Afab volume 4 issue 3

Volume 4, Issue 32014

ISSN: 2159-8967www.AFABjournal.com

158 Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 4, Issue 3 - 2014

Agric. Food Anal. Bacteriol. • AFABjournal.com • Vol. 4, Issue 3 - 2014 159

Sooyoun Ahn University of Florida, USA

Walid Q. Alali University of Georgia, USA

Kenneth M. Bischoff NCAUR, USDA-ARS, USA

Debabrata Biswas University of Maryland, USA

Claudia S. Dunkley University of Georgia, USA

Michael Flythe USDA, Agricultural Research Service

Lawrence Goodridge McGill University, Canada

Leluo Guan University of Alberta, Canada

Joshua Gurtler ERRC, USDA-ARS, USA

Yong D. Hang Cornell University, USA

Armitra Jackson-Davis Alabama A&M University, USA

Divya Jaroni Oklahoma State University, USA

Weihong Jiang Shanghai Institute for Biol. Sciences, P.R. China

Michael Johnson University of Arkansas, USA

Timothy Kelly East Carolina University, USA

William R. Kenealy Mascoma Corporation, USA

Hae-Yeong Kim Kyung Hee University, South Korea

Woo-Kyun Kim University of Georgia, USA

M.B. Kirkham Kansas State University, USA

Todd Kostman University of Wisconsin, Oshkosh, USA

Y. M. Kwon University of Arkansas, USA

Maria Luz Sanz MuriasInstituto de Quimica Organic General, Spain

Byeng R. Min Tuskegee University in Tuskegee, AL

Melanie R. Mormile Missouri University of Science and Tech., USA

Rama Nannapaneni Mississippi State University, USA

Jack A. Neal, Jr. University of Houston, USA

Benedict Okeke Auburn University at Montgomery, USA

John Patterson Purdue University, USA

Toni Poole FFSRU, USDA-ARS, USA

Marcos Rostagno LBRU, USDA-ARS, USA

Roni Shapira Hebrew University of Jerusalem, Israel

Kalidas Shetty North Dakota State University, USA

EDITORIAL BOARD


EDITOR-IN-CHIEFSteven C. RickeUniversity of Arkansas, USA

EDITORSTodd R. CallawayFFSRU, USADA-ARS, USA

Philip G. CrandallUniversity of Arkansas, USA

Janet Donaldson Mississippi State University, USA

Ok-Kyung KooKorea Food Research Institute, South Korea

MANAGING and LAYOUT EDITOREllen J. Van LooGhent, Belgium

TECHNICAL EDITORJessica C. ShabaturaFayetteville, USA

ONLINE EDITION EDITORC.S. ShabaturaFayetteville, USA

ABOUT THIS PUBLICATION

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 pend-

ing manuscripts should be addressed Ellen Van Loo,

Managing Editor, Agriculture, Food & Analytical

Bacteriology: [email protected]

Information for Potential Editors: If you are interested

in becoming a part of our editorial board, please con-

tact Editor-in-Chief, Steven Ricke, Agriculture, Food &

Analytical Bacteriology: [email protected]

Advertising: If you are interested in advertising with

our journal, please contact us at advertising@afab-

journal.com for a media kit and current rates.

Reprint Permission: Correspondence regarding re-

prints should be addressed Ellen Van Loo, Managing

Editor, Agriculture, Food & Analytical Bacteriology

[email protected]

Ordering Print Copies: print editions of this journal

may be purchased and shipped internationally from

our website order form at www.afabjournal.com

Subscription Rates: Subscriptions are not available

at this time. To be advised when subscriptions plans

are made available, please join our newsletter at

www.afabjournal.com

Mailing Address: 2138 Revere Place . Fayetteville, AR . 72701 Website: www.AFABjournal.com

EDITORIAL STAFF


Batch Culture Characterization of Acetogenesis in Ruminal Contents: Influence of Aceto-

gen Inocula Concentration and Addition of 2-Bromoethanesulfonic AcidP. Boccazzi and J. A. Patterson

177

Survival of Salmonella enterica and Listeria monocytogenes in manure-based compost mix-tures at sublethal temperatures M.C. Erickson, C. Smith, X. Jiang, I.D. Flitcroft, and M.P. Doyle

224

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

195

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

212

ARTICLES

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

164

Instructions for Authors243

Introduction to Authors

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

TABLE OF CONTENTS


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 microbiolo-gist 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 re-search 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 jour-nal articles, numerous technical/report papers, proceedings papers, and abstracts, and 2 book chapters.

NEW EDITORIAL STAFF



www.afabjournal.comCopyright © 2014

Agriculture, Food and Analytical Bacteriology

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, let-

ting 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

Correspondence: Md. Latiful Bari, [email protected]: 8801971560560 Fax: 8802-8615583

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

1Center for advanced Research in Sciences, University of Dhaka, Dhaka-1000, Bangladesh2Institute of Food and Radiation Biology, Bangladesh Atomic Energy Commission, Savar, Dhaka, Bangladesh

3Bangladesh Agricultural Research Institute, Gazipur 1701, Bangladesh4Kazi & Kazi Tea Estate Ltd. University of Liberal Arts Bangladesh (ULAB), Dhanmondi, Dhaka-1209, Bangladesh

Agric. Food Anal. Bacteriol. 4: 164-176, 2014


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 tropi-

cal 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.

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 anal-

gesic, 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 Chi-

nese 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 anti-

inflammatory (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 re-

pellents (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 labora-

tory studies particularly against Candida species (Can-

dida albicans, Candida glabrata, Candida krusei, Can-

dida parapsilosis, and Candida tropicalis) (Warnke et

al., 2009). In a preliminary study, lemongrass infusion

had beneficial effects for the treatment of oral candi-

diasis in patients with HIV/AIDS (Wright et al., 2009).

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 micro-

flora on these vegetables at harvest reflects the envi-

ronment in which they are grown, if the temperature

and humidity is relatively high then the occurrence of

enteropathogenic bacteria in this environment might

be considerable. During cultivation, use of contami-

nated water for irrigation, application of biocides,

and refreshing or washing of harvested crops, are po-

tential sources of contamination. Contamination from

contact with fresh manure used as fertilizer cannot be

excluded. Heavy rainfall may also lead to fecal con-

tamination 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; Adel-

eye 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

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 veg-

etables products, from South East Asia, and found


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 prod-

ucts from South East Asia, and now the EU accord-

ingly requires certificate of analysis for Salmonella

and E. coli before export (Olaimat and Holley, 2012).

The objectives of this present study were to evalu-

ate microbial contamination of organic herb pro-

duction at the farm level, and a case study of food

safety management in organic lemongrass produc-

tion 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-

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.

MATERIALS AND METHODS

Sample collection

Herb samples include, lemongrass, mint, neem

and jasmine were obtained from an organic farm

in Northern Bangladesh between May 15 and July

30, 2013. All samples were transported to the Food

Analysis and Research Laboratory, Center for Ad-

vanced Research in Sciences (CARS) at the Univer-

sity of Dhaka using a cool box at the earliest con-

venience 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.

Selected critical sampling locations (CSLs)

The critical sampling locations were selected

based on the production scheme presented in Fig-

ure 1a and other sources of microbiological contam-

ination 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 (Fig-

ure 1b) including the lemongrass crop.

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


Total aerobic count and total coliform count

Twenty five (25) g of each sample were homog-

enized in 225 mL of saline water (0.85% NaCl). Deci-

mal dilutions were prepared upto 10-6 mL and ap-

propriate dilutions were spread plated on Tryptic

soy agar (Oxoid Ltd., Hampshire, England) and in-

cubated at 35˚C for 24 hr for total aerobic bacterial

counts and on MacConkey agar (Oxoid Ltd., Hamp-

shire, 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


enized in 225 mL Enterobacteria enrichment broth-

Mossel pre-enrichment medium (Oxoid Ltd., Hamp-

shire, 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 con-

firm 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 indi-

cates 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 colo-

nies were then confirmed using biochemical charac-

terization (IMViC) and API 20E kit (bioMérieux, Dur-

ham, 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 homog-

enized 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 tellu-

rite 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, Ja-

pan). The colonies which gave the characteristic col-

or were serotyped using O157, O111 and O26 spe-

cific 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;


chromato VT1/2 and by polymerase chain reaction

(PCR) assay using primer 5’-CAGTTAATGTGGTG-

GCGAAGG-3’ and 5’-CACCAGACAAATGTAACC-

GCTC-3’ for stx1 and 5’-ATCCTATTCCCGGGAGTT-

TACG-3’ and 5’-GCGTCATCGTATACACAGGAGC-3’

for stx2, respectively (Vidal et al., 2004).

Salmonella spp.


enized 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 Salmo-

nella LA latex agglutination test and API 20E kits.

Hot water treatment in tea-cup & En-richment 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., Hamp-

shire, 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 non-

selective TSB medium was used to help resuscitate

these cells and enable them to grow in selective mi-

crobiological medium.

Statistical Analysis

Three samples of each category were taken from

the same farm. Reported plate count data represent-

ed 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 Mi-

crosoft 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 or-

ganic 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 investiga-

tions have raised concerns related to the microbio-

logical 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 mar-

ket as well as acceptability regardless of age or eco-

nomic 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), lemon-

grass and lemongrass blend tea were analyzed for

total aerobic population (TAB), total coliform popu-

lation (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, co-

liform 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


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

Herbs and tea Sample

Total Viable 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.pH

Black tea (Normal)

5.9 ± 0.08 6.9 ± 0.11 4.7± 0.14 3.8 ± 0.06 ND 4.84

Blend tea 6.4 ± 0.11 5.9± 0.11 4.5 ±0.11 4.5 ± 0.12 ND 5.01

Black tea (Original) 6.0± 0.14 6.0± 0.34 5.1±0.11 4.6± 0.11 ND 5.40

Neem blend herbs

4.1 ± 0.15 ND* ND ND ND 4.98

Neem Tea 3.9 ± 0.12 ND ND ND ND 5.04

Black tea (Premium) 5.4 ± 0.14 4.1 ± 0.21 4.0 ± 0.12 3.2 ± 0.11 ND 5.10

liker base 6.9 ± 0.22 6.2 ± 0.22 5.7 ± 0.13 5.2 ± 0.15 ND 7.13

Lemongrass 5.9 ± 0.11 5.8 ± 0.15 5.8 ± 0.23 4.8 ± 0.13 ND 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 ND 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 ND 7.50

Ground water 3.8 ± 0.13 3.4 ± 0.07 3.0 ± 0.11 2.1 ± 0.12 ND 6.50

Manure soil 6.0 ± 0.14 5.8 ± 0.09 5.0 ± 0.11 4.7 ± 0.11 ND 7.80

*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 de-

termined 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 deter-

mined 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.

The water and composted manure was found to be

heavily contaminated with enteric bacterial patho-

gens (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.

Salmonella spp. was not detected in the manure

sample tested (Table 1). In this study, water for irriga-

tion 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 sever-

al 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

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.

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 pro-duction & process-

ing steps

Total aerobic counts

(log CFU/g)

Total Coli-form counts (log CFU/g)

Total E. coli counts

(log CFU/g)

E. coli O157:H7 counts

(log CFU/g)

Presence of Salmonella

Spp.pH

At Harvest 5.9 ± 0.08 5.8± 0.18 5.8± 0.18 4.8 ± 0.16 ND 4.6

Cleaning & No washing

5.7 ± 0.12 5.4 ± 0.08 4.7 ± 0.08 3.7 ± 0.10 ND 5.2

Cleaning & fresh water wash

5.0 ± 0.11 4.3± 0.16 4.3 ± 0.10 3.6± 0.20 ND 5.1

Cleaning with fresh hot water

5.2 ± 0.14 4.5 ± 0.08 3.8± 0.19 3.5 ± 0.17 ND 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 ND 5.2

After grinding at room temperature

4.2 ± 0.13 4.0 ± 0.09 3.7 ± 0.18 3.7 ± 0.16 ND 5.4

After sorting at room temperature

5.6 ± 0.11 3.5± 0.20 3.3 ± 0.11 3.1 ± 0.11 ND 5.3

After final streaming 4.4 ± 0.10 4.3 ± 0.08 2.3 ± 0.10 2.3 ± 0.12 ND 5.4

*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). Nu-

merous 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 de-

tected in fresh compost (James, 2006; MAFF, 2000;

Millner, 2003; Johannessen, 2005). However, the or-

ganic fertilizer samples analyzed in the present study

were above the detection limit (3.0 log MPN/g), indi-

cating that the control of manure was not adequate.

From the same farm, lemongrass production prac-

tices were taken as a case study to determine the point

of E. coli O157:H7 contamination and to take correc-

tive measure in eliminating the risk. The average aero-

bic bacterial counts, coliform counts, E. coli and E. coli

O157: H7 counts were recorded as 5.9 log CFU /g., 5.8

log CFU /g, 5.8 log CFU/g and 4.8 log CFU /g, respec-

tively after harvest. However, Salmonella spp. was not

detected in the lemongrass sample (Table 2). After har-

vest, 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 re-

duced the microbial load by 0.5-1.0 log CFU/g of bac-

teria. 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 com-

pleted, 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-

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 find-

ing suggested that heating temperature or the contact

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.

Lemongrass pro-duction & pro-cessing steps

Total aero-bic count

(log CFU/g)

Total Coli-form count (log CFU/g)

Total E. coli Count

(log CFU/g)

E.coli O157:H7 counts

(log CFU/g)

Presence of Salmo-nella Spp.

pH

Positive / No of sample tested

Control 5.9 ± 0.08 5.8 ± 0.18 5.8 ± 0.18 4.8 ± 0.16 ND 4.6

Lemongrass (After corrective measures 1)

2.7 ± 0.18 ND* ND ND ND 5.7 0/3

Lemongrass (After corrective measures 2)

2.5 ± 0.12 ND ND ND ND 5.8 0/3

Mint (After corrective measures 1)

2.7 ± 0.14 ND ND ND ND 6.0 0/3

Mint (After corrective measures 2)

2.9 ± 0.11 1.3 ± 0.09 1.0 ± 0.11 ND ND 5.6 1/3

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


time may not be adequate to inactivate pathogens in

lemongrass samples. After that an investigation of ac-

tual 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 con-

trolled velocity conditions to create a fluidized state.

Therefore, when the sample comes in contact with

heat for few seconds, some bacteria may become in-

jured and could resuscitate in between the cycles and

in following steps, therefore, survive and subsequently

be detected in the final products. Therefore, corrective

actions of dryer temperature were undertaken and af-

ter 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 ve-

hicles 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 detect-

ed 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 hy-

giene 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 ap-

proach is not appropriate for the majority of fresh herbs

and leafy greens that are mainly consumed raw, or

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-

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 inges-

tion is a critical consideration. To solve this approach,

an experiment was designed to determine the effec-

tiveness 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 differ-

ent 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 wa-

ter 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-

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 tempera-

ture 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 with-

out 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 sec-

onds of hot water treatment despite the higher patho-

gen 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 mi-

crobiological medium. To solve this issue, an enrich-

ment 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)

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.


Tab

le 4

. Eff

ecti

vene

ss o

f p

our

ing

ho

t w

ater

ove

r co

ntam

inat

ed d

ried

(ble

nd t

ea, b

lack

tea

and

lem

ong

rass

sam

ple

s)

leav

es in

a c

up o

f te

a at

diff

eren

t ho

ldin

g t

ime.

Sam

ple

typ

eH

ot

wat

er

trea

tmen

t ti

me

(Sec

)

Rec

ove

ry o

f m

icro

org

anis

ms

(log

CFU

/g)a

Aft

er E

nric

hmen

t

Tota

l aer

o-

bic

co

unt

Tota

l Co

li-fo

rm c

oun

t E

. co

liE

. co

li O

157:

H7

Salm

one

lla

Spp

.P

rese

nce

of

Co

li-fo

rm

Pre

senc

e o

f E

.co

liP

rese

nce

of

E. c

oli

O15

7:H

7

Pre

senc

e o

f Sa

lmo

nella

Sp

p.

Ble

nded

Tea

Sa

mp

les

Co

ntro

l5.

4 ±

0.0

84.

1 ±

0.1

04.

0 ±

0.1

23.

2 ±

0.1

1N

D-

--

-

303.

5 ±

0.1

8<

1.0*

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

603.

2 ±

0.1

2<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

903.

0 ±

0.1

1<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

120

3.0

± 0

.14

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

150

2.9

± 0

.09

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

180

3.2

± 0

.09

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

5 m

in3.

1± 0

.08

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

Bla

ck T

ea

Sam

ple

sC

ont

rol

5.9

± 0

.11

6.9

± 0

.12

4.7

± 0

.11

3.8

± 0

.16

ND

--

--

301.

3 ±

0.1

4<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

601.

3 ±

0.1

3<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

901.

0 ±

0.0

9<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

120

1.0

± 0

.07

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

150

-<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

180

-<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

5 m

in-

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

lem

ong

rass

Sa

mp

les

Co

ntro

l6.

0 ±

0.1

56.

0 ±

0.1

25.

1 ±

0.1

14.

6 ±

0.1

4N

D-

--

-

303.

1 ±

0.1

6<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

603.

0 ±

0.1

2<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

903.

0 ±

0.1

1<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

120

3.0

± 0

.07

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

150

-<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

180

-<

1.0

<1.

0<

1.0

ND

Nil

Nil

Nil

Nil

5 m

in-

<1.

0<

1.0

<1.

0N

DN

ilN

ilN

ilN

il

<1.

0*=

Les

s th

an d

etec

tion

limit;

ND

= N

ot

Det

ecte

d, N

il= A

bse

nt; a R

esul

ts a

re e

xpre

ssed

in m

ean±

sta

ndar

d d

evia

tion

of t

hree

rep

licat

e sa

mp

les,

whi

ch

are

bei

ng c

alcu

late

d fr

om

dup

licat

e p

late

s.


CONCLUSIONS

In the present study it can be concluded that

the organic fertilizer and the water used for irriga-

tion and washing are critical sources of microbial

contamination that need to be controlled in the

production chain of organic produce. The contam-

ination 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 contami-

nated; nonetheless, they could very well be con-

taminated 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 organ-

ic herbs. This case study provides an overview of

the organic farms’ status in northern Bangladesh,

where good hygiene practice and GAP were intro-

duced as a part of this study.

ACKNOWLEDGEMENTS

The authors would like to thank Mr. Harun-ur

Rashid and Mr. Abul Kalam Azad for the labora-

tory assistance required to complete this task. The

authors would also like to thank the United Nations

University, Tokyo, Japan (UNU-ISP) for financial sup-

port (FY 2013-2014) in this work.

REFERENCES

Abadias, M., J. Usall, M. Anguera, C. Solsona, and

I. Viñas. 2008. Microbiological quality of fresh,

minimally-processed fruit and vegetables, and

sprouts from retail establishments. Int. J. Food

Microbiol. 123:121–129.

Abba, D., H. I. Inabo, S. E. Yakubu, and O. S. Oloni-

tola. 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. Mi-

crobial 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. 6:953-

958.

Beuchat, L. R. 1996. Pathogenic microorganisms

associated with fresh produce. J. Food Prot.

59:204–216.

Beraldo, R. M., and F. A. Filho. 2011. Bacterio-

logical quality of irrigation water from vegetable

gardens in the municipalities of Araraquara, Boa

Esperança do Sul e Ibitinga, SP. Alimentos e Nu-

trição Araraquara 22:1640.

Blumenthal, M. 1998. The Complete German

Commission E Monographs . Austin TX: Ameri-

can Botanical Council. 341-342.

Delaquis, P., S. Bach, and L. D. Dinu. 2007. Be-

havior of Escherichia coli O157:H7 in leafy veg-

etables. J. Food Prot. 70:1966–1974.

Esimone, C. O., P. O. Oleghe, and E. C. Ibezim.

2003. Effect of preservation agents on the micro-

bial stability of some indigenous herbal prepara-

tions. Niger J. Pharm. 34:37-42.

Espen Rimstad, E. A. Høiby, G. Kapperud, J. Las-

sen, B. Tore Lunestad, et al. 2008. Norwegian

Scientific Committee for Food Safety Panel on

Biological Hazards final report on Risk assess-

ment of import and dissemination of intestinal

pathogenic bacteria via fresh herbs and leafy

vegetables from South-East Asia. ISBN: 978-82-

8082-244-4 ; 07/111-final report, pages 2-32.

FAO/WHO. 2008. Microbiological hazards in fresh

fruits and vegetables. Meeting Report. Microbi-

ological risk assessment series. Available at the


following URL and accessed on August 7, 2013.

(http://www.fao.org/ag/agn/agns/jemra_riskas-

sessment_freshproduce_en.asp)

Fischer-Arndt, M., D. Neuhoff, L. Tamm, and U.

Köpke. 2010. Effects of weed management

practices on enteric pathogen transfer into let-

tuce (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. Ethno-

pharmacol. 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 inter-

vention research, published between 1990 and

2010, for microbial hazards in leafy green veg-

etables. Food Control 23:7-19.

Itohan, A. M., O. Peters, and I. Kolo. 2011. Bacte-

rial contaminants of salad vegetables in Abuja

Municipal Area Concil. Nigéria. Malaysian J. Mi-

crobiol. 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 produc-

tion 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 chromatogra-

phy- mass spectroscopy. J. Pharm. Bio Ana. 31:

401-406.

Leung, A. Y. 1980. Encyclopedia of Common Natu-

ral Ingredients Used in Food, Drugs, and Cos-

metics. New York, NY: Wiley.

Lewinsohn, E., N. Dudai, Y. Tadmor, I. Katzir, U.

Ravid, E. Putievsky, and D. M. Joel. 1998. His-

tochemical 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 Fish-

eries). 2000. A study of on-farm manure appli-

cations 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 time-

tested 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. Ruiz-

de-Castañeda. 2009. Testing the use of a citro-

nella-based repellent as an effective method to

reduce the prevalence and abundance of bit-

ing flies in avian nests. Parasitol Res. 104:1233-

1236.

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 Alter-

native Medicines. Vol. 5:74- 78.

Olaimat, A. N., and R. A. Holley. 2012. Factors in-

fluencing 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 dif-

ferent 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.


Vidal, R., M. Vidal, R. Lagos, M. Levine, and V. Pra-

do. 2004. Multiplex PCR for diagnosis of enteric

infections associated with diarrheagenic Esch-

erichia 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. 53:4633-

4636.

Warnke, P. H., S. T. Becker, R. Podschun, S. Siva-

nanthan, 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. Cranio-

maxillofac 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.




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 fermenta-

tion patterns of many ruminal microorganisms. In the rumen, methanogens consume hydrogen to gener-

ate 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 cul-

ture 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, [email protected]: +1-765-494-4826 Fax: +1-765-494-9347

(e.g., cellulose) are degraded to monomeric carbo-

hydrates (e.g., glucose) which are primarily ferment-

ed 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 fer-

mentation acids provide 60 to 80% of the daily me-

tabolizable energy intake of ruminants (Annison and

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 479072 Current address: 147 Kelton St., Allston, MA 02134



Armstrong, 1970), microbial cells provide an impor-

tant 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 an-

aerobic ecosystems (McInerney et al., 2011). Hydro-

gen 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

fermentation patterns of many ruminal microorgan-

isms because hydrogen is an end product inhibi-

tor 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 con-

verted into an increase in overall microbial cell yields

(Wolin and Miller, 1983).

The energy present in methane escapes the ru-

men 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 di-

gestible energy of the diet of ruminants (Blaxter and

Clapperton, 1965), there has been an interest to spe-

cifically 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 pro-

teins (Chalupa, 1980).

Maintaining the beneficial effects of interspecies

H2 transfer while minimizing loss of energy as meth-

ane could enhance energy provided to ruminants by

22% (Schaefer, D., personal communication). How-

ever, an alternative electron sink is required to trap

electrons into a form utilizable by the animal if meth-

anogens are to be directly inhibited. To date the

major method used to manipulate rumen fermen-

tation has been the use of ionophore antibiotics

such as monensin and lasalocid. These compounds

improve the efficiency of animal production by de-

creasing methane production and increasing ruminal

propionate concentration by 15%. Methane produc-

tion 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). Acetogen-

esis has been demonstrated to be the predominant

fate of H2 in some humans, swine, xylophagus ter-

mites, cockroaches and rats (Breznak and Blum, 1991;

Lajoie et al., 1988). Replacing methanogenesis with

acetogenesis in the rumen may have potential in de-

creasing energy losses in ruminants. Peptostrepto-

coccus productus (Bryant et al., 1958), Eubacterium

limosum (Genthner, 1981), and Acetitomaculum ru-

minis (Greening and Leedle, 1989) are chemolitho-

autotrophic acetogenic bacteria that have been iso-

lated from the bovine rumen. However they are not

considered the primary H2 consuming organisms in

this environment, since their numbers are consistent-

ly lower than methanogens.

Factors dictating whether acetogenesis or metha-

nogenesis will predominate in anaerobic environ-

ments 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

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


to have H2 thresholds 10 to 40 fold lower than ace-

togens (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 (Ty-

ler, 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 gastrointesti-

nal 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 emis-

sions from livestock and livestock waste while main-

taining interspecies H2 transfer would improve ru-

minant productivity and at the same time would be

beneficial for the environment. Certainly acetogens

offer that possibility, but rumen ecosystem condi-

tions 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 at-

tempted 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 candi-

dates. Batch culture growth experiments, while not

necessarily representative of the rumen from a pas-

sage 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 includ-

ing 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 Pat-

terson, 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.


Source of Organisms

Acetobacterium woodii (ATCC 29683) was ob-

tained 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 Pat-

terson, 2011, 2012, 2013; Jiang et al., 2012).

Media and Growth Conditions

Growth and H2 threshold experiments were con-

ducted 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 Ace-

tobacterium woodii. The prepared medium was dis-

pensed 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 auto-

claving or by injecting the solution through a 0.2 μm

filter (Nalgene, Nalge Company, Rochster, NY).

For chemolithoautotrophic growth in broth me-

dium, 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 appropri-

ate 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


Table 1. Media compositiona

Acetogen Medium

(amounts per liter)

MAC-20 Mediumb

(amounts per liter)

Rumen Fluid 50.0 ml ---

Mineral 1c 40.0 ml 40.0 ml

Mineral 2d 40.0 ml 40.0 ml

Additional Trace Min. Sol.e 10.0 ml 10.0 ml

Wolfe’s Trace Min. Sol.f 10.0 ml 10.0 ml

Vitamin Solutiong 10.0 ml 10.0 ml

Na2CO3 4.0000 g 4.0000 g

Yeast Extract 0.5400 g 2.0000 g

Casein Hydrolysate --- 1.0000 g

Betaine --- 1.0000 g

NH4Cl 0.5400 g 0.5400 g

Cysteine.HCl 0.5000 g 0.5000 g

Resazurin solution 0.0010 g 0.0010 g

Hemin solution 0.0001 g 0.0001 g

a All components, except Na2CO3 and cysteine. HCl were dissolved in distilled water and brought to a vol-ume 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

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

c Mineral 1 (g/liter): 6.00 K2HPO4

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

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

f 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

g 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


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 incuba-

tion the container was flushed for 2 min and then pres-

surized 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 anaero-

bic water. The alfalfa was dried at 60°C and ground

through a 1 mm screen. Ruminal contents were col-

lected 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 incu-

bated in a rotatory shaker at 37°C operating at 200

rpm.

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 Gas-pak 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


Acetogenic bacteria inoculum prepara-tion

Enumeration of acetogen strains to prepare inoc-

ulum was done by growing cultures in acetogen me-

dium 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 seri-

ally 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 experi-

ments, the acetogens were grown under the same

growth conditions that were used for enumeration

and diluted to the appropriate concentration. Cul-

tures were centrifuged anaerobically at 8,000 rpm for

15 minutes and resuspended in acetogen medium

to reach the CFU/ml desired. The titer for each

acetogen strain used for methanogen replacement

studies was determined before each experiment.

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

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 ace-

togen medium to reach final concentrations ranging

between 0.0 to 10 mM, respectively. Both the 5mM

and 10 mM BES effectively inhibited methanogen-

esis 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 in-

stead. Acetogenic bacteria utilized were A. woodii,

and strains A10 and G3.2a. Acetogen inocula were

prepared as described previously to give a final con-

centration of 5x108 CFU/ml. Duplicate serum bottles

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.

Table 2. Anaerobic dilution solution (ADS) compositiona

Component (amounts per liter)

Mineral 1b 75.0 ml

Mineral 2c 75.0 ml

Cysteine.HCl 0.5000 g

Resazurin solution 0.0010 g

a 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 anaerobi-cally for 12 min at 121°C and 15 psib Mineral 1 (g/liter): 6.00 K2HPO4

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


Effect of acetogenic bacteria on H2 uti-lization 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 vol-

ume 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 pro-

files of cultures were also determined at 12, 48 and

72 h by gas chromatography.

Similar procedures were used to determine ef-

ficiency of acetogen strains G1.5a, G1.5e, G2.4a,

and G3.2a except 40 mM of MES (final concentra-

tion) was added for additional buffering. A control

treatment receiving sterile anaerobic water instead

of BES was also added. Final concentrations of ace-

togens 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 modi-

fied 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 experimen-

tal medium, and a control treatment received sterile

anaerobic water instead of BES. In separate experi-

ments, the acetogen strain G2.4a was added to pro-

vide final concentrations of 5x107 and 1x108 CFU/ml.

Headspace gas volume and H2 and CH4 concentra-

tions 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

concentration of 5x108, 1x109, and 5x109 CFU/ml.

Headspace gas volume and H2 and CH4 concentra-

tions were measured from duplicate serum bottles

for each treatment at 0, 12, and 36 h.

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 acidi-

fied 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, Belle-

fonte, 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). Tempera-

tures 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 min-

ute. 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 con-

centrations. The regression line was then utilized to

calculate H2 and CH4 concentrations in experimen-

tal samples. All gas mixtures were purchased from

Airco (Indianapolis, IN).


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.


RESULTS

Functional Replacement of Methano-genesis with Acetogenesis in Ruminal Contents

The effect of 0.0 to 10 mM of BES on methano-

genesis 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

than either A. woodii or the control plus BES cultures

(Figure 5B). Hydrogen concentrations for cultures

receiving A10 and G3.2a were below 18 μmoles by

36 h (Figure 2B). Hydrogen concentrations for cul-

tures receiving A. woodii were similar to the control

plus BES cultures at 12 h, but H2 concentrations did

not decline as rapidly over time (Figure 2B), suggest-

ing 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 concentra-

tion 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 con-

centrations 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-

tions were similar for both treatments (Figure 3).

An additional experiment was conducted to de-

termine how well 4 acetogen isolates could replace

methanogens as a hydrogen sink in ruminal con-

tents. Methane was produced only in the control

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. Metha-nogenesis was inhibited by 5 mM BES in all treatments. Cultures were incubated in triplicate and one standard deviation is depicted with error bars.


Figure 4. CH4 production (Fig. A) and H2 utilization (Fig. B) by ruminal contents after the addi-tion 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.


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.


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 devia-tion is depicted with error bars.


culture where no BES was added (Figure 4A). The

acetogen isolate G2.4a had the lowest H2 concentra-

tion, 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 be-

tween 24 and 72 h of incubation (Figure 4B), indi-

cating that ruminal contents have some capability to

utilize H2.

Dose response experiments were performed to

determine the optimal acetogen dose for the func-

tional 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 con-

centrations 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 in-

cubation. The positive effect of increasing acetogen

numbers persisted even after 72 h of incubation (Fig-

ure 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 aceto-

gen isolate G3.2a with dosages increasing between

5x108 to 5x109 CFU/ml. Methane production oc-

curred 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

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 suffi-

cient to decrease H2 concentrations effectively.

DISCUSSION

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 pro-

duction (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

significantly decreased H2 concentrations at 12 hr,

compared to the 5mM BES control and Acetobac-

terium woodii. H2 concentration decreased by 40%

in the controls, indicating that there is some ability

to utilize H2 in ruminal contents. However, H2 con-

centrations 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 incuba-

tion 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 concen-

tration (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 in-

cubation bottles and 40 mM MES was added to pro-

vide additional buffering. In the first dose level ex-

periment, isolate G2.4a was inoculated at 5 x 107 cfu/

ml and 1 x 108 cfu. Under these conditions, H2 con-

centrations were significantly lower for the higher

concentration of acetogen addition at all time points

(Fig. 5). In the second dose level experiment isolate


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 accumu-

lation 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 in-

cubations where methanogenensis has been inhib-

ited 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 fer-

mentation, although Pinder and Patterson (2012)

have shown diauxic growth of isolate A10 in the pres-

ence of glucose in pure culture. The data also show

that different isolates have different capacities for uti-

lizing 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 ace-

togens in the rumen. Methanogens have lower H2

thresholds than acetogens. The acetogens we iso-

lated 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 hy-

drogen 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 di-

rectly 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.

The differences between pure culture H2 threshold

measurements and ability to reduce H2 concentra-

tions during incubation may be because H2 threshold

measurements estimate the lowest level of H2 the or-

ganism can utilize over time with no other nutrients

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 ca-

pabilities for H2 utilization in batch culture. Thus,

there is potential for acetogens to effectively utilize

H2 for interspecies H2 transfer to increase efficiency

of ruminal fermentation, trap more of the feedstuff

energy into acetate in the absence of methanogen-

esis. Feasibility of utilizing acetogens increase effi-

ciency of ruminant fermentation is dependent upon

identifying safe, cost effective methods to inhibit

methanogenesis.

REFERENCES

Adamse, A.D. 1980. New Isolation of Clostridium

aceticum (Wieringa). Antonie van Leeuwenhoek

46: 523-531.

Andreesen, J.R., A. Schaup, C. Neurauter, A. Brown,

and L.G. Ljungdahl. 1973. Fermentation of glucose,

fructose and xylose by Clostridium thermoaceti-

cum: effect of metals on growth yields, enzymes,

and synthesis of acetate from CO2. J. Bacteriol.

114:742-751.

Annison, E.F. and D.G.Armstrong. 1970. Volatile fatty

acids metabolism and energy supply. In: Phillipson

AT et al. (eds) Physiology of Digestion and Me-

tabolism in the Ruminant. Oriel Press, Newcastle,

England, pp 422-437.

Balch, W.E. and R.S. Wolfe. 1976. A new approach to

the cultivation of methanogenic bacteria: 2-mer-

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.


1977. Acetobacterium a new genus of hydrogen-

oxidazing, carbon dioxide-reducing, anaerobic

bacteria. Int. J. Sys. Bacteriol. 27:355-361.

Bernalier, A., A. Willems, M. Leclerc, V. Rochet, and

M.D. Collins 1996. Ruminococcus hydrogenotro-

phicus sp. nov., a new H2/CO2-utilizing acetogenic

bacterium isolated from human feces. Arch. Mi-

crobiol. 166:176-183.

Blaxter, K.L. and C.L. Clapperton. 1965. Prediction of

the amount of methane produced by ruminants.

Br. J. Nutr. 19: 511-522.

Boccazzi, P., and J.A. Patterson. 2013. Isolation and

initial characterization of acetogenic ruminal bac-

teria resistant to acidic conditions. Agric. Food

Anal. Bacteriol. 3: 129-144.

Boccazzi, P., and J.A. Patterson. 2011. Using hydro-

gen limited anaerobic continuous culture to iso-

late low hydrogen threshold ruminal acetogenic

bacteria. Agric. Food Anal. Bacteriol. 1:33-44.

Branine, M.E. and D.E. Johnson. 1990. Level of in-

take effects on ruminant methane loss across a

wide range of diets. J. Anim. Sci. 68 (Suppl.1):509

Braun M, Mayer F, Gottschalk. 1981. Clostridium

aceticum (Wieringa), a microorganism producing

acetic acid from molecular hydrogen and carbon

dioxide. Arch. Microbiol. 128:288-293.

Braun, M. and G. Gottschalk. 1982. Acetobacterium

wieringae sp. nov., a new species producing acetic

acid from molecular hydrogen and carbon dioxide.

Zbl. Bakt. Hyg., I abt Orig. C3 pp 368-376

Breznak, J.A., J.M. Switzer, and H.-J. Seitz. 1988.

Sporomusa termitida sp.nov., an H2/CO2-utilizing

acetogen isolated from termites. Arch. Microbiol.

150: 282-288.

Breznak, J.A. and M.D. Kane. 1990. Microbial H2/CO2

acetogenesis in animal guts: nature and nutritional

significance. FEMS Microbiol Rev. 87: 309-314.

Breznak, J.A. and J.S. Blum. 1991. Mixotrophy in the

termite gut acetogen, Sporomusa termitida. Arch.

Microbiol. 156: 105-110.

Bryant, M.P., N. Small, C. Bouma, and I. Robinson.

1958. Studies on the composition of the ruminal

flora and fauna of young calves. J. Dairy Sci .4:

1747-1767.

Bryant, M.P. and L.A. Burkey. 1953. Cultural methods

and some characteristics of some of the more nu-

merous groups of bacteria in the bovine rumen. J.

Dairy Sci. 36: 205-217.

Bryant, M.P. 1972. Commentary on the Hungate

technique for culture of anaerobic bacteria. Amer.

J. Clin. Nutr. 25:1324-1328.

Chalupa, W. 1980. Chemical control of rumen mi-

crobial metabolism. In: Ruckebusch Y, Thivend P

(eds) Digestive Physiology and Metabolism in Ru-

minants. MTP Press, Lancaster, England

Charakhch’yan, D.-I.A., A.N. Mileeva, L.L. Mityushi-

na, and S.S. Belyaev. 1992. Acetogenic bacteria

from oil fields of Tartaria and western Siberia. Mik-

robiologiya 61:306-315

Cord-Ruwisch, C., H.-J. Seitz, and R. Conrad. 1988.

The capacity of hydrogenotrophic anaerobic bac-

teria to compete for traces of hydrogen depends

on the redox potential of the terminal electron ac-

ceptor. Arch. Microbiol. 149: 350-357.

Czerkawski, J.W. and G. Brecknridge. 1971. Deter-

mination of concentration of hydrogen and some

other gases dissolved in biological fluids. LABP

20: 403-413.

Dehning, I., M. Stieb, and B. Schink. 1989. Sporomu-

sa malonica sp. nov., a homoacetogenic bacterium

growing bydecarboxylation of malonate or succi-

nate. Arch. Microbiol. 151: 421-426.

Drake, H.L. 1992. Acetogenesis and acetogenic bac-

teria. In: Ledeberg J (ed) Encyclopedia of Micro-

biology. Academic Press Inc. San Diego, V.1, pp

1-15.

Drake, H.L. 1994. Acetogenesis, acetogenic bacte-

ria, and the acetyl-CoA ‘Wood/Ljungdahl’ path-

way: past and current perspectives. In:Drake HL

(ed) Acetogenesis. Chapman & Hall, New York pp

3-60.

Eichler, B. and B. Schink. 1984. Oxidation of primary

aliphatic alcohols by Acetobacterium carbinolicum

sp. nov., a homacetogenic anaerobe. Arch. Micro-

biol. 140:147-152.

EPA 1993. Methane emission and opportunity for

control. United States Environmental Protection

Agency EPA 430/R-93/003.

Fontaine, F.E., W.H. Peterson, E. McCoy, M.A. John-

son, and G.J. Ritter. 1942. A new type of glucose


fermentation. Clostridium thermoaceticum n. sp.

J. Bacteriol. 43: 701-715.

Fuchs, G. 1986. CO2 fixation in acetogenic bacte-

ria, variation on a theme. FEMS Microbiol. Rev. 39:

181-213.

Garcia-Lopez, P.M., L. Kung Jr., and J.M. Odom.

1996. In vitro inhibition of microbial methane pro-

duction by 9,10-anthraquinone. J. Anim. Sci. 74:

2276-2284.

Geerligs, G., H.C. Aldrich, W. Harder, and G. Diekert.

1987. Isolation and characterization of carbon

monoxide utilizing strain of the acetogen Pepto-

streptococcus productus. Arch. Microbiol. 148:

305-313.

Gottwald, M, J.R. Andreesen, J. LeGall, and L.G.

Ljungdahl. 1975. Presence of cytochrome and

menaquinone in Clostridium formicoaceticum and

Clostridium thermoaceticum. J. Bacteriol. 122:

325-328.

Greening, R.C. and J.A.Z. Leedle. 1989. Enrichment

and isolation of Acetitomaculum ruminis, gen.

nov., sp. nov.: acetogenic bacteria from the bovine

rumen. Arch. Microbiol. 151:399-406.

Herman, R., M.R. Popoff, and M. Sebald. 1987.

Sporomusa paucivorans sp. nov., a methylotrophic

bacterium that forms acetic acid from hydrogen

and carbon dioxide. Int. J. Syst. Bacteriol. 37: 93-

101.

Holdeman, L.V., E.P. Cato, and W.E.C. Moore. 1977.

Anaerobic cocci. In: Anaerobe Laboratory Manual,

4th ed. Virginia Polytechnic Institute and State

University, Blacksburg, VA.

Hungate, R.E. 1966. The rumen and its microbes.

Academic Press, New York, NY.

Ivey, D.M. 1987. Generation of energy during CO2

fixation in acetogenic bacteria. Dissertation, Dept.

of Biochemistry, University of Georgia, Athens, GA.

Immig, I, D. Demeyer, D. Fiedler, C. Van Nevel, and

L. Mbanzamihigo. 1996. Attempts to induce re-

ductive acetogenesis into a sheep rumen. Arch.

Tierernahr. 49(4): 363-370.

Jiang, W., R.S. Pinder, and J.A. Patterson. 2012. Influ-

ence of growth conditions and sugar substrate on

sugar phosphorylation acitivity in acetogenic bac-

teria. Agric. Food Anal. Bacteriol.. 2: 94-102.

Kane, M.D. and J.A. Breznak. 1991. Acetonema long-

um gen. nov. sp. nov. an H2/CO2 acetogenic bac-

terium from the termite, Pterotermes occidentis.

Arch. Microbiol. 156: 91-98.

Kane, M.D, A. Brauman, and J.A. Breznak. 1991.

Clostridium mayombei sp. nov., an H2/CO2 aceto-

genic bacterium from the gut of the African soil-

feeding termite, Cubitermes speciosus. Arch. Mi-

crobiol. 156: 99-104.

Kesava, R.N.A., V.E. Worrel, R. Teal, and N.P. Nagle.

1996. The pterin lumazin inhibits growth of metha-

nogens and methane formation. Arch. Microbiol.

166: 136-140.

Kotsyurbenko, O.R., M.V. Simankova, A.N.

Norzhevnikova, T.N. Zhilina, N.P. Bolotina, A.M.

Lysenko, and G.A. Osipov. 1995. New species of

psychrophilic acetogens: Acetobacterium bakii sp.

nov., A. paludosum sp. nov., A. fimentarium sp.

nov.. Arch. Microbiol. 163: 29-34.

Krumholz, L.R. and M.P. Bryant.1985. Clostridium

pfennigii sp. Nov. uses methoxyl groups of mono-

benzenoids and produces butyrate. Int. J. Syst.

Bacteriol. 35: 454-456.

Krumholz, L.R. and M.P. Bryant. 1986. Syntropho-

coccus sucromutans sp. Nov. gen. Nov. uses car-

bohydrates as electron donors and formate, me-

thoxymonobenzenoids or Methanobrevibacter as

electron acceptor system. Arch. Microbiol. 143:

3113-318.

Lajoie, S.F., S. Bank, T.L. Miller, and M.J. Wolin. 1988.

Acetate production from hydrogen and [14] car-

bon dioxide by the microflora of human feces.

Appl. Environ. Microbiol. 54: 2723-2737.

Leedle, J.A.Z. and R.B. Hespell. 1980. Differential

carbohydrate media and anaerobic replica plating

techniques in delineating carbohydrate-utilizing

subgroups in rumen bacterial populations. Appl.

Environ. Microbiol. 39:709-719.

Leigh, J.A., F. Mayer, and R.S. Wolfe. 1981. Acetoge-

nium kivui, a new thermophilic hydrogen-oxidaz-

ing, acetogenic bacterium. Arch. Microbiol. 129:

275-280.

LeVan, T.D., J.A. Robinson, J. Ralph, R.C. Greening,

W.J. Smolenski, J.A.Z. Leedle, and D.M. Schaefer.

1998. Assessment of reductive acetogenesis with


indigenous ruminal bacterium populations and

Acetitomaculum ruminis. Appl. Environ. Micro-

biol. 64:3429-3436.

Ljungdahl, L.G. 1986. The autotrophic pathway of ac-

etate synthesis in acetogenic bacteria. Annu. Rev.

Microbiol. 40: 415-450.

Ljungdahl, L.G. 1994. The acetyl-CoA pathway and

the chemiosmotic generation of ATP during ace-

togenesis. In:Drake HL (ed) Chapman & Hall, New

York pp 63-87.

Lorowitz, W.H. and M.P. Bryant. 1984. Peptostrepto-

coccus productus strain that grows rapidly with CO

as the energy source. Appl. Environ. Microbiol. 47:

961-964.

Lupton, F.S. and J.G. Zeikus. 1984. Physiological ba-

sis for sulfate-dependent hydrogen competition

between sulfidodogens and methanogens. Curr.

Microbiol. 11: 7-12.

Mayer, F., M.D. Ivey, and L.G. Ljungdahl. 1986. Mac-

romolecular organization of FI-ATPase isolated

from Clostridium thermoaceticum as revealed by

electron microscopy. J. Bacteriol. 166: 1128-1130.

McInerney, M.J., J.R. Sieber, and R.P. Gunsalus. 2011.

Microbial syntrophy: Ecosystem-level biochemical

cooperation. Microbe 6: 479-485.

Möller, B., R. Oßmer, B.H. Howard, G. Gottschalk,

and H. Hippe. 1984. Sporomusa a new genus

of gram-negative anaerobic bacteria including

Sporomusa sphaeroides spec. nov. and Sporomu-

sa ovata spec. nov. Arch. Microbiol. 139:388-396.

Moss, A.R., J. Jouany and J. Newbold. 2000. Meth-

ane production by ruminants: its contribution to

global warming. Ann. Zootech. 49:231-253.

Ollivier, B., R. Cordruwisch, A. Lombardo, J.L. Garcia,

R. Robinson. 1985. Isolation and characterization

of Sporomusa acidovorans sp. nov., a methylotro-

phic homoacetogenic bacterium. Arch. Microbiol.

142:307-310.

Patel, B.K.C., C. Monk, H. Littleworth, H.W. Morgan,

and D.M. Daniel. 1987. Clostridium fervidus sp.

nov., a new chemoorganotrophic acetogenic ther-

mophile. Int. J. Syst. Bacteriol. 37: 123-126.

Pearman, G.I. and P.J. Fraser. 1988. Sources of in-

creased methane. Nature 332: 489-490.

Pinder, R.S. and J.A. Patterson. 2013. Growth of

acetogenic bacteria in response to varying pH, ac-

etate or carbohydrate concentration. Agric. Food

Anal. Bacteriol. 3: 6-16.

Pinder, R.S. and J.A. Patterson. 2012. Glucose and

hydrogen utilization by an acetogenic bacterium

isolated from ruminal contents. Agric. Food Anal.

Bacteriol. 2: 253-274.

Pinder, R.S. and J.A. Patterson. 2011. Isolation and

initial characterization of plasmids in an acetogen-

ic ruminal isolate. Agric. Food Anal. Bacteriol. 1:

186-192.

Ragsdale, S.W. 1991. Enzymology of the acetyl-CoA

pathway of CO2 fixation. Crit. Rev. Biochem. Mol.

Biol. 26(3/4): 261-300.

Ricke, S.C., S.A. Martin, and D.J. Nisbet. 1996. Ecol-

ogy, metabolism, and genetics of ruminal sele-

nomonads. Crit. Rev. Microbiol. 22: 27-56.

Ricke, S.C. and D.M. Schaefer. 1996. Nitrogen-lim-

ited growth response of ruminal bacterium Sele-

nomonas 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. Biore-

source Technol. 50: 253-257.

Ricke, S.C. and D.M. Schaefer. 1991. Growth inhi-

bition of the rumen bacterium Selenomonas ru-

minantium 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 an-

aerobic ecosystems: application to the rumen.

Appl. Environ. Microbiol. 41:545-548.

Russell, J.B. and R.L. Baldwin. 1978. Substrate pref-

erences in rumen bacteria: Evidence of catabolite

regulatory mechanisms. Appl. Environ. Microbiol.

36: 319-329.

Russell, J.B. and H.J. Strobel. 1989. Effect on iono-

phores on ruminal fermentation. Appl. Environ.

Microbiol. 55:1-6.

Saengkerdsub, S. and S.C. Ricke. 2014. Ecology and

characteristics of methanogenic archaea in animals

and humans. Crit. Revs. Microbiol. 40: 97–116.

Schaefer, D.M., C.L. Davis, and M.P. Bryant. 1980.


Ammonia saturation constants for predominant

species of rumen bacteria. J. Dairy Sci. 63: 1248-

1263.

Schink, B. 1984. Clostridium magnum sp. nov., a non-

autotrophic homoacetogenic bacterium. Arch. Mi-

crobiol. 137: 250-255.

Sharak-Genthner, B.R., C.L. Davies, and M.P. Bry-

ant. 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. Acetoan-

aerobium notarae gen. nov. sp. nov.: an anaerobic

bacterium that forms acetate from H2 and CO2. Int.

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 ljungda-

hlii PETC sp. nov., a new, acetogenic, gram-posi-

tive, anaerobic bacterium. Abstr R-21, p 249 Abstr

Ann Meet Am Soc Microbiol.

Tanner, R.S., L.M. Miller, and D. Yang. 1993. Clostrid-

ium 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 pro-

duction 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. Les-

sons from the cow: What the ruminant animal can

teach us about consolidated bioprocessing of

cellulosic biomass. Bioresour. Technol. 100: 5323-

5331.

Wiegel, J., M. Braun and G. Gottschalk. 1981. Clos-

tridium thermoautotrophicum species novum, a

thermophile producing acetate from molecular

hydrogen and carbon dioxide. Curr. Microbiol.

5:255-260.

Wieringa, K.T. 1936. Over het cerdwijnen van water-

stof en koolzuur onder anaerobe voorwaarden.

Antonie van Leeuwenhoek 3:263-273.

Wolin, M.J. and T.L. Miller. 1983. Interactions of mi-

crobial populations in cellulose fermentation. Fed-

eration 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. Aca-

demic Press Limited pp 201-250.

Zeikus, J.G., L.H. Lynd, J.A. Thompson, .J. Krzycki,

P.J. Weimer, and P.W. Hegge. 1980. Isolation and

characterization of a new methylotrophic, acido-

genic anaerobe, the Marburg strain. Curr. Micro-

biol. 3:381-386.

Zhilina, T.N., and G.A. Zavarzin. 1990. Extremely

halophilic, methylotrophic, anaerobic bacteria,

FEMS Microbiol Rev 87: 315-322.




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 (con-

trol), 2) chestnut extract at 100 g/d, and 3) quebracho tannin extract at 100 g/day. In experiment 1, aver-

age 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 experi-

ment 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 tan-

nins 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

Correspondence: B.R. Min, [email protected]: +1-334-524-7670 Fax: +1-334-727-8552

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

1Tuskegee University, Tuskegee, AL, USA

2Montgomery Blair High School, Silver Spring, MD, USA3Utah State University, Logan, UT, USA



INTRODUCTION

The microbial populations of the rumen, par-

ticularly 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 limit-

ed, 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 con-

densed tannins (CT; proanthocyanidins) based on

their molecular structure. (Min et al., 2003). Our un-

derstanding of interactions between rumen bacte-

ria and HT or CT in the rumen is still in its infancy.

Tannins have traditionally been considered antinu-

tritional but it is now known that their beneficial or

antinutritional properties depend upon their chemi-

cal 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 bio-

logical effects and, therefore, results obtained using

a particular type of tannins cannot be applied to oth-

ers. 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 ru-

men may affect the rumen fermentation and micro-

biome 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 fer-

mentability 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

comparing three different types of tannins (quebra-

cho, acacia and chestnut) demonstrated that chest-

nut tannins are more efficient in protecting soybean

meal from in vitro degradation by rumen bacteria.

This has been confirmed by the findings that supple-

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 flave-

faciens populations were decreased with chestnut

and mimosa tannins supplemented animals (Min et

al., 2012a). The implication of sources of plant tan-

nins (Chestnut vs. Quebracho tannins extracts or CT-

containing 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-

mance. The primary hypothesis of the in vivo grazing

research was that different sources of tannins supple-

mentation would selectively reduce rumen microbial

diversity and as a result would increase average daily

gain in meat goats. Thus, our objective was to in-

vestigate concurrent changes in ruminal bacterial

diversity and animal performance in goat response

to plant tannins using a modern pyrosequencing ap-

proach.


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 ± 2.55 kg)

were randomly assigned to 2 experimental diets (al-

falfa pellet vs. PB powder). Alfalfa pellet (no CT as a

control) or PB (11% CT) was supplemented at 0.5%

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 wa-

ter and trace mineral salt blocks grazing on winter


pea and rye grass dominant forages. Animal body

weight (BW) were measured before and after experi-

ment 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 af-

ter 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 ran-

domly assigned to 3 experimental diets: 1) no tan-

nins (control), 2) chestnut HT-extract at 100 g/d (CTE),

and 3) quebracho CT-extract at 100 g/d (QCTE). Ex-

perimental 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-

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.

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; Ar-

thur H. Thomas Co., Swedesboro, NJ), and stored

for subsequent analyses. Daily portions of ground

samples were composited for each animal and ana-

lyzed for DM, crude protein (CP), acid detergent lig-

nin, ether extract, and ash according to the methods

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

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

Experimental diet

Winter forage Alfalfa pellet BermudagrassPine bark

powderSD

Ingredients

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 50 9.57

SD = standard deviation


an ANKOM200/220 Fiber Analyzer (ANKOM Technol-

ogy, Macedon, NY). Sodium sulfate heat stable amy-

lases (Sigma Aldrich Co., St. Louis, Mo) were used in

the procedure for NDF determination and pretreat-

ment with heat stable amylase (Type XI-A from Ba-

cillus 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,

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 param-

eters: 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, detec-

tor temperature = 260 °C.

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 ND-

1000 spectrophotometer (Nyxor Biotech, Paris,

France) and run on 0.8% agarose gel with 0.5 M Tris-

Borate-EDTA (TBE) buffer. The samples were then

transported to the Research and Testing Laborato-

ry (Lubbock, TX) for PCR optimization and pyrose-

quencing analysis. Bacterial tag-encoded FLX ampli-

con 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

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 amplify-

ing 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 elonga-

tion 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 tenta-

tive consensus FASTA (A database search tool used

to compare a nucleotide or peptide sequence to a

sequence database) for each sample was then evalu-

ated 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.

Data Processing and Statistical Analysis

Statistical analyses were performed using the

SPSS package (SPSS Inc., v 17.0, Chicago, IL). Pack-

age 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 ser-

vices by the Research and Testing Laboratory (Lub-

bock, TX; (Dowd et al., 2008). Tags which did not

have 100% homology to the original sample tag

designation were not included in data analysis. Se-

quences which were less than 250 bp after quality

trimming were not also considered. The resulting se-

quences 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%.


Relative abundance data are presented as per-

centages/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 random-

ized design with the factors examined being sources

of tannins supplementation in the diets. Results are

reported as least square means.


Regardless of numerous studies (Pitta et al., 2010;

Callaway et al., 2010; Hristov et al., 2012) demon-

strating the role of the gut microbial diversity in ru-

minants associated with different sources of forages

or dried distillers grains, the response of the micro-

biome to feeding various sources of phytochemical

tannins-containing diet remain largely unknown. The

most significant findings in the present study dem-

onstrates that when goats received tannins extracts

(chestnut and quebracho extracts) in the Firmicutes

phylum populations had significant (P < 0.01) de-

creased, while Bacteroidetes populations were

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)

Exp.1 (n = 10/diet)1

PB powder 37.3 46.6 169.1

Control 36.5 46.7 185.5

SD 8.05 0.34 12.67

P-value 0.76 0.35 0.71

Exp.2 (n = 4/diet)2

Chestnut 32.7 36.6 278.6

Quebracho 33.5 35.6 150.0

Control 30.5 31.1 42.9

SD 5.88 6.98 20.02

P-value 0.85 0.26 0.05

1 Animals were grazed on winter pea and ryegrass pasture with or without PB powder supplementation dur-ing 55 days2 Animals were grazed on bermudagrass forage with or without tannin extracts supplementation during 14 days.


significantly increased. The bacterial distribution

showed that Firmicutes (56-57%) was the most domi-

nant phyla with mean relative abundance values

ranging from 56% in control to 67 % in PB diets. This

suggests that phytochemical tannins supplementa-

tion alters the microbiome and animal performance

on goats grazing fresh forage diets.

Diet Composition

Ingredients and chemical composition of experi-

mental 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 nutri-

ent 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 Fer-mentation

The animal performance and ruminal volatile fatty

acids concentrations in goats grazing fresh forages

in response to different sources of tannins supple-

mentation 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 vol-

atile 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

for CTE (275 g/d) than QCTE (145 g/d) and the con-

trol (41 g/d). Goats grazing bermudagrass pasture

with CTE had greater (P < 0.05) concentrations of

acetate, propionate, butyrate, caprionate, and to-

tal volatile fatty acids concentrations compared to

those in QCTE and control (Table 3). However, goats

grazing bermudagrass forage without tannins sup-

plementation increased (P < 0.05) concentrations of

iso-valerate and acetate: propionate ratio compared

to tannins supplemented groups.

A number of studies have demonstrated that ef-

fects 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 sup-

plementation 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) experi-

enced 82% fewer days of bloat, and had 6 and 17%

greater ADG for mimosa and chestnut tannins ex-

tracts, respectively, than animals receiving the con-

trol diet principally through reducing the rate of

rumen fermentation as well as modification of micro-

bial populations in the rumen of cattle. Our chestnut

tannins supplementation study shows a similar trend

to this. This suggests that plant tannins supplemen-

tation in high quality forage may have more impact

on mitigating rumen fermentation and improving

animal performance than low quality forages diets.

Relative Abundance of Bacterial Phyla

In this study, bacterial (Fig. 1a,b) community com-

position of the rumen fluids were examined at de-

scending levels of biological classification to deter-

mine the effect of PB powder (Exp. 1) or tannin extract

(Exp. 2) supplementation on community membership.

Detailed phylogenic analyses grouped the rumen bac-

teria associated bacterial sequences into 45 phyla (in-

cluding unknown). The relative abundances of the 19

most abundant phyla (>1%) are presented in Figure 1.

Interestingly, the gut of human and many other verte-

brae are mostly dominated by two groups of bacteria,


Bacteroidetes and Firmicutes (Backhed et al., 2004),

which is similar to the result obtained current study.

This finding agrees with the results within the cur-

rent 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 approxi-

mately 89 to 92% of the total gut population. A study

by Henderson et al. (2013) demonstrated an increase

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

Firmicutes, Bacteroidetes, Actinobacteria, and Pro-

teobacteria 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

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 Bacteroide-

tes populations were significantly increased (Fig. 1b).

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 ac-etate: propionate (A:P) ratios in meat goats grazing winter pea and ryegrass dominant forages.

Item1 C2 C3 Iso-C4 C4 Iso-C5 C5 C6Total 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

SD 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

Exp.2 (n = 4/diet)2

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

SD 7.44 1.96 0.10 0.92 0.02 0.08 0.08 9.12 0.46

P-value 0.02 0.001 0.02 0.01 0.07 0.21 0.01 0.01 0.001

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


Figure 1. Predominant bacterial phylum observed in rumen samples of healthy goats with or with-out 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 Fir-micutes (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.

0

10

20

30

40

50

60

70

80

90

100

Control Pine bark powder

33.320.5

57.367.2

ParabasaliaProteobacteriaEuryarchaeotaStreptophytaTenericutesFirmicutesElusimicrobiaCyanobacteriaAcidobacteriaNitrospiraePlanctomycetesVerrucomicrobiaFibrobacteresLentisphaeraeSpirochaetesActinobacteriaSynergistetesChloroflexiBacteroidetes

A

B

0

10

20

30

40

50

60

70

80

90

100

Control Chestnut Quebracho

35.3

51.5 52.9

55.942.6 36.7

ParabasaliaProteobacteriaEuryarchaeotaStreptophytaTenericutesFirmicutesElusimicrobiaCyanobacteriaAcidobacteriaNitrospiraePlanctomycetesVerrucomicrobiaFibrobacteresLentisphaeraeSpirochaetesActinobacteriaSynergistetesChloroflexiBacteroidetes


The gastrointestinal microbiota performs a large

number of important roles that define the physiol-

ogy 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 mam-

mals, the gastrointestinal microbiome is dominated

by four bacterial phyla that perform these tasks:

Firmicutes, Bacteroidetes, Actinobacteria and Pro-

teobacteria (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

animals, while CTE and QCTE plant extracts supple-

mentation 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 supple-

mentations.

Past research into the correlation between gut mi-

crobiota and diet had demonstrated a complex rela-

tionship 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

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-

ly, an increase in weight (Turnbaugh et al., 2006).

The Bacteroidetes spp, in particular Bacteroides

thetaiotaomicron, hydrolyzes otherwise indigest-

ible polysaccharides and accounts for 10% to 15%

of caloric requirement in humans (Xu et al. 2003).

Human colonocytes derive 50% to 70% of their en-

ergy from butyrate, which is derived from complex

carbohydrates metabolized by Firmicutes spp via

fermentation (Pryde et al., 2002). However, the cur-

rent 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

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 hy-

pothesizes 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.

Diversity and Abundance of Rumen Bac-terial species

More than 332 bacterial species (including un-

known) were classified from the ruminal fluid of

the goats in this study. However, the relative abun-

dances of the 12 most abundant species (>1%) are

presented in Tables 4 and 5. Tannins are one of the

most abundantly available plant secondary metabo-

lites, and have positive or adverse effects on rumen

microbial populations, feed digestibility and animal

performance (Min et al., 2003). The bacterial spe-

cies distribution showed that Ruminococcaceae spp.

(12-15%) and Prevotella spp. (21-40%) were the most

dominant species with mean relative abundance val-

ues ranging from 42 to 55% in control group with-

out tannins supplementation (Table 4). In Exp. 1,


Prevotella spp. was decreased (P < 0.05) in PB diet,

while Butyrivibrio spp. (P < 0.05) and Ruminococ-

caceae spp. (P = 0.08) were increased compared to

control diet. However, bacterial populations in CTE-

supplemented group in Exp. 2 were significantly in-

creased for Prevotella spp. (40.4, 33, and 32%) com-

pared with QCTE and control groups, respectively.

It has been shown that the gastrointestinal micro-

bial 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-

laway et al., 2010). Consequently, analysis of human

microbiota-associated rat feces using molecular ap-

proach has revealed that the Bacteroides/Prevotella

and Faecalibacterium species are dominant in both

humans and rats post-transfection (Licht et al., 2007).

These findings also agree with the results of a

metabolic finger print study of a rat fed CT extract

from Acacia angustissima. Condensed tannin ex-

tracts from A. angustissima altered fecal bacterial

populations in the gastrointestinal tract, resulting in

a shift in the predominant bacteria towards tannin-

resistant gram-negative Enterobacteriaceae and

Bacteroidetes (Smith et al., 2003). Presence of bac-

teria 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).

Different groups of microbes have different toler-

ance to tannin. Rumen fungi, proteolytic bacteria

and protozoa are more resistant to tannin as com-

pared to other microbes (McSweeny et al., 2001).

McSweeny et al. (1999) observed that in the animals

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 SD 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

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


fed on tannin rich Calliandra calothyrsus, the popu-

lation 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, Streptococ-

cus bovis, Eubacterium spp. and Butyrivibrio fibri-

solvenes 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-

trol) to 9.9% (15% PB) and 17.2% (30% PB). The re-

maining 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-

etary PB concentration. However, Bacteroides cap-

illosus, 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 au-

thors 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

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 hier-

archal cluster analysis of individual animal microbial

diversity within and among diets in Figures 2 and 3.

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 SD 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

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


Figure 2 (Exp. 1). Heat-map double dendrogram of the 44 most abundant bacterial genera in the ru-men 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.


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. Cluster-ing 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.


Overall, animals clustered relatively well within diet

and animals. However, one of PB powder supple-

mented goats had more dissimilarity between treat-

ments. 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 pop-

ulation (Figure 2). In Exp. 2, chestnut and quebracho

tannin extracts supplemented groups had greater

relative abundance (%) of Bacteroidia compared to

the control diet. Lower abundance of Clostridia in

tannins extract groups compared to control diet, in-

dicated that tannins extracts supplementation may

have decreased the abundance of Clostridia popula-

tion in rumen of goats.

CONCLUSIONS

In conclusion, the current results show that tannin

can exert a positive or negative effect both on ru-

men fermentation and on rumen microflora, and it

is possible that this effect is depending on sources

of tannins or tannin-containing diet. Rumen micro-

bial population is very dynamic and tannin inclusion

impacts specific members of the microbial popula-

tion. There is also possible adaptation of ruminal mi-

crobiota 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 re-

sponse to ingestion of different sources of tannin-

containing diet.

ACKNOWLEDGEMENTS

This project was supported by USDA-NIFA, The

USDA-NIFA Evans-Allen Research Program and

Tuskegee University, George Washington Carver

Agricultural Research Station. Research and Testing

Laboratory (Lubbock, TX) for PCR optimization and

pyrosequencing analysis are also acknowledged.

REFERENCES

AOAC. 1998. Association of Official Analytical Chem-

ists. Official Methods of Analysis, Gaithersburg,

Md, USA, 16th ed.

Altschul, S. F., W. Gish, W. Miller, E. W. Myers, and D.

J. Lipman. 1990. Basic local alignment search tool.

J. Mol. Biol. 215:403–410.

Backhed, F., H. Ding, T. Wang, L. V. Hooper, G. Y.

Koh, A. Nagy, C. F. Semenkovich, and J. I. Gor-

don. 2004. The gut microbiota as an environmen-

tal factor that regulates fat storage. Proc. Natl.

Acad. Sci. USA 101:15718–15723.

Brooker, J. D., L. A. O’Donovan, K. Clarke, L. Black-

all, 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 dif-

ferent levels of dried distillers grains plus soluble

using bacterial tag-encoded FLX amplicon pyro-

sequencing. 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 be-

tween rumen ciliate protozoa and bacteria. Pages

149–164 in Digestion and Metabolism in the Rumi-

nant. 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-

zotti, J. B. Poullet, S. Massart, S. Collini, and G.

Pieraccini. 2010. Impact of diet in shaping gut

microbiota revealed by a comparative study in

children from Europe and rural Africa. Proc. Natl.

Acad. Sci. 107: 14691–14696.


Dowd, S.E., T. R. Callaway, R. D. Wolcott, R. D. Wol-

cott, Y. Sun, T. McKeehan, G. R. G. Hagevoort, and

T. S. Edrington. 2008. Evaluation of the bacterial

diversity in the feces of cattle using 16S rDNA bac-

terial tag-encoded FLX amplicon pyrosequencing

(bTEFAP). BMC Microbiol. 8:125.

Fernando, S. C., H. T. Purvis, F. Z. Najar, L. O.

Sukharnikov, C. R. Krehbiel, T. G. Nagaraja, B. A.

Roe, and U. DeSilva. 2010. Rumen microbial popu-

lation dynamics during adaptation to a high-grain

diet. Appl. Environ. Microbiol. 76:7482–7490.

González, S., M. L. Pabón, and J. Carulla. 2002. Ef-

fects of tannins on in vitro ammonia release and

dry matter degradation of soybean meal. Arch.

Latinoam. Prod. Anim. 10:97–101.

Henderson, G., F. Cox, S. Kittelmann, V. H. Miri, M.

Zethof, S. J. Noel, G. C. Waghorn, and P. H. Jans-

sen. 2013. Effects of DNA extraction methods and

sampling technologies on the apparent structure

of cow and sheep rumen microbial communities.

Plos One. 8:1-14.

Hervás, G., P. Frutos, F. J. Giráldez, A. R. Mantecón,

and M. C. Álvarez del Pino. 2003. Effect of differ-

ent doses of quebracho tannins extract on rumen

fermentation in ewes. Anim. Feed Sci. Technol.

109:65-78.

Hristov, A. N., T. R. Callaway, C. Lee, and S. E. Dowd.

2012. Rumen bacterial, archaeal, and fungal diver-

sity of dairy cows in response to lauric or myristic

acids ingestion. J. Anim. Sci. 90:4449–4457.

Krueger, W.K., H. Gutierrez, G. E. Carstens, B. R.

Min, W. E. Pinchak, R. R. Gomez, R. C. Anderson,

N. A. Krueger, and T. D. A. Forbes. 2010. Effects

of dietary tannin source on performance, feed effi-

ciency, ruminal fermentation, and carcass and non-

carcass traits in steers fed high-grain diet. Anim.

Feed Sci. Technol. 159:1-9.

Kurokawa K, T. Itoh, T. Kuwahara, K. Oshima, H. Toh,

A. Toyoda, H. Takami, H. Morita, V. K. Sharma, T.

P. Srivastava, T. D. Taylor, H. Noguchi, H. Mori, Y.

Ogura, D. S. Ehrlich, K. Itoh, T. Takagi, Y. Sakaki,

T. Hayashi, and M. Hattori. 2007. Comparative

metagenomics revealed commonly enriched

gene sets in human gut microbiomes. DNA Res.

14:169-181.

Ley, R.E., F. Ba-ckhed, P. Turnbaugh, C. A. Lozupone,

R. D. Knight, and J. I. Gordon. 2005. Obesity alters

gut microbial ecology. Proc. Natl. Acad. Sci. USA

102:11070–11075.

Ley, R. E, R. Knight, and J. I. Gordon. 2007. The hu-

man microbiome: eliminating the biomedical/en-

vironmental dichotomy in microbial ecology. Envi-

ron. Microbiol. 9:3-4.

Licht, T. R., B. Madsen, and A. Wilcks. 2007. Selec-

tion of bacteria originating from a human intestinal

microbiota in the gut of previously germ-free rats.

FEMS Microbiol. Lett. 277:205-209.

Mathieu, F. and J. P. Jouany. 1993. Effect of chestnut

tannin on the fermentability of soyabean meal ni-

trogen in the rumen. Ann. Zootech. 42: 127-127.

Mazmanian, S. K., C. H. Liu, A. O. Tzianabos, and D.

L. Kasper. 2005. An immunomodulatory molecule

of symbiotic bacteria directs maturation of the

host immune system. Cell. 122:107-118.

McSweeney, C. S., B. Palmer, R. Bunch, and D. O.

Krause. 1999. Isolation and characterisation of

proteolytic ruminal bacteria from sheep and goats

fed the tannin-containing shrub legume Callian-

dra calothyrsus Appl. Environ. Microbiol. 65:3075–

3083.

McSweeney, C. S., B. Palmer, R. Bunch, and D. O.

Krause. 2001. Microbial interactions with tannins:

nutritional consequences for ruminants. Anim.

Feed Sci. Technol. 91:83–93.

Min, B.R., T. N. Barry, G. T. Attwood, and W. C.

McNabb. 2003. The effect of condensed tannins

on the nutrition and health of ruminants fed fresh

temperate forages: a review. Anim. Feed Sci. Tech-

nol. 106:3–19.

Min, B. R., G. T. Attwood, K. Reilly, J. S. Peters, T. N.

Barry, and W. C. McNabb. 2002. Lotus corniculatus

condensed tannins decrease in vivo populations

of proteolytic bacteria and affect nitrogen me-

tabolism in the rumen of sheep. Can. J. Microbiol.

48:911–921.

Min, B. R., W. E. Pinchak, K. Hernandez, C. Hernan-

dez, M. E. Hume, E. Valencia, and J. D. Fulford.

2012a. The effect of plant tannins supplemen-

tation on animal responses and in vivo ruminal

bacterial populations associated with bloat in


heifers grazing wheat forage. The Prof. Anim.

Scientist. 28:464-472.

Min, B. R., S. Solaiman, N. Gurung, J. -S. Eun, E.

Taha, and J. Rose. 2012b. Effects of pine bark sup-

plementation on performance, rumen fermenta-

tion, and carcass characteristics of Kiko crossbred

male goats. J. Anim. Sci. 90:3556–3567.

Min, B. R., S. Solaiman, R. Shange, and J.-S. Eun.

2014. Gastrointestinal bacterial and methano-

genic Archaea diversity dynamics associated with

condensed tannins-containing pine bark diet in

goats using 16S rDNA amplicon pyrosequenc-

ing. Intl. J. Microbiol. 2014:1-11. http://dx.doi.

org/10.1155/2014/141909.

Muller-Harvey, I. and A. B. McAllan. 1992. Tannins:

Their biochemistry and nutritional properties.

Adv. Plant Cell Biochem. And Biotechnol. 1:151–

217.

Nelson, K. E., M. L. Thonney, T. K. Woolston, S. H.

Zinder, and A. N. Pell. 1998. Phenotypic and phy-

logenetic characterization of ruminal tannin-tol-

erant bacteria. Appl. Environ. Microbiol. 64:3824-

3830.

National Research Council (NRC). 2007. Nutri-

ent Requirement of Sheep, Goats, Cervids and

Camelids. Academy Press, Washington, D.C.

Pages 10-200.

Odenyo, A. A. and P. O. Osuji. 1998. Tannin-toler-

ant ruminal bacteria from East African ruminants.

Can. J. Microbiol. 44:905-909.

Pitta, D. W., W. E. Pinchak, S. E. Dowd, J. Oster-

stock, V. Gontcharova, E. S. Youn, K. Dorton,

I. Yoon, B. R. Min, and J. D. Fulford. 2010. Ru-

men bacterial diversity dynamics associated with

changing from bermudagrass hay to grazed win-

ter wheat diets. Microb. Ecol. 59:511–522.

Pryde, S. E., S. H. Duncan, G. L. Hold, C. S. Stewart,

and H. J. Flint. 2002. The microbiology of butyr-

ate formation in the human colon. FEMS Micro-

biol. Lett. 217:133–139.

Rakoff-Nahoum, S, J. Paglino, F. Eslami-Varzaneh,

S. Edberg, and R. Medzhitov. 2004. Recognition

of commensal microflora by toll-like receptors is

required for intestinal homeostasis. Cell. 118:229-

241.

Schloss, P. D., S. L. Westcott, T. Ryabin, J. R. Hall,

M. Hartmann, E. B. Hollister, R. A. Lesniewski, B.

B. Oakley, D. H. Parks, C. J. Robinson, and C. F.

Weber. 2009. Introducing mothur: open-source,

platform-independent, community-supported

software for describing and comparing microbial

communities. Appl. Environ. Microbiol. 75:7537–

7541.

Smith, A. H., J. A. Imlay, and R. I. Mackie. 2003.

Increasing the oxidative stress response allows

Escherichia coli to overcome inhibitory effects

of condensed tannins. Appl. Environ. Microbiol.

69:3406–3411.

Tabacco, E., G. Borreani, G. M. Crovetto, G. Galas-

si, D. Colombo, and L. Cavallarin. 2006. Effect of

chestnut tannin on fermentation quality, prote-

olysis, and protein rumen degradability of alfalfa

silage. J. Dairy Sci. 89: 4736–46.

Terrill, T. H., G. C. Waghorn, D. J. Woolley, W. C.

McNabb, and T. N. Barry. 1994. Assay and di-

gestion of 14C-labelled condensed tannins in

the gastrointestinal tract of sheep. Brit. J. Nutr.

72:467–477.

Turnbaugh, P. J., R. E. Ley, M. A. Mahowald, V. Magri-

ni, E. R. Mardis, and J. I. Gordon. 2006. An obe-

sity-associated gut microbiome with increased

capacity for energy harvest. Nat. 444:1027-1031.

Williams, A. G. and G. S. Coleman. 1988. Rumen

ciliate protozoa. Pages 77–128 in The Rumen

Microbial Ecosystem. P. N. Hobson, ed. Elsevier,

Amsterdam, the Netherlands.

Williams, C. M., J. S. Eun, C. M. Dschaak, J. W. Mac-

Adam, B. R. Min, and A. J. Young. 2010. In vitro

ruminal fermentation characteristics of birdsfoot

trefoil (Lotus corniculatus L.) hay in continuous

cultures. The Prof. Anim. Scientist 26:570-576.

Wilson, I. D. and J. K. Nicholson. 2009. The role of

gut microbiota in drug response. Curr. Pharm.

Des. 15:1519-1523.

Wu, G. D., C. J, Hoffmann, K. Bittinger, Y. Y. Chen,

S. A. Keilbaugh, M. Bewtra, D. Knights, W. A. Wal-

ters, R. Knight, R. Sinha, E. Gilroy, K. Gupta, R. Bal-

dassano, L. Nessel, H. Li, F. D. Bushman, and J.

D. Lewis. 2011. Linking long-term dietary patterns

with gut microbial enterotypes. Sci. 334:105–108.


Xu, J., M. K. Bjursell, J. Himrod, S. Deng, L. K.

Carmichael, H. C. Chiang, L. V. Hooper, and J. I.

Gordon. 2003. A genomic view of the human-

Bacteroides thetaiotaomicron symbiosis. Sci.

299:2074–2076.

Zimmer, N. and R. Cordesse. 1996. Digestibility

and ruminal digestion of non-nitrogenous com-

pounds in adult sheep and goats: Effects of chest-

nut tannins. Anim. Feed Sci. Technol. 61:259-273.




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 obser-

vations 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 oc-

curred 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 provid-

ing energy required for biotechnological applications.

Keywords: Enterobacter cloacae, triacylglyceride, lipids, extracellular lipids, electron microscopy,

oleaginous, biofuels, biodiesel, JD6301, membrane transport

Correspondence: J. R. Donaldson, [email protected], Tel: +1-662-325-9547

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

1Department of Biological Sciences, Mississippi State University, Mississippi State, MS, USA. 2Department of Chemical Engineering, University of Louisiana Lafayette, Lafayette, LA, USA. 3Renewable Fuels and Chemicals Laboratory, Dave C. Swalm School of Chemical Engineering,

Mississippi State University, Mississippi State, MS, USA.4Institute for Imaging and Analytical Technologies, Mississippi State University, Mississippi State, MS, USA.

5 DOE Joint Genome Institute, Walnut Creek, CA, USA.



INTRODUCTION

Lipids are important energy sources in both hu-

man health and also in biofuel production, such as

biodiesel (Ranganathan et al., 2008; Rosen and Spie-

gelman, 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 (Le-

stari et al., 2009). While plant and animal fat stores

are the most common source of TAGs for biofuel

production, certain prokaryotes have been identi-

fied to accumulate TAGs as a form of energy storage

(Alvarez and Steinbuchel, 2002; Holder et al., 2011).

These microorganisms, termed oleaginous microor-

ganisms, accumulate more than 20% of their biomass

as TAGs and include the bacteria Streptomyces, No-

cardia, 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 contain-

ing 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. En-

terobacter 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 con-

structed 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 oleagi-

nous microorganisms to produce extracellular lipids

could lead to advancements in lipid biotechnology,

especially in the areas of lipid recovery and utiliza-

tion.


Culture conditions

Frozen stocks of the Enterobacter cloacae

wild type (WT) strain JD6301 and resulting isogen-

ic 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) supplement-

ed 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

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.

Mutant construction

A mutant of JD6301 capable of producing extracel-

lular lipids was constructed as previously described

for S. cerevisiae with minor modifications (Nojima et

al., 1999). A 24 h culture of JD6301 in MSM supple-

mented 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 (Sig-

ma, L9656). Cultures were incubated for an addition-

al 16 h at 30°C to allow for growth of the auxotrophic

KD115 strain. The presence of microcolony growth

of KD115 around the periphery of the Enterobacter

cloacae cells indicated the EMS introduced genetic

alterations that promoted the presence of extracel-

lular lipids, which were then available for utilization

by KD115. The potential mutants were transferred to


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 colo-

nies 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 previous-

ly described (Wilson et al., 2014), which generated

178,495,750 reads. The final draft assembly con-

tained 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 nucle-

otide 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 synony-

mous, and 3 non-coding variants (Table 1).

Growth responses and sugar consump-tion

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 glu-

cose, 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 Spec-

traMax Plus 384 plate reader at 30°C with intermit-

tent shaking.

For glucose consumption analysis, 1 mL of cells

was filtered through a 0.2 μm filter (Corning Life Sci-

ences, Amsterdam, The Netherlands). Glucose con-

centrations 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 aver-

age 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 eth-

anol series, and embedded in Spurr’s resin. Ultra-

thin sections were cut with a Reichert-Jung Ultracut

3 ultra-microtome and were stained with uranyl ac-

etate 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 ana-

lyzed 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


were fixed in 2.5% glutaraldehyde in 0.1 M cacodyl-

ate buffer, washed in 0.1 M cacodylate buffer, post-

fixed in 1% osmium tetraoxide in 0.1 M cacodylate

buffer, rewashed in distilled water, dehydrated in an

ethanol series, and finally dried in a hexamethyldisi-

lazane series as previously described (Merritt et al.,

2010). Samples were sputter coated with gold-pal-

ladium 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 1x107 cells) of JD6301 and

JD8715 was collected at 24 h of growth and viabil-

ity was determined using the LIVE/DEAD BacLightTM

bacterial viability kit consisting of SYTO 9 and prop-

idium iodide (Invitrogen, Calsbad, CA). Staining

was performed in the dark for 15 min, after which

cells were analyzed by a BD FASCaliber flow cytom-

eter using Cell Quest software (BD Biosciences, San

Jose, CA). Analyses were performed on 4,500 to

5,000 cells (gated events) using instrument param-

eters previously described by others (Gunasekera et

al., 2003). Percent viability was determined by a two-

parameter comparison of green (live cells) and red

(dead cells) fluorescent emission for individual bac-

teria using the formula: [% live green-emitting cells/

(% dead red-emitting cells + % live green-emitting

cells)] x 100. Three independent replicates were per-

formed for each strain.

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-

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 super-

natant collected. Lipids were extracted from the col-

lected supernatant and cell pellet using a standard

Bligh and Dyer lipid extraction technique (Bligh and

Dyer, 1959).

Extracted lipids were derivatized using N-Methyl-

N-(trimethylsilyl)-trifluoroacetamide (MSTFA) follow-

ing ASTM D6584, which uses tricaprin as an inter-

nal 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 al-

lowed to react for at least 20 min, after which 900

μL of n-heptane was added. Samples were then fil-

tered through a 0.45 μm PTFE filter (SUN Sri, Rock-

wood, 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 fi-

nally 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

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-

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 hy-

droxytoluene and 200 μg/mL 1,3-dichlorobenzene.


Table 1. Summary of SNPs identified.

SNP Ref-Mut

Scaffold/Pos Gene Function

Non-synonymous

G-A 2/8877 P698DRAFT_00653ABC-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_02476Phosphotransferase system IIC components, glu-cose/maltose/N-acetylglucosamine-specific

G-A 6/193118 P698DRAFT_02492 2-methylthioadenine synthetase

C-T 6/233659 P698DRAFT_02537Ribulose-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_03706SAM-dependent methyltransferases related to tRNA (uracil-5-)-methyltransferase

C-T 14/8347 P698DRAFT_03815 Protein of unknown function (DUF968)

G-A 18/63736 P698DRAFT_041987,8-dihydro-6-hydroxymethylpterin-pyrophosphoki-nase

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

C-T 2/160082 - -

C-A 4/240806 - -

C-T 7/209964 - -


The FAMEs were analyzed using an Agilent 6890N

gas chromatograph equipped with a flame ioniza-

tion detector (GC-FID) and a fused silica column Sta-

bilwax-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 cali-

bration was achieved using a 14-component FAMEs

standard mixture (C8 – C24) (Supelco, Bellefonte, PA)

containing saturated, mono-unsaturated and poly-

unsaturated 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

to analyze the significance of total biodiesel lipid

analysis.


Lipid production by Enterobacter cloa-cae JD6301 and construction of the mu-tant JD8715

A novel strain of Enterobacter cloacae previously

isolated by our group (Wilson et al., 2014) was ana-

lyzed by transmission electron microscopy, where in-

clusion 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 en-

tire 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; Al-

varez and Steinbuchel, 2002; Waltermann et al., 2005;

Waltermann and Steinbuchel, 2005). Lipids were iso-

lated 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

oleaginous isolate of Enterobacter cloacae.

A limitation to the usefulness of oleaginous mi-

croorganisms 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-

ity of lipids in water (Fischer et al., 2008). Therefore,

this bacterium was modified to produce extracellu-

lar lipids. This strain was treated with the carcinogen

EMS as previously described (Nojima et al., 1999)

and the mutant JD8715 was subsequently identified

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


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 con-

tigs 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 compo-

nents, transporter proteins, and a hook associated

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 micro-

scopic 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 chro-

matography after 24 h of growth. Mass of the re-

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

Strain%Glucose Consumed (±StDev)

Cells: glucose consumed

Yield extracellular lipid: glucose

Yield cell mass: 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) JD8715 (μg/ml, ±StDEV)

Lipid 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)

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

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

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


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 let-ter are not significantly different (p <0.05).

A

B


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

B


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 extracellu-

lar lipids (0.353 g lipid/ g sugar for JD8715 versus

0.098 g lipid/ g sugar for JD6301 WT; Table 2), sug-

gesting 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

extracellular lipids compared to WT.

Lipid analyses indicated that the extracellular lip-

ids present in the JD8715 culture consisted of mono-

glycerides, 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 com-

position from JD8715 to JD6301 indicated that the

mutant had significantly less amounts of intracellular

monoglycerides and triacylglycerides. No signifi-

cant 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 frac-

tions 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 intracellu-

lar yield of WT (Fig. 2A). The extracellular yield from

JD8715 was significantly greater than the extracel-

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 consisten-

cy between strains in regard to lipids produced (Fig.

2B). A large amount of total unknown FAMEs were

detected for JD8715 and WT, which could include

odd-numbered fatty acids.

Growth and viability of JD8715 and WT JD6301

To determine if the presence of extracellular lip-

ids was due to the JD8715 cultures containing more

lysed or dead cells as compared to JD6301, the per-

centages of live cells for both the mutant and WT

were determined using the LIVE/DEAD BacLightTM

bacterial viability kit (Invitrogen). These results indi-

cated that the JD8715 and JD6301 strains exhibited

similar viability at 24 h (77% versus 78%). The growth

of the JD8715 strain was compared to WT to deter-

mine 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 ex-

traction procedure, cells were washed with chloro-

form 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 fol-

lowing the chloroform wash (Fig. 3B), indicating that

the chloroform wash that precedes the lipid extrac-

tion procedure did not unintentionally lyse the cells,

therefore skewing the extracellular fraction.

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,

this is the first report of a genetically altered form of

an oleaginous bacterium that is capable of produc-

ing extracellular lipids. The formation of lipid inclu-

sion bodies initiates at the cell membrane (Walter-

mann et al., 2005) proposing several potential areas


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 oleag-

inous microorganisms for optimization of extracellu-

lar lipid production, providing a substantial advance-

ment 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 con-

ducted by the DOE Joint Genome Institute is sup-

ported by the Office of Science of the DOE under

contract number DE-AC02-05CH11231.

REFERENCES

Alvarez, H.M., F. Mayer, D. Fabritius, and A. Stein-

buchel. 1996. Formation of intracytoplasmic lipid

inclusions by Rhodococcus opacus strain PD630.

Arch. Microbiol. 165: 377-86.

Alvarez, H.M. and A. Steinbuchel. 2002. Triacylglyc-

erols in prokaryotic microorganisms. Appl. Micro-

biol. Biotechnol. 60: 367-76.

Bligh, E.G. and W.J. Dyer. 1959. A rapid method of

total lipid extraction and purification. Can. J. Bio-

chem. Physiol. 37: 911-917.

Fischer, C.R., D. Klein-Marcuschamer, and G. Stepha-

nopoulos. 2008. Selection and optimization of mi-

crobial hosts for biofuels production. Metab. Eng.

10: 295-304.

Grima, E.M., E.H. Belarbi, F.G. Fernandez, A.R. Me-

dina, and Y. Chisti. 2003. Recovery of microalgal

biomass and metabolites: process options and

economics. Biotechnol. Adv. 20: 491-515.

Gunasekera, T.S., D.A. Veal, and P.V. Attfield. 2003.

Potential for broad applications of flow cytometry

and fluorescence techniques in microbiological

and somatic cell analyses of milk. Int. J. Food Mi-

crobiol. 85: 269-79.

Holder, J.W., J.C. Ulrich, A.C. DeBono, P.A. Godfrey,

C.A. Desjardins, J. Zucker, Q. Zeng, A.L. Leach,

I. Ghiviriga, C. Dancel, T. Abeel, D. Gevers, C.D.

Kodira, B. Desany, J.P. Affourtit, B.W. Birren, and

A.J. Sinskey. 2011. Comparative and functional ge-

nomics of Rhodococcus opacus PD630 for biofuels

development. PLoS Genet. 7: e1002219.

Ingle, C.A. and N.R. Drinkwater. 1989. Mutational

specificities of 1’-acetoxysafrole, N-benzoyloxy-N-

methyl-4-aminoazobenzene, and ethyl methane-

sulfonate in human cells. Mutat. Res. 220: 133-142.

Lestari, S., P. Maki-Arvela, J. Beltramini, G.Q. Lu, and

D.Y. Murzin. 2009. Transforming triglycerides and fat-

ty acids into biofuels. ChemSusChem. 2: 1109-1119.

Li, H. and R. Durbin. 2009. Fast and accurate short

read alignment with Burrows-Wheeler transform.

Bioinformatics. 25: 1754-1760.

Li, H., B. Handsaker, A. Wysoker, T. Fennell, J. Ruan,

N. Homer, G. Marth, G. Abecasis, R. Durbin, and

Genome Project Data Processing Subgroup. 2009.

The Sequence Alignment/Map format and SAM-

tools. Bioinformatics. 25: 2078-2079.

Meng, X., J. Yang, X. Xu, L. Zhang, Q. Nie, and M.

Xian. 2009. Biodiesel production from oleaginous

microorganisms. Renewable Energy. 34: 1-5.

Merritt, M.E., A.M. Lawrence, and J.R. Donaldson.

2010. Comparative study of the effect of bile on the

Listeria monocytogenes virulent strain EGD-e and

avirulent strain HCC23. Arch. Clinical Micro. 1: 4-9.

Nojima, Y., A. Kibayashi, H. Matsuzaki, T. Hatano,

and S. Fukui. 1999. Isolation and characterization

of triacylglycerol-secreting mutant strain from

yeast, Saccharomyces cerevisiae. J. Gen. Appl. Mi-

crobiol. 45: 1-6.

Ranganathan, S.V., S.L. Narasimhan, and K. Muthu-

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 ho-

meostasis. Nature. 444: 847-853.


Schlegel, H.G., H. Kaltwasser, and G. Gottschalk.

1961. Ein Submersverfahren zur Kultur wasserst-

offoxydierender Bakterien: Wachstumsphysiolo-

gische Untersuchungen. Arch. Microbiol. 38: 209-

222.

Stukey, J.E., V.M. McDonough, and C.E. Martin. 1989.

Isolation and characterization of OLE1, a gene af-

fecting fatty acid desaturation from Saccharomy-

ces 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 pro-

karyotes: 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 microor-

ganisms. In Bailey’s Industrial Oil and Far Products.

Shahidi F., ed. (John Wiley and Sons, Inc.).




ABSTRACT

Aerobic composting of animal manures has been advocated as an effective management tool to inac-

tivate resident zoonotic pathogens where the time at lethal temperatures is used to determine the effec-

tiveness 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 Sal-

monella 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, persis-

tence 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 mix-

tures 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

Correspondence: M.C. Erickson, [email protected]: +1 770-412-4742 Fax: +1 770-229-3216

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



INTRODUCTION

Livestock and poultry production are major enter-

prises worldwide that in addition to the production

of food, waste by-products that include solid ma-

nure and manure slurries are also produced in large

quantities. For example, the Environmental Protec-

tion 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 pro-

portion (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-

land (15.8 million acres) in 2006 reported as having

been fertilized with livestock manure (MacDonald et

al., 2009). Zoonotic pathogens are sporadically resi-

dent 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 con-

taminate the soil, crops grown in those fields, and

waterways that collect runoff from the fields.

Aerobic composting of animal wastes can inac-

tivate zoonotic bacterial pathogens while creating

a stable amendment that improves soil quality and

fertility (Berry et al., 2013; Raviv, 2005). Heat gener-

ated from the metabolic activity of thermophilic mi-

croorganisms 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 com-

posting 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 ma-

terial must be turned a minimum of 5 times to en-

sure 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-

perature exceeds 55°C for 4 h, has also been desig-

nated 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

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 contribu-

tion of other physical factors (e.g., non-lethal tem-

peratures, light, and desiccation) to inactivation of

zoonotic pathogens in manure-based compost mix-

tures has not been elucidated because biological

(i.e., competition for nutrients) and chemical (e.g.,

ammonia, volatile acids or other antimicrobials) fac-

tors that affect pathogen inactivation are also likely

affected by the physical parameter. Such is the case

with soil systems in which increased temperatures,

despite being near the organism’s optimal growth

temperature, led to greater inactivation of Esche-

richia coli O157:H7 as a result of an accompanying

increase in competition by the dominant native mi-

crobial 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 patho-

gen 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 dur-

ing 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 in-

digenous microbial communities, it is of interest to

investigate the role of moisture levels on inactivation

of pathogens in compost mixtures that would likely

be populated with different indigenous microflora

from the different nitrogen feedstocks.

Another physical factor that has received little at-

tention for its involvement in inactivating pathogens

in manure-based compost mixtures is sunlight. Due


to its inability to penetrate compost mixtures, sun-

light 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 contami-

nation 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, Erick-

son 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

for 14 and 56 days in the summer and winter, respec-

tively. 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 un-

der 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 recom-

mended holding times for regions with a large num-

ber of overcast days compared to regions that are

dominated by sunny days.

The purpose of this study was to determine the in-

fluence 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 environmen-

tal 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 ma-

nure in compost mixture formulations from different

sources (dairy, chicken, and swine) that should have

different microbial compositions.


Pathogen Strains and Preparation

Three strains of green-fluorescent protein (GFP)-

labeled Salmonella enterica serovar Enteritidis (ME-

18, H4639, and H3353) and one strain of GFP-labeled

S.enterica serovar Newport containing an ampicillin-

resistant marker were selected from the culture col-

lection 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

the construction of these GFP strains has been de-

scribed 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. monocyto-

genes strains, respectively.

Frozen stock cultures of each GFP-labeled Sal-

monella 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. Individ-

ual Salmonella and L. monocytogenes colonies from

these plates were then inoculated into 100 ml of tryp-

tic 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 erythro-

mycin (BHIB-E), respectively. Broths were incubated

at 37°C for 20 to 24 h with agitation (150 rpm) and

bacteria were subsequently harvested by centrifuga-

tion (4,050 x g, 15 min, 4°C) with cell pellets being

washed three times in 0.1% peptone water (Difco,

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-


strain mixture of L. monocytogenes. Salmonella and

L. monocytogenes populations were determined by

plating on TSA-A and modified Oxford agar (Acu-

media Manufacturers, Lansing, MI) containing 8 μg/

ml erythromycin (MOX-E), respectively. Salmonella

transformed colonies emitted bright green fluores-

cence when viewed at 365 nm under a handheld UV

light (Fotodyne Inc., Hartland, WI); however, visual-

ization of fluorescent L. monocytogenes transformed

colonies required use of a Leica MZ16 FA stereo fluo-

rescence microscope (Bannockburn, IL).

Compost Feedstocks and Chemical Analysis

Three sources of manure including dairy cow ma-

nure, 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 labo-

ratory were frozen for at least 24 h to kill the majority

of insect eggs (Sherman et al., 2006). Carbon amend-

ments in compost formulations (i.e., wheat straw and

cottonseed meal) were purchased from a local feed

supply store. To improve the homogeneity of com-

post formulations, wheat straw was shredded using a

Flowtron Leaf Eater (Malden, MA) for lengths of 1 to

5 cm. Carbon, nitrogen, and moisture content analy-

ses 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

Each type of manure was individually sprayed in

a 28-L sanitized bowl with either GFP-labeled Sal-

monella or both GFP-labeled-Salmonella and L.

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

(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 exper-

imental 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

chicken litter) that were each obtained at two sepa-

rate times. Dairy cow manure (2 kg) or chicken litter

(1 kg) was sprayed with the Salmonella inoculum mix-

ture (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 con-

tainers were held at 25°C and another three were

held at 40°C. With the second batch of dairy cow ma-

nure, 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 contain-

ers 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 con-

ducted.

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 col-

lection being used as an independent replicate trial.

Each batch of chicken litter was sprayed with both an

inoculum of Salmonella and an inoculum of L. mono-

cytogenes before blending with wheat straw, cotton-

seed meal and sterile deionized water to obtain mix-

tures having an initial carbon:nitrogen (C:N) ratio of

40:1 and 60% moisture content Initial pathogen pop-


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). Du-

plicate samples from each inoculated mixture were

obtained for pathogen and moisture content analy-

sis 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 cy-

cle 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

Electric, Cleveland, OH) and ten 400W high pressure

sodium lamps LU400/H/ECO, LUCALOS bulbs (Gen-

eral 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 inocu-

lum of Salmonella and an inoculum of L. monocyto-

genes and, then mixed with wheat straw, cottonseed

meal, and sterile deionized water to obtain cow and

chicken compost mixtures having an initial C:N ra-

tio 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 un-

covered in an environmental chamber at 30°C with a

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

back to their initial weights and original moisture

contents by spraying the sample with a light mist

of sterile deionized water. Two cups whose mois-

ture 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

pathogen inocula.

For the fourth experimental study, three different

sources of manure (dairy cow manure, chicken litter,

and swine manure) were inoculated and incorpo-

rated into compost mixtures to examine the effect

of light on survival of Salmonella and L. monocyto-

genes. Wheat straw and cottonseed meal served as

the carbon amendments and were mixed with the

Salmonella- and L. monocytogenes-inoculated ma-

nure 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 en-

vironmental 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

chamber where only half the bulbs were turned on

to simulate daily “cloudy” conditions (12 h at 289-

359 μmol/m2/sec and 12 h in the dark). All chambers

were at 20°C. Duplicate samples from each treat-

ment at five sample times over the course of 4 weeks,


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 in-

dependent replicate trials were conducted with each

trial using manure collected at separate times and a

different batch of pathogen inoculum cultured for in-

oculation of the manure.

Microbiological and pH Analyses

Both direct plating and selective enrichment cul-

ture was used for detection of GFP-labeled Salmo-

nella and L. monocytogenes. In direct plating as-

says, 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 GFP-

labeled Salmonella or L. monocytogenes, respec-

tively. 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 ther-

mophilic 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, respec-

tively.

Measurement of pH was determined with an Acu-

met Basic pH meter (Fisher Scientific, Pittsburgh, PA)

on 5-g compost samples dispersed in 250 ml of de-

ionized 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 analy-

sis of variance (ANOVA) test using the StatGraphics

Centurion XV software package (StatPoint, Inc., Hern-

don, 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).


Several studies have previously addressed the

survival of zoonotic pathogens in manure at ambi-

ent temperatures (Himathongkham et al., 1999a, b;

2000; Sinton et al., 2007); however, this type of study

was repeated in our preliminary study (first experi-

mental study) with locally-obtained manure to give

some baseline information on the fate of Salmonel-

la and other indigenous microflora in the absence

of a carbon amendment. Different responses were

observed for Salmonella and the indigenous micro-

flora depending on the manure source and storage

temperatures. Following a 3-day storage period,

no changes in populations of the indigenous mi-

croflora (mesophilic and thermophilic bacteria) oc-

curred 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 pe-

riod. Salmonella decreases occurred in both manure

sources when held at 40°C, but reductions were sub-

stantially 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


manure held at ambient temperatures (Himathong-

kham 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 (Himathong-

kham and Riemann, 1999). For the current study, it

is plausible that the production of ammonia by the

indigenous microflora could be stimulated by the

presence of bedding material that was included dur-

ing 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-

ments, wheat straw and cottonseed meal, to create

compost mixtures prior to their storage. When com-

post mixtures were formulated to an initial moisture

content of 60% and an initial C:N ratio of 40:1, stor-

age for 2 weeks at 20°C in the dark led to L. mono-

cytogenes 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-

der lighted conditions, however, resulted in popu-

lation 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

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 Cow manure3

Day 25°C 40°C 25°C 35°C 40°C

Mesophilic bacteria populations (log CFU/g)5

0 ND4 9.38 ± 0.13 a 9.22 ± 0.10 a 6.88 ± 0.41 b 6.88 ± 0.06 b

3 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

0 ND 7.08 ± 0.02 a 8.36 ± 0.08 a 6.35 ± 0.01 b 6.35 ± 0.01 b

3 8.32 ± 0.25 5.86 ± 0.06 b 8.54 ± 0.17 a 8.46 ± 0.38 a 8.19 ± 0.83 a

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

↓7 1.21 ↓ 5.56 ↑8 1.88 ↑ 1.11 ↓ 0.89

1 Data collected from first experimental study.2 Mesophilic and thermophilic bacteria population levels are mean ± S.D., n = 3.3 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.4 Not determined.5 Values for this parameter within each column followed by a different letter are significantly different (P <

0.05). 6 Salmonella initial populations in manure samples ranged from 5 to 6 log CFU/g.7 Decrease in population.8 Increase in population.


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 investiga-

tors suggested that increases in temperature, de-

spite 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

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 in-

terest in our third experimental study in determining

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 ad-

ditions 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 con-

tainers and equilibrated to approximately the same

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 condi-

tions were therefore likely responsible for the fail-

ure of moisture content to have an effect on patho-

Table 2. Comparison of Salmonella and L. monocytogenes losses in chicken and cow manure-based 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

1 2.09 ± 0.67 a 0.59 ± 1.27 b 1.36 ± 1.62 a -0.35 ± 1.73 a-c

2 1.87 ± 1.22 a 0.60 ± 1.25 b 0.89 ± 1.54 ab -0.82 ± 1.44 bc

3 2.11 ± 0.93 a 1.15 ± 1.22 ab 1.11 ± 1.50 ab -1.31 ± 2.10 c

4 2.11 ± 0.93 a 2.21 ± 0.82 a 0.90 ± 1.64 ab -0.58 ± 1.56 bc

1 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.

2 Data collected from third experimental study.3 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.

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

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

6 Values for this pathogen (mean ± S.D.), across both rows and columns, 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 mix-

tures (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 stor-

age at 30°C and additional significant decreases oc-

curred up to 4 weeks of storage (P < 0.05). Over all

time periods, there were significantly greater reduc-

tions in Salmonella and L. monocytogenes popula-

tions in the chicken compost mixtures compared to

the cow compost mixtures (P < 0.05). Interestingly,

the decreases in pathogen populations in the chick-

en compost mixtures occurred during the first week

of storage but not after additional storage (Table 2).

Monitoring the pH of the chicken- and cow ma-

nure-based compost mixtures during storage at

30°C for 4 weeks (third experimental study) revealed

that initially the chicken compost mixtures were ap-

proximately 0.5 pH units higher than the cow com-

post 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 mix-

tures. 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

of 60%, the pH increased to values approximating

those detected in the chicken compost mixtures (Ta-

ble 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

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

0 7.25 ± 0.66 a-f 7.58 ± 1.22 a-d 6.84 ± 0.33 d-f 6.64 ± 0.34 f

1 7.06 ± 0.32 b-f 7.85 ± 0.66 a 6.80 ± 0.29 ef 7.46 ± 0.21 a-e

2 6.97 ± 0.34 c-f 7.63 ± 0.59 a-c 6.77 ± 0.22 ef 7.09 ± 0.10 b-f

3 6.99 ± 0.42 c-f 7.75 ± 0.34 ab 6.81 ± 0.16 ef 7.32 ± 0.29 a-f

4 6.87 ± 0.41 d-f 7.74 ± 0.24 ab 6.77 ± 0.08 ef 7.45 ± 0.22 a-e

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

2 Data collected from third experimental study.3 Values within the table (mean ± S.D.) followed by a different letter are significantly different (P < 0.05).


Tab

le 4

. Sal

mo

nella

and

L. m

ono

cyto

gen

es r

educ

tio

ns d

urin

g s

tora

ge

at 2

0°C

und

er d

iffer

ent

light

co

ndit

ions

in c

om

po

st m

ixtu

res

pre

par

ed w

ith

diff

eren

t m

anur

e so

urce

s1 .

Cum

ulat

ive

pat

hog

en re

duc

tions

(lo

g C

FU/g

) 2

L. m

ono

cyto

gen

es3

Salm

one

lla3

Man

ure

sour

ceW

eek

Dar

kC

loud

y4Su

nny5

Dar

kC

loud

ySu

nny

Chi

cken

20.

86 ±

0.0

1 e

1.37

± 0

.74

e1.

37 ±

0.6

1 e

0.32

± 0

.65

e0.

48 ±

0.2

6 e

1.01

± 0

.76

de

62.

70 ±

0.2

7 d

4.71

± 1

.01

c5.

83 ±

0.3

0 ab

1.10

± 1

.02

c-e

1.39

± 0

.63

b-e

3.07

± 1

.44

ab

84.

38 ±

1.0

8 c

5.83

± 0

.30

ab5.

83 ±

0.3

0 ab

1.07

± 0

.56

de

2.09

± 0

.38

a-e

3.41

± 1

.14

a

105.

01 ±

1.1

2 b

c5.

83 ±

0.3

0 ab

6.03

± 0

.13

a1.

89 ±

0.8

1 a-

e2.

11 ±

0.7

9 a-

e2.

90 ±

1.9

1 a-

c

125.

93 ±

0.2

4 ab

5.83

± 0

.30

ab5.

93 ±

0.2

8 ab

3.14

± 1

.92

ab2.

45 ±

0.7

2 a-

d2.

99 ±

1.5

3 ab

Dai

ry c

ow

20.

36 ±

0.8

1 d

1.07

± 1

.85

d2.

03 ±

2.1

2 cd

-0.3

1 ±

1.0

8 a

0.23

± 2

.04

a1.

08 ±

3.4

3 a

61.

04 ±

0.8

1 d

2.39

± 2

.90

cd2.

43 ±

1.6

1 cd

0.44

± 1

.75

a0.

58 ±

2.4

0 a

1.01

± 2

.45

a

101.

28 ±

0.2

6 d

2.87

± 2

.50

b-d

2.90

± 2

.73

b-d

0.71

± 0

.89

a1.

63 ±

2.9

4 a

1.75

± 2

.84

a

142.

25 ±

0.3

9 cd

4.09

± 1

.46

a-c

5.29

± 0

.70

ab0.

93 ±

1.9

6 a

1.78

± 2

.81

a1.

67 ±

2.0

6 a

184.

70 ±

1.7

1 a-

c4.

29 ±

1.2

7 a-

c6.

27 ±

1.0

2 a

1.61

± 2

.96

a1.

90 ±

2.7

2 a

2.09

± 2

.56

a

Swin

e0.

3-0

.57

± 0

.45

de

-0.6

4 ±

0.5

6 e

-0.0

8 ±

0.6

2 c-

e-0

.82

± 0

.37

f-0

.75

± 0

.52

ef-0

.25

± 0

.51

c-f

1-0

.26

± 0

.48

c-e

-0.2

7 ±

0.5

5 c-

e-0

.01

± 0

.46

c-e

-0.4

4 ±

0.5

3 d

-f-0

.43

± 1

.02

d-f

-0.1

1 ±

0.7

5 b

-f

20.

07 ±

0.4

1 c-

e0.

14 ±

0.5

6 b

-d0.

44 ±

0.1

1 b

c-0

.36

± 0

.69

c-f

0.02

± 0

.53

a-f

0.16

± 0

.36

a-e

30.

10 ±

0.2

5 b

-e0.

22 ±

0.2

4 b

c0.

86 ±

0.4

5 ab

0.13

± 0

.46

a-f

0.36

± 0

.62

a-d

0.74

± 0

.48

ab

40.

39 ±

0.1

7 b

c0.

39 ±

0.3

6 b

c1.

38 ±

0.6

6 a

0.42

± 0

.41

a-d

0.56

± 0

.68

a-c

0.87

± 0

.38

a

1 D

ata

wer

e co

llect

ed fr

om

four

th e

xper

imen

tal s

tud

y.

2 R

educ

tions

wer

e d

eter

min

ed re

lativ

e to

initi

al v

alue

s in

co

mp

ost

mix

ture

s.3

Valu

es (m

ean

± S

.D.)

with

in e

ach

man

ure

sour

ce fo

r th

is p

atho

gen

follo

wed

by

a d

iffer

ent

lett

er a

re s

igni

fican

tly d

iffer

ent

(P <

0.0

5).

4 C

om

po

st m

ixtu

res

wer

e ex

po

sed

dai

ly t

o li

ght

co

nditi

ons

of 2

89 t

o 3

59 μ

mo

l/m

2 /se

c fo

r 12

h a

nd t

o d

ark

cond

itio

ns fo

r 12

h.

5 C

om

po

st m

ixtu

res

wer

e ex

po

sed

dai

ly t

o li

ght

co

nditi

ons

of 5

24 t

o 5

73 μ

mo

l/m

2 /se

c fo

r 12

h a

nd t

o d

ark

cond

itio

ns fo

r 12

h.


compost mixtures initially but the low-moisture con-

ditions present in these mixtures inhibit the indig-

enous microflora from generating additional ammo-

nia. The low moisture conditions, however, do not

directly contribute to inactivation of desiccation-re-

sistant 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 pre-

pared with chicken litter, dairy cow manure or swine

manure and held at 20°C were exposed to one of

three lighting conditions simulating dark, sunny, or

cloudy conditions. Preliminary studies were con-

ducted with each of these compost mixtures to de-

termine the approximate time interval over which

samples should be taken to obtain measurable

pathogen reductions and determine whether light

conditions could significantly affect their inactiva-

tion. Unfortunately, the storage time intervals select-

ed 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 stor-

age, 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 dur-

ing 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-

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-

mal reductions that occurred during the short time

period that was examined for swine compost mix-

tures. In contrast, over all time periods, reductions in

L. monocytogenes populations were statistically sig-

nificant for both cloudy and sunny conditions com-

pared to dark conditions for chicken compost mix-

tures 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 re-

ductions in populations than dark conditions, and

these occurred midway through the storage trial. In

contrast but similar to the response in swine com-

post mixtures, light conditions did not affect the re-

ductions in Salmonella populations in cow compost

mixtures at any sampling time (Table 4).

A number of factors could contribute to light-

mediated 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 microor-

ganisms (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 inactiva-

tion of the Gram-positive, Enterococcus faecalis in

waste stabilization pond water whereas the Gram-

negative E. coli was inherently less susceptible to

this pathway (Kadir and Nelson, 2014). Although L.

monocytogenes has previously displayed some des-

iccation resistance, surviving for three months in a

simulated food processing environment (Vogel et

al., 2010), it is not as resistant as Salmonella based

on the lower recoveries of L. monocytogenes com-

pared 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.


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 com-

post mixtures (Table 5). In addition to affecting the

moisture content of the compost mixtures, light ex-

posure led to decreased pH in the chicken and cow

compost mixtures (Table 5). L. monocytogenes sur-

vival could have been improved under less alkaline

conditions; however, that response was likely to be

minimal in these compost mixtures due to the con-

current stress imposed by low moisture contents and

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

a state could also have been responsible for our in-

ability to discern an effect of light on inactivation of

Salmonella in the swine or cow compost mixtures.

In summary, both Salmonella and L. monocyto-

genes may survive in compost mixtures that are ex-

posed to sublethal temperatures for extended pe-

riods 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. At-

tempts 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. monocyto-

genes appeared to be more susceptible to desicca-

tion stress than Salmonella based on their relative re-

duction in populations in chicken and cow compost

mixtures over time. L. monocytogenes populations

also decreased more rapidly when compost mix-

tures were exposed to light conditions, described as

sunny or cloudy, compared to dark conditions, but

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 pH3

Manure source 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 ± 0.5 a 9.0 ± 0.4 a

1 Data were collected from fourth experimental study.2 Swine, chicken, and dairy cow compost mixtures were stored for 4, 12, and 18 weeks, respectively.3 Values in each row of an attribute 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.


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

The project was supported by the National Re-

search 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 tech-

nical assistance.

REFERENCES

Beck-Friis, B., S. Smårs, H. Jönsson, Y. Eklind, and H.

Kirchmann. 2003. Composting of source-separat-

ed household organics at different oxygen levels:

Gaining an understanding of the emission dynam-

ics. 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 occur-

ring 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 sub-

stances derived from swine manure-based com-

post and correlation of their characteristics with re-

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.

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 Tech-

nol. 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. Destruc-

tion of Salmonella typhimurium, Escherichia coli

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. Rie-

mann. 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. Nu-

anualsuwan, 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 Min-

istry 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.

Moore. 2005. Decline of zoonotic agents in live-

stock waste and bedding heaps. J. Appl. Micro-

biol. 99:354-362.

Jagger, J. 1985. Solar-UV action on living cells. Prae-

ger, New York.


Jiang, Y., M. Rabbi, M. Kim, C. Ke, W. Lee, R.L. Clark,

P.A. Mieczkowski, and P.E. Marszalek. 2009. UVA

generates pyrimidine dimers in DNA directly. Bio-

phys. J. 96:1151-1158.

Kadir, K. and K.L. Nelson. 2014. Sunlight mediated

inactivation mechanisms of Enterococcus faecalis

and Escherichia coli in clear water versus waste sta-

bilization pond water. Wat. Res. 50:307-317.

Lang, N.L., M.D. Bellett-Travers, and S.R. Smith.

2007. Field investigations on the survival of Esch-

erichia coli and presence of other enteric micro-

organisms in biosolids-amended agricultural soil.

J. Appl. Microbiol. 103:1868–1882.

Le Bouqin, S., V. Allain, X. Rouxel, I. Petetin, M. Pi-

cherot, V. Michel, and M. Chemaly. 2010. Preva-

lence and risk factors for Salmonella spp. contami-

nation in French broiler-chicken flocks at the end

of the rearing period. Prev. Vet. Med. 97:245-251.

LeJeune, J.T., D. Hancock, Y. Wasteson, E. Skjerve,

and A.M. Urdahl. 2006. Comparison of E. coli O157

and Shiga toxin-encoding genes (stx) prevalence

between Ohio, USA and Norwegian dairy cattle.

Int. J. Food Microbiol. 109:19-24.

Lomonaco, S., L. Decastelli, D.M. Bianchi, D. Nucera,

M.A. Grassi, V. Sperone, and T. Civera. 2009. De-

tection of Salmonella in finishing pigs on farm and

at slaughter in Piedmont, Italy. Zoonoses Public

Health 56:137-144.

Ma, L., G. Zhang, and M.P. Doyle. 2011. Green fluo-

rescent protein labeling of Listeria, Salmonella,

and Escherichia coli O157:H7 for safety-related

studies. PLoS One 6:e18083.

MacDonald, J.M., M.O. Ribaudo, M.J. Livingston, J.

Beckman, and W. Huang. 2009. Manure use for fer-

tilizer and for energy: Report to Congress. USDA-

ERS Publication No. AP-037. Available at: http://

www.ers.usda.gov/media/156155/ap037_1_.pdf.

Accessed 12 September, 2014.

Maynard, H.E., S.K. Ouki, and S.C. Williams. 1999.

Tertiary lagoons: A review of removal mechanisms

and performance. Wat. Res. 33:1-13.

Meays, C.L., K. Broersma, R. Nordin, and A. Mazum-

der. 2005. Survival of Escherichia coli in beef cattle

fecal pats under different levels of solar exposure.

Rangeland Ecol. Manage. 58:279-283.

Okraszewska-Lasica, W., D.J. Bolton, J.J. Sheridan,

and D.A. McDowell. 2014. Airborne Salmonella

and Listeria associated with Irish commercial beef,

sheep and pig plants. Meat Sci. 97:255-261.

Ongeng, D., C. Muyanja, A.H. Geeraerd, D. Spring-

ael, and J. Ryckeboer. 2011. Survival of Escherichia

coli O157:H7 and Salmonella enterica serovar Ty-

phimurium in manure and manure-amended soil

under tropical climatic conditions in Sub-Saharan

Africa. J. Appl. Microbiol. 110:1007-1022.

Park, G.W., and F. Diez-Gonzalez. 2003. Utilization

of carbonate and ammonia-based treatments to

eliminate Escherichia coli O157:H7 and Salmonella

Typhimurium DT104 from cattle manure. J. Appl.

Microbiol. 94:675-685.

Pedersen, T.B., J.E. Olsen, and M. Bisgaard. 2008.

Persistence of Salmonella Senftenberg in poultry

production environments and investigation of its

resistance to desiccation. Avian Pathol. 37:421-427.

Pell, A.N. 1997. Manure and microbes: public and

animal health problem? J. Dairy Sci. 80:2673-2681.

Raviv, M. 2005. Production of high-quality composts

for horticultural purposes: A mini-review. Hort-

technology 15:52-57.

Semenov, A.V., A.H.C. van Bruggen, L. van Overbeek,

A.J. Termorshuizen, and A.M. Semenov. 2007. In-

fluence of temperature fluctuations on Escherichia

coli O157:H7 and Salmonella enterica serovar Ty-

phimurium in cow manure. FEMS Microbiol. Ecol.

60:419-428.

Sherman, R.A., K. Goth, J. Sherman, M. Tran, and D.

Ng. 2006. Effects of food storage and handling on

blow fly (Lucilia sericata) eggs and larvae. J. Food

Sci. 71:M117-M120.

Sinton, L.W., R.R. Braithwaite, C.H. Hall, and M.L.

Mackenzie. 2007. Survival of indicator and patho-

genic bacteria in bovine feces on pasture. Appl.

Environ. Microbiol. 73:7917-7925.

Tamura, A., M. Yamasaki, A. Okutani, S. Igimi, N.

Saitoh, T. Ekawa, H. Ohta, Y. Katayama, and F.

Amano. 2009. Dry-resistance of Salmonella enteri-

ca subsp. enterica serovar Enteritidis is regulated

by both SEp22, a novel pathogenicity-related fac-

tor of Salmonella, and nutrients. Microbes Envi-

ron. 24:121-127.


United States Environmental Protection Agency [US

EPA]. 1999a. Standards for the use or disposal of

sewage sludge (40 CFR parts 403 and 503). Revised

August 4, 1999. http://www.epa.gov/EPA-WA-

TER/1999/August/Day-04/w18604.htm. Accessed

15 April, 2014.


EPA]. 1999b. Appendix B to Part 503. Pathogen

Treatment Processes. Processes to Significantly

Reduce Pathogens. (PSRP). EPA/625/R-92/013. Re-

vised October 1999. United States Environmental

Protection Agency, Office of Research and Devel-

opment, National Risk Management Laboratory,

Center for Environmental Research Information,

Cincinnati, OH. Available at: http://www.gpo.

gov/fdsys/pkg/CFR-2012-title40-vol31/pdf/CFR-

2012-title40-vol31-part503-appB.pdf. Accessed 4

February, 2014


EPA]. 2013. Literature review of contaminants in

livestock and poultry manure and implications for

water quality. http://water.epa.gov/scitech/cec/

upload/Literature-Review-of-Contaminants-in-

Livestock-and-Poultry-Manure-and-Implications-

for-Water-Quality.pdf. Accessed 15 April, 2014.

Vogel, B.F., L.T. Hansen, H. Mordhorst, and L. Gram.

2010. The survival of Listeria monocytogenes dur-

ing long term desiccation is facilitated by sodium

chloride and organic material. Int. J. Food Micro-

biol. 140:192-200.

Wichuk, K.M. and D. McCartney. 2007. A review of

the effectiveness of current time-temperature

regulations on pathogen inactivation during com-

posting. J. Environ. Engr. Sci. 6:573-586.




VOLUME 4 ISSUE 1

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

20

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

13

Behavioral Change Study at a Western Soup Production Plant

C. A. O’Bryan, J. A. Neal, and P. G. Crandall

27

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

35

The Hurricane Sandy DilemmaB. A. Almanza

43

Introduction Special IssueP. G. Crandall

8

Case Studies



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

50


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

76

Effects of Co-nutrients in Foods and Bioremediation in the Environment on Methylmercury

P. G. Crandall, C. A. O’Bryan

86

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

109

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

122

REVIEW

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

96

ARTICLES




VOLUME 4 ISSUE 2



MANUSCRIPT SUBMISSION

Authors must submit their papers electronically

([email protected]). According to instruc-

tions provided online at our site: www.afabjournal.

com. Authors who are unable to submit electroni-

cally should contact the editorial office for assistance

by email at [email protected].

INSTRUCTIONS TO AUTHORS

• Aerobic microbiology

• Aerobiology

• Anaerobic microbiology

• Analytical microbiology

• Animal microbiology

• Antibiotics

• Antimicrobials

• Bacteriophage

• Bioremediation

• Biotechnology

• Detection

• Environmental microbiology

• Feed microbiology

• Fermentation

• Food bacteriology

• Food control

• Food microbiology

• Food quality

• Food Safety

• Foodborne pathogens

• Gastrointestinal microbiology

• Microbial education

• Microbial genetics

• Microbial physiology

• Modeling and microbial kinetics

• Natural products

• Phytoceuticals

• Quantitative microbiology

• Plant microbiology

• Plant pathogens

• Prebiotics

• Probiotics

• Rumen microbiology

• Rapid methods

• Toxins

• Veterinary microbiology

• Waste microbiology

• Water microbiology

CONTENT OF MANUSCRIPT

We invite you to consider submitting your re-

search and review manuscripts to AFAB. The jour-

nal serves as a peer reviewed scientific forum for to

the latest advancements in bacteriology research

on Agricultural and Food Systems which includes

the following fields:


With an open access publication model of this

journal, all interested readers around the world can

freely access articles online. AFAB publishes origi-

nal papers including, but not limited to the types

of manuscripts described in the following sections.

Papers that have been, or are scheduled to be, pub-

lished elsewhere should not be submitted and will

not be reviewed. Opinions or views expressed in pa-

pers published by AFAB are those of the author(s)

and do not necessarily represent the opinion of the

AFAB or the editorial board.

MANUSCRIPT TYPES

Full-Length Research Manuscripts

AFAB accepts full-length research articles con-

taining four (4) figures and/or tables or more. AFAB

emphasizes the importance of sound scientific ex-

perimentation on any of the topics listed in the focus

areas followed by clear concise writing that describes

the research in its entirety. The results of experi-

ments published in AFAB must be replicated, with

appropriate statistical assessment of experimental

variation and assignment of significant difference.

Major headings to include are: Abstract, Introduc-tion, Materials and Methods, Results, Discussion (or Results and Discussion), Conclusion, Acknowl-edgements (optional), Appendix for abbreviations (optional), and References.

Manuscripts clearly lacking in language will be re-

turned to author without review, with a suggestion

that English editing be sought before the paper is

reconsidered. AFAB offers a fee based language

service upon request. Please contact [email protected] for more information about our fees

and services.

Rapid Communications

Under normal circumstances, AFAB aims for re-

ceipt-to-decision times of approximately one month or less. Accepted papers will have priority for publi-

cation in the next available issue of AFAB. However,

if an author chooses or requires a much more rapid

peer review, the journal editorial office has the capa-

bility to shorten the review timing to one week or less.

Any type of manuscript whether it be a full length

manuscript, brief communication or review paper can

be submitted as a rapid communication. There will be

additional costs for processing and page charges will

be double the normal rate. Authors who choose this

option must select Rapid Communications as the pa-

per type when submitting the paper and the editors

will judge whether a rapid review is possible and let

the author know immediately.

Brief Communications

Brief communications are short research notes giv-

ing the results of complete experiments but are con-

sidered less comprehensive than full-length articles

with three (3) figures and/or tables or less. Manuscripts

should be prepared with the same subheadings as full

length research papers. The running head above the

title of the paper is “Brief Communications.”

Unsolicited Review Papers

Review papers are welcome on any topic listed in

the focus section and have no page limits. Reviews

are assessed the same pages charges as all other

manuscripts. All AFAB guidelines for style and form

apply. Major headings to include are: Abstract, In-troduction, Main discussion topics and appropri-ate subheadings, Conclusions, Acknowledgements (optional) and References. Review papers shorter

than 20 pages of double spaced text and references

will be considered mini-reviews with the subhead-

ing above the title on the first page. The running

head above the title of the paper is either “Review”

or “Mini-review”.

Solicited Review Papers

Solicited reviews will have no page limits. The

editor-in-chief will send invitations to the authors

and then review these contributions when they are

submitted. Nominations or suggestions for potential

timely reviews are welcomed by the editors or edito-


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 origi-

nate from the editor-in-chief or one of the editors. Re-

quests 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 meet-

ings in their entirety. Conference organizers need

simply to contact AFAB at [email protected]

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. Con-

ference 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 pa-

pers and, if necessary, returning them to the authors

for revision. The conference organizer then submits

the revised manuscripts to the journal editorial of-

fice 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. Al-

though 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 jour-

nal as ordinary unsolicited review manuscripts.

Book Reviews

AFAB publishes reviews of books considered to

be of interest to the readers. The editor-in-chief ordi-

narily 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, cri-

tique, 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 par-

ticular manuscript then the author(s) of the original

paper(s) will be provided a copy of the letter and of-

fered 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 tech-

nical editor. If multiple letters on the same topic are

received, a representative set of opinions concern-

ing 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).


AFAB grants to the author the right of re-publication

in any book of which he or she is the author or edi-

tor, subject only to giving proper credit to the original

journal publication of the article by AFAB. AFAB re-

tains the copyright to all materials accepted for pub-

lication 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 au-

thority 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 Commu-

nication 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 as-

sessed upon submission. Once reviewed, the author

will be notified of the outcome and advised accord-

ingly. 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. Re-

vised versions of previously rejected manuscripts are

treated as new submissions.

PRODUCTION OF PROOFS

Accepted manuscripts are forwarded to the edito-

rial office for technical editing and layout. The manu-

script is then formatted, figures are reproduced, and

author proofs are prepared as PDFs. Author proofs

of all manuscripts will be provided to the correspond-

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 correc-

tion in Adobe Acrobat to indicate necessary chang-

es. 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 re-

sponsibility for errors that result in the final publication.

PUBLICATION CHARGES

AFAB has two publication charge options: conven-

tional 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 com-

munication, 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 quan-

tity 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.


MANUSCRIPT CONTENT REQUIREMENTS

Preparing the Manuscript File

Manuscripts must be written in grammatically

correct English. AFAB offers a fee based language

service upon request ([email protected]).

Manuscripts should be typed double-spaced, with

lines and pages numbered consecutively. All docu-

ments 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 sym-

bols 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 fol-

low these instructions will cause delays of the pro-

cessing 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 abbre-

viations 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 cor-

responding author should be indicated with an aster-

isk e.g., * Corresponding author. The title page shall

include the name and full address of the correspond-

ing author. Telephone and e-mail address must also

be provided for the corresponding author, and email-addresses must be provided for all authors.

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 au-

thors may need to provide a list of abbreviations

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 ta-

bles, 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 sepa-

rate page after the title page. The abstract should

provide a justification of your work, objectives, meth-

ods, results, discussion and implications of study or

review findings . Your abstract must consist of com-

plete 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 index-

ing. Ideally, these keywords should include signifi-

cant words from the title.

Introduction

The introduction should clearly present the foun-

dation 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 ob-

jectives must be clearly stated in the final sentence

of the last paragraphs of the introduction.

Materials and Methods

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)].


Variability, Replication, and Statistical Analysis

To properly assess biological systems indepen-

dent 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 com-

monly 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 experimen-

tal unit is the smallest unit to which an individual

treatment is imposed. For bacterial growth stud-

ies, 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. Mea-

surements on the same experimental unit over time

are also not independent and must not be consid-

ered as independent experimental units. For analy-

sis of time effects, assess as a rate of change over

time. Standard deviation refers to the variability

in the biological response being measured and is

presented as standard deviation or standard error

according to the definitions described in statistical

references or textbooks.

Results

Results represent the presentation of data in

words and all data should be described in same

fashion. No discussion of literature is included in

the results section.

Discussion

The discussion section involves comparing the

current data outcomes with previously published

work in this area without repeating the text in the

results section. Critical and in-depth dialogue is

encouraged.

Results and Discussion

Results and discussion can be under combined or

separate headings.

Conclusions

State conclusions (not a summary) briefly in one

paragraph.

Acknowledgments

Acknowledgments of individuals should include

institution, city, and state; city and country if not U.S.;

and City or Province if in Canada. Copies being re-

viewed shall have authors’ institutions omitted to re-

tain anonymity.

References

a) Citing References In Text

Authors of cited papers in the text are to be pre-

sented 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 pa-

rentheses, 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 alphabeti-

cally 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 un-

published work should be listed in the text as “(J.

Adams, unpublished data).” Personal communica-

tions 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 ci-

tations designated by et al. in the text have the same

first author, then even if the other authors are differ-

ent these references in the text and the references

section must be identified by a letter. For example


“(James et al., 2010a,b)” in text, refers to “James,

Smith, and Elliot. 2010a” and “James, West, and Ad-

ams. 2010b” in the reference section.

b) Citing References In Reference Section

In the References section, references are listed in

alphabetical order by authors’ last names, and then

chronologically. List only those references cited in the

text. Manuscripts submitted for publication, accepted

for publication or in press can be given in the refer-

ence section followed by the designation: “(submit-

ted)”, “(accepted)’, or “(In Press), respectively. If the

DOI number of unpublished references is available,

you must give the number. The year of publication fol-

lows the authors’ names. All authors’ names must be

included in the citation in the Reference section. Jour-

nals must be abbreviated. First and last page num-

bers must be provided. Sample references are given

below. Consult recent issues of AFAB for examples

not included in the following section.

Journal manuscript:

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.

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.

Book:

Examples:

Hungate, R. E. 1966. The rumen and its microbes

Academic Press, Inc., New York, NY. 533 p.

Book Chapter:

Examples:

O’Bryan, C. A., P. G. Crandall, and C. Bruhn. 2010.

Assessing consumer concerns and perceptions

of food safety risks and practices: Methodologies

and outcomes. In: S. C. Ricke and F. T. Jones. Eds.

Perspectives on Food Safety Issues of Food Animal

Derived Foods. Univ. Arkansas Press, Fayetteville,

AR. p 273-288.

Dissertation and thesis:

Maciorowski, K. G. 2000. Rapid detection of Salmo-

nella spp. and indicators of fecal contamination

in animal feed. Ph.D. Diss. Texas A&M University,

College Station, TX.

Donalson, L. M. 2005. The in vivo and in vitro effect

of a fructooligosacharide prebiotic combined with

alfalfa molt diets on egg production and Salmo-

nella in laying hens. M.S. thesis. Texas A&M Uni-

versity, College Station, TX.

Van Loo, E. 2009. Consumer perception of ready-to-

eat deli foods and organic meat. M.S. thesis. Uni-

versity of Arkansas, Fayetteville, AR. 202 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 #_____

Author(s). Year. Article title. Journal title [abbreviated].

Volume number:inclusive pages.

Author(s) [or editor(s)]. Year. Title. Edition or volume (if

relevant). Publisher name, Place of publication. Number

of pages.

Author(s) of the chapter. Year. Title of the chapter. In:

author(s) or editor(s). Title of the book. Edition or vol-

ume, if relevant. Publisher name, Place of publication.

Inclusive pages of chapter.

Author. Date of degree. Title. Type of publication, such

as Ph.D. Diss or M.S. thesis. Institution, Place of institu-

tion. Total number of pages.


Abstracts and Symposia Proceedings:

Fischer, J. R. 2007. Building a prosperous future in

which agriculture uses and produces energy effi-

ciently 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. Op-

tions and risks by using diagnostic gene chips. Pro-

gram and abstracts book , The 8th Biennieal Con-

gress 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 experi-

mental 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., Mi-

crosoft 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. Sym-

bols, lines, and bars must be made distinct as “stand

alone” black and white presentations. Stippling,

dashed lines etc. are encouraged for multiple com-

parison but shades of gray are discouraged. Color

images, micrographs, pictures are recommended

and there is no additional fee for their submission.

AFAB Online Edition is Now Available!

www.AFABjournal.com

• Free Access

• Print PDFs

• Flip Through Issues

• Search Article Archives

• Order Reprints

• Submit a Paper

Online Publication: www.AFABjournal.com

Documents

Afab volume 4 issue 3