbial load by 0.5-1.0 log CFU/g of bacteria. The lemongrass sample was subsequently dried in a fluid bed dryer for 20 minutes at a temperature recorded as 90˚C. After the drying process was completed, the microbial load was decreased substantially but not eliminated completely. Thereafter, grinding, sorting and packaging were done at the commercial settings. The contamination remaining was still evident after packaging and in the finished product. Numer-

tices were taken as a case study to determine the point of E. coli O157:H7 contamination and to take corrective measure in eliminating the risk. The average aerobic bacterial counts, coliform counts, E. coli and E. coli O157: H7 counts were recorded as 5.9 log CFU /g., 5.8

ous research reports have indicated that dry heating at 90˚C for 20 minutes is sufficient to eliminate the pathogen (Bari et al., 2009), however, in this study, pathogens were not eliminated completely. This finding suggested that heating temperature or the contact

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time may not be adequate to inactivate pathogens in lemongrass samples. After that an investigation of actual temperature and time inside the fluid bed dryer was conducted and it was discovered that the actual temperature at contact point was not homogeneous for 90˚C, and the contact time was only a few seconds because uniform conditions were not achieved by passing air through the lemongrass layer under controlled velocity conditions to create a fluidized state. Therefore, when the sample comes in contact with heat for few seconds, some bacteria may become injured and could resuscitate in between the cycles and in following steps, therefore, survive and subsequently be detected in the final products. Therefore, corrective

tice is consistent all over the world. If the herbs/tea leaves were contaminated with pathogens, whether or not hot water can reduce the risk of pathogen ingestion is a critical consideration. To solve this approach, an experiment was designed to determine the effectiveness of pouring hot water onto dried herbs/leaves in a cup for eliminating the risk of pathogen exposure. The results were presented in Table 4. Three different contaminated tea samples include black, blend and lemongrass tea were analyzed. One gram of each sample was placed in a tea-cup and 50 ml of hot water was added to each cup individually, either covered with a lid or without a lid, and held up to 5 min. At each 30 second time interval, microbiological popula-

actions of dryer temperature were undertaken and after these corrective measures, the same samples were dried in the same machine, analyzed and the results are presented in Table 3. It was found that drying at 90 ˚C for 20 min in an oven was enough to eliminate the pathogens even though the sample was contaminated initially (Table 3). To prevent further contamination, the workers was trained for personal hygiene and GAP, and hand gloves, mask, hairnet, apron, hand washing soap/ sanitizers, and a single used towel was provided, along with cleaning of utensils, machinery, and transport vehicles was conducted using steam. After these steps were taken, one batch of lemongrass was processed, dried and analyzed for pathogens. Neither generic E. coli nor any pathogenic E. coli O157:H7 were detected in the in the sample and the total viable bacteria and coliform population counts were found to be less than 100 CFU/ml, which is below the permissible limit (Table 3). These findings again showed that good hygiene practices are necessary for reducing foodborne pathogen contamination in the product. For the consumer, a common strategy to avoid foodborne disease is heating or cooking of potential risk products before consumption. However, this approach is not appropriate for the majority of fresh herbs and leafy greens that are mainly consumed raw, or

tion counts were enumerated and recorded. The hot water temperature was recorded as 82˚C when initially added in the cup and after 60 seconds the temperature was reduced to 78˚C; further reduced to 73˚C after 3 minutes holding time and at the end of 5 minutes the temperature decreased to 64˚C. It was determined that the initial viable bacterial counts were 5.4 log CFU/g, coliform counts were 4.1 log CFU/g, E. coli counts were 4.0 log CFU/g and E. coli O157:H7 counts were 3.2 log CFU/g in the blended tea samples, respectively (Table 4). After 30 seconds of treatment with hot water without a lid, a 2.0 log CFU/g reduction of viable bacterial counts was observed for the blended tea samples. The coliform bacteria, E. coli and E. coli O157:H7 counts were reduced to non-detectable levels within 30 seconds of hot water treatment despite the higher pathogen contamination levels in the initial samples. Similar experimental results were observed in black tea, and the lemongrass sample. The bacteria might be injured or severely injured when hot water was added in the cup and thus may not be able to grow in selective microbiological medium. To solve this issue, an enrichment study was done. No coliform, E. coli and E. coli O157:H7 were detected in the enrichment study after 30 seconds and above this holding time (Table 4). This finding suggested that the addition of hot water (82˚C)

added to food after the heat-treatment. For example, tea is usually made with dried tea leaves, or blended with other dried herbs and pouring the boiling water over the leaves and letting the combination remain for a few minutes and then consumed. This general prac-

in the tea leaves resulted in the reduction of pathogens below detection limits of the current study and thus the present tea making practice is potentially capable of providing safe tea for drinking even though the tea leaves were initially contaminated.

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<1.0 <1.0

3.2 ± 0.12

3.0 ± 0.11

3.0 ± 0.14

2.9 ± 0.09

3.2 ± 0.09

3.1± 0.08

5.9 ± 0.11

1.3 ± 0.14

1.3 ± 0.13

1.0 ± 0.09

1.0 ± 0.07 -

6.0 ± 0.15

3.1 ± 0.16

3.0 ± 0.12

3.0 ± 0.11

3.0 ± 0.07 -

120

150

180

5 min

Control

120

150

180

5 min

Control

120

150

180

5 min

<1.0

6.0 ± 0.12

<1.0

6.9 ± 0.12

<1.0

<1.0*

3.5 ± 0.18

4.1 ± 0.10

5.4 ± 0.08

Total Coliform count

Control

Total aerobic count

<1.0

5.1 ± 0.11

<1.0

4.7 ± 0.11

<1.0

4.0 ± 0.12

E. coli

<1.0

4.6 ± 0.14

<1.0

3.8 ± 0.16

<1.0

3.2 ± 0.11

E. coli O157:H7

Recovery of microorganisms (log CFU/g)a

Salmonella Spp.

Nil

Presence of Coliform

Nil

Presence of E.coli

Nil

Presence of E. coli O157:H7

After Enrichment

Nil

Presence of Salmonella Spp.

<1.0*= Less than detection limit; ND= Not Detected, Nil= Absent; aResults are expressed in mean± standard deviation of three replicate samples, which are being calculated from duplicate plates.

lemongrass Samples

Black Tea Samples

Blended Tea Samples

Sample type Hot water treatment time (Sec)

Table 4. Effectiveness of pouring hot water over contaminated dried (blend tea, black tea and lemongrass samples) leaves in a cup of tea at different holding time.

CONCLUSIONS In the present study it can be concluded that the organic fertilizer and the water used for irrigation and washing are critical sources of microbial contamination that need to be controlled in the production chain of organic produce. The contamination of manures also highlighted the need for a fertilizer control program in order to control the composting time and avoid the addition of fresh manure to the composted manure. Regarding the issues of irrigation and wash water, the results demonstrated the importance of using water from safe sources. It is also essential to emphasize the need for awareness and training to food handlers because even though organic vegetables may not be perceived as being chemically contaminated; nonetheless, they could very well be contaminated with pathogens and, for that reason, sanitization procedures should be developed to avoid foodborne illnesses. The use of a risk-based sampling plan in combination with corrective measures, personal hygiene and good agricultural practices (GAP) allowed us to produce safe organic herbs. This case study provides an overview of the organic farms’ status in northern Bangladesh, where good hygiene practice and GAP were introduced as a part of this study.

ACKNOWLEDGEMENTS The authors would like to thank Mr. Harun-ur Rashid and Mr. Abul Kalam Azad for the laboratory assistance required to complete this task. The authors would also like to thank the United Nations University, Tokyo, Japan (UNU-ISP) for financial support (FY 2013-2014) in this work.

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sprouts from retail establishments. Int. J. Food Microbiol. 123:121–129. Abba, D., H. I. Inabo, S. E. Yakubu, and O. S. Olonitola. 2009. Contamination of herbal medicinal products marketed in Kaduna Metropolis with selected pathogenic bacteria. Afric. J. Trad. Compl. Altern.Med. 6:70- 77. Adeleye, I. A., G. Okogi, and E. O. Ojo. 2005. Microbial contamination of herbal preparations in Lagos, Nigeria. J. Health Pop. Nutr. 23:296- 297. Bari, M. L., D. Nei, I. Sotome, Y. Nishina, S. Isobe, and S. Kawamoto. 2009. Effectiveness of sanitizers, dry heat, hot water and gas catalytic infrared heat treatments to inactivate Salmonella in raw almonds. Foodborne Pathog. Dis. 6:953958. Beuchat, L. R. 1996. Pathogenic microorganisms associated with fresh produce. J. Food Prot. 59:204–216. Beraldo, R. M., and F. A. Filho. 2011. Bacteriological quality of irrigation water from vegetable gardens in the municipalities of Araraquara, Boa Esperança do Sul e Ibitinga, SP. Alimentos e Nutrição Araraquara 22:1640. Blumenthal, M. 1998. The Complete German Commission E Monographs . Austin TX: American Botanical Council. 341-342. Delaquis, P., S. Bach, and L. D. Dinu. 2007. Behavior of Escherichia coli O157:H7 in leafy vegetables. J. Food Prot. 70:1966–1974. Esimone, C. O., P. O. Oleghe, and E. C. Ibezim. 2003. Effect of preservation agents on the microbial stability of some indigenous herbal preparations. Niger J. Pharm. 34:37-42. Espen Rimstad, E. A. Høiby, G. Kapperud, J. Lassen, B. Tore Lunestad, et al. 2008. Norwegian Scientific Committee for Food Safety Panel on Biological Hazards final report on Risk assessment of import and dissemination of intestinal pathogenic bacteria via fresh herbs and leafy vegetables from South-East Asia. ISBN: 978-828082-244-4 ; 07/111-final report, pages 2-32. FAO/WHO. 2008. Microbiological hazards in fresh fruits and vegetables. Meeting Report. Microbiological risk assessment series. Available at the

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following URL and accessed on August 7, 2013. (http://www.fao.org/ag/agn/agns/jemra_riskassessment_freshproduce_en.asp) Fischer-Arndt, M., D. Neuhoff, L. Tamm, and U. Köpke. 2010. Effects of weed management practices on enteric pathogen transfer into lettuce (Lactucasativa var. capitata). Food Control 21:1004–1010. Girón, L. M., Freire, V., Alonzo, A., and Cáceres, A. 1991. Ethnobotanical survey of the medicinal flora used by the Caribs of Guatemala. J. Ethnopharmacol. 34:173–187. Ilic, S., A. Rajic, C. Britton, E. Grasso, W. Wilkens, S. Totton, and J. LeJeune. 2012. A scoping study characterizing prevalence, risk factor and intervention research, published between 1990 and 2010, for microbial hazards in leafy green vegetables. Food Control 23:7-19. Itohan, A. M., O. Peters, and I. Kolo. 2011. Bacterial contaminants of salad vegetables in Abuja Municipal Area Concil. Nigéria. Malaysian J. Microbiol. 7:111. James, J. 2006. Microbial hazard identification in fresh fruits and vegetables. John Wiley & Sons, Hoboken, NJ. USA. p312. Johannessen, G. S. 2005. Use of manure in production of organic lettuce: Risk of transmission of pathogenic bacteria and bacteriological quality of the lettuce. Oslo, Belgic: Norwegian School of Veterinary Science. Lau, A., M. J. Holrnes, S. Woo, and H. Koh. 2003. Analysis of adulterants in a traditional herbal medicinal products using liquid chromatography- mass spectroscopy. J. Pharm. Bio Ana. 31: 401-406. Leung, A. Y. 1980. Encyclopedia of Common Natural Ingredients Used in Food, Drugs, and Cosmetics. New York, NY: Wiley. Lewinsohn, E., N. Dudai, Y. Tadmor, I. Katzir, U. Ravid, E. Putievsky, and D. M. Joel. 1998. Histochemical localization of citral accumulation in lemongrass leaves (Cymbopogon citratus (DC.) Stapf., Poaceae) Ann. Bot. 81:35–39. Levantesi, C., L. Bonadonna, R. Briancesco, E. Grohmann, S. Toze, and V. Tandoi. 2012. Salmo-

nella in surface and drinking water: occurrence and water mediated transmission. Food Res. Int. 45:587–602. MAFF (The Ministry of Agriculture Food and Fisheries). 2000. A study of on-farm manure applications to agricultural land and an assessment of the risks of pathogen transfer into the food chain. (Project Number FS2526). Retrieved from http://www.safeproduce.eu/Pics/FS2526.pdf Millner, P. 2003. Composting: Improving on a timetested technique. Agricultural Research, 51(8). Retrieved from http://www.ars.usda.gov/is/AR/ archive/aug03/time0803.pdf Martínez-de la Puente, J., S. Merino, E. Lobato, J. Rivero-de Aguilar, S. del Cerro, and R. Ruizde-Castañeda. 2009. Testing the use of a citronella-based repellent as an effective method to reduce the prevalence and abundance of biting flies in avian nests. Parasitol Res. 104:12331236. Moyne, A. L., M. R. Sudarshana, T. Blessington, S. T. Koike, M. D. Cahn, and L. J. Harris. 2011. Fate of Escherichia coli O157:H7 in field-inoculated lettuce. Food Microbiol. 28:1417-1425. Oyetayo, V. O. 2008. Microbial load and Microbial property of two Nigeria herbal remedies. African Journal of Traditional, Complimentary and Alternative Medicines. Vol. 5:74- 78. Olaimat, A. N., and R. A. Holley. 2012. Factors influencing the microbial safety of fresh produce: a review. J. Food Prot. 32:1-19. Oliveira, A. B. A., A. C. Ritter, E. C. Tondo, and M. R. de I. Cardoso. 2012. Comparison of different Washing and disinfection protocols used by Food Services in Southern Brazil for Lettuce (Lactuca sativa). Food Nutr. Sci. 3:28-33. Tawatsin, A., S. D. Wratten, R. R. Scott, U. Thavara, Y. Techadamrongsin. 2001. Repellency of volatile oils from plants against three mosquito vectors. J Vector Ecol. 26:76-82. USDA. 2008. NRCS. Cymbopogon citratus (DC. ex Nees) Stapf. The PLANTS Database. ( http:// plants.usda.gov, 10 December 2008). National Plant Data Center, Baton Rouge, LA 70874–4490 USA.

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Vidal, R., M. Vidal, R. Lagos, M. Levine, and V. Prado. 2004. Multiplex PCR for diagnosis of enteric infections associated with diarrheagenic Escherichia coli. J. Clin. Microbiol. 42: 1787 Wong, K. K., F. A. Signal, S. H. Campion, and R. L. Motion. 2005. Citronella as an insect repellent in food packaging. J. Agric. Food Chem. 53:46334636. Warnke, P. H., S. T. Becker, R. Podschun, S. Sivananthan, I. N. Springer, P. A. Russo, J. Wiltfang, H. Fickenscher, and E. Sherry. 2009. The battle against multi-resistant strains: Renaissance of antimicrobial essential oils as a promising force to fight hospital-acquired infections. J. Craniomaxillofac Surg. 37:392-397. Wright, S. C., J. E. Maree, and M. Sibanyoni. 2009. Treatment of oral thrush in HIV/AIDS patients with lemon juice and lemon grass (Cymbopogon citratus) and gentian violet. Phytomed.16:118-24.

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Batch Culture Characterization of Acetogenesis in Ruminal Contents: Influence of Acetogen Inocula Concentration and Addition of 2-Bromoethanesulfonic Acid P. Boccazzi 1,2 and J. A. Patterson1 1

Department of Animal Sciences, Purdue University. West Lafayette, IN 47907 2 Current address: 147 Kelton St., Allston, MA 02134

ABSTRACT Interspecies H2 transfer is a syntrophic interaction between H2-producing and H2-consuming organisms, that plays an important role in regulating ruminal fermentation as well as other ecosystems. Any decrease in hydrogen concentration, due to interspecies hydrogen transfer can influence volatile fatty acid fermentation patterns of many ruminal microorganisms. In the rumen, methanogens consume hydrogen to generate energy and thus serve as a hydrogen sink. However energy is lost due to eructation of methane which can not be used by the ruminant animal. Alternative hydrogen consuming organisms, such as acetogens, could be an attractive alternative hydrogen sink in rumen ecosystems because they generate actetate from hydrogen and carbon dioxide, which can be used by the host animal. However, this would require inhibiting methanogenic activity. Therefore, batch cultures were used to study acetogenesis as a functional alternative to methanogenesis in the rumen in the presence of a methanognesis inhibitor. In batch culture experiments, acetogen strains G1.5a, G2.4a, G3.2a, A10, and 3H were able to reduce H2 concentrations in ruminal contents in the presence of bromoethanesulfonic acid, an inhibitor of methanogenesis. Batch culture studies indicated that acetogens could function as an alternative electron sink to methanogens under some conditions. Keywords: Acetogen, Acetogenesis, H2, Methane, Ruminal

INTRODUCTION The productivity of ruminant domestic animals is influenced, to a large extent, by the efficiency of microbial fermentation of feedstuffs in the rumen. During rumen fermentation, complex carbohydrates Correspondence: John Patterson, jpatters@purdue.edu Tel: +1-765-494-4826 Fax: +1-765-494-9347

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(e.g., cellulose) are degraded to monomeric carbohydrates (e.g., glucose) which are primarily fermented to pyruvate via the Embden-Meyerhof-Parnas pathway (Ricke et al., 1996; Weimer, 1992; Weimer et al., 2009). Pyruvate is subsequently metabolized to volatile fatty acids (VFA; acetate, propionate, and butyrate), CO2, H2, and microbial cells. While fermentation acids provide 60 to 80% of the daily metabolizable energy intake of ruminants (Annison and

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Armstrong, 1970), microbial cells provide an important source of amino acids, vitamins, and cofactors (Hungate, 1966). Interspecies H2 transfer is a syntrophic interaction between H2-producing and H2-consuming organisms that plays an important role in regulating ruminal fermentation as well as fermentations in other anaerobic ecosystems (McInerney et al., 2011). Hydrogen produced by fermentative microorganisms is consumed by H2 utilizing microorganisms, namely, methanogens, sulfidogens, and acetogens (Lupton and Zeikus, 1984; Drake, 1992, 1994; Saengkerdsub and Ricke, 2014). The decrease in H2 concentration, due to interspecies H2 transfer, influences the VFA

major method used to manipulate rumen fermentation has been the use of ionophore antibiotics such as monensin and lasalocid. These compounds improve the efficiency of animal production by decreasing methane production and increasing ruminal propionate concentration by 15%. Methane production decreases primarily because monensin inhibits H2 producing microorganisms, therefore decreasing the amount of H2 available for methanogenesis. Chemolithoautotrophic acetogenic bacteria achieve reductive acetogenesis, utilizing CO2 and H2 as their sole carbon and energy source, respectively, fixing CO2 into acetate (Ragsdale, 1991). Acetogenesis has been demonstrated to be the predominant

fermentation patterns of many ruminal microorganisms because hydrogen is an end product inhibitor of the hydrogenase enzyme (Wolin and Miller, 1983; Ricke et al., 1996; Weimer et al., 2009). When H2 concentrations are high, pyruvate is utilized as a reducing equivalent acceptor and more reduced fermentation products (e.g., propionate, lactate, and ethanol) are produced (Wolin and Miller, 1983). When H2 concentrations are low, there is an increase in acetate and ATP production that could be converted into an increase in overall microbial cell yields (Wolin and Miller, 1983). The energy present in methane escapes the rumen through eructation and is lost to the animal. Because energy lost as methane has been estimated to be 2.4 to 7.4% of the gross energy intake (Branine and Johnson 1990) or 10 to 15% of the apparent digestible energy of the diet of ruminants (Blaxter and Clapperton, 1965), there has been an interest to specifically inhibit methanogenesis to enhance animal productivity. Direct inhibition of methanogenesis, however, also results in loss of energy in the form of H2, and reduction in production of microbial proteins (Chalupa, 1980). Maintaining the beneficial effects of interspecies H2 transfer while minimizing loss of energy as meth-

fate of H2 in some humans, swine, xylophagus termites, cockroaches and rats (Breznak and Blum, 1991; Lajoie et al., 1988). Replacing methanogenesis with acetogenesis in the rumen may have potential in decreasing energy losses in ruminants. Peptostreptococcus productus (Bryant et al., 1958), Eubacterium limosum (Genthner, 1981), and Acetitomaculum ruminis (Greening and Leedle, 1989) are chemolithoautotrophic acetogenic bacteria that have been isolated from the bovine rumen. However they are not considered the primary H2 consuming organisms in this environment, since their numbers are consistently lower than methanogens. Factors dictating whether acetogenesis or methanogenesis will predominate in anaerobic environments are not well understood. Breznak and Kane (1990) suggested several possible factors that may influence the competitiveness of acetogens with methanogens. One factor is that methanogenesis has a higher energy yield than acetogenesis (Breznak and Blum, 1991). Another important factor is that methanogens have a higher affinity for H2 than acetogens. The normal rumen hydrogen concentration is between 10-5 and 10-6 atm (Czerkawski and Breckenridge, 1971; Robinson et al., 1981). Ruminal methanogens

ane could enhance energy provided to ruminants by 22% (Schaefer, D., personal communication). However, an alternative electron sink is required to trap electrons into a form utilizable by the animal if methanogens are to be directly inhibited. To date the

have an affinity for H2 between 1 and 4x10-6 atm (Greening et al., 1989). Different acetogenic isolates have been shown to have affinities for H2 between 10-4 and 10-5 atm (Greening et al., 1989; LeVan etal., 1998). In general, methanogens have been found

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to have H2 thresholds 10 to 40 fold lower than acetogens (Greening et al., 1989; Breznak and Blum, 1991). However, in our laboratory, acetogens with H2 thresholds only 2 to 4 fold higher than those of methanogens were isolated from ruminal contents using a hydrogen limited continuous culture system (Boccazzi and Patterson, 2011). Methane contributes roughly 25% of the global green house warming and is considered the second most important “greenhouse gas” after CO2 (Tyler, 1991). Atmospheric CH4 is presently increasing by 1% per year and it has reached a concentration unprecedented in the past 160,000 years (Pearman and Fraser, 1988). Ruminal and other gastrointestinal fermentations account for some 14% of the total CH4 emissions amounting to 70 to 100 Tg per year (EPA, 1993, Moss, et al., 2000). Limiting CH4 emissions from livestock and livestock waste while maintaining interspecies H2 transfer would improve ruminant productivity and at the same time would be beneficial for the environment. Certainly acetogens offer that possibility, but rumen ecosystem conditions would need to be designed that favor not only their presence but their acetogenic activities. One approach is to administer a methane inhibitor such as BES (Immig et al. 1996), but before this can attempted in a practical application in vitro screening needs to be done to confirm that this will generally be effective and which acetogens are the best candidates. Batch culture growth experiments, while not necessarily representative of the rumen from a passage rate and rumen turnover standpoint, still offer a means to rapidly screen multiple bacterial isolate responses and have been used for a wide range of physiological studies on rumen bacteria including rumen acetogens (Russell and Baldwin, 1978; Schaefer et al., 1980; Ricke and Schaefer, 1991, 1996; Ricke et al., 1994; Jiang et al., 2012; Pinder and Patterson, 2012, 2013). The objective of this study was to conduct batch culture screening experiments to determine the feasibility of a functional replacement of methanogenesis with reductive acetogenesis in ruminal contents in the presence of a methanogen inhibitor.

MATERIALS AND METHODS Source of Organisms Acetobacterium woodii (ATCC 29683) was obtained from the American Type Culture Collection (Rockville, MD). Acetogenic bacterial strains 3H, G1.5a, G1.5e, G2.4a, G3.2a and A10 were isolated and characterized in our lab and reported previously (Boccazzi and Patterson, 2011, 2013; Pinder and Patterson, 2011, 2012, 2013; Jiang et al., 2012).

Media and Growth Conditions Growth and H2 threshold experiments were conducted with a basal rumen fluid based acetogen medium (Table 1) or with Mac-20 medium containing casein hydrolysate and no rumen fluid (Table 1). Both media were prepared as described in Table 1 with the anaerobic techniques of Hungate (1966) as modified by Bryant (1972) and Balch and Wolfe (1976). The Mac-20 medium was only used with cultures of Acetobacterium woodii. The prepared medium was dispensed anaerobically into 60 ml serum bottles (West Company, Phoenixville, PA) in an anaerobic glove box (Coy Laboratories, Ann Arbor, MI) containing a H2:CO2 (5:95) gas phase. Serum tubes and bottles were sealed with butyl rubber serum stoppers and aluminum seals (Bellco Inc., Vineland, NJ). All stock solutions utilized to formulate media were prepared anaerobically by boiling and cooling distilled water under a CO2 gas phase and sterilized either by autoclaving or by injecting the solution through a 0.2 μm filter (Nalgene, Nalge Company, Rochster, NY). For chemolithoautotrophic growth in broth medium, bacterial cultures were grown in serum bottles closed with butyl rubber stoppers and aluminum seals. After medium sterilization, cooling and inoculation, the bottles were flushed for 30 sec with an appropriate gas mixture by inserting both a sterile gassing and a release needle through the serum stoppers and then bottles were pressurized to 200 kPa by removing the release needle. Oxygen traces were removed from gas mixtures by passing the gas through a reduced

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Table 1. Media compositiona Acetogen Medium

MAC-20 Mediumb

(amounts per liter)

Rumen Fluid

50.0 ml

---

Mineral 1c

40.0 ml

Mineral 2d

40.0 ml

Additional Trace Min. Sol.e

10.0 ml

Wolfe’s Trace Min. Sol.f

10.0 ml

Vitamin Solutiong

10.0 ml

Na2CO3

4.0000 g

Yeast Extract

0.5400 g

2.0000 g

Casein Hydrolysate

---

1.0000 g

Betaine

---

1.0000 g

NH4Cl

0.5400 g

Cysteine.HCl

0.5000 g

Resazurin solution

0.0010 g

Hemin solution

0.0001 g

All components, except Na2CO3 and cysteine. HCl were dissolved in distilled water and brought to a volume to 1000 ml. These were mixed thoroughly and the pH was adjusted to 7.0 with 1 M NaOH followed by gentle heating to a boil for 1 min. Na2CO3 was added and the solution was cooled rapidly to 25°C under 100% CO2. Cysteine.HCl was added, mixed thoroughly and autoclaved anaerobically for 12 min at 121°C and 15 psi a

Modification of AC-19 medium by Breznak et al. (1988)

Mineral 1 (g/liter): 6.00 K2HPO4

Mineral 2 (g/liter): 12.00 NaCl, 6.00 K2HPO4, 6.00 (NH4)2SO4, 2.45 MgSO4.7H20, 1.60 CaCl2.2H2O

Additional Trace Mineral Solution (g/Liter): 0.10 NiCl2.6H2O, 0.01 H2SeO3

Wolfe’s Trace Mineral Solution (g/liter): 3.00 Mg SO4.7H20, 1.00 NaCl, 0.50 MnSO4.H20, 0.10 CoCl2.6H20, 0.10 FeSO4.7H20, 0.10 CaCl2.2H20, 0.18 CoSO4.6H20, 0.19 ZnSO4.7H20, 0.02 AlK(SO4)2.12H20, 0.01 CuSO4.5H20, 0.01Na2MoO4.2H20 f

Vitamin Solution (g/liter): 0.10 pyridoxine.HCl, 0.056 ascorbic acid, 0.05 choline chloride, 0.05 thiamine.HCl, 0.05 D,L-6,8-thioctic acid, 0.05b riboflavin, 0.05 D-calcium panthotenic acid, 0.05 p-amino benzoic acid, 0.05 niacinamide, 0.05 nicotinic acid, 0.05 pyridoxal.HCl, 0.05 pyridoxamine, 0.05 myo-inositol, 0.02 biotin, 0.02 folic acid, 0.001 cynocobalamin g

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Figure 1. (A) Anaerobic incubation vessels made from one gallon glass screw cap storage jars with aluminum lids, the jars were flushed with CO2 and the environment reduced with BBL Gaspak Plus Anaerobic System Envelopes with Catalyst (H2 + CO2) (Becton Dickinson & Co, Sparks, MD), once the lids were closed, they were sealed with plastacine modeling clay. These vessels will not withstand pressurization, but that could be offset with inserting swagelok fittings for gas chromatography and a septa into the lid and flushing with fresh gas on a routine basis (B). A B

copper column. Pressurized bottles, unless otherwise specified, were incubated on their side on a rotatory shaker (New Brunswick Scientific Co. Inc., Model M52) operating at 200 rpm. For growth on solid medium, 60 mm disposable petri plates were incubated in an anaerobic growth vessel (made by the Agricultural and Biological Systems Department. Purdue University, IN) able to withstand high gas pressures. Prior to incubation the container was flushed for 2 min and then pressurized to 110.34 kPa with gas mixtures specified in the text for each experiment. More recently we have used a less expensive approach using one gallon glass screw cap storage jars with aluminum lids, the jars were flushed with CO2 and the environment reduced with BBL Gaspak Plus Anaerobic System Envelopes with Catalyst (H2 + CO2) (Becton Dickinson & Co, Sparks, MD), once the lids were closed, they were sealed with plastacine modeling clay (Figure 1). These vessels will not withstand pressurization, but that could be offset with inserting swagelok fittings and a septa into the lid and flushing with fresh gas on a routine basis.

General experimental procedure Serum bottles (60 ml) were anaerobically filled with 0.35 g alfalfa, 6 ml ADS (Table 2), 4 ml ruminal contents, 4 ml of an acetogen culture (inoculum) or acetogen medium (control), and 1 ml stock solution of 2-bromoethanesolfonic acid (BES, Sigma Aldrich) was added to provide a final concentration of 5 mM. Control cultures received 1 ml sterile anaerobic water. The alfalfa was dried at 60°C and ground through a 1 mm screen. Ruminal contents were collected from a Holstein Friesian dairy cow, fed a 57:43 concentrate:forage diet, prior to morning feeding and immersed in ice during transportation to the lab, where the rumen contents were filtered through a double layer of cheesecloth under a stream of CO2 and anaerobically inoculated into the serum bottles. Serum bottles were anaerobically sealed and incubated in a rotatory shaker at 37°C operating at 200 rpm.

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Table 2. Anaerobic dilution solution (ADS) compositiona

Component

(amounts per liter)

Mineral 1b

75.0 ml

Mineral 2c Cysteine.HCl

75.0 ml 0.5000 g

Resazurin solution

0.0010 g

All components, except cysteine.HCl were added to distilled water and the volume brought to 1000 ml. These were mixed thoroughly and the pH adjusted to 7.0 with 1 M NaOH. These were gently heated and brought to a boil for 1 min. Cysteine.HCl under 100% CO2 was mixed thoroughly and autoclaved anaerobically for 12 min at 121°C and 15 psi a

Mineral 1 (g/liter): 6.00 K2HPO4

Mineral 2 (g/liter): 12.00 NaCl, 6.00 KH2PO4, 6.00(NH4)2SO4, 2.45 MgSO4.7H20, 1.60 CaCl2.2H2O

Acetogenic bacteria inoculum preparation

Effect of bromoethane sulfonic acid on H2 and CH4 production in ruminal contents

Enumeration of acetogen strains to prepare inoculum was done by growing cultures in acetogen medium plus 5 mM glucose under 200 kPa of a H2:CO2 (80:20) gas mixture for 36 h at 37°C. Cultures were then brought into a glove box where they were serially diluted to 10-8 with ADS (Table 2). Each dilution was plated (25 μl) on acetogen medium containing 2% (w/v) agar (Bacto-Agar, Difco, Fisher) and 5 mM glucose in triplicate 60 mm petri plates. Plates were incubated, at 37°C for 3 days, in an anaerobic growth vessel pressurized to 16 psi with a H2:CO2 (80:20) gas mixture. The bacterial concentration was calculated by counting colony-forming units (CFU) per ml of culture. To make the acetogen inoculum for experiments, the acetogens were grown under the same growth conditions that were used for enumeration and diluted to the appropriate concentration. Cultures were centrifuged anaerobically at 8,000 rpm for 15 minutes and resuspended in acetogen medium

To test BES as a methanogenesis inhibitor in ruminal contents with or without the addition of acetogenic bacteria, duplicate serum bottles (60 ml) were anaerobically filled with 0.15 g alfalfa, 6 ml of fresh ruminal contents, 4 ml of acetogen medium (Table 1), 4 ml of an acetogen culture, 1 ml of a BES. BES was added as sterile stock solution to the acetogen medium to reach final concentrations ranging between 0.0 to 10 mM, respectively. Both the 5mM and 10 mM BES effectively inhibited methanogenesis and only the 10 mM data is shown. Because 5 mM BES was as effective as 10 mM BES, the lower concentration was used in all other experiments. Controls that did not receive a methanogen inhibitor received an equal volume of acetogen medium instead. Acetogenic bacteria utilized were A. woodii, and strains A10 and G3.2a. Acetogen inocula were prepared as described previously to give a final concentration of 5x108 CFU/ml. Duplicate serum bottles

to reach the CFU/ml desired. The titer for each acetogen strain used for methanogen replacement studies was determined before each experiment.

for each treatment were anaerobically sealed, and incubated in a rotatory shaker at 37°C operating at 200 rpm. Headspace gas volume, and H2 and CH4 concentrations were measured at 0, 12, and 36 h of incubation.

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Effect of acetogenic bacteria on H2 utilization in ruminal contents In an initial experiment using the acetogen strain 3H to determine efficiency of H2 utilization, either 3H or acetogen medium was added to serum bottles containing ruminal contents as described above. Strain 3H was added to the experimental medium to reach the initial concentration of 5x108 CFU/ml. Cultures were incubated at 37°C in a rotatory shaker, operating at 200 rpm, for 72 h. Headspace gas volume and H2 concentrations were measured from a triplicate set of serum bottles for each treatment at 0, 12, 24, 36, 48, 60, and 72 h. Volatile fatty acid profiles of cultures were also determined at 12, 48 and 72 h by gas chromatography. Similar procedures were used to determine efficiency of acetogen strains G1.5a, G1.5e, G2.4a, and G3.2a except 40 mM of MES (final concentration) was added for additional buffering. A control treatment receiving sterile anaerobic water instead of BES was also added. Final concentrations of acetogens were 5x108 CFU/ml. Headspace gas volume and H2 and CH4 concentrations were measured from duplicate serum bottles for each treatment at 0, 12, 24, and 72 h. The experimental procedure was slightly modified to determine the effect of acetogen dose on H2 concentrations. The following three modifications were made: 0.15 g alfalfa was used instead of 0.35 g, 40 and 3 mM (final concentrations) of MES and K2CO3, respectively, were added to the experimental medium, and a control treatment received sterile anaerobic water instead of BES. In separate experiments, the acetogen strain G2.4a was added to provide final concentrations of 5x107 and 1x108 CFU/ml. Headspace gas volume and H2 and CH4 concentrations were measured from a duplicate set of serum bottles for each treatment at 0, 12, 24, and 72 h. The acetogen strain G3.2a was added to provide final

Analytical Methods Bacterial growth: optical density was measured at 660nm using a Spectronic 70 spectrophotometer (Bausch and Lomb, Rochester, NY). VFA analysis: volatile fatty acid concentrations were measured by gas-liquid chromatography (GLC; Holdeman et al., 1977). At sampling time, samples were acidified by adding 20% (v/v) of meta-phosphoric acid (25% w/v) and then frozen. Samples to be analyzed were thawed, centrifuged at 26,892 x g for 5 min, and the supernatant was analyzed. A 0.92 meter long column, packed with SP1220 (Supelco, Bellefonte, PA, USA), was used in a Hewlett Packard 5890 GLC equipped with a flame ionization detector. Oven temperature was 130°C (isothermal), injector temperature was 170°C, detector temperature was 180°C, the carrier gas was N2 flowing at a rate of 30 ml per minute. Gas Analysis: for the measurements of hydrogen and methane concentrations, gas samples were analyzed using a Varian 3700 Gas Chromatograph equipped with a thermal conductivity detector, and a 1.83 meter silica gel column (Supelco). Temperatures of the injector, oven, and detector were room temperature, 130°C, and 120°C respectively. The carrier gas was N2 flowing at a rate of 30 ml per minute. The volume of gas injected for standards and samples was 0.5 ml. The GC was standardized with 5 different concentrations of H2 (400 to 25,000 ppm) and CH4 (900 to 32,000 ppm). A regression line was obtained from the output values of the standard concentrations. The regression line was then utilized to calculate H2 and CH4 concentrations in experimental samples. All gas mixtures were purchased from Airco (Indianapolis, IN).

concentration of 5x108, 1x109, and 5x109 CFU/ml. Headspace gas volume and H2 and CH4 concentrations were measured from duplicate serum bottles for each treatment at 0, 12, and 36 h.

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Figure 2. Effect of 10 mM of BES and acetogen strains Acetobacterium woodii (Aw), A10 and G3.2a on CH4 production (Fig. A) and H2 utilization (Fig. B) by ruminal contents (C). Duplicate serum bottles (60 ml), containing 6 ml of ruminal contents, received 0.15 g of alfalfa and were incubated at 37°C. Cultures were incubated in duplicate and one standard deviation is depicted with error bars.

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RESULTS

The effect of 0.0 to 10 mM of BES on methanogenesis and H2 utilization was determined in batch culture and data for 10 mM BES is shown in Figure 2A and B. Methanogenesis was totally inhibited in all treatments that received 5 and 10 mM BES, but not with lower doses of BES (data not shown). After 12 h of incubation, cultures receiving the acetogens G3.2a and A10 had lower hydrogen concentrations

plus BES cultures at 12 h, but H2 concentrations did not decline as rapidly over time (Figure 2B), suggesting that A. woodii is not as effective at utilizing H2 as the other isolates. The efficacy of acetogen strain 3H to utilize H2 was determined by adding strain 3H to ruminal contents containing 5 mM BES to inhibit methanogenesis. After 12 and 24 h of incubation, the H2 concentration of the 3H treatment was about half of that of the control treatment (Figure 3). However, after 36 h of incubation the control treatment had similar H2 concentrations as the 3H treatment (Figure 3). While the 3H cultures had the highest acetate concentration after 12 h of incubation, by 72 h, acetate concentra-

than either A. woodii or the control plus BES cultures (Figure 5B). Hydrogen concentrations for cultures receiving A10 and G3.2a were below 18 Îźmoles by 36 h (Figure 2B). Hydrogen concentrations for cultures receiving A. woodii were similar to the control

tions were similar for both treatments (Figure 3). An additional experiment was conducted to determine how well 4 acetogen isolates could replace methanogens as a hydrogen sink in ruminal contents. Methane was produced only in the control

Functional Replacement of Methanogenesis with Acetogenesis in Ruminal Contents

Figure 3. Acetate production (bars) and H2 utilization (lines) by ruminal contents with (3H) or without (C) the addition of 5x108 CFU/ml (final concentration) of the acetogen strain 3H. Methanogenesis was inhibited by 5 mM BES in all treatments. Cultures were incubated in triplicate and one standard deviation is depicted with error bars.

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Figure 4. CH4 production (Fig. A) and H2 utilization (Fig. B) by ruminal contents after the addition of 0.35 g of alfalfa and with or without (C) the addition of 5x108 CFU/ml (final concentration) of acetogenic isolates G1.5a, G1.5e, G2.4a and G3.2a. Methanogens were inhibited by 5 mM BES in all treatments but C. Cultures were incubated in duplicate and one standard deviation is depicted with error bars.

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Figure 5. Effect of inoculum size of the acetogen G2.4a (a= 5x107 and b= 1x108 CFU/ml, final concentration) on CH4 production (Fig. A) and H2 utilization (Fig. B) in ruminal contents (C) after the addition of 0.35 g of alfalfa and in the presence (C) or absence (C – BES) of BES (5 mM) as an inhibitor of methanogenesis. Cultures were incubated in duplicate and one standard deviation is depicted with error bars.

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Figure 6. Effect of inoculum size of the acetogen G3.2a (a=5x108, b=1x109 and c=5x109 CFU/ml, final concentration) on CH4 production (Fig. A) and H2 utilization (Fig. B) in ruminal contents (C) after the addition of 0.35 g of alfalfa and in presence (C) or absence (C - BES) of BES (5 mM) as an inhibitor of methanogenesis. Cultures were incubated in duplicate and one standard deviation is depicted with error bars.

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culture where no BES was added (Figure 4A). The acetogen isolate G2.4a had the lowest H2 concentration, followed by G3.2a, G1.5a, and G1.5e after 12 h of incubation (Figure 4B). After 72 h of incubation, of the acetogen tested, G1.5e, and G2.4a had the highest (183.2 μmoles) and lowest (93.8 μmoles) H2 concentrations, respectively, and strain G1.5a had a slightly lower H2 concentration than G3.2a (Figure 4 B). Although, the control plus BES culture had the highest H2 concentration (332.2 μmoles) at all times, H2 concentrations for this treatment decreased between 24 and 72 h of incubation (Figure 4B), indicating that ruminal contents have some capability to utilize H2.

DISCUSSION

Dose response experiments were performed to determine the optimal acetogen dose for the functional replacement of methanogenesis in ruminal contents. Methane production was inhibited in all cultures receiving BES, and remained below 20 μmoles for up to 72 h (Figure 5A). Doubling the dose of the acetogen G2.4a, from 5x107 to 1x108 CFU/ml, increased H2 utilization (Figure 5B). Hydrogen concentrations for cultures containing G2.4a were lower than the control without BES at 12 h, but not after 24 h. Cultures containing the highest dose of G2.4a had a H2 concentration approximately fourfold lower than the control plus BES (Figure 4B) after 12 h of incubation. The positive effect of increasing acetogen numbers persisted even after 72 h of incubation (Figure 4 and 5). However, H2 concentrations of control plus BES cultures declined to a level similar to that of the highest dose of G2.4a added after 72 h. Similar experiments were conducted with acetogen isolate G3.2a with dosages increasing between 5x108 to 5x109 CFU/ml. Methane production occurred only in the control cultures in which BES was not added (Figure 6A). Hydrogen concentrations for all the levels of G3.2a remained below 15 μmoles for 36 h (Figure 6B). Hydrogen concentrations for control, and control plus BES cultures increased to

significantly decreased H2 concentrations at 12 hr, compared to the 5mM BES control and Acetobacterium woodii. H2 concentration decreased by 40% in the controls, indicating that there is some ability to utilize H2 in ruminal contents. However, H2 concentrations in the acetogen inoculated cultures were significantly lower than that of the control cultures. Thus, acetogens used in this study have the ability to reduce H2 concentrations during rapid fermentation and isolate G3.2a has a greater hydrogen utilization potential than isolate A10 or Acetobacterium woodii (Fig 2). Acetogenic isolate 3H significantly reduced H2 concentrations over the first 24 hours of incubation compared to the BES treated control culture and also produced more acetate during this time (Fig 3). Additon of 5 x 108 cfu /ml of isolates G1.5e, G3.2a, G1.5a and G2.4a showed a similar rapid reduction of H2 concentration over the first 24 hr, with isolate G2.4a having the greatest reduction in H2 concentration (Fig. 4). Two dose level experiments were conducted to determine the effect of the number of acetogens on H2 concentrations under slightly different conditions. Only 0.15 g ground alfalfa hay was added to the incubation bottles and 40 mM MES was added to provide additional buffering. In the first dose level ex-

148 and 88 μmoles, respectively at 12 h, and then declined to below 15 μmoles after 36 h. These data show that the lowest level of G3.2a used, which was 5 fold higher than in the previous study, was sufficient to decrease H2 concentrations effectively.

periment, isolate G2.4a was inoculated at 5 x 107 cfu/ ml and 1 x 108 cfu. Under these conditions, H2 concentrations were significantly lower for the higher concentration of acetogen addition at all time points (Fig. 5). In the second dose level experiment isolate

Methane accumulated over time during in vitro fermentations of alfalfa hay inoculated with ruminal contents (Fig 2, 4, 5, 6). The addition of 5-10 mM BES completely inhibited methane production in these fermentations, with a resultant accumulation of H2 as would be expected. Additions of less than 5 mM BES did not completely inhibit methane production (data not shown). Inoculating BES treated rumen content cultures with 5 x 108 cfu of acetogenic bacteria reduced H2 concentrations compared to the control cultures (Fig. 2) with isolate G3.2a reducing H2 concentrations most rapidly. Isolate A10 also

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G3.2a was added inoculated at 5 x 108 cfu and 1 x 109 cfu/ml. In this experiment the higher dose of Isolate G3.2a completely prevented any accumulation of H2 in the culture bottles, wherease the typical accumulation and then decrease in H2 concentrations was observed with the lower dose of acetogen (Fig 6). These experiments demonstrate that acetogens can utilize H2 produced by rumen microorganisms during fermentation of substrates during in vitro incubations where methanogenensis has been inhibited by the addition of a methanogen inhibitor, BES. Thus, these acetogens can utilize H2 in the presence of other substrates that may be available during fermentation, although Pinder and Patterson (2012)

available, whereas the mixed culture incubations measure how much and how rapidly the acetogens can utilize H2 as it is being produced. Thus, other factors such as ability to utilize multiple substrates, resistance to low pH, growth rate and maximum rate of H2 utilization may be more important for acetogen utilization of H2 in ruminal conditions. This data demonstrates that ruminal acetogens can reduce H2 concentrations in a mixed ruminal fermentation when methanogenesis is inhibited and that different acetogenic isolates have different capabilities for H2 utilization in batch culture. Thus, there is potential for acetogens to effectively utilize H2 for interspecies H2 transfer to increase efficiency

have shown diauxic growth of isolate A10 in the presence of glucose in pure culture. The data also show that different isolates have different capacities for utilizing H2 during fermentation. Acetogens can reduce H2 concentrations in relatively low numbers (5 x 107 to 1 x 109 cfu/ml), although H2 utilization is greater with the higher number of acetogens present. Hydrogen thresholds have been argued to be one possible reason that methanogens outcompete acetogens in the rumen. Methanogens have lower H2 thresholds than acetogens. The acetogens we isolated using H2 limited continuous culture have lower hydrogen thresholds than most acetogens that have been isolated. Acetobacterium woodii, isolates G1.5a, G1.5e, G2.4a, G 3.2a, A10 and 3H have hydrogen thresholds of 1007, 800, 635, 908, 960, 209 and 951ppm H2, respectively in our system (Boccazzi and Patterson 2011, 2013) whereas a methanogenic isolate had a H2 threshold of 91 ppm in our system (unpublished data). The reduction of H2 produced during in vitro incubations of alfalfa hay did not directly correlate with H2 thresholds within the range of H2 threshold differences of the acetogens used in this study, although Acetomaculum ruminis, which had the highest H2 threshold, also was less able to reduce H2 concentrations in the in vitro incubations.

of ruminal fermentation, trap more of the feedstuff energy into acetate in the absence of methanogenesis. Feasibility of utilizing acetogens increase efficiency of ruminant fermentation is dependent upon identifying safe, cost effective methods to inhibit methanogenesis.

The differences between pure culture H2 threshold measurements and ability to reduce H2 concentrations during incubation may be because H2 threshold measurements estimate the lowest level of H2 the organism can utilize over time with no other nutrients

captoethanesulfonic acid (HS-CoM)-dependent growth of Methanobacterium ruminantium in a pressurized atmosphere. Appl. Environ. Microbiol. 32: 781-791. Balch, W.E., S. Schoberth, R.S.Tanner, and R.S.Wolfe.

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Ricke, S.C., S.A. Martin, and D.J. Nisbet. 1996. Ecology, metabolism, and genetics of ruminal selenomonads. Crit. Rev. Microbiol. 22: 27-56. Ricke, S.C. and D.M. Schaefer. 1996. Nitrogen-limited growth response of ruminal bacterium Selenomonas ruminantium strain D to methylamine addition in a minimal medium. J. Rapid Methods Automation Microbiol. 4: 297-306. Ricke, S.C., D.M. Schaefer, and T.S. Chang. 1994. Influence of methylamine on anaerobic rumen bacterial growth and plant fiber digestion. Bioresource Technol. 50: 253-257. Ricke, S.C. and D.M. Schaefer. 1991. Growth inhibition of the rumen bacterium Selenomonas ruminantium by ammonium salts. Appl. Microbiol. Biotechnol. 36: 394-399. Robinson, J.A., J.M. Strayer, and J.M. Tiedje. 1981. Method for measuring dissolved hydrogen in anaerobic ecosystems: application to the rumen. Appl. Environ. Microbiol. 41:545-548. Russell, J.B. and R.L. Baldwin. 1978. Substrate preferences in rumen bacteria: Evidence of catabolite regulatory mechanisms. Appl. Environ. Microbiol. 36: 319-329. Russell, J.B. and H.J. Strobel. 1989. Effect on ionophores on ruminal fermentation. Appl. Environ.

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Ammonia saturation constants for predominant species of rumen bacteria. J. Dairy Sci. 63: 12481263. Schink, B. 1984. Clostridium magnum sp. nov., a nonautotrophic homoacetogenic bacterium. Arch. Microbiol. 137: 250-255. Sharak-Genthner, B.R., C.L. Davies, and M.P. Bryant. 1981. Features of rumen and sewage sludge strains of Eubacterium limosum, a methanol and H2-CO2-utilizing species. Appl. Environ. Microbiol. 42: 12-19. Sleat, R., R.A. Mah, and R. Robison. (1985. Acetoanaerobium notarae gen. nov. sp. nov.: an anaerobic bacterium that forms acetate from H2 and CO2. Int.

thermophile producing acetate from molecular hydrogen and carbon dioxide. Curr. Microbiol. 5:255-260. Wieringa, K.T. 1936. Over het cerdwijnen van waterstof en koolzuur onder anaerobe voorwaarden. Antonie van Leeuwenhoek 3:263-273. Wolin, M.J. and T.L. Miller. 1983. Interactions of microbial populations in cellulose fermentation. Federation Proc. 42: 109-113. Wood, H.G. and L.G. Ljungdahl. 1991. Autotrophic character of acetogenic bacteria. In: Shively JM, Banton LL (eds) Variations in autotrophic life. Academic Press Limited pp 201-250. Zeikus, J.G., L.H. Lynd, J.A. Thompson, .J. Krzycki,

J. Syst. Bacteriol. 35:10-15. Tanaka, K. and N. Pfennig. 1988. Fermentation of 2-methoxyethanol by Acetobacterium malicum sp. nov. and Pleobacter venetianus. Arch Microbiol 149:181-187 Tanner, R.S. and D.Yang. 1990. Clostridium ljungdahlii PETC sp. nov., a new, acetogenic, gram-positive, anaerobic bacterium. Abstr R-21, p 249 Abstr Ann Meet Am Soc Microbiol. Tanner, R.S., L.M. Miller, and D. Yang. 1993. Clostridium ljungdahlii sp. nov., and acetogenic species in clostridial rRNA homology group I. Int. J. Syst. Bacteriol. 43:232-236. Tyler, S.C. 1991. The global methane budget. In: Rogers JE and Whitman WB (eds) Microbial production and consumption of greenhouse gases: methane nitrogen oxides, and halomethanes. American Society for Microbiology, Washington D.C., pp 7-38. Thauer, R.K., K. Jungermann, and K. Decker. 1977. Energy conservation in chemotrophic bacteria. Bacteriol Rev 41:100-180. Weimer, P. J. 1992. Cellulose degradation by ruminal microorganisms. Crit. Rev. Biotech. 12:189-223. Weimer, P.J. J.B. Russell, and R.E. Muck. 2009. Lessons from the cow: What the ruminant animal can

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The Effect of Phytochemical Tannins-Containing Diet on Rumen Fermentation Characteristics and Microbial Diversity Dynamics in Goats Using 16S rDNA Amplicon Pyrosequencing B. R. Min1, C. Wright1, P. Ho2, J.-S. Eun3, N. Gurung1, and R. Shange1 Tuskegee University, Tuskegee, AL, USA Montgomery Blair High School, Silver Spring, MD, USA 3 Utah State University, Logan, UT, USA 1

ABSTRACT Two grazing experiments were performed to 1) investigate the effects of supplementing condensed tannins-containing pine bark powder on average daily gain, ruminal fermentation, and rumen microbial diversity dynamics (Experiment 1), and 2) to quantify the influence of different sources of extracted tannins supplementations on ruminal fermentation and rumen microbial diversity changes of goats grazing fresh forages (Experiment 2). In experiment 1, 20 Kiko-Boer cross male goats (Capra hircus; initial body weight= 39.7 ± 2.55 kg) were randomly assigned to 2 experimental diets (alfalfa pellet vs. pine bark powder). Alfalfa pellet (no tannin as a control) or pine bark powder (11% condensed tannins) was supplemented at 0.5% body weight for targeted total dry matter intake of 1.2% body weight. The remaining dry matter intake of each diet was obtained from grazing for 55 days. In experiment 2, 12 Kiko-Boer cross goats were used to measure average daily gain, ruminal fermentation, and gut microbial population in the rumen of goats grazing bermudagrass. The animals were randomly assigned to 3 experimental diets: 1) no tannins (control), 2) chestnut extract at 100 g/d, and 3) quebracho tannin extract at 100 g/day. In experiment 1, average daily gain and rumen fermentation status as measure of volatile fatty acids production were similar between diets. Bacterial population in pine bark powder-supplemented group was greater for Bacteroides (20.5 vs. 33.2%) and Firmicutes (67.2 vs. 57.3%) phylum compared with control group, respectively. In experiment 2, average daily gain was greatest (P < 0.05) for chestnut tannins extract (278.6 g/d) than quebracho tannins extract (150 g/d) and the control (42.9 g/d). Goats grazing bermudagrass pasture with chestnut tannins extract had greater (P < 0.05) concentrations of acetate, propionate, butyrate, and total volatile fatty acids compared to those in quebracho tannins extract and control. Bacterial population in chestnut tannins extract-supplemented group was greatest for Bacteroides (51.5, 52.9, and 35.3%) phylum compared with quebracho tannin extract and control group, respectively. Current study shows that tannins from plants can exhibit a positive or negative effect both on rumen fermentation and on rumen microflora, and it is possible that this effect is depending on sources of tannins or tannin-containing diet. Keywords: Goats, gut microbial diversity, plant tannins, pyrosequencing Agric. Food Anal. Bacteriol. 4: 195-211, 2014 Correspondence: B.R. Min, minb@mytu.tuskegee.edu Tel: +1-334-524-7670 Fax: +1-334-727-8552

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INTRODUCTION The microbial populations of the rumen, particularly bacteria and archaea (methanogens), have been extensively studied (Coleman, 1975; Williams and Coleman, 1988; Fernando et al., 2010). Much is known about the ruminal bacterial populations, but our understanding of interactions between ruminal bacteria and sources of plant tannins in vivo is limited, and very little data exist on the effects of sources of plant tannins or tannin-containing diet on rumen microbiome diversity in goats. Tannins are usually classified either hydrolyzable tannins (HT) or condensed tannins (CT; proanthocyanidins) based on

mentation of tannins in heifers grazing winter wheat reduced the rate of gas and biofilm production with chestnut tannin being more efficacious than mimosa tannins, but some selected rumen bacterial species such as Prevotella ruminicola and strains of both Fibrobacter succinogens and Ruminococcus flavefaciens populations were decreased with chestnut and mimosa tannins supplemented animals (Min et al., 2012a). The implication of sources of plant tannins (Chestnut vs. Quebracho tannins extracts or CTcontaining pine bark powder) likely being associated with specific rumen microorganisms led us to study the effect of two contrasting plant tannins on rumen microbial diversity associated with animal perfor-

their molecular structure. (Min et al., 2003). Our understanding of interactions between rumen bacteria and HT or CT in the rumen is still in its infancy. Tannins have traditionally been considered antinutritional but it is now known that their beneficial or antinutritional properties depend upon their chemical structure and dosage (Min et al., 2003). Structural and chemical dissimilarities between HT (chestnut tannins extract) and CT (Quebracho tannins extract) may offer an explanation for differences in their biological effects and, therefore, results obtained using a particular type of tannins cannot be applied to others. In our study, we utilized a combination of CT and HT instead of CT alone to examine if some phenolic metabolites deriving from HT degradation in the rumen may affect the rumen fermentation and microbiome diversity, giving it added value. Recent studies have demonstrated that chestnut tannins have been shown to have positive effects on silage quality in round bale silages, in particular reducing non protein nitrogen (NPN) in the lowest wilting level (Tabacco et al., 2006). Improved fermentability of soya meal nitrogen in the rumen has also been reported by Mathieu and Jouany (1993). Studies by Gonzalez et al. (2002) on in vitro ammonia release and dry matter degradation of soybean meal

mance. The primary hypothesis of the in vivo grazing research was that different sources of tannins supplementation would selectively reduce rumen microbial diversity and as a result would increase average daily gain in meat goats. Thus, our objective was to investigate concurrent changes in ruminal bacterial diversity and animal performance in goat response to plant tannins using a modern pyrosequencing approach.

comparing three different types of tannins (quebracho, acacia and chestnut) demonstrated that chestnut tannins are more efficient in protecting soybean meal from in vitro degradation by rumen bacteria. This has been confirmed by the findings that supple-

BW for targeted total dry matter intake (DMI) of 1.2% BW. The remainder DMI of each diet was obtained from grazing for 55 days (Table 1). Animals were fed once a day at 0900 h and had free access to water and trace mineral salt blocks grazing on winter

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MATERIALS AND METHODS Care and handling of all experimental animals were conducted under protocols approved by the Tuskegee University Institutional Animal Care and Use Committee.

Experimental Animals and Diets In Exp. 1, 20 Kiko-Boer cross male goats (Capra hircus; initial body weight (BW) = 39.7 Âą 2.55 kg) were randomly assigned to 2 experimental diets (alfalfa pellet vs. PB powder). Alfalfa pellet (no CT as a control) or PB (11% CT) was supplemented at 0.5%

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Table 1. Chemical compositions (%) of the pine bark powder, alfalfa pellet, winter forage (ryegrass and pea) and bermuda grass. Experimental diet Pine bark

Winter forage

Alfalfa pellet

Bermudagrass

Dry matter

91.9

91.8

92.1

92.5

0.33

Crude protein

19.0

22.0

16.9

9.1

6.17

Acid detergent fiber

32.4

37.3

30.4

49.6

9.60

Neutral detergent fiber

41.5

48.7

40.3

9.57

powder

Ingredients

SD = standard deviation

pea and rye grass dominant forages. Animal body weight (BW) were measured before and after experiment completion. Animals were drenched orally with Cydectin (1 ml/10 kg BW) when fecal egg count was over 1000 egg per gram of feces. Individual rumen fluid samples (50 ml) were collected on day 55 after slaughter at the end of the experiment for rumen volatile fatty acids (10 ml) and microbial diversity (40 ml) analyses. Rumen fluid samples from ten animals per treatment were then pooled to three samples sizes within treatment for bacterial analysis. In Exp. 2, 12 Kiko-Boer cross male goats were used to measure average daily gain (ADG), ruminal fermentation, and rumen microbial population in the rumen of goats grazing bermudagrass (Cynodon dactylon) dominant pasture. The animals were randomly assigned to 3 experimental diets: 1) no tan-

mental diets (Table 1). Rumen fluid was collected via stomach tube, fitted with a small cylindrical strainer, before the morning feeding, into 50 mL serum vials that were filled to capacity, capped immediately and stored at -20°C until analysis later that day.

nins (control), 2) chestnut HT-extract at 100 g/d (CTE), and 3) quebracho CT-extract at 100 g/d (QCTE). Experimental diets were gradually fed to animals in a stepwise increasing fashion, and at the end of week 2, all animals were fed whole, pre-assigned experi-

described by AOAC (AOAC, 1998). Nitrogen for diet sample was determined using a Kjeldahl N, and CP was calculated by multiplying N by 6.25. The neutral detergent fiber (NDF) and acid detergent fiber (ADF) concentrations were sequentially determined using

Chemical Analysis Feed and forage samples were collected daily during the collection period, dried at 60°C for 48 h, ground to pass a 1 mm screen (standard model 4; Arthur H. Thomas Co., Swedesboro, NJ), and stored for subsequent analyses. Daily portions of ground samples were composited for each animal and analyzed for DM, crude protein (CP), acid detergent lignin, ether extract, and ash according to the methods

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an ANKOM200/220 Fiber Analyzer (ANKOM Technology, Macedon, NY). Sodium sulfate heat stable amylases (Sigma Aldrich Co., St. Louis, Mo) were used in the procedure for NDF determination and pretreatment with heat stable amylase (Type XI-A from Bacillus subtilis; Sigma-Aldrich Corporation, St. Louis, MO). Acetone (70%) extractable CT in grain mixes were determined using a butanol-HCL colorimetric procedure (Min et al., 2012). For volatile fatty acids analysis, 5 mL of rumen fluid was diluted with 1 ml of 3 M meta-phosphoric acids, and samples were analyzed using a method described by Williams et al., (2011). Volatile fatty acids were analyzed via gas chromatography (Agilent 6890N, Santa Clara, CA,

samples were adjusted to 100 ng/μL. A 100 ng (1 μL) aliquot of each sample’s DNA was used for a 50 μL PCR reaction. The 16S universal eubacterial primers 530F (5’-GTG CCA GCM GCN GCG G) and 1100R (5’-GGG TTN CGN TCG TTG) were used for amplifying the 600 bp region of 16S rRNA genes. HotStar Taq Plus Master Mix Kit (Qiagen, Valencia, CA) was used for PCR under the following conditions: 94°C for 3 min followed by 32 cycles of 94°C for 30 sec; 60°C for 40 sec and 72°C for 1 min; and a final elongation step at 72°C for 5 min. The resultant individual sample after parsing the tags into individual FASTA files was assembled using CAP3. The resulting tentative consensus FASTA (A database search tool used

USA) with a 007 series bonded phase fused silica capillary column (25 m × 0.25 mm × 0.25 μm) and a flame ionizing detector with the following parameters: 1 μL injection, injector temperature = 240 °C, oven temperature = 80 °C for 1 min, ramp to 120 °C hold for 5 min, ramp to 165 °C hold for 2 min, detector temperature = 260 °C.

to compare a nucleotide or peptide sequence to a sequence database) for each sample was then evaluated using BLASTn (Altschul et al., 1990) against a custom database derived from the RDP-II database (Cole et al., 2005) and GenBank website athttp:// www.ncbi.nlm.nih.gov. The sequences contained within the curated 16S database were both >1200 bp and considered as high quality based upon RDP-II standards.

DNA Extraction Genomic bacterial DNA was isolated from 1 ml of rumen samples according to the method described in the QIAamp DNA Mini Kit (QIAGEN Inc., 27220 Turn berry Lane, Suite 200 Valencia CA). Extracted DNA (2 μL) was quantified using a Nanodrop ND1000 spectrophotometer (Nyxor Biotech, Paris, France) and run on 0.8% agarose gel with 0.5 M TrisBorate-EDTA (TBE) buffer. The samples were then transported to the Research and Testing Laboratory (Lubbock, TX) for PCR optimization and pyrosequencing analysis. Bacterial tag-encoded FLX amplicon pyrosequencing (bTEFAP) PCR was carried out according to procedure described previously (Min et al., 2012).

bTEFAP Sequencing PCR The bTEFAP and data processing were performed as described previously (Dowd et al., 2008). All DNA 198

Data Processing and Statistical Analysis Statistical analyses were performed using the SPSS package (SPSS Inc., v 17.0, Chicago, IL). Package of NCSS (NCSS, 2007, v 7.1.2, Kaysville, UT) was used for cluster analysis through which double dendrograms were generated through use of the Manhattan distance method with no scaling, and the unweighted pair technique. Quality trimmed sequences were provided with the sequencing services by the Research and Testing Laboratory (Lubbock, TX; (Dowd et al., 2008). Tags which did not have 100% homology to the original sample tag designation were not included in data analysis. Sequences which were less than 250 bp after quality trimming were not also considered. The resulting sequences were then evaluated using the classify.seqs algorithm (Bayesian method) in MOTHUR against a database derived from the Greengenes set using a bootstrap cutoff of 65%.

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Table 2. Effects of condensed tannin-containing pine bark (PB) powder and different sources of tannins extracts supplementation on the animal body weight (BW) changes and average daily gain (ADG) in meat goats grazing fresh forages

Initial BW (kg)

Final BW (kg)

ADG (g)

PB powder

37.3

46.6

169.1

Control

36.5

46.7

185.5

8.05

0.34

12.67

P-value

0.76

0.35

0.71

Chestnut

32.7

36.6

278.6

Quebracho

33.5

35.6

150.0

Control

30.5

31.1

42.9

5.88

6.98

20.02

P-value

0.85

0.26

0.05

Exp.1 (n = 10/diet)1

Exp.2 (n = 4/diet)2

Animals were grazed on winter pea and ryegrass pasture with or without PB powder supplementation during 55 days 2 Animals were grazed on bermudagrass forage with or without tannin extracts supplementation during 14 days. 1

Relative abundance data are presented as percentages/proportions, but prior to subjection to general linear model (GLM; SAS Inst., Cary, NC)), they were transformed using the arcsine function for normal distribution prior to analysis. In addition, quantification of major rumen bacterial phylum, classes and species populations was analyzed by the GLM procedure of the SAS in a completely randomized design with the factors examined being sources

RESULTS AND DISCUSSION

of tannins supplementation in the diets. Results are reported as least square means.

most significant findings in the present study demonstrates that when goats received tannins extracts (chestnut and quebracho extracts) in the Firmicutes phylum populations had significant (P < 0.01) decreased, while Bacteroidetes populations were

Regardless of numerous studies (Pitta et al., 2010; Callaway et al., 2010; Hristov et al., 2012) demonstrating the role of the gut microbial diversity in ruminants associated with different sources of forages or dried distillers grains, the response of the microbiome to feeding various sources of phytochemical tannins-containing diet remain largely unknown. The

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significantly increased. The bacterial distribution showed that Firmicutes (56-57%) was the most dominant phyla with mean relative abundance values ranging from 56% in control to 67 % in PB diets. This suggests that phytochemical tannins supplementation alters the microbiome and animal performance on goats grazing fresh forage diets.

Diet Composition Ingredients and chemical composition of experimental diet (alfalfa pellet), PB, and bermudagrass forage are presented in Table 1. Goats were provided diets that met all animals’ requirements for growth and gain according to National Research Council (NRC, 2007). Total CT concentration in the PB, alfalfa pellet, winter forages, and bermudagrass was 10.3, 0.03, 0.03, and 0.05% DM, respectively (Table 1). All the experimental treatments provided similar nutrient profiles, except CT and ADF that were higher in PB, but lower in CP compared to other diets. In our previous study, addition of PB in goat diets improved ADG and favorably modified ruminal fermentation (Min et al., 2012b).

Animal Performance and Rumen Fermentation

those in QCTE and control (Table 3). However, goats grazing bermudagrass forage without tannins supplementation increased (P < 0.05) concentrations of iso-valerate and acetate: propionate ratio compared to tannins supplemented groups. A number of studies have demonstrated that effects of tannins on ruminal fermentation is dose dependent, and a negative effect only occurs when they are fed at high concentrations (Hervás et al., 2003; Mueller-Harvey, 2006). In addition, previous studies have reported that mimosa and chestnut tannins supplementations were not affected animal performance and rumen fermentation in steers fed a high-grain diet (Krueger et al., 2010) or hay supplementation with chestnut tannins spray (Zimmer and Cordesse, 1996). However, Min et al. (2012a) reported that heifers grazing on high quality (about 28% crude protein content) winter wheat forage and supplemented with 1.5% tannins (DM basis) experienced 82% fewer days of bloat, and had 6 and 17% greater ADG for mimosa and chestnut tannins extracts, respectively, than animals receiving the control diet principally through reducing the rate of rumen fermentation as well as modification of microbial populations in the rumen of cattle. Our chestnut tannins supplementation study shows a similar trend to this. This suggests that plant tannins supplementation in high quality forage may have more impact on mitigating rumen fermentation and improving animal performance than low quality forages diets.

The animal performance and ruminal volatile fatty acids concentrations in goats grazing fresh forages in response to different sources of tannins supplementation are shown in Tables 2 and 3, respectively. In Exp. 1, Initial BW, final BW, ADG, total volatile fatty acids, acetate: propionate ratios, and individual volatile fatty acids concentration were similar between PB powder and control alfalfa pellet diets (Tables 2 and 3). In Exp. 2, Initial BW and final BW were similar among treatments (Table 2), but ADG was greatest

In this study, bacterial (Fig. 1a,b) community composition of the rumen fluids were examined at descending levels of biological classification to determine the effect of PB powder (Exp. 1) or tannin extract (Exp. 2) supplementation on community membership. Detailed phylogenic analyses grouped the rumen bac-

for CTE (275 g/d) than QCTE (145 g/d) and the control (41 g/d). Goats grazing bermudagrass pasture with CTE had greater (P < 0.05) concentrations of acetate, propionate, butyrate, caprionate, and total volatile fatty acids concentrations compared to

teria associated bacterial sequences into 45 phyla (including unknown). The relative abundances of the 19 most abundant phyla (>1%) are presented in Figure 1. Interestingly, the gut of human and many other vertebrae are mostly dominated by two groups of bacteria,

200

Relative Abundance of Bacterial Phyla

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Table 3. Effects of condensed tannin-containing pine bark (PB) powder and different sources of tannin extracts supplementation on the ruminal volatile fatty acids (VFA) concentration and acetate: propionate (A:P) ratios in meat goats grazing winter pea and ryegrass dominant forages.

Item1

Iso-C4

Iso-C5

Total VFA

A:P ratio

Exp.1 (n = 10/diet) PB powder2

14.4

3.38

0.99

1.82

1.52

0.39

0.01

22.54

4.23

Control

15.0

3.04

0.93

2.0

1.47

0.41

0.06

23.67

4.05

4.74

1.35

0.15

0.66

0.28

0.09

0.03

6.98

0.10

P-value

0.78

0.47

0.37

0.53

0.73

0.77

0.64

0.73

0.22

Chestnut3

54.4

13.55

0.23

4.4

0.09

0.5

0.15

73.4

4.02

Quebracho

44.0

12.4

0.25

3.9

0.08

0.4

0.05

61.2

3.59

Control

49.1

10.5

0.37

3.6

0.31

0.4

0.03

64.4

4.69

7.44

1.96

0.10

0.92

0.02

0.08

9.12

0.46

P-value

0.02

0.001

0.02

0.01

0.07

0.21

0.01

0.001

Exp.2 (n = 4/diet)2

Volatile fatty acids (VFA): acetic (C2), propionic acid (C3), butyric (C4), valeric (C5), and caproic acid (C6) Animals were grazed on winter pea and ryegrass pasture with or without PB powder supplementation during 55 days. 3 Animals were grazed on bermudagrass forage with or without tannin extracts supplementation during 14 days. 1 2

Bacteroidetes and Firmicutes (Backhed et al., 2004), which is similar to the result obtained current study. This finding agrees with the results within the current goat study showing that Firmicutes (56-57%) and Bacteroidetes (33-35%) were the dominant bacterial phylum in the goat rumen fluid. With the addition of Firmicutes, together these phyla constituted approximately 89 to 92% of the total gut population. A study by Henderson et al. (2013) demonstrated an increase

Firmicutes, Bacteroidetes, Actinobacteria, and Proteobacteria were reported to be dominant bacterial phyla in the goat intestine (Min et al., 2014) and human gut (Schloss et al., 2009). It has been shown that the Bacteroidetes and Firmicutes phyla comprised 35% of all sequences, followed by Proteobacteria (13% to 15%) and Fusobacterium (7% to 8%). The bacterial distribution showed that Firmicutes (56-57%) was the most dominant phylum with mean relative abundance

in the abundance of the phylum Firmicutes correlated with a decrease in the abundance of Bacteroidetes in cow (r= -0.805) and sheep (r= -0.976), which also shows similarity to the results obtained in the current study. This has been confirmed by the findings that

values ranging from 56% in control to 67 % in PB diets (Fig. 1). However, goats that received CTE and QCTE extract supplementation had significant decreases (P<0.01) in Firmicutes populations, while Bacteroidetes populations were significantly increased (Fig. 1b).

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Figure 1. Predominant bacterial phylum observed in rumen samples of healthy goats with or without tannin-containing diets supplementation based on pyrosequencing of the 16S rRNA gene. Goats with pine bark powder (A) had significantly decreases in Bacteroidetes, but increased Firmicutes (Data expressed as % of total 16S rRNA sequences). Goats with tannins extracts (B) had significant (P < 0.01) decreases in Firmicutes. Bacteroidetes were significantly increased. Parabasalia Proteobacteria Euryarchaeota Streptophyta Tenericutes Firmicutes Elusimicrobia Cyanobacteria Acidobacteria Nitrospirae Planctomycetes Verrucomicrobia Fibrobacteres Lentisphaerae Spirochaetes Actinobacteria Synergistetes Chloroflexi Bacteroidetes

A 100 90 80 57.3

67.2

60 50 40 30 33.3

20.5

10 0 Control

Pine bark powder

100 90 80 70

42.6

36.7

55.9

60 50 40 30 20

51.5

52.9

35.3

10 0 202

Control

Chestnut

Quebracho

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Parabasalia Proteobacteria Euryarchaeota Streptophyta Tenericutes Firmicutes Elusimicrobia Cyanobacteria Acidobacteria Nitrospirae Planctomycetes Verrucomicrobia Fibrobacteres Lentisphaerae Spirochaetes Actinobacteria Synergistetes Chloroflexi Bacteroidetes

The gastrointestinal microbiota performs a large number of important roles that define the physiology of the host, such as immune system maturation (Mazmanian et al., 2005), the intestinal response to epithelial cell injury (Rakoff-Nahoum et al. (2004), xenobiotic (Wilson and Nichoson, 2009), and energy metabolism (Turnbaugh et al., 2006). In most mammals, the gastrointestinal microbiome is dominated by four bacterial phyla that perform these tasks: Firmicutes, Bacteroidetes, Actinobacteria and Proteobacteria (Ley et al., 2007). The current study has shown that the number of Firmicutes population was notably greater than the number of Bacteroidetes in PB fed animals compared to alfalfa supplemented

ly, an increase in weight (Turnbaugh et al., 2006). The Bacteroidetes spp, in particular Bacteroides thetaiotaomicron, hydrolyzes otherwise indigestible polysaccharides and accounts for 10% to 15% of caloric requirement in humans (Xu et al. 2003). Human colonocytes derive 50% to 70% of their energy from butyrate, which is derived from complex carbohydrates metabolized by Firmicutes spp via fermentation (Pryde et al., 2002). However, the current study had opposite trends to human study in that the both CTE and QCTE extract supplemented groups have greater butyrate, iso-butyrate, acetate, and propionate concentrations in the rumen, and had higher Bacteroidetes population compared to

animals, while CTE and QCTE plant extracts supplementation had the opposite trend (lower abundance in Firmicutes and greater density in Bacteroidetes, respectively) compared to control group in grazing goats. The mechanism of action of tannin-resistant bacteria in animals exposed to condensed tannins is not known between two different dietary supplementations. Past research into the correlation between gut microbiota and diet had demonstrated a complex relationship between the population of the gut and fatty acid absorption. Although exact mechanisms are not yet known, it has been observed that obesity due to a high fat or high polysaccharide diet correlates with a decrease in the amount of Bacteroidetes and a proportional increase in Firmicutes. This was shown by Ley et al. (2005) in mouse models with obese and normal genotypes, and was later supported by Ley et al. (2007) in studies of human fecal matter. The number of Firmicutes was notably higher than the number of Bacteroidetes in obese mice, and vice versa for the lean mice (Ley et al., 2005). Along with increased fatty acid absorption, more energy was also found to be efficiently obtained from diet in the obese mice compared to the lean mice, illustrating the connection between Firmicutes and improved

control group. It is unclear what factors in the setting of average daily gain tip the scales in favor of the Firmicutes over Bacteroidetes in ruminants. Perhaps the Bacteroides possess may more tannins-resistant mechanisms or more diverse enzymatic capabilities (Odenyo and Osuji, 1998; Smith et al., 2003) that more efficiently extract energy when a variety of complex organic matter is available in goats. This hypothesizes that the metabolic and energy extraction functions in ruminants may be fundamentally due to microbiota, such that all are affected by alterations in nutritional state.

efficiency in energy harvesting (Turnbaugh et al., 2006). The replacement bacteria are more efficient at harvesting energy from food than the bacteria they replaced, resulting in increased calorie intake by the host (Turnbaugh et al., 2006), and ultimate-

cies distribution showed that Ruminococcaceae spp. (12-15%) and Prevotella spp. (21-40%) were the most dominant species with mean relative abundance values ranging from 42 to 55% in control group without tannins supplementation (Table 4). In Exp. 1,

Diversity and Abundance of Rumen Bacterial species More than 332 bacterial species (including unknown) were classified from the ruminal fluid of the goats in this study. However, the relative abundances of the 12 most abundant species (>1%) are presented in Tables 4 and 5. Tannins are one of the most abundantly available plant secondary metabolites, and have positive or adverse effects on rumen microbial populations, feed digestibility and animal performance (Min et al., 2003). The bacterial spe-

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Table 4. Effects of condensed tannin-containing pine bark (PB) powder supplementation (n = 10/ diet) on the rumen bacterial species population diversity (%) in meat goats grazing fresh forages1

Bacterial species

Pine bark

Control

P-value

Roseburia spp.

1.96

1.62

2.34

0.83

Ruminococcus spp.

1.03

1.89

0.85

0.31

Paraprevotella spp.

0.54

0.67

0.50

0.71

Succiniclasticum spp.

3.18

5.58

4.42

0.51

Prevotellaceae spp.

1.81

1.15

1.61

0.59

Victivallis spp.

0.38

0.58

0.41

0.56

Ruminococcaceae spp.

18.78

15.2

4.99

0.08

Prevotella spp.

11.46

21.4

3.19

0.05

Butyrivibrio spp.

13.56

5.08

3.76

0.05

Blautia spp.

4.92

4.76

5.30

0.97

Desulfovibrio spp.

0.24

0.28

0.72

0.89

Saccharofermentans spp.

4.37

6.97

3.44

0.54

Animals were grazed on winter pea and ryegrass pasture with or without PB powder supplementation during 55 days 1

Prevotella spp. was decreased (P < 0.05) in PB diet, while Butyrivibrio spp. (P < 0.05) and Ruminococcaceae spp. (P = 0.08) were increased compared to control diet. However, bacterial populations in CTEsupplemented group in Exp. 2 were significantly increased for Prevotella spp. (40.4, 33, and 32%) compared with QCTE and control groups, respectively. It has been shown that the gastrointestinal microbial population was dominated by Prevotella (18.2% of total population) in the rumen and Clostridium (19.7% of total population) in the feces of cattle (Cal-

These findings also agree with the results of a metabolic finger print study of a rat fed CT extract from Acacia angustissima. Condensed tannin extracts from A. angustissima altered fecal bacterial populations in the gastrointestinal tract, resulting in a shift in the predominant bacteria towards tanninresistant gram-negative Enterobacteriaceae and Bacteroidetes (Smith et al., 2003). Presence of bacteria able to tolerate elevated levels of condensed tannins in the rumen of animals fed forages high in tannins has been reported by Nelson et al. (1995).

laway et al., 2010). Consequently, analysis of human microbiota-associated rat feces using molecular approach has revealed that the Bacteroides/Prevotella and Faecalibacterium species are dominant in both humans and rats post-transfection (Licht et al., 2007).

Different groups of microbes have different tolerance to tannin. Rumen fungi, proteolytic bacteria and protozoa are more resistant to tannin as compared to other microbes (McSweeny et al., 2001). McSweeny et al. (1999) observed that in the animals

204

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Table 5. Effects of different sources of tannins extracts supplementation (n = 4/diet) on the rumen bacterial species population diversity (%) in meat goats grazing fresh forages1

Bacterial species

Chestnut

Quebracho

Control

P-value

Roseburia spp.

1.13

0.81

0.87

0.04

0.56

Ruminococcus spp.

2.28

1.94

2.57

0.20

0.41

Paraprevotella spp.

1.53

1.31

1.67

0.06

0.21

Succiniclasticum spp.

2.52

1.90

2.66

0.32

0.15

Prevotellaceae spp.

6.16

8.24

6.54

2.44

0.67

Victivallis spp.

1.21

0.37

1.37

0.56

0.27

Ruminococcaceae spp.

11.65

14.4

12.7

3.74

0.22

Prevotella spp.

40.37

32.8

31.76

12.77

0.05

Butyrivibrio spp.

2.36

1.62

1.93

0.28

0.38

Blautia spp.

4.15

6.91

6.95

5.14

0.29

Desulfovibrio spp.

1.13

0.69

1.12

0.12

0.71

Saccharofermentans spp.

5.2

1.45

2.30

3.98

0.88

Animals were grazed on bermudagrass forage with or without tannin extracts supplementation during 14 days 1

fed on tannin rich Calliandra calothyrsus, the population of Ruminococcus spp. and Fibrobacter spp. was reduced considerably. Min et al. (2002) reported that a decrease of 0.5-0.1 log in proteolytic ruminal bacteria Clostridium proteoclasticum, Streptococcus bovis, Eubacterium spp. and Butyrivibrio fibrisolvenes when CTs from Lotus corniculatus (3.2 % CT/kg DM) were fed to sheep. Recently, Min et al., (2014) reported that Stenotrophomonas koreensis was the most dominant bacterial species with mean relative abundance values ranging from 23.9% (con-

etary PB concentration. However, Bacteroides capillosus, Clostridium orbiscindens, and Oscillospira guilliermondii were linearly increased with increasing PB concentration. This suggests that phytochemical tannins supplementation alters microbial diversity and thereby improves animal performance. The authors observed significantly lower (P<0.05) Prevotella bacteria populations in goats fed CTE extract was increased of as compared to animals fed QCTE or control diets. For ease of presentation and interpretation, we

trol) to 9.9% (15% PB) and 17.2% (30% PB). The remaining bacterial species accounted for fewer than 10% of the relative abundance observed. Of these groups, Flavobacterium gelidilacus and Myroides odoratimimus were decreased with increasing di-

present prevalent bacterial genera (Figures 2 and 3) observed in the community based on a cutoff value of 0.9% of relative abundance for inclusion in a hierarchal cluster analysis of individual animal microbial diversity within and among diets in Figures 2 and 3.

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Figure 2 (Exp. 1). Heat-map double dendrogram of the 44 most abundant bacterial genera in the rumen of various sources of tannin extracts supplementation from a common cohort of 20 meat goats. Clustering in the Y-direction is indicative of abundance, not phylogenetic similarity. RA = relative abundance; pine bark = sample no. 9, 10, and 11; Control = 12, 13, and 14. Rumen fluid samples from ten animals per treatment were pooled to three samples sizes within treatment for bacterial analysis.

206

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Figure 3 (Exp. 2). Heat-map double dendrogram of the 44 most abundant bacterial genera in the rumen of various sources of tannin extracts supplementation from a common cohort of 12 meat goats. Clustering in the Y-direction is indicative of abundance, not phylogenetic similarity. RA = relative abundance; Chest nut = tag no. 1 and 2; Quebracho = 3 and 4; Control = 7 and 8. Rumen fluid samples from four animals per treatment were pooled to two samples sizes within treatment for bacterial analysis.

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Overall, animals clustered relatively well within diet and animals. However, one of PB powder supplemented goats had more dissimilarity between treatments. Similar trends were found in Exp. 2. Chestnut diet of goats (sample # 1 and 2) clustered more closely to the quebracho (# 3) supplementation, but one of control animal (# 7) was relatively not clustered within control diet (# 8). Goats that received PB powder in Exp. 1 had greater relative community abundance of Clostridia population compared to the control diet, and the opposite was observed for Bacterodia population (Figure 2). In Exp. 2, chestnut and quebracho tannin extracts supplemented groups had greater relative abundance (%) of Bacteroidia compared to

Agricultural Research Station. Research and Testing Laboratory (Lubbock, TX) for PCR optimization and pyrosequencing analysis are also acknowledged.

the control diet. Lower abundance of Clostridia in tannins extract groups compared to control diet, indicated that tannins extracts supplementation may have decreased the abundance of Clostridia population in rumen of goats.

Koh, A. Nagy, C. F. Semenkovich, and J. I. Gordon. 2004. The gut microbiota as an environmental factor that regulates fat storage. Proc. Natl. Acad. Sci. USA 101:15718–15723. Brooker, J. D., L. A. O’Donovan, K. Clarke, L. Blackall, and P. Muslera. 1994. Streptococcus caprinus sp. nov., a tannin-resistant ruminal bacterium from feral goats. Lett. Appl. Microbiol. 18:313-318. Callaway, T. R., S. E. Dowd, T. S. Edrington, R. C. Anderson, N. Krueger, N. Bauer, P. J. Kononoff, and D. J. Nisbet. 2010. Evaluation of bacterial diversity in the rumen and feces of cattle fed different levels of dried distillers grains plus soluble using bacterial tag-encoded FLX amplicon pyrosequencing. J. Anim. Sci. 88: 3977–3983. Cole, J. R., B. Chai, R. J. Farris, Q. Wang, S. A. Kulam, D. M. McGarrell, G. M. Garrity, and J. M. Tiedje. 2005. The Ribosomal Database Project (RDP-II): sequences and tools for high-throughput rRNA analysis. Nucleic Acids Res. 33:D294–D296. Coleman, G. S. 1975. The inter-relationships between rumen ciliate protozoa and bacteria. Pages 149–164 in Digestion and Metabolism in the Ruminant. I. W. McDonald and A. C. Warner, eds. Univ. of New England Publ. Unit, Armidale, Australia. De Filippo, C., D. Cavalieri, M. Di Paola, M. Ramaz-

CONCLUSIONS In conclusion, the current results show that tannin can exert a positive or negative effect both on rumen fermentation and on rumen microflora, and it is possible that this effect is depending on sources of tannins or tannin-containing diet. Rumen microbial population is very dynamic and tannin inclusion impacts specific members of the microbial population. There is also possible adaptation of ruminal microbiota to tannin and beneficial effect of tannin on some class of rumen microbes has been observed. However, there is need for detailed study involving effect of varying concentration of tannins on rumen bacteria, archaea and fungal diversity of goats in response to ingestion of different sources of tannincontaining diet.

ACKNOWLEDGEMENTS This project was supported by USDA-NIFA, The USDA-NIFA Evans-Allen Research Program and Tuskegee University, George Washington Carver 208

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Characterization of the Novel Enterobacter cloacae Strain JD6301 and a Genetically Modified Variant Capable of Producing Extracellular Lipids J. R. Donaldson1*, S. Shields-Menard1, J. M. Barnard1, E. Revellame2, J. I. Hall3, A. Lawrence4, J. G. Wilson1, A. Lipzen5, J. Martin5, W. Schackwitz5, T. Woyke5, N. Shapiro5, K. S. Biddle1, W. E. Holmes2, R. Hernandez2, and W. T. French3 Department of Biological Sciences, Mississippi State University, Mississippi State, MS, USA. Department of Chemical Engineering, University of Louisiana Lafayette, Lafayette, LA, USA. 3 Renewable Fuels and Chemicals Laboratory, Dave C. Swalm School of Chemical Engineering, Mississippi State University, Mississippi State, MS, USA. 4 Institute for Imaging and Analytical Technologies, Mississippi State University, Mississippi State, MS, USA. 5 DOE Joint Genome Institute, Walnut Creek, CA, USA. 1

ABSTRACT This work presents results from the phenotypic and genotypic characterization of a novel Enterobacter cloacae strain (#JD6301) recently isolated from a mixed population of oleaginous microorganisms. Lipid analysis of this strain indicated that JD6301 produces nearly 50% of its cellular weight as lipids. The yield of fatty acid methyl esters for this microorganism was 76 μg/mL. Transmission electron microscopy observations showed inclusion bodies form within this isolate. To improve the recovery of these useful lipids from this microorganism, a random mutagenesis assay was utilized to isolate an alternative form of this bacterium capable of producing extracellular lipids. The extracellular fraction of the mutant strain JD8715 had a total fatty acid methyl esters yield of 86 μg/mL, which was similar to the intracellular yield of JD6301. Furthermore, cell viability and microscopic analysis indicated that the presence of extracellular lipids was not due to cell lysis. Comparative genome analysis of JD8715 against JD6301 revealed 24 single nucleotide polymorphisms, of which 17 resulted in non-synonymous amino acid changes. Seven of these changes occurred in genes related to membrane proteins. The application of oleaginous microorganisms capable of producing extracellular lipids while still retaining cell viability represents a promising approach for providing energy required for biotechnological applications. Keywords: Enterobacter cloacae, triacylglyceride, lipids, extracellular lipids, electron microscopy, oleaginous, biofuels, biodiesel, JD6301, membrane transport Agric. Food Anal. Bacteriol. 4: 212-223, 2014

Correspondence: J. R. Donaldson, donaldson@biology.msstate.edu, Tel: +1-662-325-9547

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INTRODUCTION

Culture conditions

Lipids are important energy sources in both human health and also in biofuel production, such as biodiesel (Ranganathan et al., 2008; Rosen and Spiegelman, 2006). Biodiesel is produced through the transesterification/esterification of triacylglycerides (TAGs) to yield fatty acid alkyl esters and commonly exists as fatty acid methyl/ethyl esters, or FAMEs (Lestari et al., 2009). While plant and animal fat stores are the most common source of TAGs for biofuel production, certain prokaryotes have been identified to accumulate TAGs as a form of energy storage (Alvarez and Steinbuchel, 2002; Holder et al., 2011).

Frozen stocks of the Enterobacter cloacae wild type (WT) strain JD6301 and resulting isogenic mutant JD8715 were maintained at -80°C in 20% glycerol. Frozen stocks were cultured on nutrient agar and allowed to grow for 48 – 72 h at 30°C. For lipid analysis of JD6301 and JD8715, bacteria were cultured in mineral salts medium (MSM) supplemented with 3% (w/v) sodium gluconate in baffled culture flasks (Schlegel et al., 1961). The auxotrophic mutant Saccharomyces cerevisiae strain KD115 (MATα ole1) was purchased through the American Type Culture Collection (Stukey et al., 1989) and was cultured in

These microorganisms, termed oleaginous microorganisms, accumulate more than 20% of their biomass as TAGs and include the bacteria Streptomyces, Nocardia, Rhodococcus, Mycobacterium, Dietzia, and Gordonia (Alvarez and Steinbuchel, 2002; Wynn and Ratledge, 2005), as well as yeast and fungi, such as Yarrowia lipolytica and Mortierella isabellina (Meng et al., 2009). Recently, the novel Enterobacter cloacae strain JD6301 was isolated from a mixed culture containing oleaginous microorganisms and microorganisms from a municipal wastewater treatment facility and was sequenced (Wilson et al., 2014). The goal of this study was to further analyze this novel isolate. Enterobacter cloacae strain JD6301 was found capable of producing large quantities of lipids through both transmission electron micrograph observations and lipid analyses. A variant form of this strain was constructed following a random mutagenesis that was able to produce extracellular lipids. This strain was further analyzed through genomic comparisons to determine candidate gene mutations that resulted in the observed phenotype. The ability of oleaginous microorganisms to produce extracellular lipids could lead to advancements in lipid biotechnology, especially in the areas of lipid recovery and utiliza-

YPOD media (1% yeast extracts, 2% bacto-peptone, 2% glucose, 1% brij 58, and 0.2% oleic acid) under aerobic conditions for 24 – 48 h at 30°C in a shaking incubator (180 rpm). For plates, 2% agar was added to the YPOD medium.

tion.

of KD115 around the periphery of the Enterobacter cloacae cells indicated the EMS introduced genetic alterations that promoted the presence of extracellular lipids, which were then available for utilization by KD115. The potential mutants were transferred to

MATERIALS AND METHODS

Mutant construction A mutant of JD6301 capable of producing extracellular lipids was constructed as previously described for S. cerevisiae with minor modifications (Nojima et al., 1999). A 24 h culture of JD6301 in MSM supplemented with 3% sodium gluconate was treated for 3 h at 30°C with 3% ethyl methanesulfonate (EMS; Acros Organics), which is a mutagenic chemical that introduces predominately GC to AT base transitions (Ingle and Drinkwater, 1989). The treated cells were then plated onto nutrient agar and incubated for 48 h at 30°C. Colonies were then overlaid with 6 mL of YPD (1% yeast extracts, 2% bacto-peptone, 1% agar, and 2% glucose) agar containing 1x107 CFU/mL of S. cerevisiae KD115 and 50 U of lipoprotein lipase (Sigma, L9656). Cultures were incubated for an additional 16 h at 30°C to allow for growth of the auxotrophic KD115 strain. The presence of microcolony growth

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individual tubes with 2 mL of MSM with 3% sodium gluconate and incubated in a shaking incubator for 24 h at 30°C, plated on nutrient agar, and incubated for an additional 48 h. After incubation the mutants were tested again through an overlay with KD115 and lipase-containing YPD agar. Of the 1,000 colonies screened, the mutant JD8715 was selected for further analysis.

Genomic resequencing and analysis Resequencing of JD8715 was generated by the Department of Energy Joint Genome Institute (JGI) using the Illumina HiSeq 2000 platform as previously described (Wilson et al., 2014), which generated 178,495,750 reads. The final draft assembly contained 53 contigs and 49 scaffolds, totaling 4.8 Mb; draft assembly was deposited in NCBI (GenBank accession JDWG00000000). Alignments between the JD6301 reference genome (GenBank Accession JDWH00000000) and JD8715 were performed with Burrows-Wheeler Alignment (bio-bwa.sourceforge. net; (Li and Durbin, 2009) and putative single nucleotide polymorphisms (SNPs) and small indels were identified using samtools/mpileup/bcftools (Li et al., 2009). This analysis resulted in the identification of 24 SNPs, which included 17 non-synonymous, 4 synonymous, and 3 non-coding variants (Table 1).

Growth responses and sugar consumption JD6301 and JD8715 (25 mL) were grown overnight at 30°C in a shaking incubator (200 rpm) in flasks containing MSM supplemented with 13 g/L glucose, diluted 1:100 in fresh media, and transferred to a 96-well microtiter plate. Optical density (OD600) measurements were recorded over 24 h in triplicate using a Molecular Devices spectrophotometer SpectraMax Plus 384 plate reader at 30°C with intermittent shaking. For glucose consumption analysis, 1 mL of cells was filtered through a 0.2 μm filter (Corning Life Sci214

ences, Amsterdam, The Netherlands). Glucose concentrations were determined using an Agilent 1100 High Performance Liquid Chromatography (HPLC; Agilent Technologies, Inc., Santa Clara, CA) system. The HPLC system was coupled to a Varian 385-LC evaporative light scattering detector (ELSD; Varian Inc., Palo Alto, CA) and a Restek Pinnacle II Amino column (5 μm, 150 × 4.6 mm; Restek, Inc., Bellefonte, PA). The temperature of the nebulizer in the ELSD was set to 60°C and the drift tube was held at 80°C with a nitrogen nebulization gas flow rate of 1.8 L/ min. The mobile phase consisted of acetonitrile and water (83:17) with an injection volume of 2 μL. The flow rate was 1 mL/min. Results represent the average of three independent replicates.

Transmission electron microscopy Two mL aliquots of 24 h cultures cultured at 30°C in a shaking incubator in MSM media supplemented with 3% sodium gluconate were centrifuged (10,000 x g) for 2 min at 4°C, fixed in ½ strength Karnovsky’s fixative in 0.1 M Na cacodylate buffer at pH 7.2, rinsed with 0.1 M Na cacodylate buffer, and then post fixed in buffered 2% osmium tetraoxide. Samples were rinsed once more in buffer, en bloc stained with 2% aqueous uranyl acetate, dehydrated in a graded ethanol series, and embedded in Spurr’s resin. Ultrathin sections were cut with a Reichert-Jung Ultracut 3 ultra-microtome and were stained with uranyl acetate and lead citrate. Stained sections were viewed on a JEOL JEM-100CXII TEM at 80KV. A minimum of 50 cells from two independent replicates was analyzed by transmission electron microscopy.

Scanning electron microscopy Both JD6301 and JD8715 strains were grown for 24 h at 30°C in MSM supplemented with 3% sodium gluconate in a shaking incubator. Cells (1 mL) were pelleted by centrifugation at 8,000 x g for 10 min and washed with 1 mL of chloroform, pelleted again by centrifuging for 10 min at 8,000 x g. Bacterial pellets

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were fixed in 2.5% glutaraldehyde in 0.1 M cacodylate buffer, washed in 0.1 M cacodylate buffer, postfixed in 1% osmium tetraoxide in 0.1 M cacodylate buffer, rewashed in distilled water, dehydrated in an ethanol series, and finally dried in a hexamethyldisilazane series as previously described (Merritt et al., 2010). Samples were sputter coated with gold-palladium using a Polaron SEM coating system prior to observation with a field emission scanning electron microscope (JEOL JSM-6500F). A minimum of 50 cells from two independent replicates was analyzed.

Live/Dead BacLightTM bacterial viability One μL (approximately 1x10 cells) of JD6301 and JD8715 was collected at 24 h of growth and viability was determined using the LIVE/DEAD BacLightTM bacterial viability kit consisting of SYTO 9 and propidium iodide (Invitrogen, Calsbad, CA). Staining was performed in the dark for 15 min, after which cells were analyzed by a BD FASCaliber flow cytometer using Cell Quest software (BD Biosciences, San Jose, CA). Analyses were performed on 4,500 to 5,000 cells (gated events) using instrument parameters previously described by others (Gunasekera et al., 2003). Percent viability was determined by a twoparameter comparison of green (live cells) and red (dead cells) fluorescent emission for individual bacteria using the formula: [% live green-emitting cells/ (% dead red-emitting cells + % live green-emitting cells)] x 100. Three independent replicates were performed for each strain. 7

lected supernatant and cell pellet using a standard Bligh and Dyer lipid extraction technique (Bligh and Dyer, 1959). Extracted lipids were derivatized using N-MethylN-(trimethylsilyl)-trifluoroacetamide (MSTFA) following ASTM D6584, which uses tricaprin as an internal standard at a concentration of 100 μg/mL. This method also utilizes triolein, diolein and monoolein as reference compounds. Briefly, tricaprin (12.5 μL), MSTFA (25 μL), and pyridine (62.5 μL) were added to the lipid extracts. Samples were vortexed and allowed to react for at least 20 min, after which 900 μL of n-heptane was added. Samples were then filtered through a 0.45 μm PTFE filter (SUN Sri, Rockwood, TN) and transferred to auto-sampler vials for analysis on a Varian 3600 GC (Varian Inc., Palo Alto, CA) equipped with a flame ionization detector (FID). The GC column was a RTX®-65TG (15m × 0.25 mm ID, with a 0.10 μm film thickness) and utilized a 2 m x 0.53 mm Rxi® guard column (Restek, Bellefonte, PA). Samples were analyzed using cool-on-column injection with an initial injector temperature of 50°C and a final injector temperature of 380°C, at a ramp rate of 180°C/min. The GC oven temperature was programmed at an initial temperature of 50°C, held for 1 min, then ramped to 180°C at 15°C/min, then to 230°C at 7°C/min, to 370°C at 20°C/min, and finally held for 11.20 min. The FID was retained at 380°C for the duration of the GC analysis. Analysis was performed on three independent replicates of each extraction for each strain.

FAMES analysis Lipid analysis Cultures (20 mL) incubated at 30°C for 24 h were centrifuged for 10 min at 6,000 x g, after which 10 mL of the supernatant was extracted for lipid analy-

Lipids extracted from JD6301 and JD8715 were converted to FAMEs using 1.5 mL of 14% BF3 in methanol at 65°C for 30 min, after which 5 mL of 5% NaCl and 2% NaHCO3 in distilled water was added. FAMEs were then extracted twice with 10 mL n-hex-

sis; the remainder of the supernatant was discarded. The resulting cell pellet was rinsed gently with 1 mL of chloroform, centrifuged for an additional 5 min, and the wash was combined with the 10 mL of supernatant collected. Lipids were extracted from the col-

ane and recovered from the solvent at 45°C under 15 psi of N2 using a TurboVap LV (Caliper Sciences, Hopkinton, MA). The solid residue was re-dissolved in 1 mL toluene containing 100 μg/mL butylated hydroxytoluene and 200 μg/mL 1,3-dichlorobenzene.

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Table 1. Summary of SNPs identified. SNP Ref-Mut

Scaffold/Pos

Gene

Function Non-synonymous

G-A

2/8877

P698DRAFT_00653

ABC-type branched-chain amino acid transport systems, periplasmic component

C-T

2/82327

P698DRAFT_00726

Paraquat-inducible protein B

C-T

2/99542

P698DRAFT_00743

Predicted membrane protein

G-A

2/250263

P698DRAFT_00887

Uncharacterized protein family (UPF0259)

G-A

4/321850

P698DRAFT_01861

Mg/Co/Ni transporter MgtE (contains CBS domain)

G-A

6/39934

P698DRAFT_02343

ATPase components of ABC transporters

G-A

6/182761

P698DRAFT_02476

Phosphotransferase system IIC components, glucose/maltose/N-acetylglucosamine-specific

G-A

6/193118

P698DRAFT_02492

2-methylthioadenine synthetase

C-T

6/233659

P698DRAFT_02537

Ribulose-5-phosphate 4-epimerase and related epimerases and aldolases

C-T

6/299293

P698DRAFT_02598

Cation transport ATPase

C-T

8/199438

P698DRAFT_03078

G:T/U mismatch-specific DNA glycosylase

T-C

9/90423

P698DRAFT_03214

Hemolysin activation/secretion protein

C-T

12/90473

P698DRAFT_03655

Flagellar hook-associated protein

G-A

13/4044

P698DRAFT_03706

SAM-dependent methyltransferases related to tRNA (uracil-5-)-methyltransferase

C-T

14/8347

P698DRAFT_03815

Protein of unknown function (DUF968)

G-A

18/63736

P698DRAFT_04198

7,8-dihydro-6-hydroxymethylpterin-pyrophosphokinase

C-T

32/7612

P698DRAFT_04585

Phage-related minor tail protein

Synonymous G-A

1/491858

P698DRAFT_00477

Transcriptional regulator

C-T

2/127810

P698DRAFT_00771

Alanine racemase

C-T

4/371589

P698DRAFT_01904

Glutamine synthetase

G-A

19/23709

P698DRAFT_04231

Sugar phosphate permease

Non-Coding

216

C-T

2/160082

C-A

4/240806

C-T

7/209964

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Figure 1. Inclusion bodies form within the cytoplasm of JD6301. TEM images were acquired at 24 h and 48 h. A minimum of 50 cells was observed for each time point. Scale bars represent 1 μm

24 h

48 h

The FAMEs were analyzed using an Agilent 6890N gas chromatograph equipped with a flame ionization detector (GC-FID) and a fused silica column Stabilwax-DA (30 m × 0.25 mm, film thickness 0.25 μm) (Restek, Bellefonte, PA). The operating conditions were as follows: initial oven temperature of 50°C for 2 min to a final oven temperature of 250°C with a rate of increase of 10°C/min and was held at 250°C for 18 min with helium as the carrier gas at 1.5 mL/min and 260°C detector temperature. Instrument calibration was achieved using a 14-component FAMEs standard mixture (C8 – C24) (Supelco, Bellefonte, PA) containing saturated, mono-unsaturated and polyunsaturated fatty acids. Analysis was performed on three independent replicates of each extraction for each strain. Analysis of variance (ANOVA), followed by a Tukey post-hoc range test (p < 0.05), was used

A novel strain of Enterobacter cloacae previously isolated by our group (Wilson et al., 2014) was analyzed by transmission electron microscopy, where inclusion bodies were found to form in the cytoplasm within 24 h of growth. By 48 h, it appeared that the inclusion bodies had formed around nearly the entire inner portion of the cell membrane (Fig. 1). As this culture was in a mixed environment containing oleaginous microorganisms, it was hypothesized that these may be representative of lipid inclusion bodies similar to what has been observed in other oleaginous microorganisms (Alvarez et al., 1996; Alvarez and Steinbuchel, 2002; Waltermann et al., 2005; Waltermann and Steinbuchel, 2005). Lipids were isolated by Bligh and Dyer extraction and based on the dry weight of the cells, 50% of the cellular weight was attributed to lipids, indicating that this is a novel

to analyze the significance of total biodiesel lipid analysis.

oleaginous isolate of Enterobacter cloacae. A limitation to the usefulness of oleaginous microorganisms in industry is in recovery of the useful end products (Grima et al., 2003). However, once the lipids are outside of the cell, separation from the aqueous solution is effortless due to the insolubil-

RESULTS AND DISCUSSION Lipid production by Enterobacter cloacae JD6301 and construction of the mutant JD8715

ity of lipids in water (Fischer et al., 2008). Therefore, this bacterium was modified to produce extracellular lipids. This strain was treated with the carcinogen EMS as previously described (Nojima et al., 1999) and the mutant JD8715 was subsequently identified

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Table 2. Yield of lipids produced relative to consumption of glucose. Strain

%Glucose Consumed Cells: glucose (±StDev) consumed

Yield extracellular lipid: Yield cell mass: glucose glucose

JD6301 99.51 (0.02)

4.62X1011 CFU/g

0.098g lipid/g glucose

0.0803g cells/g glucose

JD8715 85.86 (0.31)

1.56x1011 CFU/g

0.353g lipid/g glucose

0.0588g cells/g glucose

Table 3. Extracellular and intracellular lipid concentrations.

JD6301 (µg/ml, ±StDEV) Lipid

JD8715 (µg/ml, ±StDEV)

intracella

extracellb

intracella

extracellb

Monoglycerides

17.32 (2.85)

56.15 (0.19)

12.27* (0.81)

98.23* (3.40)

Diglycerides

8.27 (0.95)

16.03 (2.32)

10.65 (1.40)

18.52 (1.44)

Triglycerides

43.28 (0.01)

3.83 (0.21)

28.74 (0.22)

5.21*(1.41)

Intracellular concentrations of lipids are based on 20mL cell pellets collected from 24hr cultures of JD6301 or JD8715. StDev represents ± standard deviations. a

Extracellular concentrations of lipids are based on 10mL of supernatant collected from 24hr cultures of JD6301 or JD8715. StDev represents ± standard deviations. b

*Indicates significant change (p < 0.05) in concentration in JD8715 compared to JD6301.

as producing extracellular lipids through a screen for growth of an auxotrophic strain of S. cerevisiae. The genome of JD8715 was resequenced using the Illumina platform. A total of 52 scaffolds and 61 contigs were generated; genome size was 4,769,233 bp and a total of 4,508 protein coding genes were identified. The genomes of JD8715 and JD6301 were compared to identify locations of SNPs that may have contributed to the observed phenotype. A total of 24 SNPs were identified, with a majority being G->A or C->T transitions (Table 1). Seventeen of the identified SNPs resulted in a non-synonymous substitution. A majority of these SNPs were linked to membrane proteins, including ATP synthase components, transporter proteins, and a hook associated 218

protein. It is possible that the resulting phenotype was due to a combination of these SNPs. Individual mutants need to be generated to determine which mutation(s) is sufficient for the release of lipids.

Lipid and sugar analysis The mutant selection procedures and microscopic observations suggested that the JD8715 mutant had extracellular lipids. Quantification of extracellular and intracellular lipids, extracted from the supernatant and cell pellets respectively, from JD6301 and JD8715 was performed using gas chromatography after 24 h of growth. Mass of the re-

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Figure 2. Total FAMEs and lipid profiles of JD6301 and JD8715. (A) Total mean FAMEs yield (± SD) of JD6301 (black) and JD8715 (white) of intracellular and extracellular fractions. (B) FAMEs profile (± SD) of JD6301 intracellular (black), JD6301 extracellular (blue), JD8715 intracellular (white), JD8715 supernatant (grey). Intracellular concentrations of lipids are based on 20 mL cell pellets collected from 24 h cultures of JD6301 or JD8715. Extracellular concentrations of lipids are based on 10 mL of supernatant collected from 24 h cultures. Means denoted by the same letter are not significantly different (p <0.05). A

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Figure 3. Enterobacter sp. JD6301 and JD8715 mutant exhibit similar growth and viability. (A) Viable growth curves of JD6301 (▀) and JD8715 (□) represent the average of three independent replicates. (B) Presence of extracellular lipids is most likely not due to lysis of JD8715. JD6301 and JD8715 cells were treated with chloroform. Scale bars represent 1 μm. A

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sulting lipid extractions was determined (in lieu of “media only” controls) to obtain the yield of lipid production relative to glucose consumption (Table 2). JD6301 produced a greater cell mass yield per gram of sugar consumed (0.0803 g/g sugar) than the mutant JD8715 (0.0588 g/g sugar). However, JD8715 consumed less sugar (85.86% versus 99.51% for WT) and also produced greater amounts of extracellular lipids (0.353 g lipid/ g sugar for JD8715 versus 0.098 g lipid/ g sugar for JD6301 WT; Table 2), suggesting that more energy may have been directed towards lipid production rather than cell replication. This is further supported by the reduced cell mass of JD8715 and therefore the greater production of

detected for JD8715 and WT, which could include odd-numbered fatty acids.

extracellular lipids compared to WT. Lipid analyses indicated that the extracellular lipids present in the JD8715 culture consisted of monoglycerides, diglycerides, and triglycerides and that the extracellular quantity of mono- and triglycerides was significantly different between the JD6301 and JD8715 strains (Table 3). The mutant had a nearly four-fold increase in extracellular lipids. However, comparing the cell pellet (intracellular) lipid composition from JD8715 to JD6301 indicated that the mutant had significantly less amounts of intracellular monoglycerides and triacylglycerides. No significant difference was observed in the concentration of diacylglycerides present in the JD8715 cells as compared to the WT intracellular lipid composition. The predominant lipids identified in the extracellular fraction were monoglycerides. It is possible that the difference in the quantity of triglycerides between the combined intracellular and extracellular fractions of JD8715 (33.95 μg/mL) and the triglyceride concentration identified in the WT (43.28 μg/mL) is due a defect in TAGs formation. The total FAMEs of JD8715 extracellular lipids were not significantly different from the intracellular yield of WT (Fig. 2A). The extracellular yield from JD8715 was significantly greater than the extracel-

similar viability at 24 h (77% versus 78%). The growth of the JD8715 strain was compared to WT to determine whether the two strains grew similarly under standard growth conditions. Both strains exhibited similar growth patterns, indicating that the external production of lipids did not affect the growth of the mutant for at least the first 24 h of growth (Fig. 3A). To further determine whether the increase in the presence of extracellular lipids was due to cell lysis occurring during the wash that precedes the extraction procedure, cells were washed with chloroform and analyzed by scanning electron microscopy (SEM). Results indicated that out of a minimum of 50 cells analyzed, 98% of WT and 96% of JD8715 cells remained intact with no structural deformities following the chloroform wash (Fig. 3B), indicating that the chloroform wash that precedes the lipid extraction procedure did not unintentionally lyse the cells, therefore skewing the extracellular fraction.

lular yield of WT and the intracellular yield of JD8715 (p < 0.05). Furthermore, the FAMEs profiles between WT and JD8715 was similar, suggesting a consistency between strains in regard to lipids produced (Fig. 2B). A large amount of total unknown FAMEs were

this is the first report of a genetically altered form of an oleaginous bacterium that is capable of producing extracellular lipids. The formation of lipid inclusion bodies initiates at the cell membrane (Waltermann et al., 2005) proposing several potential areas

Growth and viability of JD8715 and WT JD6301 To determine if the presence of extracellular lipids was due to the JD8715 cultures containing more lysed or dead cells as compared to JD6301, the percentages of live cells for both the mutant and WT were determined using the LIVE/DEAD BacLightTM bacterial viability kit (Invitrogen). These results indicated that the JD8715 and JD6301 strains exhibited

CONCLUSIONS To the author’s knowledge, this is the first report of a strain of Enterobacter cloacae that is capable of producing large quantities of lipids. Additionally,

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of mutation associated with membrane transport and integrity resulted in the observed phenotype of JD8715. The elucidation of this mechanism would allow application of similar mutations to other oleaginous microorganisms for optimization of extracellular lipid production, providing a substantial advancement to the biofuels industry.

ACKNOWLEDGEMENTS We would like to thank John Brooks, Karen Coats, Kendrick Currie, Linda MacFarland, Julie Newton, John Stokes, Darrell Sparks, and Justin Thornton for their assistance and helpful discussions with this project. This project was funded by the Northeast Mississippi Daily Journal Undergraduate Research Award to JMB and by the Mississippi State University Sustainable Energy Research Center funded through the Department of Energy to JRD. The work conducted by the DOE Joint Genome Institute is supported by the Office of Science of the DOE under contract number DE-AC02-05CH11231.

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kumar. 2008. An overview of enzymatic production of biodiesel. Bioresour. Technol. 99: 3975-3981. Rosen, E.D. and B.M. Spiegelman. 2006. Adipocytes as regulators of energy balance and glucose homeostasis. Nature. 444: 847-853.

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Schlegel, H.G., H. Kaltwasser, and G. Gottschalk. 1961. Ein Submersverfahren zur Kultur wasserstoffoxydierender Bakterien: Wachstumsphysiologische Untersuchungen. Arch. Microbiol. 38: 209222. Stukey, J.E., V.M. McDonough, and C.E. Martin. 1989. Isolation and characterization of OLE1, a gene affecting fatty acid desaturation from Saccharomyces cerevisiae. J. Biol. Chem. 264: 16537-16544. Waltermann, M., A. Hinz, H. Robenek, D. Troyer, R. Reichelt, U. Malkus, H.J. Galla, R. Kalscheuer, T. Stoveken, P. von Landenberg, and A. Steinbuchel. 2005. Mechanism of lipid-body formation in prokaryotes: how bacteria fatten up. Mol. Microbiol. 55: 750-763. Waltermann, M. and A. Steinbuchel. 2005. Neutral lipid bodies in prokaryotes: recent insights into structure, formation, and relationship to eukaryotic lipid depots. J. Bacteriol. 187: 3607-3619. Wilson, J.G., W.T. French, A. Lipzen, J. Martin, W. Schackwitz, T. Woyke, N. Shapiro, J.W. Bullard, F.R. Champlin, and J.R. Donaldson. 2014. Draft genome sequence of Enterobacter cloacae strain JD6301. Genome Announc. 2. Wynn, J.P. and C. Ratledge. 2005. Oils from microorganisms. In Bailey’s Industrial Oil and Far Products. Shahidi F., ed. (John Wiley and Sons, Inc.).

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Survival of Salmonella enterica and Listeria monocytogenes in manure-based compost mixtures at sublethal temperatures M.C. Erickson1, C. Smith2, X. Jiang3, I.D. Flitcroft4, and M.P. Doyle1 1

Center for Food Safety and Department of Food Science and Technology, University of Georgia, Griffin, GA 2 Food Safety Net Services, Atlanta Laboratory, Covington, GA 3 Department of Food, Nutrition and Packaging Sciences, Clemson University, Clemson, SC 4 Department of Crops and Soil Science, University of Georgia, Griffin, GA

ABSTRACT Aerobic composting of animal manures has been advocated as an effective management tool to inactivate resident zoonotic pathogens where the time at lethal temperatures is used to determine the effectiveness of the treatment. In the absence of meeting these process conditions, the relative contributions of other physical factors on growth and persistence of zoonotic pathogens is vague and therefore the required storage time necessary for elimination of pathogens cannot be adequately estimated. This study explored the influence of sublethal temperatures, moisture levels, and light exposure on the survival of Salmonella and Listeria monocytogenes in compost mixtures that were prepared with three different sources of manure (dairy cow, swine, and chicken). As ambient temperatures increased from 20°C to 40°C, persistence of both pathogens decreased, which was likely due to the increased competitive activity of the more dominant indigenous microflora. During storage at 30°C, evaporation of water from compost mixtures occurred rapidly. Under those conditions, populations of L. monocytogenes declined in cow compost mixtures throughout a 4-week storage period, whereas Salmonella populations increased. In chicken compost mixtures at 30°C, populations of both pathogens decreased only during the first week of storage, which was likely due to the antimicrobial properties of ammonia initially present in chicken manure. When stored at 20°C, L. monocytogenes populations decreased more rapidly when compost mixtures were exposed to more intense light conditions whereas no discernible differences in Salmonella populations occurred in swine or cow compost mixtures under the different light conditions. These results indicate that developing safety guidelines for times to hold compost mixtures at sublethal temperatures, prior to land application, will be challenging. Keywords: Salmonella, Listeria monocytogenes, compost, dairy, swine, chicken, temperature, moisture, light, manure Agric. Food Anal. Bacteriol. 4: 224-238, 2014

Correspondence: M.C. Erickson, mericks@uga.edu Tel: +1 770-412-4742 Fax: +1 770-229-3216

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INTRODUCTION Livestock and poultry production are major enterprises worldwide that in addition to the production of food, waste by-products that include solid manure and manure slurries are also produced in large quantities. For example, the Environmental Protection Agency (EPA) estimated that 1.1 billion tons of manure was produced annually within the U.S. (US EPA, 2013), with cattle contributing the greatest proportion (83%), followed by swine (10%), and poultry (7%). Land application of these wastes has been one of the most cost effective approaches to dispose of such large quantities of manure, with 5% of all crop-

occur during winter composting or if piles are not turned to expose the surface material to sufficient internal heat, the holding time of compost materials to ensure pathogen inactivation is uncertain. Compared to lethal heat exposure, the contribution of other physical factors (e.g., non-lethal temperatures, light, and desiccation) to inactivation of zoonotic pathogens in manure-based compost mixtures has not been elucidated because biological (i.e., competition for nutrients) and chemical (e.g., ammonia, volatile acids or other antimicrobials) factors that affect pathogen inactivation are also likely affected by the physical parameter. Such is the case with soil systems in which increased temperatures,

land (15.8 million acres) in 2006 reported as having been fertilized with livestock manure (MacDonald et al., 2009). Zoonotic pathogens are sporadically resident within animal manure (Le Bouquin et al., 2010; LeJeune et al., 2006; Lomonaco et al., 2009). Hence if manure is applied to land, these pathogens can contaminate the soil, crops grown in those fields, and waterways that collect runoff from the fields. Aerobic composting of animal wastes can inactivate zoonotic bacterial pathogens while creating a stable amendment that improves soil quality and fertility (Berry et al., 2013; Raviv, 2005). Heat generated from the metabolic activity of thermophilic microorganisms in manure piles that are self-insulating is the primary mechanism for inactivating zoonotic pathogens (Pell, 1997; Wichuk and McCartney, 2007). Hence, process conditions for composting manures in the U.S. are based on EPA’s regulations for composting biosolids that includes either a minimum temperature of 55°C for 3 days in aerated static piles or in-vessel systems, or 55°C for 15 days in windrow systems. Moreover, in the windrow systems, the material must be turned a minimum of 5 times to ensure that all material is subjected to the necessary thermal conditions (US EPA, 1999a). Composting at 40°C for 120 h or more, during which time the tem-

despite being near the organism’s optimal growth temperature, led to greater inactivation of Escherichia coli O157:H7 as a result of an accompanying increase in competition by the dominant native microbial community (Semenov et al., 2007). Similarly, the effect of moisture levels on the fate of pathogens (Salmonella and E. coli O157:H7) or their surrogates in soil systems has been dependent on the pathogen population levels relative to the levels of the indigenous microbial community (Lang et al., 2007; Ongeng et al., 2011) and likely play a similar role in compost mixtures. Ammonia that is generated during the composting process (Beck-Friis et al., 2003) and has been shown to be an antimicrobial agent toward Salmonella and Listeria monocytogenes in chicken and cattle manure (Himathongkham and Riemann, 1999; Park and Diez-Gonzalez, 2003) is also affected by moisture levels, with drying of manure accelerating the volatilization of ammonia (Gotaas, 1956) and inhibiting the conversion of nitrogenous compounds to aqueous ammonia (Hutchison et al., 2000). Considering the complex interactions that moisture and temperature exert on the activity of indigenous microbial communities, it is of interest to investigate the role of moisture levels on inactivation of pathogens in compost mixtures that would likely

perature exceeds 55°C for 4 h, has also been designated by EPA in Appendix B of the 503 Regulations as a process to significantly reduce pathogens (US EPA, 1999b). Unfortunately, when these EPA criteria are not met (Wichuk and McCartney, 2007), as could

be populated with different indigenous microflora from the different nitrogen feedstocks. Another physical factor that has received little attention for its involvement in inactivating pathogens in manure-based compost mixtures is sunlight. Due

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to its inability to penetrate compost mixtures, sunlight would be lethal to pathogens primarily at the surface of compost mixtures as has occurred at the surfaces of natural waterways and lagoons (Davies and Evison, 1991; Maynard et al., 1999). Hutchison et al. (2005) postulated that lack of surface contamination by Salmonella, Listeria, Campylobacter, or E. coli O157:H7 in composted static pile wastes after eight days was due to exposure of their surfaces to sunlight; however, the experiment lacked a control sample not exposed to sunlight. In contrast, Erickson et al. (2010) were able to detect both Salmonella and Listeria on the surface of static piles comprised of chicken litter and peanut hulls after composting

MATERIALS AND METHODS

for 14 and 56 days in the summer and winter, respectively. To gauge the potential impact of sunlight on pathogens in compost more specifically, results of a study on the survival of pathogens in beef cattle fecal pats is presented here for comparison (Meays et al., 2005). In that experiment, E. coli survival under 4 different levels of solar exposure (controlled by using a shade cloth) was determined. After 45 days, fecal pats under the 0% shade cloth had the least surviving E. coli, followed by the 40%, 80%, and 100% treatments. A similar response in non-turned composting systems could result in longer recommended holding times for regions with a large number of overcast days compared to regions that are dominated by sunny days. The purpose of this study was to determine the influence of several physical factors (i.e., temperature, level of light exposure, and moisture levels) on the inactivation of Salmonella and L. monocytogenes in compost mixtures that were stored in environmental chambers at temperatures ranging from 20°C to 40°C in amounts that would not be self-insulating. To account for the potential confounding influence of indigenous microflora on pathogen inactivation, this variable was addressed indirectly by utilizing manure in compost mixture formulations from different

the construction of these GFP strains has been described by Ma et al. (2011) and they also reported that the loss of the GFP-plasmid after 20 generations, indicative of its stability, has ranged from 15 to 77% and 8 to 52% for the Salmonella and L. monocytogenes strains, respectively. Frozen stock cultures of each GFP-labeled Salmonella strain and GFP-labeled L. monocytogenes strain were thawed and streaked onto tryptic soy agar (Difco, Becton Dickinson, Sparks, MD) containing 100 µg/ml ampicillin (TSA-A) and brain heart infusion agar (Becton Dickinson) containing 8 µg/ml erythromycin (BHIA-E), respectively. Following incubation at 37°C for 20 to 24 h, individual colonies from each plate were subsequently streaked onto a second plate that was incubated for another 20 to 24 h at 37°C. Individual Salmonella and L. monocytogenes colonies from these plates were then inoculated into 100 ml of tryptic soy broth (Becton Dickinson) containing 100 µg/ml of ampicillin (TSB-A) and 100 ml brain heart infusion broth (Becton Dickinson) containing 8 µg/ml erythromycin (BHIB-E), respectively. Broths were incubated at 37°C for 20 to 24 h with agitation (150 rpm) and bacteria were subsequently harvested by centrifugation (4,050 x g, 15 min, 4°C) with cell pellets being washed three times in 0.1% peptone water (Difco,

sources (dairy, chicken, and swine) that should have different microbial compositions.

Becton Dickinson). Reconstitution of the individual strains in 0.1% peptone water to an optical density of 0.5 (approximately 109 CFU/ml) was made prior to combining equal volumes of each strain to comprise one four-strain mixture of Salmonella and one five-

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Pathogen Strains and Preparation Three strains of green-fluorescent protein (GFP)labeled Salmonella enterica serovar Enteritidis (ME18, H4639, and H3353) and one strain of GFP-labeled S.enterica serovar Newport containing an ampicillinresistant marker were selected from the culture collection at the University of Georgia, Center for Food Safety (Griffin, GA). Five strains of GFP-labeled L. monocytogenes containing an erythromycin-resistant marker (12443, H7550, G3982, 101M, and F6845) were also selected from the culture collection. Details on

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strain mixture of L. monocytogenes. Salmonella and L. monocytogenes populations were determined by plating on TSA-A and modified Oxford agar (Acumedia Manufacturers, Lansing, MI) containing 8 µg/ ml erythromycin (MOX-E), respectively. Salmonella transformed colonies emitted bright green fluorescence when viewed at 365 nm under a handheld UV light (Fotodyne Inc., Hartland, WI); however, visualization of fluorescent L. monocytogenes transformed colonies required use of a Leica MZ16 FA stereo fluorescence microscope (Bannockburn, IL).

Compost Feedstocks and Chemical Analysis

(C:N) ratios of 20:1 to 40:1. Compost amendments and inoculated manure were mixed thoroughly for ca. 5 min in a Hobart mixer (model D320; ¾ h.p.) prior to distributing the mixtures into containers for experimental studies.

Experimental Design Four studies were conducted that varied in their experimental design. In the first experimental study investigating the role of temperature on survival of Salmonella in manure, no carbon amendment was added to the manure source (dairy cow manure and

Each type of manure was individually sprayed in a 28-L sanitized bowl with either GFP-labeled Salmonella or both GFP-labeled-Salmonella and L.

chicken litter) that were each obtained at two separate times. Dairy cow manure (2 kg) or chicken litter (1 kg) was sprayed with the Salmonella inoculum mixture (20 ml or 10 ml of 7 log CFU/ml, respectively) to obtain ca. 5 to 6 log CFU/g. Inoculated material (100 g) was placed into multiple square (12.7 cm2) Ziplock plastic containers (S.C. Johnson & Sons, Racine, WI). With the first batch of dairy cow manure, three containers were held at 25°C and another three were held at 40°C. With the second batch of dairy cow manure, three containers were held at 35°C and another three were held at 40°C. For inoculated chicken litter, the first batch was stored in three separate containers only at 25°C whereas the second batch was filled into three containers that were stored at 40°C only. Samples were removed from each container initially and after 3 days of storage for analysis of Salmonella and mesophilic and thermophilic bacteria. Only one replicate trial of this experimental design was conducted. The second experimental study investigated the role of temperature on survival of enteric pathogens in manure-based compost mixtures. Chicken litter was collected at six separate times, with each collection being used as an independent replicate trial. Each batch of chicken litter was sprayed with both an

monocytogenes to give initial cell populations of 3.3 to 7.5 log CFU/g. Carbon amendments and sterile deionized water were then added to the inoculated manure to comprise formulations having initial levels of 30% or 60% moisture and initial carbon:nitrogen

inoculum of Salmonella and an inoculum of L. monocytogenes before blending with wheat straw, cottonseed meal and sterile deionized water to obtain mixtures having an initial carbon:nitrogen (C:N) ratio of 40:1 and 60% moisture content Initial pathogen pop-

Three sources of manure including dairy cow manure, swine manure, and broiler chicken litter were used as the primary nitrogen source for compost mixtures. These materials were collected from farms located near Griffin, GA, and upon arrival at the laboratory were frozen for at least 24 h to kill the majority of insect eggs (Sherman et al., 2006). Carbon amendments in compost formulations (i.e., wheat straw and cottonseed meal) were purchased from a local feed supply store. To improve the homogeneity of compost formulations, wheat straw was shredded using a Flowtron Leaf Eater (Malden, MA) for lengths of 1 to 5 cm. Carbon, nitrogen, and moisture content analyses were conducted on all raw ingredients used in the compost mixtures (Erickson et al., 2010) to assist in determining recipes for formulation of compost mixtures.

Compost Mixture Formulation

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ulations in three of the batches was targeted at a low level (ca. 3.5 log CFU/g) while another three batches had a target at a higher level (ca. 6.7 log CFU/g). Duplicate samples from each inoculated mixture were obtained for pathogen and moisture content analysis prior to distributing compost mixtures into two uncovered translucent plastic cups (8.5 cm diameter x 5 cm height, ca. 45 g/cup). Cups containing the low level inoculum were placed in an environmental chamber at 30°C with a 12-h light and 12-h dark cycle whereas cups containing the high level inoculum were placed in a dark environmental chamber set to 20°C. In the lighted chamber, light was supplied by ten 400W Metal Halide MVR400/U bulbs (General

back to their initial weights and original moisture contents by spraying the sample with a light mist of sterile deionized water. Two cups whose moisture contents had been adjusted were also removed from each of the four treatments (manure source x target moisture content) at 2, 3, and 4 weeks for analysis of pathogen populations, moisture content, and pH. Three independent replicate trials in which samples were not adjusted for moisture content were conducted whereas two independent replicate trials were conducted for samples adjusted for moisture content. For each independent trial, the dairy cow manure and chicken litter were collected at separate times and were inoculated with a different batch of

Electric, Cleveland, OH) and ten 400W high pressure sodium lamps LU400/H/ECO, LUCALOS bulbs (General Electric). High pressure sodium lamps emit no ultraviolet (UV) light and while the metal halide bulbs emit a small band of long band UV light (ca. 375 nm), compost mixtures were not exposed as this UV light was filtered out by diffusive panels separating the lamps and chamber. The compost mixtures of all batches were held for two weeks at the specified temperature after which time the cups were removed and mixtures assayed for surviving pathogens. For the third experimental study, both dairy cow manure and chicken litter were used as nitrogen sources to determine the role of moisture content on the survival of Salmonella and L. monocytogenes. Manure or litter was initially sprayed with an inoculum of Salmonella and an inoculum of L. monocytogenes and, then mixed with wheat straw, cottonseed meal, and sterile deionized water to obtain cow and chicken compost mixtures having an initial C:N ratio of 20:1, initial moisture contents of either 30% or 60%, and initial pathogen populations of ca. 3 to 4 log CFU/g. The cow and chicken compost mixtures were then distributed into small translucent cups (ca. 45 g/cup) used in study 2. The cups were stored uncovered in an environmental chamber at 30°C with a

pathogen inocula. For the fourth experimental study, three different sources of manure (dairy cow manure, chicken litter, and swine manure) were inoculated and incorporated into compost mixtures to examine the effect of light on survival of Salmonella and L. monocytogenes. Wheat straw and cottonseed meal served as the carbon amendments and were mixed with the Salmonella- and L. monocytogenes-inoculated manure sources and sterile deionized water to obtain cow-, chicken-, and swine-compost mixtures having an initial C:N ratio of 30:1, an initial moisture content of 60%, and pathogen populations of ca. 6.7 to 7.5 log CFU/g. Duplicate samples from each of the three contaminated compost mixtures were analyzed for initial pathogen populations, moisture content, and pH. The remainder of each compost mixture was then distributed into the small translucent cups (ca. 45 g/cup) used in studies 2 and 3 described above. The cups for each treatment group were then divided into three groups. One group was placed in a dark environmental chamber, the second group was placed in an environmental chamber where all bulbs were turned on to simulate daily “bright” sunny conditions (12 h at 524-573 µmol/m2/sec and 12 h in the dark), and the third group was placed in an environmental

12-h light (602 µmol/m2/sec) and 12-h dark cycle for up to 4 weeks. Two cups from each treatment were removed initially and at weekly intervals and analyzed for pathogen populations, moisture content, and pH. In addition, half of the remaining cups were adjusted

chamber where only half the bulbs were turned on to simulate daily “cloudy” conditions (12 h at 289359 µmol/m2/sec and 12 h in the dark). All chambers were at 20°C. Duplicate samples from each treatment at five sample times over the course of 4 weeks,

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12 weeks, and 18 weeks for swine, chicken, and cow compost mixtures, respectively, were enumerated for pathogens and assayed for moisture content and pH. Using the above experimental design, three independent replicate trials were conducted with each trial using manure collected at separate times and a different batch of pathogen inoculum cultured for inoculation of the manure.

Microbiological and pH Analyses Both direct plating and selective enrichment culture was used for detection of GFP-labeled Salmonella and L. monocytogenes. In direct plating assays, compost sample (5 g) was added to 45 ml of 0.1% peptone water in a sterile Whirl-Pak bag and mixed in a stomacher. Ten-fold serial dilutions of this homogenate were made prior to spreading on TSA-A or MOX-E plates for enumeration of GFPlabeled Salmonella or L. monocytogenes, respectively. Selective enrichment culture for Salmonella and L. monocytogenes consisted of adding compost sample (5 g) directly to 45 ml of TSB-A or BHIB-E, respectively, and incubating at 37°C for 24 h. These enriched samples were then streaked on TSA-A or MOX-E plates to determine the presence or absence of Salmonella or L. monocytogenes, respectively, at a detection limit of 20 cells/100 g. To determine initial levels of mesophilic and thermophilic microbial populations in chicken and cow manure, ten-fold serial dilutions of the stomached homogenates were spread onto Difco plate count agar (Becton Dickinson, Sparks, MD). Colonies of mesophilic and thermophilic bacteria were counted after overnight incubation at 30°C and 55°C, respectively. Measurement of pH was determined with an Acumet Basic pH meter (Fisher Scientific, Pittsburgh, PA) on 5-g compost samples dispersed in 250 ml of deionized water. Compost mixtures were analyzed for moisture using the same procedure described for moisture analysis of feedstuffs.

Statistical Analyses Salmonella and L. monocytogenes populations in samples for each independent trial were converted to logarithmic values prior to determining differences from initial population levels. Logarithmic pathogen decreases, % moisture content, and pH values were subjected to the general linear- and one-way analysis of variance (ANOVA) test using the StatGraphics Centurion XV software package (StatPoint, Inc., Herndon, VA). When statistical differences were observed (P < 0.05) with the ANOVA test, sample means were differentiated with the least significant difference test (P = 0.05).

RESULTS AND DISCUSSION Several studies have previously addressed the survival of zoonotic pathogens in manure at ambient temperatures (Himathongkham et al., 1999a, b; 2000; Sinton et al., 2007); however, this type of study was repeated in our preliminary study (first experimental study) with locally-obtained manure to give some baseline information on the fate of Salmonella and other indigenous microflora in the absence of a carbon amendment. Different responses were observed for Salmonella and the indigenous microflora depending on the manure source and storage temperatures. Following a 3-day storage period, no changes in populations of the indigenous microflora (mesophilic and thermophilic bacteria) occurred when present in cow manure and stored at 25°C (Table 1). In contrast, the populations of both mesophilic and thermophilic bacteria increased in cow manure stored at 35°C or 40°C, but decreased in chicken litter stored at 40°C for a similar time period. Salmonella decreases occurred in both manure sources when held at 40°C, but reductions were substantially greater in chicken litter than in cow manure. Salmonella populations also decreased in chicken litter held at 25°C, but increased in cow manure held at 25°C or 35°C. Transient increases in Salmonella population have been observed previously in cow

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Table 1. Indigenous bacterial populations in chicken and cow manure and fate of Salmonella when stored for 3 days at temperatures between 25°C and 40°C1. Chicken manure2 Day

25°C

40°C

25°C

Mesophilic bacteria populations (log CFU/g)5

ND4

9.38 ± 0.13 a

9.22 ± 0.10 a

6.88 ± 0.41 b 6.88 ± 0.06 b

7.65 ± 0.14

5.86 ± 0.33 b

9.51 ± 0.16 a

8.88 ± 0.41 a 8.44 ± 0.64 a

Thermophilic bacteria populations (log CFU/g)5

7.08 ± 0.02 a

8.36 ± 0.08 a

6.35 ± 0.01 b 6.35 ± 0.01 b

8.32 ± 0.25

5.86 ± 0.06 b

8.54 ± 0.17 a

8.46 ± 0.38 a 8.19 ± 0.83 a

↓7 1.21

↓ 5.56

↑ 8 1.88

Salmonella’s fate after 3 days of storage (Δ log CFU/g)6 1 2 3

4 5

6 7 8

Cow manure3 35°C

↑ 1.11

40°C

↓ 0.89

Data collected from first experimental study. Mesophilic and thermophilic bacteria population levels are mean ± S.D., n = 3. Mesophilic and thermophilic bacteria population levels are mean ± S.D., n = 3 for 25°C and 35°C samples, n = 6 for 40°C samples. Not determined. Values for this parameter within each column followed by a different letter are significantly different (P < 0.05). Salmonella initial populations in manure samples ranged from 5 to 6 log CFU/g. Decrease in population. Increase in population.

manure held at ambient temperatures (Himathongkham et al., 1999a; Sinton et al., 2007) and a minimal water content of 80% was a prerequisite (Sinton et al., 2007). Salmonella die-off in chicken manure has also been documented and has been ascribed to the generation of ammonia (Himathongkham et al., 1999b; 2000) which is an antimicrobial (Himathongkham and Riemann, 1999). For the current study, it is plausible that the production of ammonia by the indigenous microflora could be stimulated by the

ments, wheat straw and cottonseed meal, to create compost mixtures prior to their storage. When compost mixtures were formulated to an initial moisture content of 60% and an initial C:N ratio of 40:1, storage for 2 weeks at 20°C in the dark led to L. monocytogenes reductions of 0.97 ± 0.66 log CFU/g. In contrast, Salmonella populations remained relatively constant (increase of 0.16 ± 0.56 log CFU/g) over the same time period. Storage of chicken compost mixtures at 30°C for the same time interval but un-

presence of bedding material that was included during collection of the chicken manure. For the second experimental set of studies, temperature was the main variable of interest, but chicken litter was mixed with the carbon amend-

der lighted conditions, however, resulted in population decreases for both L. monocytogenes (1.87 ± 1.22 log CFU/g loss) and Salmonella (0.89 ± 1.54 log CFU/g loss). This trend of increased pathogen inactivation with increasing ambient temperatures

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Table 2. Comparison of Salmonella and L. monocytogenes losses in chicken and cow manurebased compost mixtures1 stored for up to 4 weeks at 30°C2. Cumulative pathogen reduction (log CFU/g)3, 4 L. monocytogenes5

Salmonella6

Week

Chicken manure compost mixture

Cow manure compost mixture

Chicken manure compost mixture

Cow manure compost mixture

2.09 ± 0.67 a

0.59 ± 1.27 b

1.36 ± 1.62 a

-0.35 ± 1.73 a-c

1.87 ± 1.22 a

0.60 ± 1.25 b

0.89 ± 1.54 ab

-0.82 ± 1.44 bc

2.11 ± 0.93 a

1.15 ± 1.22 ab

1.11 ± 1.50 ab

-1.31 ± 2.10 c

2.11 ± 0.93 a

2.21 ± 0.82 a

0.90 ± 1.64 ab

-0.58 ± 1.56 bc

Compost mixtures were formulated with carbon amendments to give an initial carbon:nitrogen ratio of 20:1 and pathogen populations of ca. 3.5 log CFU/g.

Data collected from third experimental study.

Pathogen data collected from treatments evaluating compost mixtures formulated to either an initial moisture content of 30% or 60% and then either readjusted to the original moisture content on a weekly basis or left undisturbed were not significantly different. The data were therefore pooled prior to statistical analysis and displaying the data in this table by the manure source used in the compost mixture.

Pathogen reductions were calculated by subtracting the population level at each time period from the initial population level.

Values for this pathogen (mean ± S.D.), across both rows and columns, followed by a different letter are significantly different (P < 0.05).

is similar to the patterns in soil of pathogen survival previously documented (Lang et al., 2007; Semenov et al., 2007). Based on those studies, the investigators suggested that increases in temperature, despite being close to the pathogen’s optimal growth temperature, increased the competitive activity of the more dominant indigenous microflora which adversely affected the pathogen’s survival (Lang et al., 2007). This explanation may be the basis for the decreased pathogen persistence we observed in chicken compost mixtures as exposure temperatures

their influence on L. monocytogenes and Salmonella inactivation in compost mixtures stored for up to 4 weeks at 30°C. However, under these conditions, populations of either pathogen were not affected by the initial moisture contents nor did weekly additions of water to return the compost mixtures to their original moisture contents affect the reduction of pathogens (P > 0.05). Moisture analysis of the compost mixtures revealed that water was lost very quickly from the samples stored in uncovered containers and equilibrated to approximately the same

increased. Initial moisture contents (30% or 60%) and weekly adjustment of the moisture contents of chicken- and cow compost mixtures were the two variables of interest in our third experimental study in determining

percentage of moisture (9.7 ± 2.7%) regardless of the initial moisture content or when weekly additions of water were applied to the mixtures. These conditions were therefore likely responsible for the failure of moisture content to have an effect on patho-

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Table 3. pH of chicken and cow manure-based compost mixtures1 following storage at 30°C for 4 weeks when initial moisture contents were either 30% or 60%2. pH3 Chicken manure compost mixture

Cow manure compost mixture

Week

30% moisture

60% moisture

30% moisture

60% moisture

7.25 ± 0.66 a-f

7.58 ± 1.22 a-d

6.84 ± 0.33 d-f

6.64 ± 0.34 f

7.06 ± 0.32 b-f

7.85 ± 0.66 a

6.80 ± 0.29 ef

7.46 ± 0.21 a-e

6.97 ± 0.34 c-f

7.63 ± 0.59 a-c

6.77 ± 0.22 ef

7.09 ± 0.10 b-f

6.99 ± 0.42 c-f

7.75 ± 0.34 ab

6.81 ± 0.16 ef

7.32 ± 0.29 a-f

6.87 ± 0.41 d-f

7.74 ± 0.24 ab

6.77 ± 0.08 ef

7.45 ± 0.22 a-e

Compost mixtures were formulated with carbon amendments to give an initial carbon:nitrogen ratio of 20:1.

Data collected from third experimental study.

Values within the table (mean ± S.D.) followed by a different letter are significantly different (P < 0.05).

gen inactivation. Hence, the data were pooled to determine the changes that occurred in pathogen populations in the chicken and cow compost mixtures (Table 2). As with dairy manure in the absence of a carbon amendment and held at 25°C or 35°C (Table 1), the populations of Salmonella in the cow compost mixtures increased from initial populations after holding of the materials at 30°C and remained at these higher levels over the 4-week trial (Table 2). In contrast, L. monocytogenes populations declined in cow compost mixtures after only one week of storage at 30°C and additional significant decreases occurred up to 4 weeks of storage (P < 0.05). Over all time periods, there were significantly greater reductions in Salmonella and L. monocytogenes popula-

Monitoring the pH of the chicken- and cow manure-based compost mixtures during storage at 30°C for 4 weeks (third experimental study) revealed that initially the chicken compost mixtures were approximately 0.5 pH units higher than the cow compost mixtures (Table 3). Although not determined in this study, the ammonia present that has previously been associated with higher pH values in chicken manure (Himathongkham et al., 1999b; 2000) could have been the principal factor responsible for the die-off of pathogens in the chicken compost mixtures. It appears, however, that pH alone may not be used as an indicator of a compost mixture’s capacity to sustain viable pathogen populations. In the cow compost mixtures having an initial moisture content

tions in the chicken compost mixtures compared to the cow compost mixtures (P < 0.05). Interestingly, the decreases in pathogen populations in the chicken compost mixtures occurred during the first week of storage but not after additional storage (Table 2).

of 60%, the pH increased to values approximating those detected in the chicken compost mixtures (Table 3) yet Salmonella grew in those mixtures (Table 2). Based on these results, it is likely that Salmonella is susceptible to ammonia that is present in chicken

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0.07 ± 0.41 c-e 0.10 ± 0.25 b-e 0.39 ± 0.17 bc

4.70 ± 1.71 a-c

-0.26 ± 0.48 c-e

2.25 ± 0.39 cd

1.28 ± 0.26 d

-0.57 ± 0.45 de

1.04 ± 0.81 d

0.3

0.36 ± 0.81 d

0.39 ± 0.36 bc

0.22 ± 0.24 bc

0.14 ± 0.56 b-d

-0.27 ± 0.55 c-e

-0.64 ± 0.56 e

4.29 ± 1.27 a-c

4.09 ± 1.46 a-c

2.87 ± 2.50 b-d

2.39 ± 2.90 cd

1.07 ± 1.85 d

5.83 ± 0.30 ab

4.71 ± 1.01 c

1.37 ± 0.74 e

Cloudy4

1.38 ± 0.66 a

0.86 ± 0.45 ab

0.44 ± 0.11 bc

-0.01 ± 0.46 c-e

-0.08 ± 0.62 c-e

6.27 ± 1.02 a

5.29 ± 0.70 ab

2.90 ± 2.73 b-d

2.43 ± 1.61 cd

2.03 ± 2.12 cd

5.93 ± 0.28 ab

6.03 ± 0.13 a

5.83 ± 0.30 ab

1.37 ± 0.61 e

Sunny5

0.42 ± 0.41 a-d

0.13 ± 0.46 a-f

-0.36 ± 0.69 c-f

-0.44 ± 0.53 d-f

-0.82 ± 0.37 f

1.61 ± 2.96 a

0.93 ± 1.96 a

0.71 ± 0.89 a

0.44 ± 1.75 a

-0.31 ± 1.08 a

3.14 ± 1.92 ab

1.89 ± 0.81 a-e

1.07 ± 0.56 de

1.10 ± 1.02 c-e

0.32 ± 0.65 e

Dark

0.56 ± 0.68 a-c

0.36 ± 0.62 a-d

0.02 ± 0.53 a-f

-0.43 ± 1.02 d-f

-0.75 ± 0.52 ef

1.90 ± 2.72 a

1.78 ± 2.81 a

1.63 ± 2.94 a

0.58 ± 2.40 a

0.23 ± 2.04 a

2.45 ± 0.72 a-d

2.11 ± 0.79 a-e

2.09 ± 0.38 a-e

1.39 ± 0.63 b-e

0.48 ± 0.26 e

Cloudy

Salmonella3

0.87 ± 0.38 a

0.74 ± 0.48 ab

0.16 ± 0.36 a-e

-0.11 ± 0.75 b-f

-0.25 ± 0.51 c-f

2.09 ± 2.56 a

1.67 ± 2.06 a

1.75 ± 2.84 a

1.01 ± 2.45 a

1.08 ± 3.43 a

2.99 ± 1.53 ab

2.90 ± 1.91 a-c

3.41 ± 1.14 a

3.07 ± 1.44 ab

1.01 ± 0.76 de

Sunny

Data were collected from fourth experimental study. Reductions were determined relative to initial values in compost mixtures. 3 Values (mean ± S.D.) within each manure source for this pathogen followed by a different letter are significantly different (P < 0.05). 4 Compost mixtures were exposed daily to light conditions of 289 to 359 µmol/m2/sec for 12 h and to dark conditions for 12 h. 5 Compost mixtures were exposed daily to light conditions of 524 to 573 µmol/m2/sec for 12 h and to dark conditions for 12 h.

Swine

Dairy cow

5.93 ± 0.24 ab

4.38 ± 1.08 c

8 5.01 ± 1.12 bc

2.70 ± 0.27 d

0.86 ± 0.01 e

Chicken

Dark

Week

Manure source

L. monocytogenes3

Cumulative pathogen reductions (log CFU/g) 2

Table 4. Salmonella and L. monocytogenes reductions during storage at 20°C under different light conditions in compost mixtures prepared with different manure sources1.

compost mixtures initially but the low-moisture conditions present in these mixtures inhibit the indigenous microflora from generating additional ammonia. The low moisture conditions, however, do not directly contribute to inactivation of desiccation-resistant Salmonella (Pedersen et al., 2008; Tamura et al., 2009), whereas a substantial proportion of the L. monocytogenes population is susceptible to either ammonia or desiccation stress. The last study addressed the influence of light in the visible and infrared spectrum on inactivation of pathogens in manure-based compost mixtures held at sublethal temperatures. Compost mixtures prepared with chicken litter, dairy cow manure or swine

mal reductions that occurred during the short time period that was examined for swine compost mixtures. In contrast, over all time periods, reductions in L. monocytogenes populations were statistically significant for both cloudy and sunny conditions compared to dark conditions for chicken compost mixtures stored for 6 weeks or in cow compost mixtures stored for 14 weeks (Table 4, P < 0.05). A completely different set of responses to light was observed for Salmonella in chicken- or cow compost mixtures. In chicken compost mixtures, only sunny conditions led to statistically greater reductions in populations than dark conditions, and these occurred midway through the storage trial. In

manure and held at 20Â°C were exposed to one of three lighting conditions simulating dark, sunny, or cloudy conditions. Preliminary studies were conducted with each of these compost mixtures to determine the approximate time interval over which samples should be taken to obtain measurable pathogen reductions and determine whether light conditions could significantly affect their inactivation. Unfortunately, the storage time intervals selected for swine compost mixture were underestimated and the greatest pathogen reduction was only slightly greater than 1 log CFU/g (Table 4). Despite this limitation, significant trends were determined for the swine compost mixture data. In particular, both pathogen populations increased during the first week of storage under all lighting conditions in swine compost mixtures. Following 2 weeks of storage, Salmonella remained at elevated populations for the swine compost mixtures that were held in the dark, whereas under cloudy or sunny conditions, Salmonella populations decreased slightly. Further reductions in Salmonella populations occurred during the next two weeks of storage, but there were no statistical differences in response to light exposure (P > 0.05). For L. monocytogenes in swine compost mixtures, only sunny conditions at week 4 had signifi-

contrast but similar to the response in swine compost mixtures, light conditions did not affect the reductions in Salmonella populations in cow compost mixtures at any sampling time (Table 4). A number of factors could contribute to lightmediated inactivation of L. monocytogenes in the compost mixtures. As a component of sunlight, both long wave (UVA, 315 to 400 nm) and medium wave (UVB, 280 to 315 nm) ultraviolet light has been shown to damage the genetic material of microorganisms (Davies and Evison, 1991; Jagger, 1985; Jiang et al., 2009); however, in our environmental chambers, ultraviolet light was filtered out by the diffusive ceiling light panels and hence had no role. Alternatively, exogenous sensitizers in the compost materials such as humic substances (Chien et al., 2007) may be activated by visible light energy. Such a mechanism has been demonstrated for inactivation of the Gram-positive, Enterococcus faecalis in waste stabilization pond water whereas the Gramnegative E. coli was inherently less susceptible to this pathway (Kadir and Nelson, 2014). Although L. monocytogenes has previously displayed some desiccation resistance, surviving for three months in a simulated food processing environment (Vogel et al., 2010), it is not as resistant as Salmonella based

cantly greater reductions of this pathogen compared to mixtures held under cloudy or dark conditions (P < 0.05). The inability to detect significant differences in the reduction of either pathogen under dark and cloudy conditions is likely due to the relatively mini-

on the lower recoveries of L. monocytogenes compared to Salmonella in aerosols of meat processing plants (Okraszewska-Lasica et al., 2014). Hence, a third mechanism by which increased intensities of light may have led to increased inactivation of the L.

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Table 5. % Moisture content and pH in compost mixtures held at 20°C and exposed to different light conditions across all five storage time periods examined with each type of manure1, 2.

% Moisture3 Manure source

pH3

Dark

Cloudy4

Sunny5

Dark

Cloudy

Sunny

Chicken

45.6 ± 24.4 a

33.8 ± 17.1 b

25.0 ± 16.8 b

9.5 ± 0.3 a

9.3 ± 0.3 ab

9.2 ± 0.6 b

Dairy cow

32.1 ± 19.9 a

21.4 ± 13.4 b

12.3 ± 5.7 c

8.7 ± 0.6 a

8.1 ± 0.8 b

8.3 ± 0.5 b

Swine

35.1 ± 17.2 a

29.9 ± 12.5 a

19.9 ± 13.5 b

9.0 ± 0.5 a

9.0 ± 0.4 a

Data were collected from fourth experimental study.

Swine, chicken, and dairy cow compost mixtures were stored for 4, 12, and 18 weeks, respectively.

Values in each row of an attribute followed by a different letter are significantly different (P < 0.05).

Compost mixtures were exposed daily to light conditions of 289 to 359 µmol/m2/sec for 12 h and to dark conditions for 12 h.

Compost mixtures were exposed daily to light conditions of 524 to 573 µmol/m2/sec for 12 h and to dark conditions for 12 h.

monocytogenes isolates used in this study may be through localized heating, additional dehydration at surface locations, and in turn increased desiccation stress. In support of this explanation, the moisture content in the compost mixtures decreased as the mixtures were exposed to higher levels of light and the highest levels of dehydration occurred in the dairy compost mixtures followed by the swine compost mixtures (Table 5). In addition to affecting the moisture content of the compost mixtures, light exposure led to decreased pH in the chicken and cow compost mixtures (Table 5). L. monocytogenes survival could have been improved under less alkaline conditions; however, that response was likely to be minimal in these compost mixtures due to the concurrent stress imposed by low moisture contents and

a state could also have been responsible for our inability to discern an effect of light on inactivation of Salmonella in the swine or cow compost mixtures. In summary, both Salmonella and L. monocytogenes may survive in compost mixtures that are exposed to sublethal temperatures for extended periods of time. As ambient temperatures increased, the persistence of pathogens decreased which may be attributed to increased competitive activity by the more dominant indigenous microflora. Attempts to maintain the moisture content of compost mixtures on a weekly basis was challenging because rapid evaporation resulted in very dry mixtures in most cases. Under these conditions, L. monocytogenes appeared to be more susceptible to desiccation stress than Salmonella based on their relative re-

light exposure. In the case of Salmonella, however, it is known that it is extremely resistant to desiccation (Pedersen et al., 2008; Tamura et al., 2009). Given that dehydration has induced cross-tolerance to a number of other stressors (Gruzdev et al., 2011), such

duction in populations in chicken and cow compost mixtures over time. L. monocytogenes populations also decreased more rapidly when compost mixtures were exposed to light conditions, described as sunny or cloudy, compared to dark conditions, but

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this response did not occur with Salmonella in cow or swine compost mixtures. It is suggested that the drier conditions encountered in light-exposed cow and swine compost mixtures may have induced a cross-tolerance response by Salmonella to the light stress. If cross-tolerance responses by Salmonella are generated in low moisture compost mixtures held at sublethal temperatures, it will therefore be important to apply an intervention treatment to those compost mixtures prior to the activation of that response.

ACKNOWLEDGEMENTS

Erickson, M.C., J. Liao, G. Boyhan, C. Smith, L. Ma, X. Jiang, and M.P. Doyle. 2010. Fate of manure-borne pathogen surrogates in static composting piles of chicken litter and peanut hulls. Bioresource Technol. 101:1014-1020. Gotaas, H.B. 1956. Composting. Sanitary disposal and reclamation of organic wastes. World Health Organization, Geneva, Switzerland. Gruzdev, N., R. Pinto, and S. Sela. 2011. Effect of desiccation on tolerance of Salmonella enterica to multiple stresses. Appl. Environ. Microbiol. 77:1667-1673. Himathongkham, S. and H. Riemann. 1999. Destruction of Salmonella typhimurium, Escherichia coli

Beck-Friis, B., S. Smårs, H. Jönsson, Y. Eklind, and H. Kirchmann. 2003. Composting of source-separated household organics at different oxygen levels: Gaining an understanding of the emission dynamics. Compost Sci. Util. 11:41-50. Berry, E.D., P.D. Millner, J.E. Wells, N. Kalchayanand, and M.N. Guerini. 2013. Fate of naturally occurring Escherichia coli O157:H7 and other zoonotic pathogens during minimally managed bovine feedlot manure composting processes. J. Food Prot. 76:1308-1321. Chang Chien, S.W., M.C. Wang, C.C. Huang, and K. Seshaiah. 2007. Characterization of humic substances derived from swine manure-based compost and correlation of their characteristics with re-

O157:H7 and Listeria monocytogenes in chicken manure by drying and/or gassing with ammonia. FEMS Microbiol. Lett. 171:179-182. Himathongkham, S., S. Bahari, H. Riemann, and D. Cliver. 1999a. Survival of Escherichia coli O157:H7 and Salmonella typhimurium in cow manure and cow manure slurry. FEMS Microbiol. Lett. 178:251-257. Himathongkham, S., S. Nuanualsuwan, and H. Riemann. 1999b. Survival of Salmonella enteritidis and Salmonella typhimurium in chicken manure at different levels of water activity. FEMS Microbiol. Lett. 172:159-163. Himathongkham, S., H. Riemann, S. Bahari, S. Nuanualsuwan, P. Kass, and D.O. Cliver. 2000. Survival of Salmonella typhimurium and Escherichia coli O157:H7 in poultry manure and manure slurry at sublethal temperatures. Avian Dis. 44:853-860. Hutchison, M.L., F.A. Nicholson, K.A. Smith, C.W. Keevil, B.J. Chambers, and A. Moore. 2000. A study on farm manure applications to agricultural land and an assessment of the risks of pathogen transfer into the food chain. Report to The Ministry of Agriculture Fisheries and Food. Available at: http://www.safeproduce.eu/Pics/FS2526.pdf. Accessed 21 April, 2014. Hutchison, M.L., L.D. Walters, S.M. Avery, and A.

activities with heavy metals. J. Agric. Food Chem. 55:4820-4827. Davies, C. M. and L. M. Evison. 1991. Sunlight and the survival of enteric bacteria in natural waters. J. Appl. Bacteriol. 70:265-274.

Moore. 2005. Decline of zoonotic agents in livestock waste and bedding heaps. J. Appl. Microbiol. 99:354-362. Jagger, J. 1985. Solar-UV action on living cells. Praeger, New York.

The project was supported by the National Research Initiative of the USDA Cooperative State Research, Education, and Extension Service, grant # 2008-35201-18658. We gratefully thank Derrick Huckaby, Lindsey Davey, and Jessica Colvin for technical assistance.

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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

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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

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rial board members and should be sent to submit@ afabjournal.com. There will be no page charges for solicited review papers but the solicitation must originate from the editor-in-chief or one of the editors. Requests from authors will automatically be classified as unsolicited review papers. The running head above the title of the paper will be “Invited Review.”

Conference and Special Issues Reviews AFAB welcomes opportunities to publish papers from symposia, scientific conference, and/or meetings in their entirety. Conference organizers need simply to contact AFAB at submit@afabjournal.com and a rapid decision is guaranteed. If in agreement, the conference organizers must guarantee delivery of a set number of peer reviewed manuscripts within a specified time and submitted in the same format as that described for unsolicited review papers. Conference papers must be prepared in accordance with the guidelines for review articles and are subject to peer review. The conference chair must decide whether or not they wish to serve as Special Issue Editor and conduct the editorial review process. If the conference chair/organizer chooses to serve as special issue editor, this will involve review of the papers and, if necessary, returning them to the authors for revision. The conference organizer then submits the revised manuscripts to the journal editorial office for further editorial examination. Final revisions by the author and recommendations for acceptance or rejection by the chair must be completed by a mutually agreed upon date between the editor and the conference organizer. Manuscripts not meeting this deadline will not be included in the published symposium proceedings but if submitted later can still be considered as unsolicited review papers. Although offprints and costs of pages are the same as for all other papers, the symposium chair may be asked to guarantee an agreed upon number of hard copies to be purchased by conference attendees. If the decision is not to publish the symposium as a special issue, the individual authors retain the right to submit their papers for consideration for the journal as ordinary unsolicited review manuscripts.

Book Reviews AFAB publishes reviews of books considered to be of interest to the readers. The editor-in-chief ordinarily solicits reviews. Book reviews shall be prepared in accordance to the style and form requirements of the journal, and they are subject to editorial revision. No page charges will be assessed solicited reviews while unsolicited book reviews will be assigned the regular page charge rate.

Opinions and Current Viewpoints The purpose of this section will be to discuss, critique, or expand on scientific points made in articles recently published in AFAB. Drafts must be received within 6 months of an article’s publication. Opinions and current perspectives do not have page limits. They shall have a title followed by the body of the text and references. Author name(s) and affiliation(s) shall be placed between the end of the text and list of references. If this document pertains to a particular manuscript then the author(s) of the original paper(s) will be provided a copy of the letter and offered the opportunity to submit for consideration a reply within 30 days. Responses will have the same page restrictions and format as the original opinion and current viewpoint, and the titles shall end with “Opinions.” They will be published together. Letters and replies shall follow appropriate AFAB format and may be edited by the editor-in-chief and a technical editor. If multiple letters on the same topic are received, a representative set of opinions concerning a specific article will be published. A disclaimer will be added by the editorial staff that the opinion expressed in this viewpoint is the authors alone and does not necessarily represent the opinion of AFAB or the editorial board.

COPYRIGHT AGREEMENT The copyright form is published in AFAB as space permits and is available online (www.afabjournal.com).

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AFAB grants to the author the right of re-publication in any book of which he or she is the author or editor, subject only to giving proper credit to the original journal publication of the article by AFAB. AFAB retains the copyright to all materials accepted for publication in the journal. If an author desires to reprint a table or figure published from a non-AFAB source, written evidence of copyright permission from an authority representing that source must be obtained by the author and forwarded to the AFAB editorial office.

PEER REVIEW PROCESS Authors will be requested to provide the names and complete addresses including emails of five (5) potential reviewers who have expertise in the research area and no conflict of interest with any of the authors. Except for manuscripts designated as Rapid Communication each reviewer has two (2) weeks to review the manuscript, and submit comments electronically to the editorial office. Authors have three (3) weeks to complete the revision, which shall be returned to the editorial office within six (6) weeks after which the authors risk having their manuscript removed from AFAB files if they fail to ask the editorial office for an extension by email. Deleted manuscripts will be reconsidered, but they will have to be processed as new manuscripts with an additional processing fee assessed upon submission. Once reviewed, the author will be notified of the outcome and advised accordingly. Editors handle all initial correspondence with authors during the review process. The editor-in chief will notify the author of the final decision to accept or reject. Rejected manuscripts can be resubmitted only with an invitation from the editor or editor-in chief. Revised versions of previously rejected manuscripts are treated as new submissions.

PRODUCTION OF PROOFS Accepted manuscripts are forwarded to the editorial office for technical editing and layout. The manuscript is then formatted, figures are reproduced, and author proofs are prepared as PDFs. Author proofs of all manuscripts will be provided to the correspond246

ing author. Author proofs should be read carefully and checked against the typed manuscript, because the responsibility for proofreading is with the author(s). Corrections must be returned by e-mail. Changes sent by e-mail to the technical editor must indicate page, column, and line numbers for each correction to be made on the proof. Corrections can also be marked using “track changes” in Microsoft Word or using e-annotation tools for electronic proof correction in Adobe Acrobat to indicate necessary changes. Author alterations to proofs exceeding 5% of the original proof content will be charged to the author. All correspondence of proofs must be agreed to by the editorial office and the author within 48 hours or proof will be published as is and AFAB will assume no responsibility for errors that result in the final publication.

PUBLICATION CHARGES AFAB has two publication charge options: conventional page charges and rapid communication. The current charge for conventional publication is $25 per printed page in the journal. There is no additional charge for the publication of pages containing color images, micrographs or pictures. For authors who wish to have their papers processed as a rapid communication, authors will pay the rapid communication fee when proofs are returned to the editorial office in addition to twice the conventional page charges. Charges for rapid communications are $1000 per manuscript for guaranteed peer review within one week and $100 per journal page.

HARD COPY OFFPRINTS If you are wishing to obtain a physical hard copy of the AFAB journal, offprints are available in any quantity at an additional charge: $100/page for black-white and $150/page for color prints. You may order your offprints at any time after publication on our website. Scientific conference organizers may be expected to agree to a set number of offprints as a part of their agreement with AFAB.

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MANUSCRIPT CONTENT REQUIREMENTS Preparing the Manuscript File Manuscripts must be written in grammatically correct English. AFAB offers a fee based language service upon request (language@afabjournal.com). Manuscripts should be typed double-spaced, with lines and pages numbered consecutively. All documents must be submitted in Microsoft Word (.doc or .docx, PC or Mac). All special characters (e.g., Greek, math, symbols) should be inserted using the symbols palette available in this font. Tables and figures should be placed in separate sections at the end of the manuscript (not placed in the text). Failure to follow these instructions will cause delays of the processing and review of the manuscript.

Title Page At the very top of the title page, include a title of not more than 100 characters. Format the title with the first letter of each word capitalized. No abbreviations should be used. Under the title, the authors names are listed. Use the author’s initials for both first and middle names with a period (full-stop) between initials (e.g., W. A. Afab). Underneath the authors, a list affiliations must be listed. Please use numerical superscripts after the author’s names to designate affiliation. If an authors address has changed since the research was completed, this new information must be designated as “Current address:”. The corresponding author should be indicated with an asterisk e.g., * Corresponding author. The title page shall include the name and full address of the corresponding author. Telephone and e-mail address must also be provided for the corresponding author, and emailaddresses must be provided for all authors.

at the beginning of the manuscript. In vivo, in vitro and bacterial names must be italicized (obligatory). Authors must avoid single sentence paragraphs and merge such paragraphs appropriately. Authors must not begin sentences with “Figure or Table shows…” as these are inanimate objects and cannot “show” anything. When number are reported in text or in tables, always put a zero in front of decimal numbers: “0.10” instead of “.10”.

MANUSCRIPT SECTIONS Abstract The abstract provides an abridged version of the manuscript. Please submit your abstract on a separate page after the title page. The abstract should provide a justification of your work, objectives, methods, results, discussion and implications of study or review findings . Your abstract must consist of complete sentences without references to other work or footnotes and must not exceed 250 words. On the same page as your abstract, please provide at least ten (10) keywords to be used for linking and indexing. Ideally, these keywords should include significant words from the title.

Introduction The introduction should clearly present the foundation of the manuscript topic and what makes the research or the review unique. The introduction should validate why this topic is important based on previously published literature, and the relevance of the current research. Overall goals and project objectives must be clearly stated in the final sentence of the last paragraphs of the introduction.

Materials and Methods Editing Author-derived abbreviations should be defined at first use in the abstract and again in the body of the manuscript. If abbreviations are extensive authors may need to provide a list of abbreviations

Information on equipment and chemicals used must include the full company name, city, and state (country if outside the United States or Province if in Canada) [i.e., (Model 123, ACME Inc., Afab, AR)].

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Variability, Replication, and Statistical Analysis To properly assess biological systems independent replication of experiments and quantification of variation among replicates is required by AFAB. Reviewers and/or editors may request additional statistical analysis depending on the nature of the data and it will be the responsibility of the authors to respond appropriately. Statistical methods commonly used in the bacteriology do not need to be described in detail, but an adequate description and/or appropriate references should be provided. The statistical model and experimental unit must be designated when appropriate. The experimental unit is the smallest unit to which an individual treatment is imposed. For bacterial growth studies, the average of replicate tubes per single study per treatment is the experimental unit; therefore, individual studies must be replicated. Repeated time analyses of the same sample usually do not constitute independent experimental units. Measurements on the same experimental unit over time are also not independent and must not be considered as independent experimental units. For analysis of time effects, assess as a rate of change over time. Standard deviation refers to the variability in the biological response being measured and is presented as standard deviation or standard error according to the definitions described in statistical references or textbooks.

Results Results represent the presentation of data in words and all data should be described in same fashion. No discussion of literature is included in the results section.

Discussion The discussion section involves comparing the current data outcomes with previously published work in this area without repeating the text in the results section. Critical and in-depth dialogue is encouraged.

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Results and Discussion Results and discussion can be under combined or separate headings.

Conclusions State conclusions (not a summary) briefly in one paragraph.

Acknowledgments Acknowledgments of individuals should include institution, city, and state; city and country if not U.S.; and City or Province if in Canada. Copies being reviewed shall have authors’ institutions omitted to retain anonymity.

References a) Citing References In Text Authors of cited papers in the text are to be presented as follows: Adams and Harry (1992) or Smith and Jones (1990, 1992). If more than two authors of one article, the first author’s name is followed by the abbreviation et al. in italics. If the sentence structure requires that the authors’ names be included in parentheses, the proper format is (Adams and Harry, 1982; Harry, 1988a,b; Harry et al., 1993). Citations to a group of references should be listed first alphabetically then chronologically. Work that has not been submitted or accepted for publication shall be listed in the text as: “G.C. Jay (institution, city, and state, personal communication).” The author’s own unpublished work should be listed in the text as “(J. Adams, unpublished data).” Personal communications and unsubmitted unpublished data must not be included in the References section. Two or more publications by the same authors in the same year must be made distinct with lowercase letters after the year (2010a,b). Likewise when multiple author citations designated by et al. in the text have the same first author, then even if the other authors are different these references in the text and the references section must be identified by a letter. For example

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

Book Chapter: Examples:

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.

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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Online Publication: www.AFABjournal.com