EFFECTS OF MYCOTOXIN ADSORBENTS ON ...ufdcimages.uflib.ufl.edu/UF/E0/05/08/73/00001/OGUNADE_I.pdfEFFECTS OF MYCOTOXIN ADSORBENTS ON AFLATOXINS AND OF MICROBIAL ADDITIVES ON Escherichia

EFFECTS OF MYCOTOXIN ADSORBENTS ON AFLATOXINS AND OF MICROBIAL

ADDITIVES ON Escherichia coli O157:H7 IN FEEDS FOR DAIRY COWS

By

IBUKUN MICHAEL OGUNADE

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL

OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2017

© 2017 Ibukun Michael Ogunade

To my wife, Adeola, and my daughter, Tomisin

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ACKNOWLEDGMENTS

My sincere gratitude goes to my advisor, Dr. Adegbola Adesogan, for believing in me

despite my very slow and poor start to the program. Your encouragement and words of wisdom

kept me going throughout the program. My appreciation goes to my supervisory committee

members, Dr. José Santos, Dr. Charles Staples, Dr. Casey Jeong, Dr. Patricio Munoz, and Dr.

Diwakar Vyas, for their considerable contributions to and constructive criticisms of my projects

and guidance in my professional life.

I acknowledge my lab mates, Yun Jiang and Andres Pech Cervantes for their support and

friendship throughout the program. I would not be presenting this dissertation without their

pivotal roles in my experiments. My sincere thanks goes to Dr. Kathy Arriola for helping and

coordinating my first experiment. I learned a lot from your professionalism. Special thanks to Dr.

Kim Donghyeon, who spent a lot of his time helping me closely on two of the projects.

I was privileged to work with visiting professors who contributed greatly to my

professional life. Special thanks goes to Dr. Bamikole Musibau from the University of Benin,

Nigeria; Dr. Zwika Weinberg from The Volcani Center, Israel; Dr. Thiago Bernardes from the

Federal University of Lavras, Brazil; and Dr. André Oliveira from the Federal University of

Mato Grosso, Brazil. My most sincere thanks to all my friends in the department including

Marcos Zenobi, Chengcheng Li, Carlos Martinez, Zaira Estrada, Felipe Amaro, Rafael Martins,

Guan Yang, Zhengxin Ma, and Olutoye Olowolafe.

I would like to thank the Dairy Research Unit farm crew for their help with my projects

and for allowing me to use the farm’s facilities.

To my Mom and siblings, Tayo, Lanre and Kehinde, though we are far apart, we have

always been in touch. Thank you for your love and care.

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My special appreciation goes to my wife, Adeola and daughter, Tomisin. I sincerely

thank you for your love and patience. Your words of encouragement kept me going when I

almost gave up. I sincerely thank you for your endurance during the unpleasant times.

Lastly, I thank the almighty God for giving me the strength and preserving my life to

witness this day; all glory and adoration to His name.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...............................................................................................................4

LIST OF TABLES .........................................................................................................................10

LIST OF FIGURES .......................................................................................................................12

LIST OF ABBREVIATIONS ........................................................................................................14

ABSTRACT ...................................................................................................................................16

CHAPTER

1 INTRODUCTION ..................................................................................................................18

2 LITERATURE REVIEW .......................................................................................................20

Silage Fermentation ................................................................................................................20

Phases of Silage Fermentation .........................................................................................20 Aerobic Phase ..................................................................................................................21 Fermentation Phase .........................................................................................................21

Stable Phase .....................................................................................................................22 Feed-out Phase .................................................................................................................22

Silage Additives ......................................................................................................................22 Homofermentative Bacteria .............................................................................................23

Heterofermentative Bacteria ............................................................................................24 Ferulate Esterase-producing Bacteria ..............................................................................25

Inoculants Containing Saccharomyces Strains ................................................................26 Silage Microbial Community Analysis by Molecular Techniques.........................................27

Terminal Restriction Fragment Length Polymorphism ...................................................28

Length Heterogeneity Polymerase Chain Reaction .........................................................29 Denaturing Gradient Gel Electrophoresis .......................................................................29 Next-generation Sequencing Methods ............................................................................30

Roche 454 sequencing pyrosequencing ...................................................................30

Single-molecule real-time (SMRT) sequencing .......................................................31

Ion torrent personal genome machine platform .......................................................31

Illumina MiSeq platform. .........................................................................................32 Pathogenic Escherichia coli ...................................................................................................32 Cattle as a Reservoir of Escherichia coli O157:H7 ................................................................35 Pathogenic Escherichia coli in Ensiled Forages .....................................................................36 Control of Fecal Shedding of EC by Ruminants ....................................................................38

Diet Change .....................................................................................................................38 Vaccinations ....................................................................................................................40 Direct-Fed Microbials .....................................................................................................40

Bacteriophage Therapy ....................................................................................................41

7

Mycotoxins in Ensiled Forages ..............................................................................................42

Prevalence of Mycotoxins in Foods and Feeds ......................................................................43 Factors that Predispose to Mold and Mycotoxin Production in Silage ..................................43 Types of Mycotoxins in Ensiled Forages ...............................................................................45

Fusarium Toxins .............................................................................................................45 Deoxynivalenol ........................................................................................................45 Fumonisins ...............................................................................................................47 Zearalenone ..............................................................................................................48 T-2 Toxin ..................................................................................................................49

Aspergillus Toxins ...........................................................................................................50 Penicillium Toxins ...........................................................................................................52

Ochratoxin A ............................................................................................................53 Mycophenolic acid and Roquefortine C ...................................................................53

Ruminal Degradation of Mycotoxins .....................................................................................54 Co-occurrence of Mycotoxins in Livestock Feeds .................................................................55

Dietary Contribution of Corn Silage to Mycotoxin Intake in Cattle ......................................56 Preventing Mycotoxin Contamination of Feeds .....................................................................57

Field or Pre-harvest Phase ...............................................................................................57 Harvest Phase ..................................................................................................................57 Ensiling Phase .................................................................................................................58

Mycotoxin Detoxification Strategies ......................................................................................60

3 CONTROL OF Escherichia coli O157:H7 IN CONTAMINATED ALFALFA

SILAGE: EFFECTS OF SILAGE ADDITIVES ....................................................................67

Background .............................................................................................................................67

Materials and Methods ...........................................................................................................68 Harvesting, Ensiling and Inoculation ..............................................................................68

Survival of E. coli O157:H7 in Aerobically Exposed Silages .........................................69 Laboratory Analyses ........................................................................................................70 Antibacterial Activity of the Microbial Inoculants and Silage Extracts ..........................71

Statistical Analysis ..........................................................................................................71 Results.....................................................................................................................................72

Anaerobic Phase ..............................................................................................................72 Aerobic Phase ..................................................................................................................73

Antibacterial Activity of Microbial Inoculants and Silage Extracts ...............................74 Discussion ...............................................................................................................................74

Anaerobic Ensiling Phase ................................................................................................75 Aerobic Post-ensiling Phase ............................................................................................77

Conclusions.............................................................................................................................78

4 BACTERIAL DIVERSITY AND COMPOSITION OF ALFALFA SILAGE AS

ANALYZED BY ILLUMINA MISEQ SEQUENCING: EFFECTS OF E. coli O157:H7

AND SILAGE ADDITIVES ..................................................................................................88

Background .............................................................................................................................88 Materials and Methods ...........................................................................................................90

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Metagenomic DNA Extraction and Purification .............................................................90

PCR Amplification of the V3 and V4 Hypervariable Regions of Bacterial 16S

rRNA Genes .................................................................................................................90 Sequencing and Diversity Analysis .................................................................................91

Results.....................................................................................................................................92 Sequencing Results and Bacterial Diversity ....................................................................92 Comparisons at the Phylum Level ...................................................................................93 Comparisons at the Family Level ....................................................................................93 Comparisons at the Genus Level .....................................................................................94

Comparisons at the Species Level ...................................................................................94 Correlations between Abundance of Bacteria and Fermentation Quality Indices ..........95

Discussion ...............................................................................................................................97 Bacterial Diversity and Composition ..............................................................................97

Correlations between Bacterial Abundance and Fermentation Quality Indices ............100 Bacterial Candidates for Future Research .....................................................................103

Conclusions...........................................................................................................................104

5 FATE OF Escherichia coli O157:H7 AND BACTERIAL DIVERSITY IN CORN

SILAGE CONTAMINATED WITH THE PATHOGEN AND TREATED WITH

MICROBIAL AND CHEMICAL ADDITIVES ..................................................................116

Background ...........................................................................................................................116

Materials and Methods .........................................................................................................118 Forage and Treatments ..................................................................................................118

Survival of E. coli O157:H7 in Aerobically Exposed Silages .......................................120

Laboratory Analyses ......................................................................................................120

Antibacterial Activity of the Silage Extracts .................................................................122 DNA Extraction and Sequencing ..................................................................................122

Data and Statistical Analysis .........................................................................................123 Results...................................................................................................................................124

Anaerobic Phase ............................................................................................................124

Aerobic Phase ................................................................................................................125 Antibacterial Activity ....................................................................................................126

Bacterial Community Composition and Diversity ........................................................126 Discussion .............................................................................................................................127

Anaerobic Phase ............................................................................................................127 Aerobic Post-Ensiling Phase .........................................................................................129

Bacterial Community Composition and Diversity ........................................................130 Conclusions...........................................................................................................................134

6 EFFECTS OF THREE SEQUESTERING AGENTS ON MILK AFLATOXIN M1

CONCENTRATION AND THE PERFORMANCE AND IMMUNE STATUS OF

DAIRY COWS FED DIETS ARTIFICIALLY CONTAMINATED WITH

AFLATOXIN B1 ..................................................................................................................146

Background ...........................................................................................................................146 Materials and Methods .........................................................................................................147

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Cows, Treatments, and Design ......................................................................................147

Experimental Measurements and Analytical Procedures ..............................................149 Statistical Analysis ........................................................................................................151

Results and Discussion .........................................................................................................152

Milk AFM1 Concentration .............................................................................................152 Animal Performance ......................................................................................................156 Blood Parameters ...........................................................................................................157

Conclusions...........................................................................................................................159

7 GENERAL SUMMARY ......................................................................................................167

LIST OF REFERENCES .............................................................................................................174

BIOGRAPHICAL SKETCH .......................................................................................................214

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LIST OF TABLES

Table page

2-1 Estimated dietary contributions of whole plant corn silage to total dietary mycotoxin

in diets ................................................................................................................................66

3-1 Chemical composition and pH of alfalfa forage treated without (Control) or with

Escherichia coli O157:H7 (EC) alone, or EC and either of 2 bacterial inoculants or

propionic acid before ensiling ............................................................................................80

3-2 Fermentation indices, chemical composition and yeast counts of alfalfa silage treated

without (Control) or with Escherichia coli O157:H7 (EC) alone, or EC and either of

2 bacterial inoculants or propionic acid and ensiled for 100 d ..........................................81

4-1 Relative abundance (%) of phyla isolated from alfalfa silage treated without

(Control) or with Escherichia coli O157:H7 (EC) alone, or EC and either of two

bacterial inoculants or propionic acid and ensiled for 100 d ...........................................105

4-2 Relative abundance (%) of the 13 most abundant families isolated from alfalfa silage

treated without (Control) or with Escherichia coli O157:H7 (EC) alone, or EC and

either of two bacterial inoculants or propionic acid and ensiled for 100 d ......................106

4-3 Relative abundance (%) of 13 most abundant genera isolated from alfalfa silage


either of two bacterial inoculants or propionic acid and ensiled for 100 .........................107

4-4 Relative abundance of bacterial species isolated from alfalfa silage treated without


bacterial inoculants or propionic acid and ensiled for 100 d ...........................................108

4-5 Relative abundance (%) of bacterial species isolated from alfalfa silage treated


two bacterial inoculants or propionic acid and ensiled for 100 d ....................................110

4-6 Pearson’s correlation between abundance of bacterial genus and fermentation indices

(% DM) of alfalfa forage ensiled for 100 d .....................................................................111

4-7 Pearson’s correlation between abundance of bacterial species and fermentation

indices (% DM) of alfalfa forage ensiled for 100 d .........................................................112

4-8 Pearson’s correlation between abundance of unknown bacterial species and

fermentation indices (% DM) of alfalfa forage ensiled for 100 d....................................113

5-1 Characteristics of whole-plant corn forage inoculated with Escherichia coli O157:H7

(EC) alone or EC and bacterial inoculants or propionic acid before ensiling .................136

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5-2 Fermentation indices and chemical composition and microbial counts (log cfu/g) of

corn silage that was inoculated with Escherichia coli O157:H7 (EC) alone or with

bacterial inoculants or propionic acid after 120 d of ensiling ..........................................137

5-3 Bacterial community composition and diversity of whole-plant corn silage inoculated

with Escherichia coli O157:H7 (EC) alone or with bacterial inoculants or propionic

acid ...................................................................................................................................138

6-1 Ingredient and chemical composition of the experimental diet .......................................161

6-2 Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents on the

concentration, transfer and excretion of aflatoxin M1 (AFM1) in the milk of dairy

cows during toxin-dosing and withdrawal periods ..........................................................162


performance of cows during Predosing (d 16 to 20), Toxin-dosing (d 21 to 25) and

Withdrawal (d 26 to 28) periods ......................................................................................163

6-4 Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents on blood

cell counts of dairy cows..................................................................................................164

6-5 Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents on immune

response indices of dairy cows ........................................................................................165

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LIST OF FIGURES

Figure page

3-1 Changes in lactate concentration of alfalfa forage treated without (Control) or with


propionic acid during ensiling. ..........................................................................................82

3-2 Changes in acetate concentration of alfalfa forage treated without (Control) or with



3-3 Changes in pH of alfalfa forage treated without (Control) or with Escherichia coli

O157:H7 (EC) alone, or EC and either of 2 bacterial inoculants or propionic acid

during ensiling. ..................................................................................................................84

3-4 Escherichia coli O157:H7 (EC) of alfalfa forage treated without (Control) or with



3-5 Effect of reinoculation with 1 × 105 cfu/g of Escherichia coli O157:H7 (EC) at silo

opening on counts of EC and yeasts (log cfu/g) on aerobically exposed alfalfa silages

that were treated without (Control) or with EC alone or EC and either of 2 bacterial

inoculants or propionic acid at ensiling. ............................................................................86

3-6 Effect of reinoculation with 1 × 105 cfu/g of Escherichia coli O157:H7 (EC) at silo

opening on pH of aerobically exposed alfalfa silages that were treated without

(Control) or with EC alone or EC and either of 2 bacterial inoculants or propionic

acid at ensiling. ..................................................................................................................87

4-1 Estimate of alpha diversity and species richness calculated as the Chao1 index.. ..........114

4-2 Unweighted UniFrac PCoA plot of individual samples in each treatment.. ....................115

5-1 Changes in the pH of whole-plant corn forage inoculated with 1 × 105 cfu/g of

Escherichia coli O157:H7 (EC) alone or with bacterial inoculants or propionic acid

and ensiled for different durations.. .................................................................................139

5-2 Changes in Escherichia coli O157:H7 (EC) counts of whole-plant corn forage

inoculated with 1 × 105 cfu/g of Escherichia coli O157:H7 (EC) alone or with

bacterial inoculants or propionic acid and ensiled for different durations.. .....................140

5-3 Changes in lactate concentration of whole-plant corn forage inoculated with 1 × 105

cfu/g of Escherichia coli O157:H7 (EC) alone or with bacterial inoculants or

propionic acid and ensiled for different durations.. .........................................................141

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5-4 Changes in acetate concentration of whole-plant corn forage inoculated with 1 × 105


propionic acid and ensiled for different durations.. .........................................................142

5-5 Escherichia coli O157:H7 (EC) counts and pH of whole-plant corn silage (d 128) re-

inoculated with 5 × 105 cfu/g of EC 168 h after silo opening..........................................143

5-6 The aerobic stability (h) of whole-plant corn forage inoculated with 1 × 105 cfu/g of


and ensiled for 120 d.. ......................................................................................................144

5-7 Unweighted UniFrac principal coordinate analysis (PCoA) plot of individual samples

in each treatment. Individual sample was represented with blue (Control), red

(ECCH), orange (ECLP), purple (ECLB), and yellow (ECA).. ......................................145


concentration of aflatoxin M1 (AFM1) in the milk of dairy cows.. ...................................166

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LIST OF ABBREVIATIONS

ADF Acid-detergent fiber

AFB1 Aflatoxin B1

AFM1 Aflatoxin M1

ATP Adenosine triphosphate

BPW Buffered-peptone water

CDC Centers for Disease Control and Prevention

CP Crude protein

DAEC Diffusely-adherent E. coli

DFM Direct-fed microbial

DIM Days-in-milk

DM Dry matter

DMI Dry matter intake

DON Deoxynivanenol

EAEC Entero-aggregative E. coli

EC Escherichia coli O157:H7

EIEC Entero-invasive E. coli

EPEC Entero-pathogenic E. coli

ETEC Entero-toxigenic E. coli

FCM Fat-corrected milk

FDA Food and Drug Administration

HPLC High-performance liquid chromatograph

HUS Hemolytic uremic syndrome

LAB Lactic acid bacteria

LB Lactobacillus buchneri

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LP Lactobacillus plantarum

MFI Mean florescence intensity

MRS Man-Rogosa-Sharpe

NDF Neutral-detergent fiber

NEL Net energy of lactation

NH3-N Ammonia nitrogen

OTU Operational taxonomic unit

PC Principal coordinate

PCoA Principal coordinate analysis

SCC Somatic cell counts

SE Standard error

SEQ Sequestering agent

SMAC Sorbitol-macConkey agar

STEC Shiga toxin-producing E. coli

TMR Total mixed ration

US United States

ZEA Zearalenone

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Abstract of Dissertation Presented to the Graduate School

of the University of Florida in Partial Fulfillment of the

Requirements for the Degree of Doctor of Philosophy

EFFECTS OF MYCOTOXIN ADSORBENTS ON AFLATOXINS AND OF MICROBIAL

ADDITIVES ON Escherichia coli O157:H7 IN FEEDS FOR DAIRY COWS

By

Ibukun Michael Ogunade

May 2017

Chair: Adegbola T. Adesogan

Cochair: Diwakar Vyas

Major: Animal Sciences

Three experiments were conducted to evaluate the effects of two microbial cultures and a

chemical additive on the survival of pathogenic Escherichia coli O157:H7 (EC) and bacterial

diversity in corn and alfalfa silages. A fourth experiment examined the effects of sequestering

agents on the performance and health of dairy cows challenged with dietary aflatoxin B1.

Experiment 1 examined whether adding Lactobacillus plantarum (LP), Lactobacillus

buchneri (LB) or propionic acid to alfalfa silage contaminated with EC would inhibit the growth

of the pathogen during or after ensiling. The bacterial inoculants hastened the inhibition of EC

during ensiling and, like propionic acid, they also prevented its growth after ensiling during

aerobic exposure.

Experiment 2 assessed the effects of silage additives on bacterial diversity of alfalfa

silage analyzed via Illumina MiSeq sequencing. Inoculation with LP or LB reduced the silage

bacterial diversity by increasing the abundance of the respective bacteria. Relative to the

uninoculated forage, inoculation with LP increased the abundance of Lactobacillus,

Sphingomonas, Pantoea, Pseudomonas, and Erwinia, but reduced that of genera Pediococcus,

Weissella, and Methylobacterium. Abundance of Weissella and Methylobacterium also were

17

reduced in the EC+LB silage compared to uninoculated alfalfa. Application of propionic acid did

not affect the abundance of genera Lactobacillus, Weissella, and Pediococcus. Strong

correlations between some unknown bacteria and indices of silage fermentation quality, such as

lactate, acetate, and ammonia-N, were detected.

Experiment 3 examined the fate of EC and bacterial diversity in corn silage contaminated

with the pathogen and treated with a chemical additive or two microbial inoculants prior to

ensiling. The pathogen was eliminated in all silages within 7 d of contamination at ensiling

because of rapid drops in pH. Propionic acid prevented the growth of the pathogen during the

feed-out stage by keeping the pH of the silage below 4.0. The composition of the bacterial

community of the silages was modified by additive treatment, notably, the abundance of

Lactobacillus was greater in silages treated with LP or propionic acid. Silage treated with LB at

ensiling contained greater abundance of Acinetobacter and Weissella than other silages.

Experiment 4 examined if adding three mycotoxin-sequestering agents to diets

contaminated with aflatoxin B1 would reduce milk aflatoxin M1 concentration and improve the

performance and immune status of dairy cows. The sequestering agent mixtures contained

varying proportions of Saccharomyces cerevisiae, clay, and chlorophyll-based products. The

sequestering agents did not prevent the toxin from being transferred to milk but they reduced

toxin-induced inflammatory stress in the dairy cows.

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CHAPTER 1

INTRODUCTION

The safety of animal feeds has emerged as an important issue because of the recent

deaths and illnesses because of feed and food-borne pathogens and toxins in the United States

(US) and other parts of the world. Escherichia coli is a facultative anaerobic bacterium

commonly found in the mammalian intestinal tract (Drasar and Barrow, 1985) and E. coli

O157:H7 (EC) is a Shiga-toxin-producing bacterium that causes hemorrhagic colitis, bloody

diarrhea, and hemolytic uremic syndrome (HUS) in humans (Kaper et al., 2004). This pathogen

has been isolated from both dairy and beef cattle and cattle are considered the primary reservoir

of EC (Cernicchiaro et al., 2013; Mir et al., 2015). Manure from cattle containing viable EC can

contaminate the water supply on farms such that EC subsequently can be consumed directly in

drinking water or contaminated irrigation water or manure used as fertilizer used on harvested

forages (Weinberg et al., 2004; Mir et al., 2015) that are later fed to cows. Furthermore,

pathogens also can be found in improperly fermented silages. Fenlon and Wilson (2000) reported

that the number of EC in poorly fermented silage grew from 3 to 6 log cfu/g during the first 7 d

of ensiling when the pH stayed above 5. Consequently, livestock feeds have been suggested to be

the vehicle of transmission of EC on the farm (Hancock et al., 2001; Davis et al., 2003).

Therefore, inhibiting the growth of the pathogen may help reduce its cycling and spread on the

farm.

Fungal contamination of animal feeds is a common occurrence on farms and the

production of mycotoxins by some fungal species poses a significant threat to the livestock

industry because of the detrimental effects of fungal mycotoxins on all classes of farm animals.

Aflatoxin is the most studied mycotoxin because of its carcinogenic potential and transmission

into milk of dairy cows (Diaz et al., 2004; Liu and Wu, 2010). In fact, up to 160,000 cases of

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cancer in the US have been traced to aflatoxin (Liu and Wu, 2010). Clay-based sequestering

agents have reduced the transfer of aflatoxins into milk but may interfere with absorption of

vitamins and minerals (Chestnut et al., 1992). Yeast-based sequestering agents have reduced the

transfer of dietary aflatoxins into milk (Diaz et al., 2004); however, the efficacy of yeast-based

binders is greater at lower levels of dietary aflatoxin contamination (Xiong et al., 2015).

Therefore, more research is needed on alternative aflatoxin-sequestering agents that reduce the

transfer of dietary aflatoxins to milk and prevent or ameliorate negative effects of the toxin on

the performance of livestock. Therefore, the studies presented in this dissertation investigated the

protective effects of sequestering agents and silage additives against aflatoxin and EC,

respectively, in feeds for dairy cows.

The studies presented in Chapters 3 and 4 examined the efficacy of microbial and

chemical additives at inhibiting the growth of EC in alfalfa silage during and after ensiling, and

examined if the additives influence bacterial diversity in silage. The study presented in Chapter 5

examined if addition of microbial inoculants or propionic acid would inhibit the growth of the

EC in corn silage contaminated with EC at ensiling, at silo opening, or after aerobic exposure, as

well as how additive treatment affected the composition of the bacterial community in silage.

The study in Chapter 6 examined if adding a yeast cell wall product, clay, or a chlorophyll-based

product to diets contaminated with aflatoxin B1 (AFB1) would reduce milk aflatoxin M1 (AFM1)

concentration and improve the performance and immune status of dairy cows.

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CHAPTER 2

LITERATURE REVIEW

Silage Fermentation

Anaerobic fermentation of fresh forage is a preservation method that involves the

conversion of plant sugars into organic acids, mainly lactic acid, by epiphytic lactic acid bacteria

(Weinberg and Muck, 1996; Kung, 2010). Production of these organic acids lowers the pH of the

ensiled forage and inhibits the growth of spoilage organisms (Weinberg and Muck, 1996;

Driehuis, 2013). Forages constitute a significant proportion of the feed costs of livestock

production, and preserved forages, such as silage, form the bulk of the ration of dairy cows and

many beef cattle in the US, therefore, production of high quality silage is paramount (Addah,

2014). The main aim of preserving forages is to make them available at the same nutritive value

all-year-round, especially, during the winter period when forages experience little or no growth.

In addition to oxygen restriction and moisture content of forages, successful preservation of

forages as silage is dependent on the activities of the silage microbial population, especially the

bacterial population (Muck, 2013), which often needs to be dominated by inoculated bacteria or

restricted by additives to optimize the fermentation. Improvements in ensiling processes by rapid

acidification, maintenance of anaerobic conditions during ensiling, and reducing aerobic spoilage

after opening the silo, also have helped ensure production of quality silage (Kaiser and Evans,

1997; Muhlbach, 1998; Kung, 2010).

Phases of Silage Fermentation

Forage fermentation involves the following phases (Weinberg and Muck, 1996; Kung,

2010): aerobic, fermentation, stable, and feed-out phases.

21

Aerobic Phase

This phase occurs for a few hours after sealing the silo. During this phase, respiration of

plants and aerobic organisms continues because of the presence of oxygen within and between

the forage plants. This phase continues until the oxygen is used up. Aerobic respiration utilizes

the soluble carbohydrate mass needed by lactic acid bacteria for fermentation. Furthermore,

aerobic respiration produces heat which can reduce protein digestibility when the temperature

reaches 32oC (Collins and Owens, 2003; Pahlow et al., 2003). In addition, plant proteases are

still active during this phase and they metabolize protein through proteolysis and deamination

resulting in ammonia accumulation.

For efficient silage fermentation, the aerobic phase must be shortened as much as

possible to reduce the growth of unwanted and spoilage microorganisms, such as yeast, mold and

aerobic bacteria which can deplete soluble carbohydrates in the ensiled forage (Kung, 2010).

Wilting of the forage to recommended dry matter concentrations, chopping to proper lengths,

and proper and quick packing or compaction are some of the practices that help to shorten the

duration this phase (McDonald et al., 1991; Kung, 2010).

Fermentation Phase

This phase begins when the oxygen in the silo is used up and anaerobic conditions

prevail. During this phase, the lactic acid bacteria convert plant sugars to organic acids, primarily

lactic acid. This reduces the pH of the silage to 3.6 to 5.0, depending on the forage type and

ensiling conditions. The drop in pH helps to preserve the nutritive value and reduce shrinkage of

the silage by preventing the growth of spoilage bacteria, yeasts, and molds (Addah et al., 2014).

This phase can last for several days or weeks depending on the forage type and ensiling

conditions.

22

Stable Phase

Once the pH is low and air ingress into the silo is prevented, the silage environment

becomes relatively stable. However, some acid-tolerant enzymes and micro-organisms, e.g. L.

buchneri, still function at a slow rate during this stage (Schmidt et al., 2008), whereas other

organisms such as clostridia and bacilli survive the acidic environment as spores (Muck, 2010).

Feed-out Phase

Immediately after silo opening, aerobic conditions prevail, therefore dormant aerobic

organisms such as yeasts, are resuscitated. Aerobic spoilage occurs because of depletion of

lactate by lactate-utilizing yeasts and acetic acid bacteria (McDonald et al., 1991). This increases

the pH, thereby, creating a conducive environment for the growth of molds and pathogenic

organisms (McDonald et al., 1991). Yeasts and molds utilize sugars, nutrients and lactate as

substrates, producing heat in the process. This denatures protein and depletes nutrients in the

silage (Kung, 1998). In addition, molds can produce mycotoxins which are detrimental to the

health of livestock and humans (Campbell et al., 2004; Fung and Clark, 2004).

Silage Additives

Production of high quality silage is dependent on enhancing the fermentation processes

during the different phases of forage preservation. Silage additives have been employed for this

purpose for many years (Henderson, 1993). Some of the main classes of silage additives are

fermentation stimulants (microbial inoculants, enzymes, and fermentable carbohydrate sources),

fermentation inhibitors (acids and formaldehyde), aerobic deterioration inhibitors (lactic acid

bacteria, acids and acid salts), and nutrients (McDonald et al., 1991; Kaiser, 1999).

Bacterial inoculants have been the subject of considerable research because, relative to

chemical additives, they are less expensive, less hazardous, and non-corrosive to farm machinery

23

(Yitbarek and Tamir, 2014). The following are the main bacterial inoculants currently used for

silage preservation.

Homofermentative Bacteria

These are commonly referred to as first-generation bacterial additives (Muck, 2010;

Addah et al., 2014). Lactobacillus plantarum, L. casei, Pediococcus species, and Enterococcus

faecium are the species most commonly used as first generation inoculants to rapidly reduce

silage pH by increasing production of lactic acid. These bacteria ferment 1 mole of glucose to 2

moles of lactic acid and 2 moles of adenosine triphosphate (ATP) via the Embden-Meyerhof

pathway. Lactobacillus plantarum is also capable of fermenting pentoses to lactic acid, CO2, and

acetic acid via the heterofermentative pathway in absence of glucose. Consequently, it is now

classified as a facultative heterofermenter (Holzer et al., 2003). Homofermentative bacterial

strains used as silage inoculants are chosen for their rapid growth and ability to compete against

and dominate the silage microbial community. Previous studies have confirmed greater lactic

acid concentrations along with reduced pH and acetic acid, ammonia-N and butyric acid

concentrations in silage inoculated with homofermentative bacteria (Hu et al., 2009; Queiroz et

al., 2013). Similar findings were reported in a review of 230 studies conducted by Muck and

Kung (1997). Furthermore, a recent meta-analytic study from our laboratory confirmed that

inoculation with these bacteria improved silage fermentation (Oliveira et al., 2017).

The effectiveness of homofermentative bacteria at improving silage fermentation is

affected by factors such as population of epiphytic bacteria, plant sugar concentration, and

buffering capacity (Muck and Kung, 1997; Filya et al., 2007). The benefits of inoculation are

more pronounced in forages with low epiphytic bacterial counts, high sugar concentrations,

and/or high buffering capacity (Filya et al., 2007). However, inoculation with homofermentative

24

bacteria may increase aerobic spoilage because of the poor antifungal effects of lactate (Kung,

1998; Johnson et al., 2002) as well as the utilization of lactate by spoilage yeasts (Kung, 2010).

Heterofermentative Bacteria

The problem of aerobic deterioration at feed-out led to the development of

heterofermentative or second-generation bacterial additives (Addah et al., 2014).

Heterofermentative bacteria produce lactic acid, ethanol, CO2, and acetic acid during

fermentation (Oude Elferink et al., 2001) because they lack the fructose diphosphate aldolase

enzyme. Consequently, during the fermentation, heterofermentative bacteria ferment glucose-6-

phosphate into 6-phosphogluconate rather than fructose-6-phosphate (Kung et al., 2003). These

bacteria use the heterofermentative pathway to generate antifungal compounds, such as acetic

and propionic acids (Oude Elferink et al., 2001; Krooneman et al., 2002) which inhibit the

growth of spoilage yeasts (Moon, 1983). Lactobacillus buchneri is the most commonly used

heterofermentative bacterial inoculant; others include L. brevis, Propionibacterium

freudenreichii, and P. acidipropionici. Lactobacillus buchneri converts lactic acid to acetic acid,

1,2-propanediol, and ethanol (Oude Elferink et al. 2001). Silage inoculation with L. buchneri

reduced the lactate and increased acetate concentrations (Kleinschmit and Kung, 2006b),

subsequently reducing the number of yeasts and improving aerobic stability of corn, barley and

sorghum silages (Huisden et al., 2009). However, heterofermentation in silage by L. buchneri

involves DM losses (1 to 3%; Kleinschmit and Kung, 2006b) because of release of one mole of

CO2 for every mole of acetic acid produced.

Lactobacillus brevis is another heterofermentative bacterium often used as a silage

additive as it can generate acetic acid from carbohydrate fermentation (Holzer et al., 2003).

However, this bacterium is not as effective as L. buchneri in producing acetic acid and

improving aerobic stability of silages. Danner et al. (2003) reported that on a DM basis, corn

25

forage treated with L. brevis had greater acetate concentration (2.86 vs. 1.65%) and aerobic

stability (72 h vs. 40 h) compared to the uninoculated silage. Similarly, silage treated with L.

buchneri increased acetate concentration (5.53 vs. 1.65%) and aerobic stability (274 h vs. 40 h)

compared to the uninoculated silage. The greater efficacy of L. buchneri at improving acetate

concentration and subsequently aerobic stability, explains why it is used more commonly as a

bacterial inoculant than L. brevis.

Propionibacteria produce propionic acid which is the strongest antifungal agent of all the

short chain fatty acids (Moon, 1983); however, because of their inability to compete with and

dominate the epiphytic bacterial population at low pH, silage inoculants containing P.

freudenreichii and P. acidipropionici have not affected the propionic acid concentration and

aerobic stability of corn silages (Taylor et al., 2002; Pedroso et al., 2010; Arriola et al., 2011).

Ferulate Esterase-producing Bacteria

The quest for microbial inoculants that have several synergistic benefits led to the

development of third generation inoculants containing strains like L. buchneri PTA 6138 and L.

crispatus NRRL B-30868 that can improve fiber digestibility as well as fermentation and aerobic

stability. The improved fiber digestion occurs because of production of ferulate esterase, an

enzyme that targets ferulic acid, which limits the digestion of forage fiber in ruminants (Yu et al.

2005; Addah et al., 2014) by inhibiting the attachment of fibrolytic bacteria in the rumen (Varel

and Jung 1986). Ferulic acid esters of arabinoxylans impede access of xylanases to xylan

(Gorbacheva and Rodinova, 1977). In addition, ferulic acid forms linkages with lignin and

proteins (Rawel and Rohn, 2010) which further reduce the nutritional value of forages. Ferulate

esterases release ferulic acid from arabinoxylans, thereby increasing susceptibility of the cell

wall to enzymatic degradation (Bartolome et al., 1995). Nsereko et al. (2008) evaluated effects of

adding eight lactic acid bacteria that produce ferulate esterase (L. buchneri PTA 6138, NNRL B-

26

30866; L. crispatus NRRL B-30868, 30869 and 30870; L. reuteri NRRL B-30867, L. brevis

NRRL B-30865; and an unidentified Lactobacillus strain NRRL B-30871) at the rate of 1 × 105

colony forming units (cfu)/g on fermentation and neutral detergent fiber (NDF) digestibility of

perennial ryegrass. All treated silage samples had approximately 10% greater NDF digestibility

compared with the control. However, only silages treated with L. buchneri strain 6138 and

NRRL B-30866 had lower pH and greater acetate concentration relative to the control. In the

same experiment, inoculation of corn forage with a combination of L. buchneri 6138 and L.

paracasei tolerans 6135 at ensiling improved the aerobic stability by up to 128 h and improved

NDF digestibility by 7% relative to the control. In another study, Kang et al. (2009) evaluated

the effects of an esterase-producing inoculant containing 1.0 × 104 cfu/g of L. casei strain

PTA6135 and 1.0 × 105 cfu/g of L. buchneri strain PTA6138 on two corn silage hybrids

(Croplan Genetics 851RR2 and Vigoro 61R36). The authors reported that inoculation of the first

hybrid decreased lactate to acetate ratio, increased propionate concentration, improved the

aerobic stability, and increased the potentially degradable DM fraction. However, inoculation of

the second hybrid did not affect fermentation, but decreased aerobic stability and increased total

degradable DM fraction, 24-h and 48-h DM digestibility, and 48-h NDF digestibility. These

results suggest that responses to these inoculants may also depend on properties of the forage

such as differences in their epiphytic bacterial population, water activity, water soluble

carbohydrates, and cell-wall component concentrations (McDonald et al., 1991; Kang et al.,

2009).

Inoculants Containing Saccharomyces Strains

Advancements in silage inoculant science that focus on delivering silage with probiotic

properties that enhance the performance and health of animals led to the development of fourth

generation of silage inoculants. These inoculants contain Saccharomyces cerevisiae yeast strains

27

that synergistically can improve silage fermentation, digestibility, and aerobic stability, and alter

microbial ecology within the gastrointestinal tract of ruminants to benefit health and/or

production efficiency. Some S. cerevisiae strains have been documented widely as direct-fed

microbials (DFM) that are effective at improving feed efficiency, decreasing ruminal acidosis,

and mitigating methane emissions (Desnoyers et al., 2009; McAllister et al., 2011). A recent

study examined if probiotic yeasts can persist in silage. Duniere et al. (2015) evaluated the

viability of three unnamed S. cerevisiae strains added at the rate of 1.0 × 103 cfu/g fresh weight

to whole-plant corn forage ensiled for 90 d. They confirmed survival of the inoculated strains

during ensiling and noted that they did not affect the nutritional quality or aerobic stability of the

silage. This study suggests that silage can be used as a vehicle to supply yeasts to cattle. A

recent study from our laboratory examined the effects of the dose (5.7 × 107 or 6.0 × 108

cfu/cow/day) and viability (live or killed) of a S. cerevisiae strain isolated from corn silage on

the ruminal fermentation and bacterial population of dairy cows (Jiang et al., 2017). Adding the

low dose of live yeast to the diet increased the relative abundance of ruminal cellulolytic

bacteria, such as Ruminococcus and Fibrobacter, and amylolytic bacteria, such as Ruminobacter

and Bifidobacterium and increased milk production by dairy cows. The results from the latter

two studies suggest that fourth generation inoculants containing Saccharomyces could be used to

deliver probiotics that enhance the performance and health of dairy cows. However, more studies

are needed to validate this theory.

Silage Microbial Community Analysis by Molecular Techniques

The microbial ecology of ensiling is complex and a better understanding of the silage

microbial community may improve silage fermentation and preservation. The microbial

population of silages has been described previously using culture-based methods, which

characterize microbial ecology using commercial growth media such as Luria–Bertani medium,

28

nutrient agar, and tryptic soy agar (Kirk et al. 2004). Culture-based techniques are relatively

quick and inexpensive (Kirk et al., 2004) but their main limitation is that they grossly

underestimate the bacterial diversity (Ercolini, 2004) because of the inability to cultivate 99% of

naturally-occurring species (Temmerman et al., 2003).

In recent years, molecular approaches have increased our understanding of the silage

microbial community (Rossi and Dellaglio, 2007; McEniry et al., 2008; Dolci et al., 2011).

These techniques have allowed detection and enumeration of strains that cannot grow on agar

and identified new lactic acid-producing bacterial species (Muck, 2013). Techniques that have

been used to study microbial communities in silage include length heterogeneity polymerase

chain reaction (PCR) (Brusetti et al., 2006), terminal restriction fragment length polymorphism

(McEniry et al., 2008), denaturing gradient gel electrophoresis (Parvin et al., 2010; Santos et al.,

2015), ion torrent sequencing platform (Kraut-Chen et al., 2016), and single molecule, real-time

sequencing technology (Bao et al., 2016). Most of these techniques utilize PCR to amplify a

portion, especially the 16S rRNA gene, of microbial DNA (Muck, 2013).

Terminal Restriction Fragment Length Polymorphism

Terminal restriction fragment length polymorphism (T-RFLP) is based on variations in

the DNA sequence present in PCR-amplified 16S rRNA genes (Smit et al. 1997). The PCR

products are digested with restriction enzymes and terminal restriction fragments (T-RFs) are

separated using a DNA sequencer (Thies, 2007; Rastogi and Sani, 2011). Microbial diversity is

estimated by analyzing the size, numbers, and peak heights of resulting T-RFs. Each T-RF

represents a single Operational Taxonomic Unit (OTU) (Rastogi and Sani, 2011). The limitation

of the T-RFLP method is that it underestimates community diversity because less than 100 bands

per gel can be resolved and different species can have the same T-RF length (Rastogi and Sani,

2011). McEnry et al. (2008) studied the temporal diversity of bacteria in grass silage using T-

29

RFLP technique, which revealed a reduction in the abundance of species with time (0, 2, 6, 14,

35, and 98 d of ensiling). However, identification of species was not possible because multiple

species share the same fragment length.

Length Heterogeneity Polymerase Chain Reaction

Length heterogeneity polymerase chain reaction (LH-PCR) utilizes the variation in the

length of a gene to differentiate microbial species (Mills et al., 2007; Muck, 2013). The

technique interrogates the hypervariable regions present in 16S rRNA genes. It utilizes a

fluorescent dye-labeled forward primer. A fluorescent internal size standard is run with each

sample to measure the amplicon lengths in base pairs (Rastogi and Sani, 2011). The area under

the peak in the electropherogram is used to indicate the relative abundance of that amplicon.

Unlike T-RFLP, one advantage of using LH-PCR is that it does not require any restriction

digestion and therefore PCR products are directly analyzed by a fluorescent detector (Rastogi

and Sani, 2011). The main limitation of the LH-PCR technique includes underestimation of

diversity as taxa may produce same-length amplicons, which cannot be differentiated by LH-

PCR (Mills et al. 2007). Brusetti et al. (2006) studied the development of various lactic acid

bacterial species during ensiling of whole-plant corn forage using LH-PCR and up to 58% of the

peaks could not be identified. In addition, species of Weissella and Enterobacter could not be

differentiated.

Denaturing Gradient Gel Electrophoresis

In this technique, PCR products are obtained from DNA using 16S rRNA primers and

separated on a polyacrylamide gel containing a linear gradient of DNA denaturant, such as a

mixture of urea and formamide (Muyzer et al. 1993). The movement of each DNA segment is

based on the composition of the DNA nucleotides rather than the length of the DNA sequence

(Muck, 2013). The main limitation is that Denaturing Gradient Gel Electrophoresis (DGGE)

30

underestimates the microbial diversity because not all sequence changes in 16S rDNA fragments

can be detected by DGGE, and partial 16S ribosomal DNA sequences differing by more than one

base pair can migrate to identical positions in DGGE gels (Jackson et al., 2000). Santos et al.

(2015) reported variations in the DGGE-based microbial structure of corn silages inoculated with

three strains of L. buchneri (UFLA SLM11, UFLA SLM103 and UFLA SLM108) by using

DGGE technique.

Next-generation Sequencing Methods

Deoxyribonucleic acid sequencing is a method that determines the order of occurrence of

nucleotides in a DNA strand (Raza and Ahmad, 2016). The first-generation sequencing method,

called Sanger sequencing, was developed by Edward Sanger in 1975. The basis of Sanger

sequencing is rapid DNA synthesis with DNA polymerase and the use of 2', 3'-dideoxy

nucleotide (ddNTP) as a specific inhibitor for the process (Sanger et al., 1977). Sanger

sequencing was considered the gold standard of DNA sequencing for several years (Grada and

Weinbrecht et al., 2013) because of its high quality, long read length and low error rate (Liu et

al., 2012). However, because of limitations in throughput and speed, Next-generation sequencing

(NGS) methods were developed to cater for high demand for cheaper and faster sequencing

technology (Hall, 2007; Raza and Ahmad, 2016). The NGS is a powerful biological tool that

provides greater through-put, and it is a cheaper and faster alternative to traditional Sanger

sequencing. Some of the existing NGS platforms include Roche 454 pyrosequencing, the Pacific

Biosciences real-time sequencing, ion torrent personal genome machine and Illumina MiSeq

platform.

Roche 454 sequencing pyrosequencing

Roche 454 was the first commercially available NGS that was first developed in 1996

(Ronaghi et al., 1996; Liu et al., 2012). This sequencer uses pyrosequencing technology which

31

relies on the detection of pyrophosphate released during nucleotide incorporation (Ronaghi et al.,

1996). Pyrosequencing is a commonly used DNA sequencing method because of its long read

length and fast sequencing speed (Liu et al., 2012). One of the limitations is the relatively high

error rate, particularly for poly-bases longer than 6 base pairs (Liu et al.,2012). Currently, no

studies have utilized this technique to analyze the microbial diversity of silage.

Single-molecule real-time (SMRT) sequencing

The SMRT sequencing platform was developed by Pacific Bioscience (Menlo Park, CA,

USA). This is based on detection of a fluorescent pulse in real time as sequencing proceeds

(Quail et al., 2012). Advantages of SMRT include a short read time (4 to 6 hours) and long read

length (average of 1300 base pairs). Also, it does not need the PCR preparation step, which

reduces bias and error caused by PCR (Liu et al., 2012). However, SMRT has lower throughput

than other NGS methods (Liu et al., 2012). Bao et al. (2016) applied the SMRT platform to

examine the microbial community composition of alfalfa silage because it generates much longer

sequence read lengths. The authors found that the relative abundance of bacterial species

reduced after fermentation, and Pediococcus acidilactici and L. plantarum were the most

prevalent species.

Ion torrent personal genome machine platform

This platform uses semiconductor sequencing technology (Quail et al., 2012). It is based

on detection of released protons during nucleotide base incorporation (Rothberg et al., 2011;

Quail et al., 2012). It is quick, inexpensive and involves a small-sized instrument (Quail et al.,

2012). The sample preparation time is less than 6 hours for 8 samples and it can read 200 base

pairs in 2 hours (Liu et al., 2012). A recent study that applied 16S ribosomal RNA gene

sequencing using the Ion Torrent Personal Genome Machine revealed that 99% of the sequence

32

reads detected were L. reuteri and L. pontis in corn, sorghum, and wheat silages (Kraut-Cohen et

al., 2016).

Illumina MiSeq platform.

The Illumina sequencing platform allows for deeper sequencing at much lower costs than

other sequencers (Degnan and Ochman, 2012) and is currently one of the most popular methods

for microbial community profiling. It integrates all steps of sequencing together in the Illumina

MiSeq desktop. The process from sample preparation to analyzed data can be completed within 8

hours (Degnan and Ochman, 2012; Liu et al., 2012). Illumina MiSeq has great advantages over

other sequencers in terms of the cost to sequence 1 megabase pair (Quail et al., 2012). Though,

Illumina Miseq platform generates short read length (150 bases, Quail et al., 2012), low error

rate (0.80%, Quail et al., 2012) and high throughput (55.5 megabase pairs/h, Loman et al., 2012)

makes it one of the most accurate and widely used sequencers (Degnan and Ochman, 2012).

Currently, no studies have utilized this technique to analyze the microbial diversity of silage.

Therefore, one of the objectives of this dissertation was to examine the diversity of silage

bacteria in corn and alfalfa silages via Illumina Miseq sequencing in order to better characterize

the complex microbial community of silages and to suggest roles of unknown silage bacteria by

investigating if their abundance correlates with silage quality indices.

Pathogenic Escherichia coli

Escherichia coli is a gram-negative, facultative anaerobic bacterium that accounts for

approximately 1% of the gut microbial population in mammals (Winfield and Groisman, 2003;

Callaway et al. 2009; Tchaptchet and Hansen, 2011). Most strains of E. coli are commensal

organisms. However, some strains, often referred to as intestinal pathogens, cause diarrhea and

associated diseases (Robins-Browne et al., 2016). Under normal conditions, out of 8 - 9 L of

fluid presented to the intestine daily from intake and other metabolic processes in the body,

33

approximately 200 mL are excreted in the stool (Hodges and Ravinder, 2010). These enteric

pathogens can alter this balance leading to greater excretion of fluid in the stool (diarrheal

disease) (Hodges and Ravinder, 2010). According to Farrokh et al. (2013), the six types of

pathogenic E. coli, based on their virulence genes and phenotypic characteristics, are STEC (also

called verocytotoxin-producing E. coli), enteropathogenic E. coli (EPEC), enterotoxigenic E. coli

(ETEC), enteroaggregative E. coli (EAEC), enteroinvasive E. coli (EIEC), and diffusely adherent

E. coli (DAEC). The STEC strains possess a phage-encoded potent cytotoxin, called Shiga toxin,

as the main virulence factor (Nataro and Kaper, 1998). In North America, Japan and parts of

Europe, the most commonly identified serotype is O157:H7, a strain belonging to the STEC

group (Farrokh et al., 2013; Centers for Disease Control and Prevention (CDC), 2015), which is

a zoonotic pathogen of significant worldwide public health concern (Chase-Topping et al.,

2008). The high virulence of STEC serotype is because of its ability to resist stress conditions,

such as low pH, and its low infectious dose (Viazis and Diez-Gonzalez, 2011; Jafari et al., 2013).

This pathogen was first discovered in 1977 (Konowalchuk et al., 1977) and was first detected as

a food-borne pathogen in 1982 (Riley et al., 1983). It has been implicated in several illness

outbreaks in more than 50 countries including the US (Chase-Topping et al., 2008). Escherichia

coli O157:H7 causes gastrointestinal illness, leading to bloody diarrhea, haemolytic uremic

syndrome (HUS), renal failure, and death, especially among children and elderly people (Siegler,

2003; Thorpe, 2004; Razzaq, 2006). Escherichia coli O157:H7 is responsible for more than 60

deaths and 70,000 illness cases in the US annually (Mead et al., 1999) and the annual cost to the

economy is approximately $1 billion (USDA-ERS, 2001). According to the CDC, two multistate

outbreaks of STEC were reported in the US in 2016.

34

Classification of EPEC is based on the presence of specific virulence genes known as the

locus of enterocyte effacement (LEE) pathogenicity island (Croxen et al., 2013). Expression of

LEE is associated with attaching and effacing lesions that enable the attachment of bacteria to

the intestinal epithelial cells, mediated by a protein called intimin, which is encoded by the eae

gene, type 3 protein secretory system and Tir protein (Jafari et al., 2013; Ingle et al., 2016). The

exact pathogenesis of these bacteria is not yet fully understood, and may involve factors other

than attaching and effacing lesions (Jafari et al., 2013; Ingle et al., 2016).

Enterotoxigenic E. coli strains attach to intestinal epithelial cells via proteinaceous

surface structures called colonization factors (Jafari et al., 2013). Following attachment and

colonization, ETEC strains cause diarrhea by producing enterotoxins. Strains of ETEC are the

leading causes of diarrhea or cholera-like disease in all age groups in areas with poor sanitation

and inadequate clean water, especially in developing countries (Jafari et al., 2013).

Enteroaggregative E. coli is a newly discovered pathotype that has been associated with

diarrhea in children and adults in developing countries (Robins-Browne et al., 2016). The EAEC

strains colonize the intestinal mucosa via aggregative adherence fimbrae. However, the

virulence-associated determinants of EAEC are not yet known and therefore, there is no clear

definition of EAEC strains (Croxen et al., 2013; Robins-Browne et al., 2016).

Diffusely adherent E. coli is a heterogenous group of strains that originally were

identified by their adherence to tissue culture cells (Nataro et al., 1987). They possess an

autotransporter protein that generates a diffuse adherence pattern (Jafari et al., 2013). However,

Robins-Browne et al. (2016) reported that ingestion of up to 1010 cfu of two DAEC strains failed

to cause diarrhea in humans, thereby casting doubts on the role of DAEC in diarrhea.

35

Adherent-Invasive E. coli strains are not associated with diarrhea and are believed to be

associated with the development of Crohn’s disease because they are isolated more commonly

from patients with the disease than from healthy people. However, it is not yet known whether

AIEC strains contribute to the pathogenesis of Crohn’s disease (Robins-Browne et al., 2016).

Cattle as a Reservoir of Escherichia coli O157:H7

Cattle are considered to be the main reservoir of EC and the main source from which this

pathogen enters the food chain (Callaway et al., 2009). In cattle, Escherichia coli represent less

than 1% of the ruminal microbial population (Min et al., 2007) because of the competition with

other ruminal microbes and the bactericidal effect of the volatile fatty acids produced in the

rumen (Wallace et al., 1989). However, the high pH and low volatile fatty acid concentrations of

the lower gut, particularly the colon, are favorable for E. coli; however, the terminal end of the

colon and the lymphoid tissue of the recto-anal junction are the major primary sites of EC

colonization (Lim et al., 2007). Cattle are asymptomatic carriers of EC because they lack

Globotriaosylceramides (Gb3) and Globotetraosylceramides (Gb4), the vascular receptors for

Shiga toxins (Samuel et al., 1990; Obrig et al. 1993).

Reinstein et al. (2007) reported that the fecal prevalence of EC in beef cattle was 7.1% in

a study with 933 cows that were delivered to two commercial US abbatoirs for slaughter.

However, EC was isolated from 13.5% of the fecal samples collected from cows in five

commercial dairies (Stanford et al., 2005). In fact, the population of EC in wet feces ranged from

101 to 109 cfu/g (Stephens et al., 2009; Munns et al., 2015). Cattle that shed at least 104 cfu/g of

wet feces are termed super-shedders (Matthews et al., 2006a, b). The prevalence of super-

shedders is dependent on several factors, such as size and method of sample collection (fecal or

recto-anal swab sampling), sampling time (summer versus winter), source of measure (farm, pen,

truck, or animal), and method of enumeration (Munns et al., 2015). Up to 7% of a given

36

population of beef cattle are super-shedders (Cobbold et al., 2007; Stanford et al., 2012) that can

be significant contributors to transmission of EC in their environments. Up to 90% of EC

shedding within feedlot pens was linked to super-shedders (Omisakin et al., 2003; Chase-

Topping et al., 2008; Stephens et al., 2009). Therefore, research towards reducing outbreaks of

EC has focused on pre-harvest control of the pathogen by inhibiting the growth and viability of

super-shedders (Matthews et al., 2006a).

Pathogenic Escherichia coli in Ensiled Forages

Dairy and beef producers are more concerned about the nutritive value and occasionally

the toxin concentration of feeds and forages than on their pathogen load as the former are more

likely to directly affect animal performance (Diaz and Trujillo, 2007). However, feeds and

forages also can play a role in disease transmission, or as a source of disease agents (Diaz and

Trujillo, 2007). Feeds, especially ensiled forages, can harbor pathogens that can compromise the

health and performance of animals and humans and cause fatalities (Driehuis, 2013). The

following section summarizes the current scientific knowledge on the occurrence and control of

pathogenic E. coli in ensiled forages.

Species of Enterobacteriaceae, such as EC, are part of the epiphytic microbial population

of most forage crops (Driehuis, 2013) and they compete with lactic acid bacteria for nutrients

during the early phase of ensiling. However, as fermentation progresses, the abundance of E. coli

is reduced because their survival decreases at low silage pH (< 4.0). A rapid drop in pH was

shown to eliminate E. coli in corn silage (Bach et al., 2002; Byrne et al., 2002). Pedroso et al.

(2010) evaluated the effectiveness of 3 commercial bacterial inoculants at controlling EC in corn

silages. The pathogen was eliminated within 3 d of ensiling with or without silage inoculation

when the pH dropped below 4.0. In a similar study, EC was eliminated from ensiled, artificially

contaminated wheat and corn forages when the pH dropped below 5.0 (Chen et al., 2005).

37

Similar findings were observed in corn silage contaminated with E. coli O26, a different strain of

STEC (Duniere et al., 2011). The elimination of this pathogen in these studies was caused by the

inhibitory low pH and/or the enhanced antimicrobial activities of organic acids at low pH

(Bjornsdottir et al., 2006).

Chen et al. (2005) reported that inoculated E. coli survived longer in wilted versus

unwilted wheat and corn silages because the pH of the former decreased more slowly than the

latter. In theory, pathogens like E. coli will persist and survive for longer periods in ensiled

forages with high buffering capacity, like alfalfa silage, because of slow decline in pH. However,

no published studies on the persistence or survival of pathogens in legume or high buffering

capacity silages were found. Consequently, one of the objectives of this dissertation was to

examine the survival of EC in alfalfa silage and to determine if microbial inoculants can inhibit

the survival of EC in alfalfa silage, thereby reducing its cycling on farms.

Laboratory mini silos have a more controlled environment than farm silos which are

more prone to air penetration and dirt contamination (Johnson et al., 2005). Studies have shown

that the presence of oxygen prolongs the survival of pathogenic E. coli such that it can survive

the ensiling phase (Duniere et al., 2011; Driehuis, 2013) and grow during the feedout stage when

the pH increases (Duniere et al., 2011). For instance, up to 107 cfu/g of E. coli were found in the

shoulders of commercial wheat and corn silages stored in bunker silos because the shoulders of

such silages are the most susceptible sites for air penetration, subsequent less acidic pH values

and greater spoilage (Weinberg et al. 2004). Pedroso et al. (2010) monitored the survival of EC

in laboratory corn silage samples aerobically exposed for 7 days and then inoculated with the

pathogen to mimic its survival during the feedout stage on farms. Up to 6 log10 cfu/g of the

pathogen were detected after 24 hours of inoculation in all silages except those inoculated with

38

L. buchneri alone or L. buchneri and P. pentosaceus (1.97 log10 cfu/g or none, respectively) at

ensiling, which maintained a lower pH than the control silage (3.91 or 3.66 vs. 4.71).

Other pathogens that occur in silage include Listeria monocytogenes, Clostridium

botulinum, Neospora caninum, Bacillus cereus, and Salmonella organisms. These organisms

cause listeriosis, botulism, neosporosis, emesis, and salmonellosis, respectively in animals. Just

like EC, proper silage management practices to ensure effective fermentation, and minimization

of contamination with soil, manure, and animal carcasses will effectively control the occurrence

of these agents on farms. The major sources of most of these pathogenic organisms are either

dirt, manure and carcass contamination, or improperly managed silage, which create conducive

conditions (high pH and aerobic condition) for the development of pathogens (Vissers et al.

2006; Driehuis, 2013). Like E. coli, L. monocytogenes, C. botulinum, N. caninum, B. Cereus, and

Salmonella require oxygen and high pH for growth and are therefore likely to persist and grow in

aerobically-deteriorating silage (Weinberg et al., 2004; Vissers et al. 2006; Driehuis, 2013).

Control of Fecal Shedding of EC by Ruminants

Research studies have focused mainly on pre-harvest control of the EC because of the

severity of infections in humans. The following approaches have been employed to reduce

shedding of EC in ruminants:

Diet Change

Changing from a grain to a forage diet has affected fecal shedding of EC (Callaway et al.,

2009). Kudva et al. (1995) reported an increase in the number of sheep shedding EC when the

diet was suddenly switched from alfalfa pellets to a sagebrush grass (Artemisia tridentate). In

another study, cattle fed a corn-soybean-based ration shed more generic E. coli (108 vs. 105

cfu/g) than those fed medium-quality timothy hay (14% crude protein, 40% NDF) (Diez-

Gonzalez et al., 1998). These authors suggested that switching the diet of feedlot cattle from a

39

grain-based diet to a hay-based diet a few days before slaughter might help reduce carcass

contamination in the abattoir. However, this approach reduces the final bodyweight of cattle and

thus might not be economical, particularly in feedlot cattle (Keen et al., 1999; Stanton and

Schutz, 2000). When cattle were switched from an 85% corn diet to a diet containing 30% millet

hay and 62% corn 5 days before slaughter, they lost an average of 1 kg/head/day but had fewer

(1 × 106 vs. 3.2 × 107) total generic E. coli counts in the feces (Stanton and Schutz, 2000).

Similarly, Keen et al. (1999) reported a decrease in body weight (0.6 kg/head/day) and a

reduction (18 vs. 52%) in the proportion of cattle shedding EC when they were abruptly switched

from a diet of corn grain to alfalfa hay. The food safety benefits in these studies should be

weighed against potential reduced income from lighter carcasses. However, food safety concerns

may be more important because slaughter houses and farms can be shut down if they are found

to be the source of EC outbreaks (USDA, 1998).

The effects of changes in diet on EC counts may be attributed to the change in microbial

population caused by the altered availability and concentrations of fermentable substrates in the

lower intestinal tract (Callaway et al., 2009). The greater abundance of EC in feces with grain-

based diets is probably because of higher starch supply from the rumen to the colon. A switch

from a grain-based to hay-based diet shifts the site of digestion from the colon to the rumen,

which leads to reduced starch availability in the colon (Callaway et al., 2009). It is suggested that

this change in nutrient availability causes a shift in the microbial population of the gut. In

particular, organisms that are dominant on hay diets, such as Enterococcus, are thought to

competitively exclude (Gregory et al., 2000), displace and prevent colonization by EC (Callaway

and Martin, 2006). Furthermore, tannins and lignin in forages can reduce fecal shedding of

generic E. coli and kill EC in vitro and in manure (Wells et al., 2005; Min et al., 2007).

40

Vaccinations

Vaccination is a recent approach that reduces fecal shedding of EC (Snedeker et al.,

2011). Escherichia coli O157:H7 vaccines exploit the antigenic components important for

bacterial adherence to the intestinal epithelium or virulence of the pathogen, such as type III

secreted proteins, intimin protein, and the O157 lipopolysaccharide (Dean-Nystrom et al., 2002;

Potter et al., 2004). Siderophore receptor and porin (SRP) protein-based vaccine is a

commercially available vaccine that uses SRP proteins as antigens so immunized animals

produce anti-SRP antibodies that block iron transport by binding to the outer membrane proteins

of bacterial cells (Cull et al., 2012). Blocking iron transport reduces the survival of the gram-

negative bacteria, such as EC, in a mixed microbial environment (Cull et al., 2012). The SRP-

based vaccine reduced the prevalence of EC, fecal shedding of the pathogen, and the number of

high-shedders in feedlot cattle (Fox et al., 2009). In a similar study, vaccinated cattle had less

prevalence of EC and less prevalence of high shedders relative to the unvaccinated cattle (Cull et

al., 2012). However, despite the proven effectiveness of vaccines, their high cost has hindered

the adoption of this approach by producers.

Direct-Fed Microbials

Direct-fed microbials (DFM) are utilized primarily to increase feed efficiency in feedlot

cattle, enhance milk production in dairy cows, and improve health and performance of young

calves (Fuller, 1989; Krehbiel et al., 2003). Some DFM have reduced shedding of EC by

ruminants (Lema et al., 2001; Ohya et al., 2001). However, the method of monitoring the EC can

affect the results. Bach et al. (2003) reported that a Saccharomyces cerevisiae DFM culture

reduced counts of EC in a batch culture but not in a continuous flow culture system (Rusitec

apparatus). This was because the populations of EC in the continuous culture system declined

41

steadily from the onset of inoculation, perhaps because of the high dilution rate of the system

(0.32 mL/min).

In recent years, probiotic products that are aimed specifically at reducing fecal shedding

of EC have been developed (Callaway et al., 2009). A microbial culture containing L.

acidophilus alone or in combination with P. freudenreichii reduced fecal shedding of EC in

cattle (Elam et al., 2003; Younts-Dahl et al., 2004). Microbial products containing Streptococcus

faecium alone or in combination with L. acidophilus, L. casei, L. fermentum, and L. plantarum

were effective, whereas L. acidophilus alone was not effective at reducing fecal shedding of EC

in sheep (Lema et al., 2001). Direct-fed microbials might be an effective strategy for reducing

the abundance of EC when they improve the performance of ruminants, which can economically

justify their inclusion in rations (Cull et al., 2012). However, animal responses to DFM have

been inconsistent because of differences in the types, strains, and combinations of organisms that

have been used (Kenney et al., 2015), or lack of understanding of the exact modes of action of

many DFM.

Bacteriophage Therapy

Some viruses, called bacteriophages, can lyse and kill bacteria commonly found in the

gut of food-producing animals and environments (Klieve and Swain, 1993). Lysis of bacterial

cells can occur because of phage replication in the host cell. The bacteriophages release a

hydrophobic polypeptide that forms pores in the cell membrane, thus, provoking the lysis of the

bacterial cell (Andreoletti et al., 2009). Lysis also can occur in the absence of phage replication.

In this case, phages act like DNA or RNA viruses that inject their genetic material into the

bacterium, therefore disrupting bacterial expression of genes and proteins, resulting in impaired

metabolism (Andreoletti et al., 2009). Some studies have shown that bacteriophages specific for

EC reduced shedding of the pathogen in cattle (Callaway et al., 2008; Niu et al., 2009). However,

42

regulatory approval is needed for use of phage therapies (LeJeune and Wetzel, 2007), partly

because long term exposure to a specific phage preparation can result in bacterial resistance

(Jassim and Limoges, 2014; Kateryna and Rai, 2016). Therefore, phage therapy requires clinical

testing of every component of the phage preparation (Kateryna and Rai, 2016). There is currently

no regulatory approval required for the use of phages to control pathogenic organisms in live

food animals. However, the US Food and Drug Administration (FDA) recently approved the use

of a Listeria-specific bacteriophage preparation on Ready-to-Eat meat and poultry products

(FDA, 2014).

Mycotoxins in Ensiled Forages

Mycotoxins are a group of secondary metabolites secreted by fungal organisms mostly

belonging to genera Aspergillus, Fusarium, Claviceps, and Penicillium (Yiannikouris and

Jouany, 2002; CAST, 2003; Kabak et al., 2006). Fungi often thrive well in environments with

high humidity, high temperature, and oxygen access at all stages of plant production and material

storage (Egal et al., 2005). Toxic effects, such as reduced feed intake and milk production,

reproductive problems, immunosuppression, and death can occur when animals are fed

mycotoxin-contaminated diets (Jouany, 2001; Whitlow and Hagler, 2005).

Approximately 25,000 to 166,000 new cases of liver cancer are annually caused by

aflatoxins (Liu and Wu, 2010). The ubiquitous nature of mycotoxins and the severity of their

effects on human health make them a major food safety concern. In addition to the severity of

their effects on livestock and human health, direct costs of disposal of condemned food and feed

ingredients, and indirect costs of regulatory enforcement and quality control measures caused by

fungal toxin contamination in the USA were estimated at approximately $1 billion (CAST, 2003;

Milicevic et al., 2010).

43

Prevalence of Mycotoxins in Foods and Feeds

The Food and Agriculture Organization estimates that 25% of all crops worldwide are

contaminated with mycotoxins annually (CAST, 1989). In addition, Pittet (1998) reported that

40% of 25,000 food and feed samples collected in 30 countries around the world were

contaminated with one or more mycotoxins. A three-year (2009 to 2011) survey on the

worldwide occurrence of mycotoxins revealed that 81% of 7,049 livestock feed samples

collected from the Americas, Europe and Asia were positive for at least one mycotoxin

(Rodrigues and Naehrer, 2012). Mycotoxins are present in concentrates, forages and in preserved

feeds such as hay and silage (Biomin, 2015). Most research studies have focused primarily on

mycotoxin occurrence in grains and cereals (Binder, 2007; Zinedine et al., 2007; Bhat et al.,

2010), perhaps caused by their greater consumption by humans than other parts of forages. Other

studies have focused on exposure of cows to mycotoxins via forages (Boysen et al., 2000;

O’Brien et al., 2006) because forages account for a high proportion of the ruminant diet (Cheli et

al., 2013). A survey by Driehuis et al. (2008) revealed that the contribution of ensiled forage to

mycotoxin ingestion in dairy cows is approximately three times more than that of finished feed.

Factors that Predispose to Mold and Mycotoxin Production in Silage

Ensiled forages may contain a mixture of mycotoxins, originating from pre-harvest

contamination by Fusarium species (Baath et al., 1990), and/or from post-harvest contamination

with toxigenic molds that are common in silage, such as Aspergillus, Monascus, Rhizopus, and

Geotrichum (Boysen et al., 2000; Garon et al., 2006; Fink-Gremmels, 2008). Anaerobic

conditions and low pH are required to restrict the growth of these molds during ensiling and low

pH after ensiling curtails their resuscitation after aerobic exposure. Except for Penicillium,

typical silage molds, such as Fusarium, Aspergillus, Monascus, Rhizopus, and Geotrichum

species are intolerant of low oxygen and low pH conditions (Pahlow et al., 2003). Therefore,

44

during ensiling, molds are often found at the top layer or sides of silos that are commonly prone

to oxygen infiltration. In agreement, Gonzalez-Pereyra et al. (2008) noted that improperly

managed silage promotes aerobic conditions and less extensive fermentation that favors the

growth of toxigenic molds that are normally less tolerant of acidic or anaerobic conditions. Once

silos are opened, the aerobic conditions allow the growth of lactate-utilizing yeasts that

metabolize lactate to CO2 and thereby, increase the pH. The increased pH and exposure to

oxygen allow toxigenic fungi to grow aggressively during the feed-out phase (McDonald et al.,

1991), particularly in poorly-managed silages.

Although the most notorious negative effect of molds is the secretion of mycotoxins,

feeding moldy feeds has been reported to expose animals to respiratory problems, decreased

reproductive function, kidney damage, and skin and eye irritation (Di Costanzo et al., 1995;

Scudamore and Livesey, 1998; Alm et al., 2002). In fact, Aspergillus fumigatus has been

proposed to be associated with mycotic hemorrhagic bowel syndrome, a fatal intestinal disease

of adult dairy cows in the US (Puntenney et al., 2003). Many common molds do not produce

mycotoxins, and the presence of molds in silage does not indicate the presence of mycotoxins

nor does their absence confirm mycotoxins are absent (Zain, 2011).

Temperature and insect activity are the major factors influencing mycotoxin

contamination of feedstuffs (Coulombe, 1993). Molds can grow at a temperature range of 10° to

40°C, a pH range of 4 to 8, and above 0.7 aw (Whitlow, 2003). In wet feeds, such as silage, high

moisture levels allow mold growth if oxygen is available (Whitlow, 2003). Delayed harvesting,

slow or delayed filing of silos, inadequate packing and sealing of silos, slow feedout rates,

bridging in silage bags, and damaged plastic wrap, bags, or silo covers can create a conducive

microclimate for mold proliferation and mycotoxin production. (Whitlow, 1993). Other factors

45

that may predispose to mold growth and mycotoxin production in feeds include rodent, rain, hail,

drought, and lodging damage.

Types of Mycotoxins in Ensiled Forages

More than 400 mycotoxins occur naturally, however, only a few have been studied

extensively (Whitlow and Hagler, 2005). Mycotoxins that are frequently present in ensiled

forages include deoxynivanenol, fumonisin, aflatoxin, ochratoxin A, zearalenone, T2-toxin,

ergot, and roquefortine C (Driehuis et al., 2008; Whitlow, 2009).

Fusarium Toxins

Fusarium species are field fungi because they proliferate in the field during plant growth

(Christensen et al., 1977; Storm et al., 2008). Their activities are favored at high humidity

(>70%) and temperatures that fluctuate between hot days and cool nights (CAST, 2003).

Ensiling conditions are not favorable for Fusarium species, as they do not tolerate low pH and

anaerobic conditions. Nevertheless, Fusarium toxins can tolerate ensiling (Lepom et al., 1990).

Hence, concentrations of Fusarium toxins detected in silage usually reflect the contamination

levels at the time of harvesting.

Deoxynivalenol

Deoxynivalenol (DON) is a toxin produced by Fusarium species such as F.

graminearum, F. sporotrichioides, F. culmorum, F. poae, F. roseum, and F. tricinctum. Cold,

wet periods followed by a short, dry period favor their secretion (Diekman and Green, 1992).

The toxin also is prevalent when wet conditions coincide with warm days and cool nights (De

Wolf et al., 2005). Deoxynivalenol also is known as vomitoxin because it is associated with

vomiting in swine, as well as feed refusal, diarrhea, reproductive problems, and eventual death

(Canady et al., 2001).

46

Deoxynivalenol is the most commonly detected mycotoxin in preserved forages at very

high concentrations (Rotter et al., 1996). A three-year survey of 7,049 livestock feed samples to

estimate the worldwide occurrence of mycotoxins revealed that DON represents one of the most

frequent threats to livestock with a prevalence rate of 59% and an average contamination level of

1,104 µg/kg (Rodrigues and Naehrer, 2012). In another survey on the presence of mycotoxins in

feed ingredients in the Netherlands (Driehuis et al., 2008), corn and wheat silages were found to

be the main sources of DON in diets of dairy cattle, with average concentrations of 854 and 621

μg/kg, respectively, and maximum concentrations 3,142 and 1,165 μg/kg, respectively.

Ruminants are relatively resistant to DON (Pestka, 2007), because rumen

microorganisms can extensively convert DON to non-toxic, de-epoxy DON (Swanson et al.,

1987; Marczuk et al., 2012). Charmley et al. (1993) reported only a numerical loss in milk yield

(- 1.4 kg/d) in first lactation dairy cows consuming a DON-contaminated diet (2.6 – 6.5 mg/kg).

Likewise, no reduction in milk production was observed when DON was fed to dairy cows at 66

mg/kg or 6.4 mg/kg for 5 days or 6 weeks, respectively (Trenholm et al., 1985; Cote et al.,

1986). A similar result was obtained by Korosteleva et al. (2009) who reported no reduction in

milk production by dairy cows consuming diets contaminated with 2.6 or 6.5 mg/kg of DON.

However, altered rumen fermentation and reduced protein flow to the duodenum (Danicke et al.,

2005; Seeling et al., 2005) were observed in dairy cows when 8.21 mg/kg DM of DON and 0.09

mg/kg of ZEA were fed daily for 4 weeks, possibly because of the synergistic toxic effects of

ZEA and DON (Grenier and Oswald, 2011).

The guidance level for DON in ruminant feeds in Western Europe is 5 mg/kg DM

(European Commission, 2006) and the advisory guidelines stipulated by the US FDA are 5 and

10 mg/kg of diet DM for dairy and beef cattle, respectively (FDA, 2011).

47

Fumonisins

Fumonisin B1 and B2 are the most important of a group of at least 28 silage toxins

produced by two Fusarium species, F. verticillioides and F. prolferatum and a strain of F.

nygamai (Marasas, 1996; Whitlow and Hager, 2005). Hot, dry periods followed by humid

conditions and insect damage are the major predisposing factors that favor the secretion of

fumonisins by Fusarium species (DeWolf et al., 2005). Fumonisin represents the most frequent

threat to livestock among mycotoxins with a 64% prevalence rate and an average concentration

of 1,965 µg/kg in 7,049 livestock feed samples collected from the Americas, Europe, and Asia

(Rodrigues and Naehrer, 2012). Gonzalez-Pereyra et al. (2008) reported that fumonisin levels in

corn silage varied from 340 to 2,490 µg/kg of DM, with higher concentrations detected in

samples taken from the top layer and side walls of the silo, which are normally prone to air

infiltration and less extensive fermentation.

The structural formula of fumonisin is similar to sphingosine, which is a component of

sphingolipids found at high concentrations in nerve tissues (Soriano et al., 2005). Fumonisin

interrupts sphingolipid biosynthesis (Diaz and Boermans, 1994), thereby, causing

leucoencephalomalacia, also known as moldy corn poisoning, in horses (Kellerman et al. 1990),

pulmonary edema in swine (Colvin and Harrison 1992), and hepatotoxicity in rats, mice and

rabbits (Gumprecht et al., 1995; Voss et al., 1995). Because it is rumen degradable (60 to 90%),

the toxin tends to be less potent in ruminants (WHO, 2000); however, fumonisin has been

nephrotoxic to calves fed 1 mg/kg BWof the toxin (Mathur et al., 2001). Similar results were

observed in beef calves supplemented with 148 mg/kg of total fumonisin in the diet for 31 d

(Osweiler et al., 1993). Diaz et al. (2000) reported lower milk yield in Holstein and Jersey cows

fed diets containing 100 mg/kg of fumonisin from 7 days before calving to 70 d after parturition.

Nevertheless, carry-over of the toxin into milk is believed to be negligible (Scott et al., 1994),

48

and hence the toxin poses no serious food safety concern for people. The US FDA advisory

guideline for fumonisin is 30 mg/kg of diet DM for lactating dairy cows, 30 mg/kg of diet DM

for breeding ruminants, and 60 mg/kg of diet DM for calves over 3 months that are intended for

slaughter (FDA, 2011).

Zearalenone

Zearalenone (ZEA) is an estrogenic metabolite produced by several species of Fusarium

such as F. graminearum, F. culmorum, and F. crookwellense (Saeger et al., 2003). Fusarium

graminearum occurs naturally in high-moisture corn and it has also been detected in moldy hay

and pelleted feeds (CAST, 2003). It also co-occurs with DON in grain crops, like corn, barley

and sorghum at very low concentrations (CAST, 2003). High humidity conditions with

alternating low (11-14°C) and moderate (27°C) temperatures favor its production (Vigier et al.

1997; De Wolf et al., 2005).

Several studies have reported the occurrence of ZEA in ensiled forages. Whitlow and

Hagler (2005) reported an average ZEA concentration of 525 µg/kg of DM and 30% incidence of

ZEA in 461 corn silage samples in the US. Driehuis et al. (2008) reported ZEA contamination in

13 and 50% of grass and corn silage samples in Denmark with average concentrations of 180 and

146 µg/kg, respectively. Reed and Moore (2009) reported that sorghum and alfalfa silages

contained 660 µg/kg and 79.8 mg/kg of ZEA, respectively. Zearalenone was the most common

mycotoxin detected in a study that assessed the exposure of Danish cattle to mycotoxins from

maize silage (Storm et al., 2014). A recent survey reported that approximately 45% of 7,049

livestock feed samples collected from the Americas, Europe, and Asia contained ZEA, with an

average concentration of 233 µg/kg (Rodrigues and Naehrer, 2012).

The structural similarity of ZEA to estrogen enables the toxin to mimic the hormone

(Saeger et al., 2003). This causes numerous reproductive problems, including hyper-estrogenism,

49

vaginitis, and mammary gland enlargement, particularly in swine (Diekman and Green, 1992;

Marczuk et al., 2012). Ruminants are less susceptible than swine because ZEA is converted to its

hydroxyl-metabolites, α and β-zearalenol, by ruminal flora, predominantly by ruminal protozoa

(Kennedy et al., 1998). Although α-zearalenol has higher affinity for estrogen receptors than

ZEA, its lower absorption rate and or interconversion to β zearalenol in the liver decrease its

negative effects (Malekinejad et al., 2006; Fink-Gremmels, 2008). Nevertheless, the capacity of

ruminal microbes may be exceeded by high intakes of ZEA. Weaver et al. (1986) reported

reduced conception rates in dairy heifers fed 12.5 mg/kg of ZEA. Furthermore, depressed intake

and milk production, diarrhea, and reproductive failure were reported in heifers fed diets

contaminated with 660 µg/kg ZEA and 440 µg/kg DON (Coppock et al., 1990).

The major concern with ZEA toxicity is its negative effect on animal health and

reproduction because, like fumonisins, the extent of transfer into milk is negligible (Seeling et

al., 2005). Consequently, there are no FDA action limits, guidance, or advisory levels for ZEA.

However, the guidance value for ruminant feeds in Europe is 500 µg/kg (European Commission,

2006).

T-2 Toxin

T-2 is a potent Fusarium-produced mycotoxin (Saito and Okubo, 1974). It is more

prevalent in cereal grains than ensiled forages (Whitlow and Hagler, 2005) and is produced

during wet and warm conditions (Xu et al., 2014). Approximately 5% of 388 corn silage samples

collected from different countries contained 65 µg/kg of T-2 toxin (Biomin, 2015). The T-2 toxin

is degraded by ruminal flora to its less toxic metabolites (Whitlow, 1993), nevertheless, T-2 is

associated with gastroenteritis and intestinal hemorrhages (Hsu et al., 1972), immunosuppression

(Gentry et al., 1984; Black et al., 1992), and reduced performance and reproductive problems in

cattle (Kegl and Vanyi, 1991). A study noted that it is toxic to cattle at high intake level of 0.7 to

50

1.5 mg/kg (Li et al., 2011). Calves consuming 10 to 50 mg/kg of T-2 toxin experienced abomasal

ulcers and sloughing of the ruminal papillae. However, no death was recorded (Cheeke, 1998).

Guidelines for T-2 toxin have not been established by US FDA or European legislative agencies;

nevertheless, feeding levels close to the toxic dose of Li et al. (2011) is not recommended.

Aspergillus Toxins

Aspergillus molds are considered storage fungi because usually, they do not infect crops

before harvest. Some species, such as Aspergillus flavus may infect crops and produce aflatoxins

in the field (Wicklow, 1983; Storm et al., 2008) when temperatures are high (> 32°C) and during

drought stress (Whitlow and Hagler, 2005).

Aflatoxins

Aflatoxins are a group of toxic, mutagenic and carcinogenic compounds produced by

Aspergillus flavus, A. parasiticus, and A. nominus (Deiner et al., 1987; Kurtzman et al., 1987;

Creepy, 2002). High humidity (> 80%) and temperature (> 32°C), insect damage and drought

stress (DeWolf et al., 2005; Tsitsigiannis et al., 2012) are conducive conditions for aflatoxin

production. Natural forms of aflatoxins such as B1 and G1, and their dihydroderivatives, B2 and

G2, are naturally present in food and feed ingredients because of the ability of Aspergillus to

grow over a broad range of temperatures and humidities (Dutton, 1988; Phillips, 1999; Tulayakul

et al., 2005). Aflatoxin B1 (AFB1) is the major aflatoxin and the most potent natural carcinogen

produced by toxigenic Aspergillus (Squire, 1981). The incidence of aflatoxin is relatively low in

silages compared to other mycotoxins, probably because A. flavus and A. parasiticus are less

tolerant of acidic and anaerobic silage environments. Aflatoxins were present in 33% of 7049

livestock feed samples at an average concentration of 63 µg/kg (Rodrigues and Naehrer, 2012).

In a survey conducted in Denmark by Driehuis et al. (2008), no AFB1 was detected in 140 corn

silage samples. However, high concentrations of aflatoxin can occur in poorly made silages or in

51

silages made in hot, humid areas or with diseased corn plants (Gonzalez-Pereyra et al. 2008;

Queiroz et al., 2012). In addition, aflatoxins are more prevalent in trench silos because

Aspergillus is one of the main fungal contaminants in such silos (Gónzalez Pereyra et al., 2011).

Up to 51 µg/kg DM of AFB1 was detected in corn silage collected in a trench silo in France

(Richard et al., 2009). Furthermore, Gonzalez-Pereyra et al. (2008) reported an AFB1

concentration of 156 µg/kg of DM in corn silage stored in a trench type silo without proper

sealing.

Aflatoxins have been the most studied mycotoxin because of the severe public health

concern associated with their carcinogenicity (CAST, 2003). Aflatoxins do not have a lactone

ring and consequently, are relatively resistant to ruminal microbial degradation; moreover, low

concentrations can inhibit ruminal bacterial growth (Yiannikourisa and Jouany, 2002). Long

exposure to aflatoxins in diets of lactating cows caused adverse effects such as reduced health

and performance of cows, impaired liver function, (Garrett et al., 1968), suppressed immune

status and increased susceptibility to diseases despite vaccinations (Diekman and Green, 1992).

The symptoms associated with acute aflatoxin ingestion (aflatoxicosis) include inappetence,

ataxia, rough hair coat, and enlarged liver (Guthrie and Bedell, 1979; Whitlow and Hagler,

2005). Precise diagnosis of aflatoxicosis is difficult because the symptoms are not specific to the

condition (Coulombe, 1993). Increased liver weight and reduced performance were observed in

beef cattle fed a diet containing 100 µg/kg of aflatoxin (Garret et al., 1968; Patterson and

Anderson, 1982). Furthermore, reduced feed efficiency because of compromised ruminal

function was reported in steers fed 600 µg/kg (Diekman and Green, 1992). Queiroz et al. (2012)

noted decreased milk yield and milk protein concentration, as well as, altered immune response

in dairy cows fed 75 µg/kg of aflatoxin daily for 5 d. However, other studies reported no changes

52

in milk yield when 170 (Kutz et al., 2009) or 112 µg/kg (Stroud, 2006) of AFB1/kg of TMR DM

were fed for 7 or 11 d, respectively. This indicates that several factors, other than the amount

ingested, may influence the response of cows to the toxin; such factors may include heat or

acidotic stress, subclinical disease, etc.

After ingestion of contaminated feeds, a portion of the ingested aflatoxin B1 is degraded

by the ruminal microbes, resulting in the formation of aflatoxicol (Upadhaya et al., 2010). In

humans and susceptible animals, the toxin is metabolized by cytochrome P-450 enzymes into

aflatoxin-8,9-epoxide (highly toxic, mutagenic and carcinogenic), which causes chrosomosome

damage by binding with and forming adducts on DNA (Jouany et al., 2009). Other metabolic

fates of AFB1 in the liver include O-dealkylation to AFP1, ketoreduction to aflatoxicol, and

hydroxylation to AFM1 (highly toxic and carcinogenic), AFQ1, or AFB2a (Kuilman et al., 2000;

Wu et al., 2009; Jafarian-Dehkordi and Pourradi, 2013). Aflatoxin B1 is excreted in milk of dairy

cows as AFM1 (Diaz et al., 2004; Battacone et al., 2005), which represents a safety risk of milk

and milk products because both AFB1 and AFM1 are Group 1 human carcinogens (International

Agency for Research on Cancer, IARC, 2002). The rate of transmission of aflatoxin from feed

into milk can be as high as 6% (EFSA, 2004). Therefore, aflatoxin is the only mycotoxin that has

government legislation for its maximum permissible level in feedstuffs and milk products. The

Action Levels established by the US FDA are 0.5 and 20 µg/kg for fluid milk and feed

ingredients offered to dairy cattle, respectively (FDA, 2011), while the maximum level permitted

in Europe for fluid milk is 0.05 µg/kg (European Commission, 2006).

Penicillium Toxins

Penicillium molds can grow in typical silage environments with low water activity (0.79

– 0.83aw), low oxygen concentration (1%) and low pH (3.0 – 6.0) (Boysen et al., 2000; Nevarez

53

et al., 2009). Penicillium molds are more common during storage, but they also can grow on

plants under very wet conditions (Christensen et al., 1977).

Ochratoxin A

Ochratoxins are structurally-related secondary metabolites produced by Aspergillus

ochraceus, A. clavatus and Penicillium verrucosum, particularly when temperatures range

between 20 and 25°C. Ochratoxin A, which is carcinogenic, immunotoxic, and which inhibits

glucose metabolism, is considered the most toxic of the group (Whitlow, 1993; Yiannikourisa

and Jouany, 2002). However, this toxin is not currently classified as being carcinogenic to

humans. Veldman et al. (1992) reported 27% incidence of ochratoxin A in 89 randomly selected

feed ingredient samples, especially in grains, with an average concentration of 120 µg/kg.

Ochratoxin A is extensively and rapidly degraded in the rumen to a less toxic product,

ochratoxin-alpha (Hult et al., 1976; Fink-Gremmels, 2008), which explains tolerance of

ruminants to the toxin. However, it also may be metabolized to ochratoxin C, which is similarly

toxic (Chu, 1974). Bioavailability of dietary ochratoxin may be increased with high-grain diets,

probably because of acidotic conditions (Hohler et al., 1999). Furthermore, the detoxifying

capacity of the rumen may be exceeded at high intake, in which case the toxin can reduce milk

production and be transfered into milk (Whitlow, 1993; Yiannikourisa and Jouany, 2002).

Overall, the occurrence of ochratoxins in feed ingredients, especially in ensiled forages, is low

because the toxin is not stable in silos and it poses no significant threat to food safety and supply.

Mycophenolic acid and Roquefortine C

These are secondary metabolites of Penicillium roqueforti (Miller and Trenholm, 1994;

Schneweis et al., 2000). Penicillium roqueforti is commonly referred to as the silage mold

(Pahlow et al., 2003) because it is acid-tolerant and it can grow at low oxygen concentrations.

54

Consequently, it is the most frequently isolated Penicillium mold in silage (Sumarah et al.,

2005).

Roquefortine C and mycophenolic acid have been frequently detected in silage. In

Netherlands, Roquefortine C was detected in 19% of 47 corn and grass silage samples and 7% of

29 samples of ensiled by-products. Mycophenolic acid was detected in 13 and 10% of samples of

these feed classes, respectively. (Driehuis et al., 2008). A survey of silage samples in Germany

showed prevalence estimates of 56 and 32% of roquefortine C and mycophenolic acid,

respectively, (Schneweis et al., 2000).

Symptoms, such as reproductive disorders, mastitis, and lack of appetite were detected in

cattle herds fed silages containing 0.2 to 1.5 mg/kg of roquefortine C in Northern Germany

(Auerbach et al., 1998). Also, paralytic effects were reported in cows fed 4 to 8 mg/kg of

Roquefortine C (Haggblom, 1990). However, no clinical signs of intoxication were observed in

sheep fed 25 mg/kg of Roquefortin C for 18 days (Tuller et al., 1998), indicating that sheep are

more tolerant to Roquefortine C than cattle.

Mycophenolic acid is considered an immunosuppressive agent; it blocks the proliferative

response of T and B lymphocytes, and inhibits antibody formation and production of cytotoxic T

cells (Eugui et al., 1991). Research data on the effects of mycophenolic acid in cattle are lacking.

However, Mohr et al. (2007) found no effects in sheep exposed to 10 to 300 mg/kg daily for 44

days.

Ruminal Degradation of Mycotoxins

Ruminants are more protected from toxic effects of mycotoxins than monogastrics

because rumen microbes degrade or inactivate some of the toxic molecules (Pestka et al., 2007;

Cheli et al., 2013). However, the capacity of rumen detoxification can be saturated and it varies

with diet and or metabolic disease infestation (Fink-Gremmels, 2008). In addition, starch-

55

induced low ruminal pH (5.5 to 5.8) and high ruminal passage rates of high-producing cows may

greatly reduce microbial detoxification (Fink-Gremmels, 2008; Pantaya et al., 2016). Therefore,

some mycotoxins, such as ochratoxins and aflatoxins, can partially escape ruminal degradation

and be transferred to animal products, particularly milk, posing a severe health hazard to

humans.

Kiessling et al. (1984) investigated the ruminal degradation of different mycotoxins and

reported that whole rumen fluid degraded ZEA, T-2 toxin, and ochratoxin to their less toxic

metabolites (zearalenol, HT-2 toxin, ochratoxin alpha, respectively), but had no effect on AFB1

and DON. The authors also demonstrated that 90 to 100% of the metabolism of ochratoxin A,

ZEA, and T-2 toxin were achieved by the rumen protozoa and, therefore, they are considered

important agents of ruminal microbial mycotoxin biodegradation (Upadhaya et al., 2010).

However, some studies have demonstrated that ruminal bacteria also are involved in mycotoxin

degradation (Schatzmayr et al., 2006; Yang, 2010).

Co-occurrence of Mycotoxins in Livestock Feeds

Contamination of livestock feeds by multiple mycotoxins is a common occurrence on

farms because most fungi can produce several mycotoxins, and several fungi can simultaneously

contaminate feed commodities (Bottalico, 1998; Sweeney and Dobson, 1998; Alassane-Kpembi

et al., 2016). Therefore, studying the occurrence and effects of a single mycotoxin will more

likely give incomplete and/or false information about the associated risk of mycotoxin exposure.

An extensive literature review revealed that more than 75% of feed samples in Europe contain

one or more mycotoxins (Streit et al., 2012). In addition, almost half of 7049 animal feed

samples collected from the Americas, Asia, and Europe simultaneously contained at least two of

DON, aflatoxins, ZEA, fumonisins, and ochratoxin A (Rodrigues and Naehrer 2012). Aflatoxin

B1 was found to co-occur with ochratoxin A, ZEA, or the combination of both toxins in 31, 12,

56

or 27% of 123 Spanish barley samples, respectively (Ibáñez-Vea et al., 2012). In another study in

the Netherlands, Driehuis et al. (2008) reported that DON and ZEA co-occurred in 44% of dairy

diets, and corn silage was the major source of both mycotoxins in the feeds.

Synergistic negative effects of several toxins in the diet can worsen health and production

problems of cattle consuming mycotoxin-infested feeds. A meta-analytic study summarizing 112

publications reported negative synergistic or additive effects of different mycotoxins on animal

performance (Grenier and Oswald, 2011), which emphasizes the need to study co-occurrence of

mycotoxins in feeds to get a clearer picture of their adverse effects on the performance and

health of livestock.

Dietary Contribution of Corn Silage to Mycotoxin Intake in Cattle

Whole-plant corn silage is an important component of dairy and beef diets in the US,

Europe and some other parts of the world as approximately 120 million tons of corn silage are

produced and fed to ruminants annually in the US (USDA, 2016). Corn forage and silage can be

contaminated with mycotoxins in the field or during or after ensiling, respectively. The

percentage contribution of corn silage mycotoxins to total dietary intake of mycotoxins varies

widely among studies (Table 2-1). In the worst case scenario, the contributions may be as high as

81, 134, 107, 68, 23, and 270% of dietary concentrations for AFB1, ZEA, DON, fumonisin,

mycophenolic acid, and roquefortine C, respectively (Table 2-1). However, the actual range

varies widely because of factors such as location, sampling technique, method of mycotoxin

analysis, prevailing weather conditions, etc. Nevertheless, these data suggest that affordable

mycotoxin analysis should be conducted on silages to increase feed safety.

57

Preventing Mycotoxin Contamination of Feeds

Field or Pre-harvest Phase

Pre-harvest control of mycotoxins is the first step to ensure that feeds are safe. Mycotoxin

contamination can be reduced by minimizing environmental stress on the plant through

appropriate agronomic practices such as use of varieties or hybrids that are adapted to local

conditions and are naturally resistant to fungal infection, control of insect infestation, proper

management of crop residues and weeds, application of fungicides and pesticides, use of

irrigation, timely crop rotation, and proper fertilization (Edwards, 2004; Whitlow and Hagler,

2005). Insect management is the most important practice for reducing the risk of mycotoxins,

particularly fumonisins and aflatoxins, in temperate regions because they act as vectors of fungal

spores and insect damage predisposes crop plants, such as maize, to fungal infection (Munkvold,

2014).

The public health concern over the use of pesticides and fungicides as well as

development of resistance to fungicides by fungal organisms has led to development of bio-

agents to control mycotoxin contamination on the field (Tsitsigiannis et al., 2012). Application of

atoxigenic strains of A. flavus or A. parasiticus to crops is a successful approach that has been

used to reduce aflatoxin contamination of the field (Tsitsigiannis et al., 2010). Two

atoxigenic strains of A. flavus, AF36 and NRRL 21882, are widely used in the US to minimize

aflatoxin contamination of crops. An in-depth review on biological control of mycotoxins was

published by Tsitsigiannis et al. (2012) and is an excellent supplement to this section.

Harvest Phase

Careful harvest timing is important in controlling the level of mycotoxin contamination.

The risk of mycotoxin contamination is reduced significantly with earlier harvests (Jones et al.,

58

1981). However, harvest should be timed to ensure optimum yield, DM concentration, and

nutritive value.

The cutting height of the crop harvesters should be set to minimize soil contamination

because of the widespread occurrence of Fusarium spores in soil (Jean Pierre, 2007). In addition,

harvesting equipment should be properly maintained and calibrated to avoid physical damage to

crops at harvest because contact with soil and kernel damage predisposes forage to

contamination with fungal spores (Whitlow and Hagler, 2005). Furthermore, immediate storage

of harvested feeds is necessary to minimize the risk of mycotoxin contamination, because

delayed storage may expose the crop plants to temperature and humidity conditions that favor

continued growth of toxigenic fungi and mycotoxin contamination (Munkvold, 2014).

Ensiling Phase

Controlling mycotoxin production at the pre-harvest stage via physical, chemical, and

chemical methods is best; however, problems about feed safety, potential losses in quality of

commodities, limited efficacy and cost implications (Köhl et al., 2011) limit the success at this

stage (Köhl et al., 2011). Hence, mycotoxin control during storage is often critical.

Low silage pH and anaerobic conditions during ensiling inhibit the growth of spoilage

and toxigenic fungi, and reduce production of mycotoxins in silage (Pahlow et al., 2003). Plants

should be harvested at the recommended DM of 65 – 70% for pit silage and 45–50% for baled

silage, and silos should be filled rapidly, tightly packed and completely sealed to preserve

anaerobic conditions (Jouany, 2007). Cavallarin et al. (2011) reported that using an oxygen-

barrier plastic film for ensiling limits aflatoxin production. A similar result was reported by

Amaral et al. (2010). To reduce mold infestation and or mycotoxin contamination during feed--

out, aerobic stability-enhancing measures should be taken including maintaining a straight firm

59

silo face, feeding out the silage at a minimum rate of 10 to 16 cm per day, and feeding the silage

immediately after removal from the silo (Jouany, 2007).

The potential of silage lactic acid bacteria to mitigate aflatoxin contamination of silage

has been evaluated in vitro (Gourama and Bullerman, 1995; Ahlberg et al., 2015). Lactic acid

bacterial inoculants have been used successfully to enhance silage fermentation, increase aerobic

stability and reduce fungal populations in silages (Mari et al., 2009; Pedroso et al., 2010).

However, research reports on using additives to reduce forage aflatoxin contamination are

limited. Inoculation of harvested corn plant infested with southern rust with a mixture of

Pediococcus pentosaceus and L. buchneri at ensiling increased aerobic stability and prevented

production of aflatoxins (5.2 vs. 0 mg/kg) (Queiroz et al., 2012). In another study, L. buchneri

inoculation reduced aflatoxin concentration but had no effects on concentrations of DON or ZEA

in corn silage (Iglesias et al., 2005). Cavallarin et al. (2011) reported that limiting aerobic

spoilage by inoculating corn forage with L. buchneri at ensiling also reduced aflatoxin

production. Teller et al. (2012) studied the effects of additive application and damaging ears of

corn in the field before harvest on production of selected mycotoxins. Concentrations of DON

and fumonisins were increased in damaged ears but not in undamaged ears and inoculation with

L. buchneri 40788 (400,000 cfu/g of fresh forage) and Pediococcus pentosaceus (100,000 cfu/g),

or treatment with potassium sorbate, reduced mold and yeast counts and aerobic spoilage, but

had no effect on the concentration of DON, ZEA, and fumonisins in the corn silage. The main

deduction from these studies is that proper silage management and use of mold-inhibiting

additives or microbial inoculants can decrease aerobic spoilage and reduce the growth of

toxigenic fungi in silage (Kung et al., 2003), but these measures may not always reduce

concentrations of mycotoxins existing on plants before ensiling. In summary, good silage

60

management principles and practices at all stages of silage making and feeding are essential to

reduce mycotoxin contamination of silages.

Mycotoxin Detoxification Strategies

Strategies for detoxification of mycotoxins are aimed at destroying or reducing the

concentration of a mycotoxin, or making it non-toxic and or unavailable for absorption in the gut

of ruminants (CAST, 2003). Despite several efforts to control mycotoxins at pre and post-harvest

stages, frequent instances of feed contamination with mycotoxins occur, emphasizing the need

for effective detoxification of feed mycotoxins. Such measures should target detoxification of

silage mycotoxins because silage represents the bulk of dairy cattle diets in the US and many

other parts of the world, and it is prone to mycotoxin contamination particularly when poorly

managed.

Various mycotoxin decontamination strategies, such as manual removal, cleaning, or

milling of contaminated grains (Bata and Lastity, 1999; Siwela et al., 2005), thermal irradiation

(Rater and Matissek, 2008), ammoniation (Hogeboom et al., 2001), and ozonation (McKenzie,

1997) are considered effective; however, in addition to the high cost and safety issues, these

strategies are not suitable for ensiled forages probably because of their high volume. Dilution of

contaminated ingredients with others in rations is also an effective strategy for reducing the

ingestion of mycotoxins, though this does not involve detoxification of the toxin. Though

effective, it is not an ideal strategy for contaminated silage because it reduces the feed-out rate

which may promote further mold growth (Whitlow and Hagler, 2005) and mycotoxin production.

A feasible, safe, and relatively cost effective alternative strategy to reduce problems

associated with ingestion of mycotoxins is to decrease their bioavailability in the gut. This

involves inclusion of sequestering agents in the diets (CAST, 2003). The use of clay minerals as

sequestering agents in the feeds of livestock has been well documented (Stroud, 2006). Clay

61

minerals are fine-grained components of geological materials, occurring as particles with

a phyllosilicate structure that are formed usually as products of chemical weathering (Veldt and

Meniere, 2008). Clay minerals can hold water and dissolved nutrients because of the presence of

unbalanced electrical charges (Veldt and Meniere, 2008). Therefore, they can adsorb organic

substances on their external surfaces or within their inter-layer spaces by the interaction or

substitution of the exchanged cations within these spaces (Veldt and Meniere, 2008). Aflatoxins

can be adsorbed into this porous structure and be trapped by electric charges (Jouany, 2007).

Adsorbent clay minerals are effective at binding AFB1 in liquid media (Moschini et al., 2008).

Kutz et al. (2009) examined the efficacy of two calcium montmorillonite clay-based sequestering

agents, soils (Novus International Inc., St. Charles, MO) and overspills (Engelhard Corp.,

Cleveland, OH), in mid lactating cows fed 100 μg/kg of aflatoxin B1. Addition of soils and

overspills reduced the milk AFM1 concentration by 45% and 48%, respectively. Queiroz et al.

(2012) evaluated the effects of dietary addition of 0.5 and 1% of diet DM of a montmorillonite-

based mycotoxin sequestering agent on milk AFM1 concentrations and reported that only the

high dose reduced the concentration of AFM1 in the milk. Furthermore, a recent study by Maki

et al. (2016) revealed that transfer rate of aflatoxin into milk was reduced in cows fed overspills.

Although, clay-based products have been effective at adsorbing aflatoxin in diets, their

composition varies significantly by lot and type; therefore, the efficacy and safety of each

product should be evaluated before inclusion in animal diets (CAST, 2003). The most

extensively studied clay minerals are a range of products known as hydrated sodium calcium

aluminosilicate (HSCAS) and montmorillonite is one of those frequently used (Jouany, 2007).

This clay mineral binds with aflatoxin by chelating the β-dicarbonyl moiety in aflatoxin with

uncoordinated metal ions in the clay materials (Phillips et al., 1991; Whitlow, 2006). Hydrated

62

sodium calcium aluminosilicate has high affinity for AFB1 (Phillips et al., 1988). In fact, the

aflatoxin-HSCAS binding complex formed is stable over a wide pH (2 to 10) and temperature

(25 to 37°C) range (Sarr et al., 1990, 1991). Other clay products that have been studied include

bentonites, zeolites, clinoptilolites, and others that are not characterized (Whitlow, 2006). Clay

minerals differ in terms of ion exchange capacity, surface area, and particle size, shape and

distribution (Grim, 1968). Montmorillonite clay is believed to have the greatest cation exchange

capacity (Grim, 1968). Due to such compositional differences, it is important for research

studies to report the characteristics of clay products being tested. Some of the limitations of clay-

based products as mycotoxin sequestering agents are that they accumulate in manure, may be

contaminated with toxic metals and dioxins, and they are only effective at binding aflatoxins

(Bursian et al., 1992; Phillips et al., 1999). For instance, the inclusion of HSCAS in the DON-

contaminated diet of pigs did not mitigate the effects of DON (Patterson and Young, 1993).

Also, HSCAS reduced the toxicity of aflatoxin, but did not prevent that of ochratoxin A alone or

combined with aflatoxin in poultry (Huff et al., 1992). In addition, clay-based sequestering

agents may bind with dietary minerals and vitamins. Chestnut et al. (1992) reported impaired

absorption of zinc, magnesium, and manganese in sheep fed HSCAS. Similar findings were

observed in poultry (Chung et al., 1990) and swine (Schell et al., 1993). No published reports

indicating that clay-based sequestering agent ingestion impaired mineral absorption in dairy and

beef cattle were found. However, a recent study reported an increased loss in milk yield as

concentration of clay product increased from 0.5 to 2% in dairy cows fed 100 µg/kg of dietary

aflatoxins (Sulzberger et al., 2017). The clay product used in this study contained vermiculite,

nontronite, and montmorillonite.

63

To ensure protection against other mycotoxins and reduce the need for clay products in

diets, yeast and yeast cell wall products were evaluated for their ability to bind aflatoxins and

other mycotoxins (Devegowda et al., 1998). A yeast cell wall product (MTB-100) reduced the

AFM1 concentration of milk by 59% when fed at 0.05% of a diet contaminated with 55 µg/kg of

AFB1 (Diaz et al., 2004). Xiong et al. (2015) reported that adding a blend of yeast extract and

yeast culture reduced transfer of AFB1 into milk from diets contaminated with 20 μg/kg of AFB1

but had no effect when diets were contaminated with 40 μg/kg of AFB1. Furthermore, no effects

of yeast products have been found in diets contaminated with at least 60 μg/kg of AFB1

(Waltman et al., 2008; Kutz et al., 2009; Kissell et al., 2013), indicating that the efficacy of yeast

products is dependent upon the extent of mycotoxin contamination.

Glucans and mannans are major components of yeast cells (Fruhauf et al., 2012).

Sequestration of mycotoxins by yeast is caused by adhesion of the toxin to the cell wall

components (mannan and β-glucan) (Pfliegler et al., 2015). Yeast cell wall glucans, besides

being nutritional aids and growth promoters, can bind mycotoxins in vitro and in vivo (Fruhauf et

al., 2012). Beta-glucans (glucomannan) fed at 0.05 or 0.1% of diet to dairy cow or poultry,

respectively, bound aflatoxins, ochratoxin, and T-2 toxin (Raju and Devegowda, 2000; Diaz et

al., 2004). Furthermore, Freimund et al. (2003) noted that a glucan polymer can bind both T-2

toxin and ZEA in vitro. An additional benefit of adding yeast or yeast products to diets is that β-

glucan can improve the performance of animals by stimulating both nonspecific and specific

immunological responses (Keller et al., 2015).

Activated charcoal is a mycotoxin sequestering agent with a large surface area and

excellent adsorptive capacity (Whitlow, 2006). However, the effects of activated charcoal have

been variable. Galvano et al. (1996) reported that two activated charcoals differing in surface

64

area and iodine number were fed at 2% of the diet reduced the transfer of toxin into the milk of

Friesian cows fed 11 µg/kg of AFB1 by 22 to 45%, but these responses were not better than that

with HSCAS. However, when cows were fed 55 µg/kg of AFB1, feeding 45 g/cow daily of

activated charcoal did not reduce milk aflatoxin residues, whereas bentonite clay binder (225

g/cow daily) or an esterified glucan (10 g/cow daily) reduced milk aflatoxin (Diaz et al., 2004).

This suggests that the efficacy of activated charcoal is dependent on the level of contamination.

Studies with broilers (Edrington et al., 1997) and turkey poults (Edrington et al., 1996) also

suggest that charcoal is not as effective as clay-based sequestering agents.

A synthetic water-soluble polymer, polyvinlypyrrolidone (PVP), also has been evaluated

as a mycotoxin sequestering agent. Alegakis et al. (1999) reported binding of ZEA by PVP

ranging from 33.5 to 66.2% per 25 mg of PVP in an in vitro medium. However, no studies on the

binding efficacy of PVP in animals were found.

The binding effects of chlorophyll-based products on AFB1 also have been studied in

vitro (Hsu et al., 2008). The mechanism of action of chlorophyll-based product involves trapping

AFB1 and other carcinogens via binding of their planar ring structures to that of chlorophyll

(Arimoto et al., 1993), thereby, preventing the toxin from forming adducts with DNA, and hence

facilitating the clearance of the toxin from the body (Hsu et al., 2008). In humans challenged

with 30 ng of AFB1, two chlorophyl products fed at 150 mg daily produced a significant 40% to

60% reduction in excretion of urinary aflatoxin equivalents (Jubert et al., 2009).

Summary. This review discussed the phases of silage fermentation, the importance of

silage additives, the types and modes of action of different silage additives, different techniques

for silage microbial evaluation, and the critical need for proper understanding of silage microbial

composition and its relationship to the metabolites in silage. In addition, the review highlighted

65

the role of feeds and ruminant livestock in cycling of pathogenic EC on farms, and emphasized

the need for effective strategies to prevent the growth of EC on feeds. Chapters 3, 4, and 5

summarize experiments that examined the efficacy of strategies to inhibit the growth of EC in

corn and alfalfa silage. These experiments also characterized the microbial composition of the

treated and untreated silages and examined relationships between the abundance of known and

unknown silage microbes and indices of silage quality.

This review also described the role, importance and danger of mycotoxins for animals

and humans, and their prevalence. It also showed that equivocal results have been achieved when

sequestering agents were used to bind mycotoxins and reduce their transfer from the diet to the

milk of dairy cows. This emphasizes the need to develop more effective sequestering agents and

to continue to evaluate their effects on milk AFM1 concentration, as well as on milk yield and

quality. Chapter 6 describes an experiment aimed at examining whether dietary addition of three

novel Saccharomyces cerevisiae fermentation products could act as mycotoxin-sequestering

agents that would reduce transfer of aflatoxin to milk and improve the performance and health of

dairy cattle challenged with dietary aflatoxin.

66

Table 2-1. Estimated dietary contributions of whole plant corn silage to total dietary mycotoxin

in diets.*

Reference Mycotoxin Mean

Concentration

(µg/kg)

Maximum

concentration

(µg/kg)

% mean

contribution

in dairy diet

% maximal

contribution in

dairy diet

Richard et al. 2009 AFB1 28.0 NA 42 (168)# NA

Keller et al. 2013 AFB1 33.0 54 49.5 (198)# 81.0 (324)#

Driehuis et al. 2008 ZEA 174 943 13.05 70.7

Storm et al. 2014 ZEA 66.0 311 3.96 23.3

Schollenberger et al.

2006

ZEA 432 1790 25.92 134.3

Eckard et al. 2011 ZEA 180 430 10.8 32.3

Storm et al. 2010 DON 1056 5094 6.336 38.2

Driehuis et al. 2008 DON 854 3142 5.124 23.6

Storm et al. 2014 DON 1629 2974 9.774 22.3

Gallo et al. 2015 DON 280 560 1.68 4.20

Schollenberger et al.

2006 DON 2919 3944 17.514 29.6

Eckard et al. 2011 DON 1356 2990 8.136 22.4

Keller et al. 2013 DON 1610 3420 9.66 25.7

Oldenburg et al. 2005 DON 2100 14290 12.6 107

Mansfield et al. 2005 DON 1400 5100 8.4 38.3

Driehuis et al. 2008 Fumonisin B1 +

B2 NA 34,000 NA 68 (20.4)#

Kim et al. 2004 Fumonisin B1 615 1824 1.23 (0.37)# 3.65 (1.09)#

Mansfield et al. 2007 Fumonisin B1 +

B2 3000 30400 6.00 (1.8)# 60.8 (18.2)#

Schneweis et al., 2000 Mycophenolic

acid 690 23000 0.69 23.0

Storm et al. 2014 Mycophenolic

acid 9.00 52.0 0.01 0.05

Gallo et al. 2015 Mycophenolic

acid 1760 48000 1.76 48.0

Auerbach et al. 1998 Roquefortine C 17000 36000 127.5 270

Auerbach et al. 1998 Roquefortine C 5470 28000 41.025 210

Mansfield et al. 2008 Roquefortine C 380 5710 2.85 42.8

Storm et al. 2014 Roquefortine C 173 189 1.30 1.42

Gallo et al. 2015 Roquefortine C 740 32000 5.55 240 *Estimated daily corn silage intake of 7.5 kg; Estimated concentrations of mycotoxins in the

diets; aflatoxins (AFB1; 20 µg/kg, FDA, 2010; 5 µg/kg, European commission, 2006),

Deoxynivalenol (DON; 5 mg/kg, FDA, 2011 and European Commission, 2006), fumonisin (30

mg/kg, FDA, 2011; 50 mg/kg, European commission, 2006), ZEA (0.5 mg/kg, European

commission, 2006), and mycophenolic acid (300 mg/kg, Mohr et al., 2007) and Roquefortine C

(4 mg/kg; Haggblom, 1990) #Values in parenthesis are estimates based on maximum permissible level of the mycotoxin in

Europe.

NA = Not given

67

CHAPTER 3

CONTROL OF Escherichia coli O157:H7 IN CONTAMINATED ALFALFA SILAGE:

EFFECTS OF SILAGE ADDITIVES

Background

Escherichia coli O157:H7 is a Shiga toxin-producing food-borne pathogen associated

with hemorrhagic colitis and hemolytic uremic syndrome (HUS) in humans (USDA-APHIS,

2001). This pathogen has been implicated in outbreaks of these illnesses in the US, Canada and

Europe and accounts for about 95% of HUS cases (Griffin, 1995). Escherichia coli O157:H7 is

estimated to cause about 75,000 illness cases and 50 deaths yearly in the US (Mead et al., 1999).

Cattle are considered the primary reservoir of EC (Cernicchiaro et al., 2013) and they shed the

pathogen in their manure (Lahti et al., 2003; Mir et al., 2015).

Animal feeds, such as silage, have been suggested to be the vehicle for transmission of

the pathogen among livestock (Hancock et al., 2001; Davis et al., 2003). Additive treatments of

silages may reduce the prevalence and spread of the pathogen on farms and reduce the risk of

human exposure (Matthews et al., 2013). Corn silage and alfalfa hay or silage are the most

commonly fed forages in dairy cow diets in the US and they can be contaminated with EC

through the manure and saliva of cows (Russel et al., 2000). The rapid drop in pH during ensiling

kills EC in corn silage contaminated with the pathogen before ensiling. Lactobacillus plantarum

hastened this decrease in pH (Chen et al., 2005), but it did not hasten or increase inhibition of the

pathogen during ensiling (Pedroso et al., 2010). However, L. buchneri prevented the growth of

the pathogen on contaminated aerobically exposed corn silage (Pedroso et al., 2010). Despite its

widespread usage in dairy cow diets due to its high nutritional value (Albrecht and Beauchemin,

2003), no studies have examined the use of microbial inoculants to inhibit the spread of EC via

alfalfa silage. The high buffering capacity and low water-soluble carbohydrate concentration of

alfalfa hinder the rapid pH decline necessary for efficient fermentation. Therefore, pathogens

68

eliminated by the rapid pH decline and sustained acidic conditions in ensiled corn may not be

eliminated in ensiled alfalfa. Whether microbial inoculants or acid additives can reduce the

transmission of EC on dairy farms by inhibiting the growth of the pathogen on alfalfa forage

during or after ensiling is unknown. The objective of our study was to examine the effectiveness

of obligate or facultative heterofermentative lactic acid bacteria (LAB) inoculants or propionic

acid at controlling EC in alfalfa silage during and after ensiling.

Materials and Methods

The Biological Safety Office, Division of Environmental Health and Safety, and the

Institutional Animal Care and Use Committee (IACUC) of the University of Florida approved all

procedures for this study (protocol no. 201408675).

Harvesting, Ensiling and Inoculation

Alfalfa forage was harvested at the early bloom stage with a mower (John Deere 946,

Moline, IL), wilted to a DM of 54%, and chopped to a theoretical length of cut of 19 mm with a

forage harvester (John Deere 7950, Moline, IL). The following treatments were applied in

quadruplicate with a hand-held garden sprayer to 1.5 kg of the forage: (1) distilled water

(Control); (2) 1 × 105 cfu/g of Escherichia coli O157:H7 EDL933 alone (EC); (3) EC + 1 × 106

cfu/g of Lactobacillus plantarum R2014 (EC+LP); (4) EC + 1 × 106 cfu/g of L. buchneri R1102

(EC+LB); and (5) EC + 2.2 g/kg (fresh weight) of propionic acid containing 99.9% of the acid

(EC+PA). The EC was supplied by the University of Florida Emerging Pathogens Institute

(Gainesville) and the silage inoculant bacteria were supplied by Lallemand Animal Nutrition,

Milwaukee, WI. The number of bacteria in the inoculants was verified by serial dilutions in

Buffered Peptone Water (BPW; Oxoid, Basingstoke, UK) followed by plating on de Man-

Rogosa-Sharpe (MRS) agar (Difco, Detroit, MI) and incubating at 37°C for 48 h. The EC was

grown in Luria-Bertani (LB) broth (Sigma-Aldrish, Saint Louis, MO) overnight at 35°C for 12 h.

69

The cells in the suspension were adjusted to an optical density of 0.5 at 600 nm using a

GENESYS-20 spectrophotometer (Thermo Scientific, Waltham, MA). The number of EC in the

suspension was verified by spread plating on sorbitol MacConkey agar (SMAC; Oxoid,

Basingstoke, UK) supplemented with cefixime (0.05 mg/L) and potassium tellurite (2.5 mg/L)

(CT-SMAC), and incubated at 35°C for 24 hours. The pathogen inoculum was diluted in distilled

water (1:5 ratio) and applied to the forage to supply 5 log cfu/g at ensiling. The bacterial

inoculants and propionic acid were also diluted in distilled water (1:2 and 1:0.3 ratio,

respectively) and applied to the forage at 3 mL/kg of fresh forage (Pedroso et al., 2010) to give

application rates of 1 × 106 cfu/g for each inoculant and 0.22% for the acid. Control silages were

also treated with the 3 mL/kg fresh forage of distilled water. After proper mixing, the treated

forages were packed into 14 × 21-cm, thick-walled plastic bags that were subsequently

vacuumed to remove air and sealed with plastic ties. Each treatment was ensiled for 0, 3, 7 and

16 d at 20°C. Samples from these ensiling durations were analyzed for EC counts, pH, organic

acids and ammonia-N. In addition, each treatment was applied to 4.5 kg of forage in

quadruplicate, packed into 20-L plastic mini silos lined with thick-walled plastic bags and

ensiled for 100 d at 20°C. On d 100, samples from these silos were analyzed for EC, pH, organic

acids, ammonia-N, aerobic stability, and yeast and mold counts. In addition, samples from days 0

and 100 were analyzed for chemical composition.

Survival of E. coli O157:H7 in Aerobically Exposed Silages

The objective of this phase of the experiment was to determine whether prior treatment

with the silage additives at ensiling would enhance the aerobic stability and prevent the growth

of EC in silages contaminated with the pathogen after silos were opened. Approximately 1 kg of

d 100 silages from each replicate was transferred to a 20-L plastic mini silo and re-inoculated

with 5 log cfu/g of EC. The silages in each silo were covered with 2 layers of cheese cloth to

70

avoid dehydration and stored at 20°C for 7 d. Silage temperature was monitored with wireless

temperature sensors (Onset Computer Corporation, Bourne, MA) that were placed in the

geometrical center of the silage mass in each silo. The sensors recorded silage temperatures at

30-min intervals. Two additional sensors were placed in the room to record ambient temperature.

Aerobic stability was defined as the number of hours the temperature in the silages remained

stable before rising more than 2°C above room temperature. On d 107, samples from these silos

were analyzed for EC, yeast and mold counts, pH, an aerobic stability.

Laboratory Analyses

Silage extracts for microbial enumeration, VFA, pH, and ammonia-N analyses were

prepared by blending 20 g of silage with 180 mL of distilled water in a stomacher (UL Lab-

Blender 400, Seward Laboratory, London, UK) for 1 minute and filtering through 2 layers of

cheese cloth. The filtrate was immediately analyzed for EC, yeast and mold counts, and pH. For

EC enumeration, serial 10-fold dilutions were prepared in BPW followed by direct plating on

duplicate plates of CT-SMAC agar (Zadik et al., 1993). Plates were incubated for 24 h at 35°C

and sorbitol-negative colonies were enumerated as EC. Colonies were confirmed for the O157

and H7 antigens by latex agglutination (Remel, Lenexa, KS) (March and Ratnam, 1989). Yeasts

and molds were enumerated after spread-plating 10-fold serial dilutions on malt extract agar

(Difco, Detroit, MI) and incubated at 32°C for 72 h. The pH of silage extracts was measured

using a digital pH meter (Accumet AB15, Fisher Scientific, Pittsburgh, PA). An aliquot of the

silage extract was centrifuged at 1,800 × g for 15 min at 4°C and the supernatant was analyzed

for ammonia-N using an adaptation for the Technicon Auto Analyzer of the method of Noel and

Hambleton (1976). Organic acids were quantified using a Merck Hitachi Elite La-Chrome High

Performance Liquid Chromatograph system (Hitachi L2400, Tokyo, Japan) fitted with a Bio-Rad

Aminex HPX-87H column (Bio-Rad Laboratories, Hercules, CA) with a 0.015-M sulfuric acid

71

mobile phase and a flow rate of 0.7 mL/min at 50°C (Muck and Dickerson, 1988). The DM

concentrations of the fresh forage and silage were determined by drying samples in a forced-air

oven at 55°C for 48 h. Dried samples were ground to pass the 1-mm screen of a Wiley mill (A.

H. Thomas, Philadelphia, PA). Concentrations of NDF and ADF were measured in an ANKOM

200 Fiber Analyzer (Ankom Technologies, Macedon, NY) using the procedures of Van Soest et

al. (1991). Heat-stable amylase was used in the NDF analysis and the results were expressed on a

DM basis inclusive of residual ash. Crude protein was calculated by multiplying N

measurements obtained after Kjeldahl digestion by 6.25.

Antibacterial Activity of the Microbial Inoculants and Silage Extracts

Antibacterial activity against EC of pure cultures of the microbial inoculants and d 100

silage extracts was determined by the agar disk diffusion method of Bauer et al. (1966). The EC

isolates were cultured in LB and adjusted to an optical density of 0.5 at 600 nm and the inoculum

was prepared on CT-SMAC agar. Bacteria in the commercial inoculants were grown in MRS

broth at 35°C for 24 h. Cell-free supernatants were prepared by centrifuging the cultures at 3,000

× g for 15 min at 4°C. The pH of the supernatants was adjusted to 5.5. The MRS broth was also

adjusted to a pH of 5.5 and used as the Control. Silage extracts were similarly centrifuged and

the cell-free supernatants were adjusted to pH 5.5. Two sterile cloning discs (6.0-mm diameter;

cat. no. 231039, Becton Dickinson Co.) were immersed in the supernatants for 20 s using flame-

sterilized forceps and then placed on the surface of CT-SMAC agar plates containing EC. Plates

were prepared in triplicate and incubated for 24 h at 35°C. The diameter of zones of inhibition

around the paper discs were measured with a ruler.

Statistical Analysis

The experiment had a completely randomized design with 5 treatments and 4 replicates

per treatment. All microbial data were transformed to log10. Data were analyzed using the

72

GLIMMIX procedure of SAS (SAS Institute Inc., Cary, NC). The model for analyzing data

collected repeatedly over time (pH, EC counts and organic acid concentrations) included effects

of treatment, day and treatment × day. When treatment × day interactions were significant,

effects of treatment on each day were examined. The model for analyzing the data collected once

included the treatment effect. Differences between means were determined using the Tukey

procedure. Significance was declared at P < 0.05.

Results

Anaerobic Phase

The chemical composition of the fresh alfalfa forage before ensiling was similar for all

treatments except that as expected, the pH of EC+PA silage was lower (P = 0.01) than those of

the other treatments (Table 3-1). The EC+LP silage had the fastest (P = 0.01) increase in lactate

concentration and it had greater concentrations than other treatments for the entire ensiling

duration (Figure 3-1). The EC+LB silage consistently had greater lactate concentrations than EC,

EC+PA or Control silages after 3 d of ensiling. None of the silages contained acetate until d 7

when it was only detected in the EC+LP silage (Figure 3-2). Yet the EC+LB silage had the

greatest acetate concentrations after 16 and 100 d of ensiling and the EC+LP silage had

intermediate concentrations. Propionic acid was detected in only the EC+PA silage on d 3, 7 and

16 (0.17 ± 0.03, 0.24 ± 0.06 and 0.18 ± 0.01 % DM, respectively).

Within 7 d of ensiling, the pH of the EC+LP silage had decreased below 5.0 (Figure 3-3),

which is the minimal pH value in well preserved, high-DM alfalfa silage (Kung et al., 1987). In

contrast, it took 16 d for the pH of the EC+LB silage to decrease below 5.0 and a longer period

for a similar decrease in the other silages.

The fresh, unensiled Control forage contained 2.3 log cfu/g of EC (Figure 3-4), possibly

because of contamination from the liquid dairy manure applied to the forage approximately 50-d

73

before it was harvested. Nevertheless, the EC counts of the Control silage were about 10,000-

fold lower than those of the silages spiked with the pathogen. The pathogen was detected in all

silages on d 3 and 7 but by d 16, it was no longer detected in inoculant-treated silages, whereas it

persisted for longer periods in the other silages. Nevertheless, no EC was detected in any of the

silages by d 100 when all their pH values were below 5.0 (Table 3-2).

The pH was lower (P < 0.05) in d 100 EC+LP and EC+LB silages (4.57 and 4.64,

respectively) than the corresponding Control and EC silages (4.88 and 4.82, respectively; Table

3-2). This was caused by the greater (P < 0.05) concentrations of lactate and acetate in EC+LP

and EC+LB silages, respectively than in Control and EC silages. The EC+PA silage had a

similar pH value to silages treated with the microbial inoculants partly because it had a higher

propionic acid concentration (0.22% DM) than the other silages, as it had less lactate and acetate

than EC+LP and EC+LB, respectively. None of other silages contained propionic acid and no

butyrate was detected in the silages. The EC+LB and EC+PA silages had lower lactate-acetate

ratios than the EC+LP silage. Yeast counts were lower (P < 0.01) in the EC+LB and EC+PA

silages (3.06 and 2.65 log10 cfu/g, respectively) than in Control, EC and EC+LP silages (5.22,

4.85 and 4.50 log10 cfu/g, respectively).

Aerobic Phase

After 168 h of re-inoculation and aerobic exposure, 2.3 log10 cfu/g of EC were detected in

the EC silage but no EC was detected in the Control, inoculant or propionic acid- treated silages

(Figure 3-5). The pH values of the inoculant and propionic acid-treated silages were below 5.0

(Figure 3-6) after the aerobic exposure period, whereas those of the Control and EC silages were

greater (P < 0.05) as they had increased to 5.4 and 5.6, respectively.

74

The EC+PA silage had the lowest (P < 0.05) yeast counts, followed by the silages treated

with the inoculants (Figure 3-5). All silages were stable for the entire duration of aerobic

exposure (168 h).

Antibacterial Activity of Microbial Inoculants and Silage Extracts

The pH-corrected supernatants from pure cultures of L. plantarum and L. buchneri

produced 3.20 ± 0.30 and 3.07 ± 0.25-mm zones of inhibition against EC, respectively, but no

inhibition zone was produced by cell-free supernatants from extracts of d 100 silages that had

been treated with the inoculants at ensiling.

Discussion

Silages form the bulk of the ration of dairy cows and many beef cattle in the US.

Therefore, they are integral initial components of the human food chain. Consequently, the

hygienic quality or feed safety of silage is of paramount importance. In this context, the current

experiments studied the effects of two LAB inoculants or propionic acid on the survival of EC in

alfalfa silage during and after the ensiling periods as part of a broader effort to understand the

probiotic and food-safety enhancing effects of such inoculants.

The presence of EC in the fresh Control alfalfa forage confirmed the results of various

studies and surveys on the potential risk of pathogenic E. coli contamination of forages and

silages because of fertilization with manure slurry or irrigation water (Beuchat and Ryu, 1997).

Dunn et al. (2004) reported that 38.5% of herds of dairy cows and 6.5% of dairy cows per herd

had E. coli O157 in the manure. The population of E. coli in dairy manure varies between 5 to 8

log cfu/g (Zhao et al. 1995; Walczak and Xu, 2011) and the viability of the bacteria in such

organic wastes after land application can last for as long as 60 d (Avery et al., 2005).

75

Anaerobic Ensiling Phase

The elimination of pathogenic and non-pathogenic E. coli in well preserved corn and

grass silages because of the rapid drop in pH during ensiling is well known (Byrne et al., 2002;

Chen et al., 2005; Pedroso et al., 2010, Duniere et al., 2011). To our knowledge, only our

previous study (Pedroso et al. 2010) and that of Bach et al. (2002) have investigated the survival

of EC in corn and barley silages, respectively, during aerobic exposure. No study seems to have

examined the survival of this serotype during the ensiling and aerobic phases of alfalfa silage

production. Yet as shown in this current study, the pathogen is more persistent in untreated

ensiled alfalfa silage than in untreated ensiled corn, grass or barley silages (Bach et al., 2002;

Pedroso et al. 2010). This is because of the high buffering capacity of alfalfa which decreases the

rate of pH decline during ensiling.

This study shows that adding L. plantarum or L. buchneri to alfalfa at ensiling reduced

the time (16 vs. 100 d) required for elimination of EC in alfalfa silages contaminated with the

pathogen at ensiling. This suggests that inoculant treatment is needed to eliminate EC in

contaminated forage that is ensiled for relatively short periods. This was likely caused by the

faster and greater production of organic acids by the LAB, which rapidly reduced the pH of the

silages to values below 5.0. Chen et al. (2005) reported elimination of a recombinant Escherichia

coli strain in wheat and corn silages when the pH decreased below 5.0 but noted that it persisted

longer in wilted silages with a slow pH decline. A microbial inoculant containing Pediococcus

pentosaceus and Propionibacterium jenzenii accelerated the accumulation of lactate and

reduction in pH of barley silage and thereby hastened (7 vs. 15 d) the elimination of EC that had

been added to the barley forage at ensiling (Bach et al., 2002). However, EC is rapidly

eliminated during ensiling of corn and grass silages, even in the absence of microbial inoculants,

76

because of the inherent rapid accumulation of organic acids and the subsequent rapid decrease in

pH in these forages (Duniere et al., 2011; Pedroso et al., 2010).

The rate and extent of silage fermentation is reduced with increasing DM content (Kung

et al., 1984; Muck, 1990; Jones et al., 1992). The high DM at ensiling and high buffering

capacity of the alfalfa silages in this study caused their pH to decrease relatively slowly.

Application of the inoculants resulted in a faster decrease in pH despite the high DM content of

the unensiled forages and also decreased the pH at d 100 relative to the value for the Control

silage (Figure 3-3).

The inhibitory effect of the inoculants on EC may have been partly because of the

organic acids they produced. Such acids are generally considered to have strong antimicrobial

activity (Buchanan et al., 2004). Within the pH range of 4.0 to 5.5, lactate, the main acid

produced during homofermentative ensiling pathways, had the greatest activity in vitro against

enterohemorrhagic E. coli (Buchanan and Edelson, 1999). Whereas, Abdul-Raouf et al. (1993)

reported that relative inhibitory activity against EC was higher for acetate than lactate. In

addition, Breidt et al. (2004) demonstrated that acetic acid reduced the survival of E. coli strains

when compared with effects attributed to pH alone.

The inhibitory effect of the inoculants may have also been partly because of their

antibacterial effects on EC. This is evident from the inhibition zones caused by pure cultures of

the inoculants on the Kirby bauer disks coated with EC. These inhibitory effects may have been

caused by the production of bacteriocins (Jack et al., 1995) or other inhibitory compounds.

However, the antibacterial effect was reduced or lost during ensiling because extracts of the d

100 inoculant-treated silages produced no inhibition zones around the EC, probably because of

77

loss of metabolic activity of the LAB after 100 d of ensiling. These results are similar to those

reported by Pedroso et al. (2010).

Aerobic Post-Ensiling Phase

Lactobacillus buchneri increases the concentration of acetic acid in silages via anaerobic

degradation of lactic acid to acetic acid (Oude Elferink et al., 2001), and thereby prevents

accumulation of yeasts and improves the aerobic stability of silages (Weinberg et al., 2002;

Kleinschmit et al., 2005; Pedroso et al., 2010; Queiroz et al., 2013). In this study, inoculation

with L. buchneri resulted in higher concentration of acetic acid and about 1000-fold reduction in

the number of yeasts compared with the Control silage. The antifungal characteristics of acetic

acid are well known (Danner et al., 2003; Schmidt and Kung, 2010). A meta-analysis on the

effect of L. buchneri in corn silages showed that it caused a 10 to 100-fold decrease in yeast

counts in corn silage (Kleinschmit and Kung, 2006a).

Propionic acid has the greatest antimycotic characteristics of all the short chain fatty

acids (Woolford, 1975) and its undissociated form is effective at reducing the growth of yeasts

and molds, which are responsible for causing and maintaining or enhancing aerobic deterioration

in silages, respectively (Kleinschmit et al., 2005). The fraction of propionic acid that is

undissociated increases as the pH decreases (Lambert and Stratford, 1999). In this study,

treatment with propionic acid predictably increased the concentration of the acid and prevented

the growth of yeasts. Kleinschmit et al. (2005) reported that buffered-propionic acid (0.1% of

fresh forage) did not reduce counts of yeasts in corn silage. This may be caused by the relatively

low application rate and the concentration of the acid in the product. Rates of application of

buffered propionic acid-based products can be as high as 0.25 % of fresh forage (Kung et al.,

1998) and higher application rates are more effective (Kung et al., 1998, 2000).

78

Aerobic instability is not a common problem in alfalfa silages with good silo

management (Muck, 2008). In this study, the alfalfa silages were stable for the entire aerobic

exposure duration. This agrees with the results of Pahlow et al. (2001), who showed that about

90% of legume silages surveyed were stable on exposure to air for the same duration used in this

study (168 h).

When the silages contaminated with EC on d 100 were exposed to air for 168 h, 2.3 log

cfu/g of the pathogen were detected in the EC silage yet the pathogen was not detected in silages

that had been treated with the inoculants or propionic acid at ensiling. This is probably because

the additive-treated silages seem to have kept the pH below 5.0 during the aerobic exposure

period. Pedroso et al. (2010) reported that L. buchneri prevented the growth of EC on aerobically

exposed silages because it maintained an acidic pH (4.0) for 144 h after silo opening and

exposure to air. In a similar study, Duniere et al. (2011) showed that E. coli O26 was not

detected in corn silages when contamination occurred at silo opening because of low pH in those

silages. However, when EC contamination occurred after 144 h when the pH had increased

above 6.0, about 2 to 6 log10 cfu/g of the pathogen were recovered from the silages. This

indicates that the risk of survival of pathogenic E. coli can be reduced markedly or curtailed with

silage additives that maintain acidic pH conditions in aerobically exposed silages. In addition,

the more rapid elimination of the pathogen by obligate or facultative LAB may help to curtail the

spread of pathogen by preventing it from cycling on dairy farms.

Conclusions

Escherichia coli O157:H7 was eliminated from alfalfa silages treated with L. plantarum

or L. buchneri within a shorter time (16 vs. 100 d of ensiling) than that required for elimination

of the pathogen from Control, EC and EC+PA silages. Therefore, application of certain

inoculants is needed to prevent survival of the pathogen in alfalfa ensiled for short periods. The

79

elimination of EC during ensiling was partly because the inoculants produced more lactate and

acetate, caused a faster reduction in silage pH than the other treatments, and exhibited

antibacterial effects on EC. Application of L. buchneri and propionic acid curtailed the

accumulation of yeasts and molds during ensiling and after aerobic exposure. The bacterial

inoculants and propionic acid kept the pH below 5.0 for the entire duration of the 168-h aerobic

exposure period. Consequently, the pathogen did not survive in aerobically exposed additive-

treated silages contaminated with the pathogen at silo opening but it did survive and persist in

silages treated with EC alone, which had a pH of 5.4. This suggests that keeping silage pH below

5.0 during the feedout stage with an appropriate silage additive will help curtail the growth of EC

in alfalfa silages contaminated with the pathogen after silo opening.

80

Table 3-1. Chemical composition (% of DM except as noted) and pH of alfalfa forage treated

without (Control) or with Escherichia coli O157:H7 (EC) alone, or EC and either of 2

bacterial inoculants or propionic acid before ensiling.

Item Treatment1 SE P-value

Control EC EC+LP EC+LB EC+PA

pH 6.23a 6.20a 6.21a 6.20a 5.85b 0.04 0.01

DM (%) 56.8 56.3 54.8 56.2 56.7 0.87 0.17

CP 22.9 23.6 23.1 23.5 23.8 0.43 0.25

NDF 36.4 35.8 37.0 35.5 37.5 1.27 0.58

ADF 29.3 28.3 28.9 28.3 30.5 1.00 0.30

a,bMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; LP = 1 × 106 cfu/g of L.

plantarum; LB = 1 × 106 cfu/g of L. buchneri; PA = 2.2 g/kg of propionic acid

81

Table 3-2. Fermentation indices, chemical composition (% of DM except as noted) and yeast

counts (log cfu/g) of alfalfa silage treated without (Control) or with Escherichia coli

O157:H7 (EC) alone, or EC and either of 2 bacterial inoculants or propionic acid and

ensiled for 100 d.

a–cMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; LP = 1 × 106 cfu/g of L.

plantarum; LB = 1 × 106 cfu/g of L. buchneri; PA = 2.2 g/kg of propionic acid



pH 4.88a 4.82ab 4.57c 4.64c 4.67bc 0.05 0.01

Lactate 2.14b 1.77b 4.47a 2.59ab 1.14b 0.93 0.03

Acetate 0.95cb 0.67c 1.27b 2.20a 0.98cb 0.23 0.01

Lactate: acetate 2.27ab 2.83ab 3.93a 1.15b 1.28b 0.92 0.05

Propionate 0.00b 0.00b 0.00b 0.00b 0.22a 0.01 0.01

Yeast 5.22a 4.85a 4.50a 3.06b 2.65b 0.53 0.01

Aerobic stability (h) 166 160 >168 >168 >168 5.25 0.39

DM (%) 54.5 54.1 54.7 54.6 55.6 0.53 0.12

NDF 40.4 42.1 40.5 42.5 42.5 1.38 0.41

ADF 32.6 34.4 32.2 34.3 33.0 1.47 0.52

CP 24.3 23.1 23.6 24.5 24.0 0.82 0.47

Ammonia-N 0.34 0.42 0.30 0.36 0.36 0.10 0.76

82

Figure 3-1. Changes in lactate concentration of alfalfa forage treated without (Control) or with


propionic acid during ensiling. Control = distilled water; EC = 1 × 105 cfu/g of E. coli

O157:H7; LP = 1 × 106 cfu/g of L. plantarum; LB = 1 × 106 cfu/g of L. buchneri; PA

= 2.2 g/kg of propionic acid; Treatment × day P-value = 0.01; *Concentrations

differed at the day of ensiling specified (P < 0.05). Error bars represent SE.

0

1

2

3

4

5

0 3 7 16 100

Lac

tate

, %

DM

Day of ensiling

Control EC

EC+LP EC+LB

EC+PA

/ /

83

Figure 3-2. Changes in acetate concentration of alfalfa forage treated without (Control) or with






0

0.5

1

1.5

2

2.5

0 3 7 16 100

Ace

tate

, %

DM

Day of ensiling

Control EC

EC+LP EC+LB

EC+PA

/ /

84

Figure 3-3. Changes in pH of alfalfa forage treated without (Control) or with Escherichia coli

O157:H7 (EC) alone, or EC and either of 2 bacterial inoculants or propionic acid

during ensiling. Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; LP

= 1 × 106 cfu/g of L. plantarum; LB = 1 × 106 cfu/g of L. buchneri; PA = 2.2 g/kg of

propionic acid; Treatment × day P-value = 0.01; *Concentrations differed at the day

of ensiling specified (P < 0.05). Error bars represent SE.

4.0

4.5

5.0

5.5

6.0

6.5

0 3 7 16 100

pH

Day of ensiling

Control EC

EC+LP EC+LB

EC+PA

/ /

85

Figure 3-4. Escherichia coli O157:H7 (EC) of alfalfa forage treated without (Control) or with






0

1

2

3

4

5

6

7

0 3 7 16 100

EC

, lo

g1

0cf

u/g

Day of ensiling

Control

EC

EC+LP

EC+LB

EC+PA

/ /

86

Figure 3-5. Effect of reinoculation with 1 × 105 cfu/g of Escherichia coli O157:H7 (EC) at silo

opening on counts of EC and yeasts (log cfu/g) on aerobically exposed alfalfa silages

that were treated without (Control) or with EC alone or EC and either of 2 bacterial

inoculants or propionic acid at ensiling. Control = distilled water; EC = 1 × 105 cfu/g

of E. coli O157:H7; LP = 1 × 106 cfu/g of L. plantarum; LB = 1 × 106 cfu/g of L.

buchneri; PA = 2.2 g/kg of propionic acid; Similarly shaded bars with different letters

differed (P < 0.05); Treatment SE values for EC and yeasts were 0.23 and 0.50,

respectively

0

1

2

3

4

5

6

7


Log 1

0cf

u/g

Treatment

Yeast and Mold E. coli O157:H7

B

b

C

B

AA

bbb

a

87

Figure 3-6. Effect of reinoculation with 1 × 105 cfu/g of Escherichia coli O157:H7 (EC) at silo

opening on pH of aerobically exposed alfalfa silages that were treated without

(Control) or with EC alone or EC and either of 2 bacterial inoculants or propionic

acid at ensiling. Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; LP

= 1 × 106 cfu/g of L. plantarum; LB = 1 × 106 cfu/g of L. buchneri; PA = 2.2 g/kg of

propionic acid; Bars with different letters differed (P < 0.05); Treatment SE = 0.29

4

4.5

5

5.5

6


pH

Treatment

a

a

b b b

88

CHAPTER 4

BACTERIAL DIVERSITY AND COMPOSITION OF ALFALFA SILAGE AS ANALYZED

BY ILLUMINA MISEQ SEQUENCING: EFFECTS OF E. coli O157:H7 AND SILAGE

ADDITIVES

Background

Ensiling is a forage preservation method based on lactic acid fermentation that occurs

spontaneously under anaerobic conditions because of activities of a complex community of

forage epiphytic microbes, which are highly variable in composition and species (Eikmeyer et

al., 2013; Muck, 2013). The low pH achieved during ensiling, caused by the production of lactic

acid and other organic acids, inhibits or prevents the growth of spoilage and pathogenic

organisms (Woolford, 1975). Microbial inoculants, such as homofermentative and

heterofermentative bacteria, are added to silage to dominate the natural population of bacteria in

the forage and influence the dynamics of the microbial population during ensiling (Kung, 1998;

Wang et al., 2006). Homofermentative bacteria are mainly used to improve silage preservation

by accelerating lactic acid fermentation and causing a rapid drop in pH (Kristensen et al., 2010;

Muck, 2010; Nkosi et al., 2010). Heterofermentative bacteria produce high concentrations of

acetic or propionic acid, thereby inhibiting the growth of spoilage yeasts and molds and thus,

improving the shelf life of silage during the feedout stage (Filya, 2003; Kleinschmit and Kung,

2006b; Queiroz et al., 2013). Chemical additives, such as propionic acid, are added mainly as

antifungal agents to enhance the shelf life of silages during the feedout stage (Kung et al., 1998,

2000).

The microbial population of silages have been previously described using culture-based

methods, which are notorious for underestimating the bacterial diversity (Ercolini, 2004) because

of their inability to cultivate most naturally occurring species (Temmerman et al., 2003). In

recent years, better understanding of the microbial populations in silages has been achieved

89

through molecular approaches, such as denaturing gradient gel electrophoresis (Dolci et al.,

2011), random amplified polymorphic DNA PCR (Rossi and Dellaglio, 2007), and terminal

restriction fragment length polymorphism (McEniry et al., 2008), that do not rely on culturing.

However, these methods can only identify a few of the abundant Operational Taxonomic Units

(OTUs) (Temmerman et al., 2003). The application of 454 pyrosequencing to simultaneously

sequence thousands of 16S rDNA sequences has increased our understanding of complex

microbial communities (Degnan and Ochman, 2012). Furthermore, Illumina MiSeq sequencing

platform, which generates higher throughput per run with lower error rates, and greater depth and

breadth of coverage than the 454 pyrosequencing system, at much lower costs (Frey et al., 2014),

has enhanced our understanding of complex microbial communities. Recent studies applied Ion

Torrent and single molecule real-time sequencing technologies to examine the microbial

communities of corn, alfalfa and sorghum silages (Bao et al., 2016; Kraut-Chen et al., 2016),

however, these platforms have lower throughput (Loman et al., 2012) and higher error rate

(Quail et al., 2012), respectively, than Illumina MiSeq sequencing.

Forages and silages may be contaminated with various micro-organisms, including

pathogenic Escherichia coli when manure and irrigation slurry are applied as fertilizers (Beuchat

and Ryu, 1997). Dairy manure can contain up to 8 log cfu/g of E. coli (Walczak and Xu, 2011)

and counts of more than 2 log cfu/g have been detected on forages and silages (Chen et al., 2005;

Ogunade et al., 2016). Effective control measures are needed to inhibit the growth of the

pathogen and prevent its cycling on farms or entry into the food chain. The first objective was to

examine effects of adding E. coli O157:H7 with or without chemical or microbial additives on

the bacterial diversity and composition of alfalfa silage. The second objective was to examine

90

associations between the abundance of known and unknown bacterial species and indices of

silage fermentation quality.


Metagenomic DNA Extraction and Purification

Approximately 100 g of each silage sample from Chapter 3 was added to 200 mL of PBS

+ 0.01% Tween-80 in a sterile stomacher bag. The mixture was sonicated for 30 min using

Sonicor SC-150 (Sonicor Instrument Corporation, Copiague, NY). The wash was centrifuged at

9000 × g for 1 h to create a pellet from which DNA was extracted using a PowerLyzer

PowerSoil DNA isolation kit (MO BIO Laboratories, Inc., Carlsbad, CA) according to the

manufacturer's instructions. Bead beating (Bullet Blender Storm 24, Next Advance, Averill Park,

NY) was used for homogenizing the suspension and for mechanical disruption of bacterial cells.

The purity and concentration of gDNA were measured with a spectrophotometer (Nanodrop 115

1000, Thermo Fisher Scientific-Oxoid) and the Integrity of the DNA was verified by agarose

(0.7%) gel electrophoresis.

PCR Amplification of the V3 and V4 Hypervariable Regions of Bacterial 16S rRNA Genes

Samples for sequencing were prepared according to the Illumina 16S Metagenomic

Sequencing Library protocols to amplify the V3 - V4 region. The DNA quality was measured by

PicoGreen and Nanodrop and 10 ng of gDNA was amplified using the following PCR

conditions: 94°C for 3 minutes followed by 35 cycles of 94°C for 15 seconds, 55°C for 45

seconds, and 72°C for 1 minute, followed by a final elongation step of 8 minutes at 72°C. The

barcoded fusion primer sequences used for amplifications were 519F:

5’ CCTACGGGNGGCWGCAG 3’ and 806R: 5’ GACTACHVGGGTATCTAATCC 3’. The

final purified product was quantified using qPCR according to the qPCR Quantification Protocol

Guide (KAPA Library Quantificatoin kits for Illumina Sequencing platforms) and qualified

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using the LabChip GX HT DNA High Sensitivity Kit (PerkinElmer, Massachusetts, USA). The

paired-end (2 × 300 bp) sequencing was done by Macrogen (Seoul, Korea) on the Illumina

MiSeq platform (Illumina, San Diego, CA).

Sequencing and Diversity Analysis

Overlapping reads were merged using fast length adjustment of short reads (Magoc and

Salzberg, 2011). Pre-processing and clustering of sequences to identify OTUs was done using

CD-HIT-OTU software (Li et al., 2012). Taxonomic composition of the samples from phylum to

species was generated using QUIIME UCLUST (Edgar 2010) against the Ribosomal Database

Project Release 11. Alpha diversity (rarefaction curve for OTUs, Chao1, and Shannon-Weaver

diversity indices) and beta diversity (unweighted principal coordinate analysis (PCoA) UniFrac

distance) were generated using the QIIME software package with a script

core_diversity_analyses.py (Caporaso et al., 2010). A nonparametric Monte Carlo test with no

Bonferroni correction was used to compare the within-treatment and between-treatment UniFrac

distances, which represent phylogenetic distances based on the presence/absence of OTUs

among samples (Park et al., 2016).


The experiment was a completely randomized design with 5 treatments and 4 replicates

per treatment. Continuous variables, such as relative abundance of bacteria, number of reads, and

species richness and diversity were analyzed using the GLIMMIX procedure of SAS. The model

included the treatment effect and means were separated using the PDIFF procedure of SAS,

which is based on the Fisher’s F-protected least significant difference test. Significant

differences were declared at P < 0.05. Pearson’s correlation coefficients were generated using

the R software (http://www.r-project.org). Significant correlations were declared at P < 0.10.

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Results

Fermentation quality indices, counts of E. coli O157:H7, and chemical composition of

the silages at silo opening (d 100) were reported previously in Chapter 3. Briefly, the pH was

lower (P < 0.05) in EC+LP and EC+LB silages (4.57 and 4.64, respectively) than in the Control

and EC silages (4.88 and 4.82, respectively), and the EC+PA silage had a similar (P > 0.10) pH

to inoculated silages. Compared to the Control, silages treated with EC+LP and EC+LB had

greater (P < 0.05) concentrations of lactate and acetate, respectively. Propionate was only

detected in EC+PA silage. Escherichia coli O157:H7 was eliminated from the silages once the

pH dropped below 5.0. This occurred earlier (16 vs. 100 d of ensiling) in EC+LP and EC+LB

silages because they produced more lactate and acetate throughout the fermentation than other

silages.

Sequencing Results and Bacterial Diversity

Sequencing of the V3 and V4 regions of the bacterial 16S rRNA gene of the 20 samples

yielded 1,702,729 reads with an average of 85,136 ± 4522 reads per sample after quality

filtering. Rarefaction curves plateaued in all the samples sequenced (figure not shown),

indicating that the number of reads was sufficient for identifying OTU. The average Good’s

coverage for all the samples was greater than 99%, indicating that the depth of sequencing was

adequate for reliable analysis of the bacterial community. Compared to the others, silages treated

with microbial inoculants had lower (P < 0.05) Shannon index values (Figure 4-1A), which

measures species diversity based on the number and evenness of species. The Chao1 index, a

measure of species richness based on the number of rare species was unaffected (P > 0.10) by

treatment (Figure 4-1B). The unweighted PCoA UniFrac plot revealed compositional differences

in the bacterial community of the 20 samples (Figure 4-2). The Control samples seemed to

cluster apart from most other samples, except one EC sample, and the Monte Carlo test revealed

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that the between-treatment variation (0.12) was greater (P < 0.01) than that (0.07) of within

treatment.

Comparisons at the Phylum Level

The microbial community structure of alfalfa silage was represented by 5 phyla (Table 4-

1). Phylum Firmicutes (73.3 ± 6.05) was more abundant than others, followed by Proteobacteria

(20.4 ± 3.80). Compared to the Control, EC treatment did not affect the abundance and

composition of bacteria at the phylum level. The abundance of Firmicutes in the Control and

EC+LB silages (79.5 and 77.4%, respectively) was greater (P < 0.05) than that in the EC+LP or

EC+PA silages (69.2 or 68.8%, respectively), whereas, that of Proteobacteria was greater in the

EC+LP and EC+PA silages (24.0 and 24.9%, respectively) than in the Control, EC and EC+LB

(16.5, 18.0, and 18.5% respectively) silages. The abundance of Bacteroidetes, Actinobacteria and

Cyanobacteria were similar for all treatments (P > 0.10).

Comparisons at the Family Level

Treatment effects on the percentage distribution of the 13 most abundant families that

represented 91.7% of the silage bacterial community are shown in Table 4-2. Lactobacillaceae

was the most abundant family followed by Leuconostocaceae. Compared to the Control, EC

treatment did not affect the abundance of bacteria at the family level, but treatment with EC+LP

and EC+LB increased (P < 0.05) the abundance of Lactobacillaceae and reduced (P < 0.05)

those of Leuconostocaceae and Enterococcaceae. Compared to the Control, EC+PA did not

affect (P > 0.10) the abundance of Lactobacillaceae but decreased (P = 0.01) those of

Leuconostocaceae and Enterococcaceae. Among the families belonging to phylum

Proteobacteria, the abundance of Methylobacteriaceae was less (P = 0.01) in EC+LP and EC+LB

silages whereas those of Enterobacteriaceae, Pseudomonadaceae and Xanthomonadaceae were

greater (P < 0.05) in EC+LP and EC+PA silages, compared with the Control. Also, the

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abundance of Sphingomonadaceae was greater (P =0.02) in EC+LP, EC+LB and EC+PA silages

relative to the Control.

Comparisons at the Genus Level

Treatment effects on the percentage distribution of the 16 most abundant genera,

accounting for approximately 90.3% of the silage bacterial community are shown in Table 4-3.

Pediococcus and Weissella were the most abundant genera in the Control silage. Compared to

the Control, EC treatment did not affect the abundance of bacteria at the genus level. Treatment

with EC+LP resulted in the greatest (P = 0.01) abundance of Lactobacillus (62.6%), followed by

EC+LB (47.8%), whereas Control, EC and EC+PA silages had similar (P > 0.50) and smaller (P

< 0.50) values than EC+LB silage (16.7, 16.8, and 8.98%, respectively). Relative to the Control,

EC+LP silage had lower (P < 0.05) abundance of Pediococcus (29.9 vs. 0.55%), Weissella (27.7

vs. 3.89%) and Methylobacterium (1.50 vs. 0.60%), and increased (P < 0.05) abundance of

Sphingomonas (1.86 vs. 2.93%), Pantoea (4.14 vs. 8.28%), Pseudomonas (1.26 vs. 2.44%) and

Erwinia (0.35 vs. 0.57%); whereas EC+LB silage contained less (P < 0.05) abundance of

Weissella (29.7 vs. 9.19%), and Methylobacterium (1.50 vs. 0.94%) relative to the Control.

Compared to the Control, treatment with EC+PA resulted in greater (P < 0.05) abundance of

Sphingomonas (1.86 vs. 3.61%), Pantoea (4.14 vs. 7.21%), Pseudomonas (1.26 vs. 2.75%), and

Erwinia (0.35 vs. 0.63%) but EC+PA did not affect the abundance of three most abundant

genera; Lactobacillus, Weissella and Pediococcus.

Comparisons at the Species Level

The relative abundance of bacterial species that accounted for more than 0.1% of the

bacterial community is shown in Table 4-4. Treatment effects on the abundance of the bacterial

species that were affected by treatments are shown in Table 4-5. Unknown Pediococcus and

Weissella species were the most abundant species in Control silages and the abundance of

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bacterial species was similar (P > 0.10) in Control and EC silages. The abundance of L.

plantarum and L. buchneri were greatest (P = 0.01) in EC+LP and EC+LB silages, respectively.

Relative to the Control, the abundance of L. acidipiscis, Weissella koreensis, Methylobacterium

gossipiicola, M. populi and unknown Lactobacillus and Weissella species, were reduced (P <

0.05), whereas that of unknown Pseudomonas species was increased in EC+LP and EC+LB

silages. Furthermore, EC+LP silage contained greater (P < 0.05) abundance of Pseudomonas

fulva and unknown Rhodococcus and Sphingomonas species, and less (P < 0.05) abundance of

an unknown Pediococcus species compared to the Control. The EC+PA silage contained lower

(P < 0.05) abundance of L. acidipiscis, unknown Lactobacillus and Weissella species, W.

koreensis, and M. populi, and increased abundance of an unknown Sphingomonas and

Pseudomonas species compared to the Control.

Correlations between Abundance of Bacteria and Fermentation Quality Indices

The correlations between the abundance of bacterial genera and silage fermentation

indices are presented in Table 4-6. Lactate concentration correlated positively with the

abundance of Lactobacillus (r = 0.56, P = 0.01), and negatively with the abundance of

Pediococcus (r = -0.41, P = 0.08). Abundance of genera Sphingomonas (r = -0.51), Pantoea (r =

-0.46), Pseudomonas (r = -0.45), and Stenotrophomonas (r = -0.38) were negatively correlated

(P < 0.10) with ammonia-N concentration. Silage pH was negatively correlated (P < 0.10) with

the abundance of Lactobacillus (r = -0.59), Sphingomonas (r = -0.66), Pantoea (r = -0.69),

Pseudomonas (r = -0.69), and Stenotrophomonas (r = -0.50), and positively correlated (P < 0.10)

with the abundance of Pediococcus (r = 0.44), Methylobacterium (r = 0.53), Weissella (r = 0.60),

and Erwinia (r = 0.40). Abundance of Sphingomonas (r = -0.49), Pantoea (r = -0.48),

Pseudomonas (r = -0.45), and Stenotrophomonas (r = -0.37) were negatively correlated (P <

0.10) with yeast and mold counts.

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Correlations between the abundance of known and unknown bacterial species and silage

fermentation quality indices are presented in Tables 4-7 and 4-8, respectively. As expected, the

abundance of L. buchneri was positively correlated (r = 0.42, P = 0.06) with acetate

concentration and negatively (r = -0.41, P = 0.06) correlated with yeast and mold counts. The

abundance of L. plantarum and Pseudomonas fulva were positively correlated with lactate

concentration (P ≤ 0.10) and negatively correlated with silage pH (P ≤ 0.10). The abundance of

L. acidipiscis (r = 0.52), Methylobacterium gossipiicola (r = 0.48), M. populi (r = 0.50) and

Weissella koreensis (r = 0.43) were positively correlated (P ≤ 0.10) with silage pH. The

abundance of L. acidipiscis (r = 0.41), M. populi (r = 0.50), and W. koreensis (r = 0.43) were

positively correlated (P ≤ 0.10) with yeast and mold counts. Acetate concentration was positively

correlated (r = 0.51, P = 0.02) with an unknown species and genus in family Xanthomonadaceae.

Lactate concentration was negatively correlated (P ≤ 0.10) with the abundance of unknown

species belonging to genera Rathayibacter (r = -0.37) and Pediococcus (r = 0.51), and positively

correlated (P ≤ 0.10) with those of unknown species belonging to genera Rhodococcus (r = 0.43)

and Pseudomonas (r = 0.53), and an unknown genus in family Xanthomonadaceae (r = 0.39).

Ammonia-N concentration was negatively correlated (P ≤ 0.10) with abundance of unknown

species belonging to genera Salana (r = -0.47), Frigoribacterium (r = -0.41), Pseudorhodobacter

(r = -0.44), Pantoea (r = -0.45), Stenotrophomonas (r = -0.44), Rhodococcus (r = -0.40),

Pseudomonas (r = -0.40) and Sphingomonas (r = -0.51). Abundance of unknown species

belonging to genera Saguibacter (r = -0.38), Pseudorhodobacter (r = -0.44), Pantoea (r = -0.79),

Stenotrophomonas (r = -0.72), Rhodococcus (r = -0.39), Sphingomonas (r = -0.68) and

Pseudomonas (r = -0.78) were negatively correlated (P ≤ 0.10) with pH. Whereas, unknown

species belonging to Pediococcus (r = 0.40), Erwinia (r = 0.48), Weissella (r = 0.60) and

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Lactobacillus (r = 0.53) had positive correlations (P ≤ 0.10) with pH. Abundance of unknown

species belonging to family Xanthomonadaceae and genera Stenotrophomonas and Pantoea were

negatively correlated (P ≤ 0.10) with yeast and mold counts.

Discussion

This experiment was conducted as part of a broader study aimed at understanding the

probiotic and food safety - enhancing effects of lactic acid bacterial silage inoculants. Silages

were inoculated with E. coli O157:H7 to monitor its fate during ensiling of alfalfa that had been

ensiled with or without additives. The pathogen was not detected in any silage at silo opening

because of the inhibitory effect of the low pH resulting from silage acidification (Chapter 3).

Bacterial Diversity and Composition

This study revealed the abundance and diversity of bacteria in alfalfa silage treated with

EC with or without microbial or chemical additives. The Illumina Miseq sequencing platform

used for the analysis is a powerful tool for studying phylogeny and taxonomy of complex

microbial communities of environmental samples (Gloor et al., 2010; Degnan and Ochman,

2012).

It is well known that majority of the bacterial community involved in lactic acid

fermentation in silage belong to phylum Firmicutes and to the genera Lactobacillus, Pedicoccus,

Lactococcus, Weissella and Leuconostoc (Pahlow et al., 2003; Pang et al., 2011). Our study

revealed that approximately 74.1 ± 3.28% of the alfalfa silage bacterial community belong to

phylum Firmicutes, consisting mainly of families Lactobacillaceae and Leuconostocaceae, and

genera Lactobacillus, Pediococcus, and Weissella. Approximately 20.4 ± 1.65% belonged to

phylum Proteobacteria, consisting mainly of families Enterobacteriaceae, Xanthomonadaceae,

Sphingomonadaceae and Aurantimonadaceae, and genera Sphingomonas, Pantoea,

Pseudomonas and Stenotrophomonas. Bao et al. (2016) assessed the bacterial community of

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alfalfa silage via single molecule real-time sequencing technology and reported that majority of

the species detected belong to genera Lactobacillus, Weissella, Pediococcus and Pantoea. In a

similar study, McGarvey et al. (2013) monitored the bacterial population before and after

ensiling of alfalfa forage and found that the abundance of Proteobacteria reduced from 89.6 to

26.9%, while that of Firmicutes increased from 8.1 to 70.6% after 40 days of ensiling, indicating

that silage fermentation involves a shift in the bacterial community from Proteobacteria to

Firmicutes.

The fact that the bacterial diversity and taxonomic composition of the EC silage did not

differ from that of the Control silage suggests that the microbiome shifts in the additive-treated

silages reflect the effects of the respective additives alone. Inoculation with microbial inoculants

has been employed for several years to enhance fermentation of silage (Stokes 1992; Whiter and

Kung, 2001). Microbial inoculants are added to dominate and control microbial events during

silage fermentation (Kung, 1998; Wang et al., 2006). Our results revealed that silage inoculation

with L. plantarum or L. buchneri increased the abundance of the respective bacteria, thereby,

reducing the diversity of the bacterial community (Polley et al., 2007; Allen et al., 2009). Similar

results were obtained by Eikmeyer et al. (2013) and Ogiy et al. (2015), who reported dominance

of inoculated L. buchneri and L. plantarum in grass and wheat silages, respectively. However, in

some cases, inoculated strains may be outcompeted by the indigenous microbial population

(Stevenson et al., 2006).

Silage inoculation with L. plantarum reduced the abundance of Weissella and

Pediococcus after 100 d of ensiling probably because of the faster rate of pH decline during

ensiling and lower pH at silo opening (Ogunade et al., 2016). Most species of Weissella are

obligate heterofermentative bacteria that produce lactate and acetate as major end products of

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sugar metabolism (Graf et al., 2016). Weissella species are considered early colonizers (Dellaglio

and Torriani, 1986) as they are outcompeted by acid-tolerant Lactobacillus species because of

the pH drop as fermentation progresses (Graf et al., 2016). That the abundance of L. buchneri

and species of Weissella, both of which are obligate heterofermenters, were reduced in EC+LP

silage partly explain greater lactate to acetate ratio usually observed in alfalfa silage treated with

L. plantarum compared with the untreated silage (Whiter and Kung, 2001; Kizilsimsek et al.,

2007; Ogunade et al., 2016).

Species of Pediococcus, such as Pediococcus pentosaceus, are used as silage inoculants

because they are acid-tolerant and are capable of synthesizing lactate (Garvie, 1986; Axelsson,

1998). Pediococcus species grow rapidly during the early stages of fermentation when the pH is

between 5 - 6.5 (Kung et al., 2003; McDonald et al., 1991). These characteristics allow

Pediococcus species to dominate the early stages of ensiling and promote a pH decrease, thereby,

stimulating the dominance of Lactobacillus species, which get more active and grow vigorously

as the pH drops. Lin et al. (1992) studied the succession of lactic acid bacteria during ensiling

periods of alfalfa and corn, and reported that P. pentosaceus was dominant at early stages of

fermentation, however, after 7 d of ensiling, L. homohiochii, L. brevis and L. gasseri dominated

the community. Therefore, some inoculants are formulated to contain both P. pentosaceus and L.

plantarum in order to increase the rate of acidification during the early and later stages of silage

fermentation, respectively (Lin et al., 1992).

Lactobacillus buchneri is primarily added to silage to ferment lactate to acetate and 1,2-

propanediol (Driehuis et al., 2001). As a potent antimycotic agent, acetate inhibits the growth of

spoilage yeasts, thereby, improving the aerobic stability of silage (Danner et al., 2003; Tabacco

et al., 2009). Treatment with L. buchneri improved the aerobic stability of corn, sorghum, wheat,

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barley, and alfalfa silages (Kung and Ranjit 2001; Weinberg et al. 2002; Kung et al., 2003).

However, acetic acid production has often been associated with DM loss in silage (Wilkinson

and Davies, 2012). In this study, irrespective of treatment applied, the alfalfa silage was stable

for the entire duration of the aerobic phase (168 h; Ogunade et al., 2016), however DM loss was

not measured. Like L. plantarum, silage treatment with L. buchneri increased the abundance of

Lactobacillus and reduced that of Weissella, but had no effect on abundance of Pediococcus. In a

similar study, Eikmeyer et al. (2013) studied the taxonomic profiles of ensiling microbial

communities of grass silage inoculated with L. buchneri. The authors observed greater

abundance of Lactobacillus genus and lower abundance of Weissella genus relative to the

untreated silage.

Chemical additives, such as formic acid, propionic acid-based products, and

formaldehyde are added as antifungal agents to improve the aerobic shelf life of silage by

reducing the population of yeasts and molds, which cause aerobic deterioration (Kung et al.,

1998, 2000). As expected, propionic acid treatment had no effects on the fermentation. In

addition, it did not affect bacterial diversity or abundance of Lactobacillus, Weissella and

Pediococcus, the three most abundant genera responsible for driving silage fermentation in this

study. However, treatment with propionic acid increased the abundance of bacteria belonging to

genera Pantoea, Pseudomonas and Sphingomonas and Erwinia.

Correlations between Bacterial Abundance and Fermentation Quality Indices

In support of previous findings (Whiter and Kung, 2001; Kizilsimsek et al., 2007),

abundance of L. plantarum was positively (r = 0.57) and negatively correlated (r = -0.49) with

lactate and pH, respectively. Abundance of bacteria belonging to genera Pantoea, Pseudomonas

and Sphingomonas were greater in EC+LP versus Control silages and their abundance correlated

negatively with pH (r = -0.69, -0.69, and -0.66, respectively) and ammonia-N concentration (r = -

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0.46, -0.45, and -0.51, respectively). High concentrations of ammonia-N in silage is usually

indicative of protein breakdown, and is typically caused by extensive plant protease activity

and/or clostridial fermentation (Kung and Shaver, 2001). Tao et al. (2012) showed that plant

proteolytic enzymes in alfalfa silage are active at pH 4.0 to 8.0, however, most of them had

higher activities at pH 5.0 to 6.0. The rate of pH decline during ensiling of alfalfa is slow

because of the high buffering capacity and low water-soluble carbohydrate content in this forage

(Kung et al., 2003; Muck, 1998). Consequently, alfalfa is particularly susceptible to extensive

proteolysis and clostridial fermentation (Kung et al., 2003). True protein degradation in alfalfa

silage has been successfully reduced by L. plantarum inoculation because of the rapid drop in pH

which inhibits proteolytic bacteria (Weinberg and Muck, 1996; Filya et al., 2007). While L.

plantarum inoculation may play a major role in true protein preservation in alfalfa, our results

suggest that Pantoea, Pseudomonas and Sphingomonas may also contribute to the effect because

their abundances were negatively correlated with ammonia-N concentration. Studies on waste

water treatment revealed greater abundance of bacterial species belonging to genera

Pseudomonas and Sphingomonas towards the onset of nitrification (Bal-Krishna et al., 2013), a

microbiological process whereby ammonia is oxidized to nitrite and then, to nitrate (Bal-Krishna

et al., 2013); however, silage studies have not involved these bacteria. Fitzgerald et al. (2015)

monitored ammonia-oxidizing microbial communities in reactors under low dissolved oxygen

conditions and reported that some species of Pseudomonas, Rhodococcus, and Sphingomonas are

involved in nitrification. Therefore, the negative correlations between Pseudomonas and

Sphingomonas, and ammonia-N concentration may be partly due to nitrification. However,

previous studies have not shown that nitrification by these bacteria can occur under anaerobic

conditions, and in this study, nitrate-N concentrations were well below the detection limit of 300

102

mg/kg or the recommended safe level in ruminant diets (< 1000 mg/kg nitrate-N, Rasby, 1988),

possibly because of the low abundances of the bacteria. Therefore, more research is needed to

elucidate the role of these bacteria in reducing silage ammonia-N concentrations.

The negative correlations of these same bacteria (Sphingomonas, Pantoea, and

Pseudomonas) with yeast and mold counts may reflect the inhibitory effect of propionic acid on

yeast and mold growth (Kung et al., 1998, 2000). Notably, compared to that for Control silage,

these bacteria were also more abundant (P ≤ 0.05) in EC+PA silage which also had the lowest (P

= 0.01) yeast and mold counts (Ogunade et al., 2016).

Methylobacterium and Erwinia were among the dominant genera in pre-ensiled alfalfa

forage (McGarvey et al., 2013; Ni et al., 2016). However, in agreement with our results, their

abundance was low (approximately 1%) after ensiling in the study of Ni et al. (2016). Species of

Methylobacterium are strictly aerobic and neutrophilic (Doronina et al., 2002; Knief et al., 2012),

supporting their positive correlation (r = 0.53) with pH and reduced (P ≤ 0.05) abundance in

silages treated with the microbial additives in this study. Certain species of Erwinia do not grow

below pH 5.0 (Shrestha et al., 2005), supporting their low abundance in the silage and the

positive correlation (r = 0.40) with pH. Although, the abundance of Erwinia was increased (P ≤

0.05) by EC+LP and EC+PA treatments, but this difference may not have practical significance

because of their low abundance (< 0.6%) and the small differences among the treatments.

Using 16S gene sequences for taxonomic assignment of lactobacilli to the species level

may be misleading because of the relatively little sequence diversity among lactobacilli (Berthier

and Elrhich, 1998; Felis and Dellaglio, 2007). Therefore, the species-level taxonomic assignment

in these and similar studies should be cautiously interpreted.

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Bacterial Candidates for Future Research

Silage inoculation with L. plantarum increased the abundance of the added L. plantarum

and that of Pseudomonas fulva by approximately 750 and 524%, respectively, relative to the

Control. Both L. plantarum and P. fulva had strong positive correlations with lactate

concentration. Pseudomonas fulva belongs to the Pseudomonas putida group (Anzai et al.,

2000), which, like L. plantarum, possesses lactate dehydrogenase (Nelson et al., 2002), a key

enzyme for homolactic fermentation (Gaspar et al., 2014). While L. plantarum has been

extensively studied for its role in silage homolactic fermentation, P. fulva has not, possibly

because of the associated public health and safety concerns. Human infections with P. fulva have

been documented in previous studies (Almuzara et al., 2010; Seok et al., 2010) and the bacterium

has caused meningitis in a young girl (Almuzara et al., 2010) and bacteremia in a 56-year-old

man (Seok et al., 2010), but it is susceptible to a wide range of antibiotics (Liu et al., 2014).

Adequate control measures and strict regulations will be required for evaluation or use of this

bacterium as a silage inoculant because of the possible health and safety risks.

This study shows perhaps for the first time the associations of some unknown silage

bacteria and key silage quality indices. For instance, an unknown bacterial species belonging to

family Xanthomonadaceae was positively correlated with lactate and acetate concentrations and

negatively correlated with yeast and mold counts. Furthermore, unknown bacterial species

belonging to genera Rhodococcus and Pseudomonas had strong positive correlations with lactate

concentration. In addition, unknown species belonging to genera Salana, Frigoribacterium,

Stenotrophomonas, Pantoea and Pseudorhodobacter had strong negative correlations with

ammonia-N concentration. These unknown bacteria should be identified, cultured and

investigated further to determine their roles in silage fermentation. To our knowledge, no

member of the aforementioned genera and/or families has been evaluated as a microbial additive.

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Further studies on these bacterial species are needed as at least some of them may be candidates

for development into new silage inoculants that can improve the preservation of forages.

Conclusions

This study revealed that 74.1 ± 3.28% and 20.4 ± 1.65% of the alfalfa silage bacterial

community belong to phylum Firmicutes and Proteobacteria, respectively. Lactobacillus,

Weissella and Pediococcus were the most abundant genera detected. Relative to the Control,

adding E. coli O157:H7 alone at ensiling did not affect the bacterial community composition of

the silage. The abundance of Firmicutes was reduced (115 and 116%) and that of Proteobacteria

was increased (145 and 151%) by EC+LP and EC+PA treatments, respectively. Treatment with

EC+LP resulted in the greatest abundance of Lactobacillus, but reduced those of Pediococcus,

Weissella, and Methylobacterium. Abundance of Weissella and Methylobacterium were also

reduced in the EC+LB silage compared to the Control. Application of propionic acid did not

affect the abundance of Lactobacillus, Weissella and Pediococcus. Unknown bacterial species

that were positively correlated with lactate and acetate concentrations, and negatively correlated

with ammonia-N concentration were detected. These unknown bacteria should be speciated and

examined for their respective potential roles in fermentation and true protein preservation in

alfalfa silage.

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Table 4-1. Relative abundance (%) of phyla isolated from alfalfa silage treated without (Control)

or with Escherichia coli O157:H7 (EC) alone, or EC and either of two bacterial

inoculants or propionic acid and ensiled for 100 d.

abMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; EC+LP = EC + 1 × 106 cfu/g

of L. plantarum R2014 (Lallemand Animal Nutrition, Milwaukee, WI); EC+LB = EC + 1 × 106

cfu/g of L. buchneri R1102 (Lallemand Animal Nutrition, Milwaukee, WI); EC+PA = EC + 2.2

g/kg of propionic acid

Treatment1 SE P-value


Actinobacteria 1.52 2.27 2.03 1.18 2.00 0.58 0.21

Bacteroidetes 1.76 2.81 3.67 2.33 2.91 1.20 0.63

Cyanobacteria 0.79 1.33 1.12 0.59 1.38 0.32 0.12

Firmicutes 79.5a 75.6ab 69.2b 77.4a 68.8b 3.28 0.02

Proteobacteria 16.5b 18.0b 24.0a 18.5b 24.9a 1.65 0.01

106

Table 4-2. Relative abundance (%) of the 13 most abundant families isolated from alfalfa silage


either of two bacterial inoculants or propionic acid and ensiled for 100 d.

Treatment1

SE

P-value Control EC EC+LP EC+LB EC+PA

Firmicutes

Lactobacillaceae 38.5b 45.5b 63.2a 67.9a 44.7b 4.03 0.01

Leuconostocaceae 29.7a 24.8ab 3.89d 9.19c 23.4b 2.31 0.01

Bacillaceae 0.69 3.21 1.20 0.11 0.30 1.57 0.27

Enterococcaceae 0.48a 0.64a 0.04b 0.07b 0.01b 0.12 0.01

Proteobacteria

Caulobacteraceae 0.12 0.18 0.08 0.09 0.19 0.07 0.13

Aurantimonadaceae 1.94 2.55 1.87 1.60 2.23 0.49 0.24

Hyphomicrobiaceae 0.14 0.19 0.10 0.10 0.12 0.05 0.16

Methylobacteriaceae 1.53ab 1.61a 0.61d 0.96c 1.26bc 0.16 0.01

Rhizobiaceae 1.80 1.80 1.55 1.33 1.84 0.29 0.27

Sphingomonadaceae 1.93c 2.35bc 3.00ab 2.78ab 3.68a 0.33 0.02

Enterobacteriaceae 4.55c 4.60c 9.70a 5.94c 8.02b 0.76 0.01

Pseudomonadaceae 1.26b 1.29b 2.44a 1.97ab 2.75a 0.50 0.03

Xanthomonadaceae 2.65c 2.75bc 3.78ab 3.21abc 4.00a 0.49 0.05

a–dMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; EC+LP = EC + 1 × 106 cfu/g



g/kg of propionic acid.

107

Table 4-3. Relative abundance (%) of 13 most abundant genera isolated from alfalfa silage


either of two bacterial inoculants or propionic acid and ensiled for 100 d.

Treatment1

SE

P-value Control EC EC+LP EC+LB EC+PA

Firmicutes

Lactobacillus 14.7c 16.8c 62.6a 47.8b 8.98c 4.43 0.01

Pediococcus 29.9ab 28.7ab 0.55c 20.1b 35.8a 5.17 0.01

Weissella 29.7a 24.8a 3.89c 9.19b 23.4a 4.32 0.01

Exigobacterium 0.29 0.48 0.30 0.12 0.37 0.14 0.17

Bacillus 0.14 0.41 0.20 0.11 0.26 0.14 0.25

Proteobacteria

Aureimonas 1.94 2.15 1.87 1.60 2.22 0.29 0.19

Methylobacterium 1.50ab 1.60a 0.60d 0.94dc 1.24bc 0.16 0.01

Rhizobium 1.80 1.79 1.55 1.33 1.84 0.29 0.27

Sphingomonas 1.86c 2.26bc 2.93ab 2.71abc 3.61a 0.45 0.02

Pantoea 4.14b 4.04b 8.28a 5.40b 7.21a 0.59 0.01

Pseudomonas 1.26b 1.29b 2.44a 1.97ab 2.75a 0.50 0.03

Stenotrophomonas 2.43 3.50 2.95 3.56 3.32 0.63 0.27

Erwinia 0.35c 0.35c 0.57ab 0.44bc 0.63a 0.09 0.02

a–dMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; EC+LP = EC + 1 × 106 cfu/g


cfu/g of L. buchneri R1102 (Lallemand Animal Nutrition, Milwaukee, WI); ECA = EC + 2.2


108

Table 4-4. Relative abundance of bacterial species isolated from alfalfa silage treated without


bacterial inoculants or propionic acid and ensiled for 100 d.

Genus Species Average abundance (%)

Salana Unknown 0.21 ± 0.09

Frigoribacterium Unknown 0.36 ± 0.13

Rathayibacter Unknown 0.13 ± 0.06

Sanguibacter Unknown 0.38 ± 0.17

Sphingobacterium Unknown 2.43 ± 1.65

Exiguobacterium acetylicum 0.31 ± 0.21

Kroppenstedtia Unknown 0.36 ± 0.70

Pediococcus Unknown 23.0 ± 14.2

Brevundimonas Unknown 0.13 ± 0.07

Aureimonas Unknown 2.04 ± 0.49

Devosia Unknown 0.13 ± 0.06

Rhizobium Unknown 1.66 ± 0.31

Rhodobacteteraceae* Unknown 0.18 ± 0.08

Pseudorhodobacter Unknown 0.36 ± 0.10

Sphingomonas Unknown 0.16 ± 0.06

Erwinia Unknown 0.47 ± 0.16

Pantoea Unknown 5.81 ± 2.02

Xanthomonadaceae* Unknown 0.16 ± 0.07

Pseudoxanthomonas Unknown 0.13 ± 0.07

Stenotrophomonas Unknown 2.91 ± 0.75

Lactobacillus diolivorans 10.8 ± 18.9

Lactobacillus plantarum 16.0 ± 24. 1

Lactobacillus acidipiscis 0.31 ± 0.53

Lactobacillus Unknown 1.73 ± 2.06

Rhodococcus Unknown 0.34 ± 0.20

109

Table 4-4. Continued.

*Unknown genus in the indicated family

Genus Species Average abundance (%)

Methylobacterium gossipiicola 0.92 ± 0.35

Weissella Unknown 17.4 ± 9.96

Weissella Koreensis 0.81 ± 0.87

Methylobacterium Populi 0.26 ± 0.09

Sphingomonas Unknown 2.52 ± 0.80

Pseudomonas Unknown 3.26 ± 1.67

Pseudomonas Fulva 0.39 ± 0.36

110

Table 4-5. Relative abundance (%) of bacterial species isolated from alfalfa silage treated


two bacterial inoculants or propionic acid and ensiled for 100 d.

Treatment1

SE

P-

value Control EC EC+LP EC+LB EC+PA

Lactobacillus buchneri 1.84b 2.24b 0.19b 46.8a 3.19b 2.27 0.01

Lactobacillus plantarum 8.03b 10.8b 61.8a 0.47c 5.14b 3.09 0.01

Lactobacillus acidipiscis 1.13a 1.21a 0.00b 0.01b 0.22b 0.26 0.01

Lactobacillus* 4.59a 2.58ab 0.63bc 0.46c 0.42c 0.95 0.01

Rhodococcus* 0.23b 0.40ab 0.55a 0.26b 0.31b 0.09 0.02

Methylobacterium gossipiicola 1.27a 1.44a 0.51c 0.75bc 1.11ab 0.17 0.01

Weissella* 27.8a 23.4ab 3.77d 8.89c 23.1b 2.19 0.01

Weissella koreensis 1.89a 1.41a 0.12b 0.30b 0.32b 0.39 0.01

Methylobacterium populi 0.35a 0.35a 0.16b 0.21b 0.21b 0.03 0.01

Sphingomonas* 1.71c 2.08bc 2.78ab 2.58abc 3.43a 0.42 0.01

Pseudomonas* 0.06c 0.09c 0.41ab 0.23b 0.16bc 0.06 0.01

Pseudomonas fulva 0.17b 0.27b 0.89a 0.37b 0.31b 0.20 0.02

Pseudomonas* 1.04b 0.93b 1.14b 1.40b 2.28a 0.33 0.01

a–cMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; EC+LP = EC + 1 × 106 cfu/g




*Unknown species in the indicated genus

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Table 4-6. Pearson’s correlation between abundance of bacterial genus and fermentation indices

of alfalfa forage ensiled for 100 d.

Genus Acetate

(% DM)

Lactate (%

DM)

Ammonia-N

(% DM)

pH Yeast and mold

(log cfu/g)

Lactobacillus 0.09 0.56* -0.27 -0.59* -0.01

Pediococcus 0.07 -0.41* 0.22 0.44* -0.10

Weissella 0.04 -0.29 0.04 0.60* 0.12

Aureimonas -0.01 -0.03 -0.24 0.03 0.24

Methylobacterium 0.06 -0.26 -0.03 0.53* 0.34

Rhizobium -0.26 -0.04 -0.12 0.16 0.16

Sphingomonas 0.31 0.22 -0.51* -0.66* -0.49*

Pantoea 0.07 0.32 -0.46* -0.69* -0.48*

Pseudomonas 0.25 0.20 -0.45* -0.69* -0.45*

Exigobacterium -0.09 -0.06 -0.18 0.08 0.14

Bacillus -0.22 -0.04 -0.06 0.02 0.24

Erwinia 0.13 0.05 -0.06 0.40* -0.12

Stenotrophomonas 0.26 0.35 -0.38* -0.50* -0.37*

*Significant (P ≤ 0.10) Pearson’s correlation coefficient

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Table 4-7. Pearson’s correlation between abundance of bacterial species and fermentation indices

of alfalfa forage ensiled for 100 d.

Acetate

(% DM)

Lactate

(% DM)

Ammonia-

N (% DM) pH

Yeast and mold

(log cfu/g)

Lactobacillus buchneri 0.42* -0.02 -0.03 -0.18 -0.41*

Lactobacillus plantarum -0.24 0.57* -0.23 -0.49* 0.19

Lactobacillus acidipiscis 0.06 -0.06 0.01 0.52* 0.41*

Methylobacterium

gossipiicola 0.07 -0.28 -0.07 0.48* 0.29

Methylobacterium populi 0.01 -0.14 0.15 0.63* 0.50*

Weissella koreensis -0.05 -0.04 -0.19 0.46* 0.43*

Pseudomonas fulva -0.05 0.54* -0.32 -0.58* -0.14*

Exiguobacterium acetylicum -0.16 -0.11 -0.18 0.02 -0.21


113

Table 4-8. Pearson’s correlation between abundance of unknown bacterial species and

fermentation indices of alfalfa forage ensiled for 100 d.

Genus1

Acetate

(% DM)

Lactate

(% DM)

Ammonia-N

(% DM) pH

Yeast and

mold (log

cfu/g)

Salana -0.14 0.03 -0.47* -0.15 0.03

Frigoribacterium -0.23 -0.06 -0.41* -0.11 0.04

Rathayibacter -0.2 -0.37* -0.26 0.02 0.09

Sanguibacter -0.31 0.23 -0.12 -0.38* 0.07

Sphingobacterium -0.02 0.12 -0.29 -0.32 -0.27

Kroppenstedtia -0.18 -0.05 0.61* 0.33 0.09

Pediococcus -0.01 -0.51* 0.35 0.40* -0.22

Brevundimonas -0.01 -0.33 -0.16 0.11 -0.16

Aureimonas -0.12 -0.15 -0.16 -0.03 0.15

Devosia -0.33 -0.25 0.14 0.34 0.28

Rhizobium -0.20 -0.25 0.05 0.11 0.04

Rhodobacteteraceae+ -0.04 -0.09 -0.09 0.07 0.19

Pseudorhodobacter -0.17 0.01 -0.44* -0.38* -0.09

Erwinia 0.08 -0.05 -0.03 0.48* -0.14

Pantoea 0.03 0.20 -0.45* -0.79* -0.48*

Xanthomonadaceae+ 0.51* 0.39* -0.50* -0.59 -0.46*

Pseudoxanthomonas 0.03 -0.20 -0.17 0.09 -0.17

Stenotrophomonas 0.28 0.25 -0.44* -0.72* -0.68*

Lactobacillus -0.01 0.01 -0.15 0.53* 0.22

Rhodococcus -0.18 0.43* -0.40* -0.39* 0.29

Weissella 0.05 -0.31 0.07 0.60* 0.25

Sphingomonas 0.30 0.23 -0.51* -0.68* 0.03

Pseudomonas 0.15 0.53* -0.40* -0.78* -0.11

1Unknown species in this genus +Unknown genus in the indicated family


114

A

B

Figure 4-1. A. Estimate of alpha diversity, Treatment P-value = 0.01. B. Species richness

calculated as the Chao1 index, Treatment P–value = 0.51. Bars with different letters

differed (P < 0.05); Control = distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7;

EC+LP = EC + 1 × 106 cfu/g of L. plantarum R2014 (Lallemand Animal Nutrition,

Milwaukee, WI); EC+LB = EC + 1 × 106 cfu/g of L. buchneri R1102 (Lallemand

Animal Nutrition, Milwaukee, WI); ECA = EC + 2.2 g/kg of propionic acid. Error

bars represent SE.

2.5

2.7

2.9

3.1

3.3

3.5

3.7

3.9


Shan

non i

ndex

Treatment

a a

b

b

a

60

65

70

75

80

85


Chao

index

Treatment

115

Figure 4-2. Unweighted UniFrac Principal Coordinate Analysis (PCoA) plot of individual

samples in each treatment. This PCoA plot indicates the phylogenetic distance

(variation) between samples using two principal coordinates (PC1 and PC2). The

percentage variation explained by each PC is indicated on each axis. Control =

distilled water; EC = 1 × 105 cfu/g of E. coli O157:H7; EC+LP = EC + 1 × 106 cfu/g

of L. plantarum R2014 (Lallemand Animal Nutrition, Milwaukee, WI); EC+LB = EC

+ 1 × 106 cfu/g of L. buchneri R1102 (Lallemand Animal Nutrition, Milwaukee, WI);

ECA = EC + 2.2 g/kg of propionic acid.

116

CHAPTER 5

FATE OF Escherichia coli O157:H7 AND BACTERIAL DIVERSITY IN CORN SILAGE

CONTAMINATED WITH THE PATHOGEN AND TREATED WITH MICROBIAL AND

CHEMICAL ADDITIVES

Background

Escherichia coli O157:H7 is a foodborne pathogen that causes various forms of enteritis

in humans ranging in severity from bloody diarrhea to hemolytic uremic syndrome and even

death (Kaper et al., 2004). This pathogen is implicated in about 100,000 illness cases and 90

deaths annually in the US (Mead et al., 1999, Scallan et al., 2011). The estimated annual cost

associated with EC illnesses can be as high as $405 million (Frenzen et al., 2005).

Cattle harbor and shed EC in their manure (Lahti et al., 2003; Mir et al., 2015) and are

considered the main reservoirs of the pathogen (Cernicchiaro et al., 2013). Up to 30% of all

cattle are asymptomatic carriers of EC (Callaway et al., 2006, Reinstein et al., 2007) and

livestock feeds are considered to be the vehicle of transmission among livestock (Hancock et al.,

2001; Davis et al., 2003). Corn silage forms the bulk of most dairy cattle rations in the US and it

can be contaminated with EC when untreated manure is used to fertilize corn plants that are

subsequently ensiled and fed to cattle and or after silo opening via the manure and saliva of cattle

(Russell et al., 2000; Keen and Elder, 2002; Smith et al., 2005).

The low pH achieved during ensiling of corn eliminates pathogenic EC that is present at

ensiling (Pedroso et al., 2010; Duniere et al., 2011). At silo opening, aerobic conditions cause

increases in pH (Weinberg et al., 1995) via stimulation of the growth of lactate-utilizing yeasts.

The resulting elevated pH can facilitate growth and spread of EC (Pedroso et al., 2010).

Whereas, in silages contaminated with the pathogen after aerobic exposure, maintenance of a

low pH (below 4.0) because of application of L. buchneri at ensiling curtailed the growth of EC

(Pedroso et al., 2010). In contrast, Duniere et al. (2011) showed that application of L. buchneri at

117

the time of ensiling did not prevent the growth of EC when aerobically exposed silages were

contaminated with a pathogenic EC. Therefore, more studies are needed to ascertain the

inhibitory effect of L. buchneri or other additives on EC-contaminated aerobically exposed

silage. Such information is critically needed to reduce or prevent cycling of EC among animals

and or people.

Propionic acid is a strong antifungal agent (Woolford, 1975) that can inhibit the growth

of silage yeasts and molds, which are responsible for the rise in pH of silages during aerobic

exposure (Kleinschmit et al., 2005). We hypothesized that by maintaining a low pH because of

inhibition of the growth of lactate-utilizing yeasts that increase the pH after silo opening,

propionic acid would be effective at killing the pathogen during the aerobic phase. To our

knowledge, no experiments have examined the use of propionic acid to prevent the growth of EC

during or after ensiling. Furthermore, no studies have simultaneously compared the efficacy of

inhibiting the growth of EC on silage with propionic acid and obligate or facultative

heterofermentative lactic acid bacteria (LAB) inoculants. Therefore, this study was aimed at

examining the effectiveness of propionic acid and microbial inoculants containing either L.

plantarum or L. buchneri at controlling EC in corn silage during the ensiling and feedout stages.

Most studies that assessed the effects of silage additives on the microbial community

composition of silage have either been based on culture-based techniques (Ercolini, 2004), real

time PCR (Stevenson et al., 2006), or characterization of denaturing gradient gel electrophoresis

(DGGE) bands, which all underestimate the microbial community composition (Kraut-Cohen et

al., 2016). The application of 454 pyrosequencing has increased our understanding of complex

microbial communities (Degnan and Ochman, 2012) such as the rumen. Furthermore, the

Illumina MiSeq sequencing system, which gives higher throughput per run, lower error rates, and

118

greater depth and breadth of coverage than the 454 pyrosequencing system, at much lower costs

(Degnan and Ochman, 2012; Frey et al., 2014), has enhanced our understanding of microbial

communities. A recent study applied a newer sequencing technology, called single molecule

real-time sequencing technology, to examine the bacterial profile of alfalfa silage because of its

power to generate long sequence reads (Bao et al., 2016), however, this platform suffers from

high error rate (Quail et al., 2012). To our knowledge, no studies have used 454 pyrosequencing

or Illumina MiSeq sequencing system to examine the silage microbiome. In addition, no studies

that examined inhibitory effects of silage additives on pathogens have characterized the

microbiome of silage. Therefore, the second objective of this study was to understand how

propionate and inoculants affected the bacterial community composition in corn silage using the

Illumina sequencing platform.


The Biological Safety Office, Division of Environmental Health and Safety, and the

Institutional Animal Care and Use Committee (IACUC) of the University of Florida approved all

procedures for this study (protocol no. 201408675).

Forage and Treatments

Whole-plant corn forage (D598R50, Dyna Gro, East Lansing, MI), harvested with a

forage chopper (Claas 940, Claas of America Inc., Omaha, NE) at approximately 35% DM and

chopped to a theoretical length of cut of 10 mm, was treated with the following: (1) Distilled

water (Control); (2) 1 × 105 cfu/g Escherichia coli O157:H7 EDL933 alone (ECCH); (3) ECCH

+ 1 × 106 cfu/g Lactobacillus plantarum R2014 (ECLP); (4) ECCH + 1 × 106 cfu/g Lactobacillus

buchneri R1102 (ECLB); (5) ECCH + 2.2 g/kg (fresh weight) of propionic acid, containing

99.5% of the acid (ECA). Escherichia coli O157:H7 EDL933 was supplied by University of

Florida Emerging Pathogens Institute (Gainesville, Florida), the silage inoculant bacteria were

119

supplied by Lallemand Animal Nutrition (Milwaukee, WI), and the propionic acid was

purchased from Sigma-Aldrich (St. Louis, MO). Bacterial counts and their viability in the

inoculants were verified before silage inoculation by plating on de Man-Rogosa-Sharpe (MRS)

agar (Difco Laboratories, Inc., Detroit, MI, 288210), at 37°C for 48 h, and appropriate amounts

of the inoculants were added to achieve the desired application rate. The E. coli O157:H7 was

grown on Luria-Bertani (LB; Sigma-Aldrich, St. Louis, MO, L3022) broth overnight for 18 h at

35°C. The next day, the culture was diluted (1:100) in fresh LB broth and again incubated at

37°C for 4 h. The cells in the suspension within 16-mm pathlength tubes were adjusted to an

optical density of 0.50 at 600 nm (GENESYS 20 spectrophotometer; Thermo Fisher Scientific-

Oxoid Ltd., Basingstoke, UK, 4001-000). Bacterial counts in the suspension were confirmed by

direct plating on Sorbitol-MacConkey agar (SMAC; Sigma-Aldrich, St. Louis, MO, 83339)

supplemented with cefixime (0.05 mg/L, Sigma-Aldrich, St. Louis, MO, 18588) and potassium

tellurite (2.5 mg/L, Sigma-Aldrich, St. Louis, MO, 60539) to make CT-SMAC after incubation

at 35°C for 24 hours. The bacterial inoculants and propionic acid were diluted in distilled water

and applied to the forage at 3 mL/kg of fresh forage resulting in application of 1 × 106 cfu/g and

2.2 g/kg of bacterial inoculants and propionic acid, respectively. Before inoculation, the EC

suspension was re-plated on CT-SMAC to ensure accurate application rates. The EC suspension

was mixed with 5 mL of water and applied to the fresh forage at 1 × 105 cfu/g. Equal quantities

of water (9.5 mL each) were applied to the treated and control forages. Treatments were applied,

using a hand-held garden sprayer, to each of quadruplicate samples per treatment separately (1.5

kg each). Each treated sample was hand-mixed, manually packed into a 14 × 21 cm, thick-

walled plastic bag (International Plastics Inc., Greenville, SC, PB61430) which was vacuumed

(Canister Vacuum 4122, Bissell Inc., Grand Rapids, MI) and sealed with silo ties (Uline Corp.,

120

Pleasant Prairie, WI, S12356). The silages in the bags were ensiled for 3 and 7 d at 20°C after

which subsamples were analyzed for EC, pH, organic acids and ammonia-N. Additionally, 4.5

kg of the forage was treated with each additive or nothing (Control) in quadruplicate and packed

into 20-L plastic buckets lined with thick-walled plastic bags and ensiled for 120 d at ambient

temperature (25°C). Subsamples of these silages were analyzed for EC, pH, organic acids,

ammonia-N, LAB, yeast and mold counts, and aerobic stability, Furthermore, subsamples from

day 0 and 120 were analyzed for chemical composition.

Survival of E. coli O157:H7 in Aerobically Exposed Silages

After the bucket silos were opened on d 120, about 1 kg of silage from each replicate was

transferred to an open-top 20 L plastic bucket, re-inoculated with 5 × 105 cfu/g of EC and stored

at 20°C for 24 h. Subsamples were taken after 6, 12, 18 and 24 h to assess changes in EC counts.

Furthermore, 2 kg of silage from each d 120 replicate was placed in a different open-top 20 L

plastic bucket. Wireless sensors (Onset Computer Corporation, Cape Cod, MA) that recorded

silage temperatures at 30 min intervals were placed in the geometrical center of the silage mass

in each bucket. Aerobic stability was defined as the time (h) for the silage temperature to exceed

2.0°C above the ambient temperature. After 168 h of aerobic exposure, silages were re-

inoculated with 5 × 105 cfu/g of EC on d 127 and 24 h later (d 128), the population of the

pathogen on the silages was counted.

Laboratory Analyses

Silage subsamples (20 g each) taken on d 120 were diluted with 180 mL of distilled water

and homogenized for 1 minute in a stomacher (UL Lab-Blender 400, Seward Laboratory,

London, UK) to obtain silage extracts for further analyses. The suspension was filtered through 2

layers of cheesecloth and immediately analyzed for pH and counts of EC, LAB, yeasts and

molds. Enumeration of EC was done by serial dilution in buffered peptone water (BPW; peptone

121

(10 g/L) and sodium chloride (5 g/L); Thermo Fisher Scientific-Oxoid Ltd., Basingstoke, UK,

CM0009) followed by pour plating on duplicate plates of CT-SMAC agar (Zadik et al., 1993).

Plates were incubated for 24 h at 35°C and sorbitol-negative colonies were presumptively

identified as E. coli O157:H7 and confirmed by latex agglutination (Thermo Fisher Scientific-

Oxoid Ltd., Basingstoke, UK, DR0120M; March and Ratnam, 1989). The detection limit for EC

counts was 10 cfu/g. An aliquot of the serially diluted extract was also plated on malt extract

(Thermo Fisher Scientific-Oxoid, CM0059B) and de Man, Rogosa, Sharpe agars at 32°C for 72

and 48 h for enumeration of yeasts and molds, and LAB, respectively. The pH of silage extracts

was measured using a digital pH meter (Accumet AB15, Fisher Scientific). For organic acid and

ammonia-N analyses, an aliquot of the silage extract was centrifuged at 1,800 × g for 15 min at

4°C. The supernatant was analyzed for ammonia-N using an Auto Analyzer (Technicon,

Tarrytown, NY) and an adaptation of the Noel and Hambleton (1976) procedure that involved

colorimetric N quantification, and for organic acids using a High-Performance Liquid

Chromatograph system (Hitachi L2400, Tokyo, Japan) coupled to a UV detector set at 210 nm

and an Aminex HPX-87H column (Bio-Rad Laboratories, Hercules, CA) with 0.015 M sulfuric

acid mobile phase and a flow rate of 0.7 mL/min at 45°C. The DM content of the fresh forages

and silages was determined by drying samples in a forced-air oven at 55°C for 48 h. Dried

samples were ground to pass the 1-mm screen of a Wiley mill (Thomas Scientific, Swedesboro,

NJ). The procedures of Van Soest et al. (1991) were used for NDF and ADF analyses. Heat-

stable amylase was used in the NDF analysis and the results were expressed on a DM basis

inclusive of residual ash. Crude protein was calculated by multiplying N measurements obtained

from Kjeldahl digestion by 6.25.

122

Antibacterial Activity of the Silage Extracts

Antibacterial activity against EC of the silage extracts was determined by the agar disk

diffusion method of Bauer et al. (1966). The EC isolates were cultured in LB broth and adjusted

to an optical density of 0.5 at 600 nm and the inoculum was prepared on CT-SMAC agar. Silage

extracts were centrifuged at 5,000 × g for 30 min at 4°C and the cell-free supernatants were

collected. Two sterile cloning discs (6.0-mm diameter; cat. no. 231039, Becton Dickinson Co.,

Franklin Lakes, NJ) were immersed in the supernatants for 20 s using flame-sterilized forceps

and then placed on the surface of CT-SMAC agar plates containing EC. Plates were prepared in

duplicate and incubated for 24 h at 35°C. The diameters of zones of inhibition around the paper

discs were measured with a ruler. In addition, the pH of the cell-free supernatants of the silage

extracts was adjusted to pH 5.0 using 2 N NaOH and tested for antibacterial activity against EC.

DNA Extraction and Sequencing

To understand the influence of EC with or without silage additives on the composition

and diversity of the silage microbial community, bacterial diversity analysis of the d 120 silage

samples was performed using the Illumina MiSeq platform (Illumina, San Diego).

Approximately 100 g of each sample was added to 200 mL of PBS + 0.01% Tween-80 in a

sterile 1-L Erlenmeyer flask. The mixture was sonicated for 45 minutes using the Sonicor SC-

150 (Sonicor Instrument Corporation, Copiague, NY) followed by centrifugation at 9000 × g for

15 minutes to create a pellet for subsequent DNA extraction.

Total metagenomic DNA was extracted from the resulting pellet using a PowerLyzer

PowerSoil DNA isolation kit (MO BIO Laboratories, Inc., Carlsbad, CA). Bead beating (0.1 mm,

glass beads; Bullet Blender Storm 24, Averill Park, NY) was used for mechanical disruption of

bacterial cells. The purity and concentration of gDNA were measured with a spectrophotometer

123

(Nanodrop 115 1000, Thermo Fisher Scientific-Oxoid Ltd., Basingstoke, UK) and the integrity

of DNA was verified by agarose (0.7%) gel electrophoresis.

Each replicate sample was prepared for sequencing according to the Illumina 16S

metagenomic sequencing library protocols to amplify the V3 and V4 regions (Klindworth et al.,

2013). The DNA quality was measured by using PicoGreen (Invitrogen, Carlsbad, USA) and

Nanodrop (Thermo Scientific, Waltham, MA). Ten nanograms of the gDNA was PCR-amplified

with the following barcoded fusion primer sequences: 519F (5’-CCTACGGGNGGCWGCAG-

3’), 806R (5’-GACTACHVGGGTATCTAATCC-3’). The purified product was quantified using

qPCR according to the qPCR Quantification Protocol Guide (KAPA Library Quantification kits

for Illumina sequencing platforms, Illumina, San Diego) and qualified using the LabChip GX HT

DNA High-Sensitivity Kit (PerkinElmer, Massachusetts, USA). The paired-end (2 × 300 bp)

sequencing was performed by Macrogen Inc. (Seoul, Korea) using the Illumina MiSeq platform

(Illumina, San Diego).

Data and Statistical Analysis

The experiment had a completely randomized design with 5 treatments and 4 replicates

per treatment. All microbial data were transformed to log units and were presented on a wet

weight basis. Data were analyzed using the GLIMMIX procedure of SAS (SAS Institute Inc.,

Cary, NC) and the model for analyzing data collected over time included effects of treatment,

day and treatment × day interaction. When treatment × day interactions were significant, effects

of treatment on each day were examined. The model for analyzing data collected at one-time

point only included the treatment effect. Means were separated using the Tukey test. Significant

differences were declared at P < 0.05.

For the bacterial community composition data, CD-HIT was used for clustering of

sequences to identify Operational Taxonomic Units (OTUs) (Li et al., 2012). The taxonomic

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composition of each sample was generated using QIIME UCLUST (Edgar, 2010) against the

Ribosomal Database Project Release 11 (Cole et al., 2014). Alpha diversity (rarefaction curve for

OTUs, Chao1, and Shannon indices) and beta diversity (unweighted principal coordinate

analysis (PCoA) UniFrac distance) were generated using the QIIME software package with a

script core_diversity_analyses.py (Caporaso et al., 2010). A nonparametric Monte Carlo test was

used to compare the within-treatment and between-treatment UniFrac distances. Variables, such

as relative abundance of bacteria, number of reads, and species richness and diversity, were

analyzed using the GLIMMIX procedure of SAS and a model that included the treatment effect.

Differences between means were determined using the Tukey test. Significant differences were

declared at P < 0.05.

Results

Anaerobic Phase

The DM, CP, NDF, and ADF concentrations of the forage used for each treatment before

ensiling were similar and averaged 35.2 ± 0.44 (%, P = 0.75), 8.54 ± 0.23 (% DM, P = 0.16),

41.7 ± 0.78 (% DM, P = 0.68) and 25.1 ± 0.31 (% DM, P = 0.98) (Table 5-1). The pH of the

ECA corn forage was lower (P = 0.01) than those of other silages after the treatments were

applied.

After 3 d of ensiling, all silages had pH values below the threshold of 4.0. The pH values

remained low for the entire ensiling duration (Figure 5-1), such that the pH at final silo opening

(120 d) was similar for all silages (Table 5-2). Escherichia coli O157:H7 was below the

detection limit after 3 d of ensiling in ECCH, ECLP and ECLB silages suggesting that it had

been eliminated. Although it was detected in the ECA silage on d 3, EC was below the detection

limit in all d 7 silages (Figure 5-2), though it is possible that selective enrichment of samples

may have led to recovery of EC.

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By d 7 of ensiling, the ECLP silage had greater (P = 0.01) lactate concentration than

other silages (Figure 5-3), however, at final silo opening (d 120), its lactate concentration was

similar to those of Control, ECCH and ECA silages (Table 5-2). The acetate concentration was

similar for all silages at the initial stages of ensiling (on d 3 and 7), however, the concentration

was higher in the ECLB silage than other silages at d 120 (Figure 5-4; Table 5-2). Propionate

was detected only in the ECA silage (0.41% DM), reflecting the addition at ensiling, whereas

butyrate was not detected in any silage. Ammonia-N concentration was higher (P < 0.05) in

ECLB silage than ECLP silage at final silo opening (Table 5-2). Yeast and mold counts were

lower (P = 0.01) in the ECLB and ECA d 120 silages (3.43 and 2.92 log cfu/g, respectively) than

in corresponding Control, ECCH and ECLP silages (5.08, 5.30 and 4.91 log cfu/g, respectively).

Lactic acid bacterial counts were unaffected (P = 0.52) by treatment.

Aerobic Phase

Escherichia coli O157:H7 was below the detection limit in silages within 6 h of re-

inoculation with the pathogen at silo opening (d 120). Except for the ECA silage, EC was

detected on d 128 in all silages that had been re-inoculated with the pathogen on d 127 (Figure 5-

5). In addition, the d 128 ECCH and ECLP silages had greater pH values (5.67 and 6.13,

respectively; P < 0.05) and EC counts (5.39 and 5.30 log cfu/g, respectively; P = 0.01) than the

ECLB and ECA silages, which had pH values of 4.24 and 3.96, and had approximately 10,000-

fold lower and no EC, respectively. These responses suggest that the lower pH of the ECLB and

ECA silages prevented the growth of EC, which was facilitated by pH values that exceeded 5.0.

The ECLB and ECA silages also had greater (P = 0.001) aerobic stability than Control, ECCH

and ECLP silages (Figure 5-6).

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Antibacterial Activity

The cell-free supernatants from all silage extracts that were not pH-corrected produced a

similar zone of inhibition (4.15 ± 0.71 mm) against EC, whereas no zone was detected for pH-

corrected supernatants of the extracts.

Bacterial Community Composition and Diversity

Sequencing of the V3 to V4 regions of the bacterial 16S rRNA gene resulted in 1,250,072

reads, with an average of 62,504 ± 2,642 reads per sample after quality filtering. Coverage

values were greater than 99% for all the samples. Rarefaction curves plateaued in all samples

sequenced, indicating that the number of reads used in analysis was sufficient in identifying

OTU (figure not shown). The unweighted PCoA UniFrac plot showed that compositional

differences (P = 0.05) existed among the bacterial communities of the different silages (Figure 5-

7), and the Monte Carlo test revealed that the between-treatment variation (0.19) in the

unweighted UniFrac distance was greater (P < 0.001) than the within-treatment distance (0.09).

Relative to the Control, the ECLP silage resulted in a lower (P < 0.05) Shannon index, which

measures diversity based on the number and evenness of species (Table 5-3). The Chao1 index, a

measure of the richness of the bacterial species based on number of rare species, was unaffected

by treatment (Table 5-3).

Five phyla (Actinobacteria, Bacteroidetes, Cyanobacteria, Firmicutes, and

Proteobacteria) were detected in all samples. Firmicutes and Proteobacteria accounted for

approximately 99.9% of the bacterial communities (Table 5-3). Relative to the Control (97.8%

Firmicutes and 2.07% Proteobacteria), ECLP and ECA silages had greater (P < 0.05) OTU

abundance of Firmicutes (99.0 and 98.7%, respectively) and lower abundance of Proteobacteria

(0.98 and 1.18%, respectively). It is important to note that sequences that seemed to represent

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Cyanobacteria may represent chloroplasts, whose 16S genes have much homology to those of

Cyanobacteria (Hanshew et al., 2013).

At the genus level, six genera (Lactobaccillus, Sphingobacterium, Weissella,

Acinetobacter, Stenotrophomonas, Serratia) accounted for approximately 99% of the bacterial

community (Table 5-3). Compared to the Control (94.9%), ECLP and ECA silages had greater

(P < 0.05) abundance of Lactobacillus (98.3 and 97.8%, respectively). The ECLB silage

contained greater (P < 0.01) abundance of Acinetobacter and Weissella (1.16% and 2.39%,

respectively) than other silages.

Among the Lactobacillus species, the ECLP silage contained greater (P < 0.05)

abundance of L. diolivorans (96.8%) and L. plantarum (0.98%) when compared to the ECLB

(93.8 and 0.35%, respectively) and control silages (94.3 and 0.52%, respectively).

Discussion

Anaerobic Phase

The elimination of EC from the corn silage during the early part of the ensiling phase is

largely attributable to the low pH achieved because of rapid accumulation of organic acids.

Previous studies have shown that EC was eliminated in corn and barley silages, and alfalfa silage

when the pH was below 4.0 and 5.0, respectively (Chapter 3; Bach et al., 2002; Chen et al.,

2005). While the disappearance of EC was hastened by microbial inoculation of alfalfa silage

because of the slow pH decline, EC was rapidly eliminated during ensiling of corn, with or

without inoculation, because of a rapid drop in pH (Pedroso et al., 2010; Duniere et al., 2011). A

microbial inoculant containing Pediococcus pentosaceus and Propionibacterium jensenii also

hastened (7 vs. 15 d) the elimination of EC in barley silages because of a relatively slow pH

decline (Bach et al., 2002).

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Although antibacterial activity against EC was evident in the low pH silage extracts, it

was not observed in the pH-corrected extracts. This supports the notion that the elimination of

EC during ensiling is most probably because of inhibitory effect of the low pH. Nevertheless, the

pKa (acid dissociation constant) may also be implicated because at low pH, organic acids are

mostly in protonated states, and the protons can diffuse into bacterial cells, and cause cell death

(Bjornsdottir et al., 2006). Previous studies have shown that antibacterial activity was not

detected in extracts of inoculant-treated alfalfa and corn silages adjusted to pH 5.5 and 5.0,

respectively, whereas the pH-corrected cell-free supernatants from pure cultures of inoculant

bacteria showed activity against EC (Pedroso et al., 2010). Gollop et al. (2005) reported that pure

cultures of L. plantarum, Enterococcus faecium, and L. buchneri inhibited the growth

of Micrococus luteus and Pseudomonas aeruginosa, but very few of the extracts of silages

treated with these bacteria showed activity independent of pH.

More time (7 d) was required for elimination of EC in ECA silages than in others

probably because of the lower initial pH of the ECA silage before ensiling. Goodson and Robury

(1989) reported that E. coli strains that were previously habituated in a medium with a pH of 5.0

survived longer after short periods of exposure to pH 3.5, whereas those that were previously

habituated in a medium with a pH of 7 failed to grow. Likewise, strains of EC habituated in a

sub-lethal medium with a pH of 5.0 had greater resistance than those habituated in a medium

with higher pH values when they were subsequently exposed to a medium with an acidic pH of

3.85 (Gregory et al. 1995). Similarly, in this study, EC persisted longer in the ECA silage than in

other silages and the respective pH values were 5.14 and 5.95 ± 0.03 at ensiling (Table 5-1).

Addition of L. buchneri to silage enhances anaerobic degradation of lactate to acetate

(Oude Elferink et al., 2001; Kleinschmit and Kung, 2006a; Queiroz et al., 2013). In accordance,

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in this study, silage inoculation with L. buchneri increased the concentration of acetate. The

antimycotic properties of acetate (Danner et al., 2003; Schmidt and Kung, 2010) and propionate

(Woolford, 1975) resulted in the lower yeast counts of ECLB and ECA silages, respectively

compared to those in the other silages. Lactobacillus buchneri-inoculated silage had higher

ammonia-N concentration compared with ECLP silage. This agrees with previous studies on

corn and sorghum silages (Driehuis et al., 2001; Filya, 2003). A high concentration of ammonia-

N (>10 % of total N) in corn silage is a marker of excessive protein breakdown (Ward, 2008),

typically caused by a slow drop in pH and or clostridial fermentation (Kung and Shaver, 2001).

However, the concentrations observed in these silages are within the range for well-preserved

corn silage (Ward, 2008). Also, the fact that no butyrate was detected supports the deduction that

spoilage because of clostridial fermentation was not evident.

Aerobic Post-Ensiling Phase

This aspect of the experiment was conducted to determine whether treatment with the

silage additives at ensiling would enhance the aerobic stability of the silages and inhibit the

growth of EC in silages contaminated at silo opening (d 120) and after 168 h of aerobic exposure

(d 127). The elimination of EC within 6 h of re-inoculation and aerobic exposure in all silages

on d 120 is probably because the low pH value (3.93 ± 0.04) prevented their survival. Bach et al.

(2002) reported that EC was not detected in barley silage 24 h after re-inoculation of the

pathogen at silo opening because the pH value of the silage was below 4.0. Likewise, EC was not

detected in corn silage after 24 h of re-inoculation at silo opening (Pedroso et al., 2010) because

the pH values of the silages were still below 4.0.

The inhibitory effects of acetate and propionate on spoilage-causing fungi (Ranjit et al.,

2002; Kleinschmit et al., 2005) in silages treated with L. buchneri and propionic acid, are

responsible for the greater aerobic stability of such silages. Aerobic spoilage occurs because of

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degradation of lactic acid by lactate-utilizing yeasts in the presence of air (Pahlow et al., 2003;

Kung, 2010). A meta-analysis showed that L. buchneri inoculation decreased yeast counts and

improved aerobic stability of corn silages (Kleinschmit and Kung, 2006a). Similarly, in three

separate experiments, addition of propionic acid-based products to corn silages at 0.2% of fresh

weight improved the aerobic stability of the silages (Kung et al., 1998).

Whereas no or low counts of EC were detected in ECA and ECLB silages reinoculated

with the pathogen after 168 h of aerobic exposure, high counts were detected in ECCH and

ECLP silages probably because of their higher pH values (pH > 5) compared to the other silages.

Previous studies have shown that EC grows well in corn silage with pH above 4.0 (Fenlon and

Wilson, 2000; Pedroso et al., 2010; Duniere et al., 2011), whereas its growth is curtailed by

additives that keep the silage pH below 4.0 after silo opening (Pedroso et al., 2010; Duniere et

al., 2011). Likewise, our results showed that keeping the silage pH below 4 after aerobic

exposure improved the silage shelf-life and also helped to curtail or prevent the growth and

spread of EC. This indicates that keeping the corn silage pH below 4.0 during the feedout stage

with appropriate silage additives may help reduce the risk of survival and/or prevalence of EC or

other low pH-sensitive pathogens in aerobically exposed silages. Nevertheless, aerobically

stable silage may permit survival of EC that contaminates the silage during the aerobic exposure

phase, as shown in this study and our previous study on alfalfa silage (Chapter 3).

Bacterial Community Composition and Diversity

Principal coordinate analysis of diversity based on UniFrac (Unweighted) distances

indicates compositional differences based on the presence or absence of OTUs among samples

(Park et al., 2016). The fact the bacterial diversity and taxonomic composition of the ECCH

silage did not differ from that of the Control silage suggests that the microbiome shift observed

in the ECLP, ECLB and ECA silages reflect the effects of the respective additives alone.

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Notable features of such microbiomes were greater abundance of Lactobacillus in ECLP and

ECA silages, and greater abundances of Acinetobacter and Weissella in ECLB silage compared

to the Control silage. Studies that utilized denaturing gradient gel electrophoresis (DGGE)

technique reported little or no change in the bacterial community structure of corn silage treated

with L. buchneri, L. plantarum or L. rhamnosus (Li and Nishino, 2011; Santos et al., 2015).

The taxonomic classification of lactobacilli to the species level with 16S gene sequences

may sometimes misrepresent the microbial population because of the relatively little sequence

diversity among lactobacilli (Berthier and Elrhich, 1998; Felis and Dellaglio, 2007). This may

partly explain the unexpected low abundance of L. plantarum in ECLP silage, absence of L.

buchneri in ECLB silage, and high abundance of L. diolivorans in all silages. Pang et al. (2011)

identified Weissella cibaria and W. confusa as the dominant bacterial species in corn silage, L.

plantarum as the dominant species in sorghum and rice silages, and L. pseudomesenteroides, L.

paraplantarum, and L. plantarum as the dominant species in alfalfa silage. A recent study that

applied 16S ribosomal RNA gene amplicon sequencing revealed that 99% of the sequence reads

detected were L. reuteri and L. pontis in corn, sorghum and wheat silages (Kraut-Cohen et al.,

2016). Lactobacillus reuteri and L. pontis have previously been detected in silages, but had never

been identified as dominant bacteria (Li and Nishino, 2011; Wu et al. 2014). These dominance

reports are also likely to be affected by the ensiling duration or time of sampling, which is known

to influence the bacterial community of silage (Beck, 1972; Graf et al., 2016).

Lactobacillus diolivorans, which phylogenetically belongs to L. buchneri group, has

previously been isolated from corn silage (Krooneman et al., 2002), but we found no study that

attempted to quantify the abundance of the bacterium in silage. This bacterium is not regarded as

a dominant Lactobacillus species in silage (Krooneman et al., 2002), but it can degrade 1,2-

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propanediol, a fermentation product of L. buchneri, to 1-propanol and propionic acid

(Krooneman et al., 2002). In corn silage treated with L. diolivorans alone or with L. buchneri, no

1-propanol or propionate was detected, and the authors suggested that L. diolivorans converted

1,2-propanediol that was present into an unknown intermediate compound (Charley and Kung,

2005). In contrast, Zhang et al. (2010) reported that a higher concentration of propionate was

formed when L. diolivorans was grown with 1,2-propanediol as the sole carbon source than

when it was grown with 1,2-propanediol, fructose and maltose in sourdough. Both 1,2-

propanediol and 1-propanol were not measured in this study and propionate was only detected in

ECA silage because of treatment with propionic acid at ensiling. Therefore, no conclusive

deductions can be drawn about the presence or role of L. diolivorans in the silages. More

research is needed to confirm the dominance of L. diolivorans in corn silage and to elucidate its

role. Such studies should consider other approaches for characterizing the microbial community.

For instance, previous studies showed that recA gene sequences are more precise than 16S rRNA

for differentiation of LAB species in mixed culture (Felis et al. 2001; Torriani et al. 2001).

Furthermore, recent studies claimed that the Pacific Biosciences (PacBio) single molecule, real-

time sequencing technology (SMRT) can predict the bacterial profile of target samples to species

level because of its power to generate long sequence reads (Zhang et al., 2015; Bao et al., 2016).

The low bacterial diversity observed in this study, as shown by the low shannon index

(0.44 ± 0.09), is as a result of the high abundance (96.5 ± 1.63%) of Lactobacillus species due to

the low pH value of corn silage (< 4.0). The more the abundance of a dominant bacterium, the

less diverse the microbial community (Polley et al., 2007; Allen et al., 2009).

In this study, we detected two genera, Weissella and Acinetobacter, whose roles in corn

silage have not been extensively studied. Bacteria assigned to genus Weissella are strictly

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heterofermentative, producing a mixture of lactate and acetate as the major end products of sugar

metabolism (Fusco et al., 2015, Graf et al., 2016). Inoculation of alfalfa silage with

heterofermentative LAB strain, Weissella paramesenteroides, resulted in increased acetate

concentration (Cai et al., 1998). In addition, lactate and acetate are the major acids produced by

W. confusa during fermentation of rice cake (Baek et al., 2012). Furthermore, Ndagano et al.

(2011) reported the production of acetate as well as other antifungal compounds, such as 3-

hydroxy fatty acids and phenyllactate, by W. paramesenteroides isolated from fermented cassava

(Manihot esculenta). Thus, the greater abundance of Weissella in the ECLB silage may partly

explain its greater acetate concentration and greater aerobic stability relative to other silages

except ECA. Other silages contained equal or greater abundance of Lactobacillus, which also

produce lactate and acetate (Charley and Kung, 2005) and no other bacteria that are known to

produce lactate or acetate responded to the inoculant treatments. Future studies should

characterize the antifungal compounds produced by lactic acid bacteria belonging to genus

Weissella and examine their effects on the fermentation and aerobic stability of corn silage.

Acinetobacter species are aerobic, non-fermenting bacteria, which can be found in

different environments (Kämpfer and Glaeser, 2012) but are supposedly absent in silage (Li and

Nishino, 2011). Some Acinetobacter species can survive in an anaerobic environment in the

presence of acetate as a substrate (Fuhs and Chen, 1975). The uptake of acetate by Acinetobacter

in an anaerobic condition requires energy from carbohydrate degradation (Satoh et al., 1996).

Silage DM loss during ensiling is typically caused by silage decay as a result of depletion of

forage carbohydrate mass by activities of aerobic micro-organisms at the initial stage of ensiling

(Pitt, 1986). Aerobic respiration utilizes readily available carbohydrates and produces heat and

the consequent silage DM and energy losses during ensiling (Muck and Holmes, 2000). The

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increased abundance of Acinetobacter in ECLB silage may have resulted from the increased

acetate concentration and this may partly explain the small, though important, DM losses

sometimes observed in silages that had been treated with L. buchneri at ensiling (Filya, 2003;

Kleinschmit and Kung, 2006a). The practical significance of the seemingly small statistical

differences (0.71 vs. 1.16%) in the abundance of Acinetobacter is unknown. Future studies

should examine if Acinetobacter is responsible for some or all of the DM losses that occur in

silages that were treated with L. buchneri at ensiling. The abundances of unknown or

unculturable bacteria that were not described were either too low (< 0.01%) or did not respond to

silage treatment.

Conclusions

This study shows that when the pH dropped below 4.0 within 3 d of ensiling, EC counts

were below the detection limit of 10 cfu/g in all silages contaminated at ensiling with 1 × 105

cfu/g of the pathogen. The exception was the ECA silage, which required 7 d of ensiling for

elimination of EC, likely because of habituation of EC to acidic conditions when the forage was

treated with propionic acid. Application of L. buchneri and propionic acid reduced yeast counts,

improved aerobic stability, and maintained a low pH for the entire duration of aerobic exposure.

Re-inoculation of silages with EC after 168 h of aerobic exposure resulted in relatively high EC

counts of 5.39 and 5.30 cfu/g in ECCH and ECLP silages, respectively, whereas the pathogen

was 10,000-fold lower in ECLB silage, which had a pH of 4.24 and it was below the detection

limit in the ECA silage, which had a pH 3.96. This suggests that maintenance of pH of

approximately 4.0 or lower with additives can curtail or prevent the growth of EC in

contaminated aerobically exposed silages, and inhibit cycling of EC on farms or limit its

introduction into the food chain. The fact that extracts of d 120 silages exhibited activity against

EC before but not after they were pH-adjusted, confirms that EC elimination from the silages

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was mediated by low silage pH. Bacterial diversity analysis of the silage revealed that the

bacterial community composition in the silages was modified by additive treatment. Our results

revealed that exactly 98.3 ± 0.65% of the bacterial community involved in lactic acid

fermentation in corn silage belong to phylum Firmicutes and 96.5 ± 1.63% belong to genus

Lactobacillus. Furthermore, the potential respective roles in corn silage fermentation and DM

losses of Weissella and Acinetobacter should be determined because their abundances increased

with inoculation with L. buchneri.

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Table 5-1. Characteristics of whole-plant corn forage inoculated with Escherichia coli O157:H7

(EC) alone or EC and bacterial inoculants or propionic acid before ensiling.


Control ECCH ECLP ECLB ECA

pH 5.94a 5.99a 5.93a 5.93a 5.14b 0.04 0.01

DM (%) 34.7 35.7 35.5 35.4 34.8 0.88 0.75

CP (% of DM) 8.75 8.77 8.37 8.26 8.55 0.23 0.16

NDF (% of DM) 41.6 41.4 40.6 42.5 42.4 1.40 0.68

ADF (% of DM) 24.9 24.8 25.5 25.4 25.0 1.41 0.98

abMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; ECCH = 1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen

Institute, Gainseville, FL); ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014 (Lallemand

Animal Nutrition, Milwaukee, WI); ECLB = ECCH + 1 × 106 cfu/g of L. buchneri R1102

(Lallemand Animal Nutrition, Milwaukee, WI); ECA = ECCH + 2.2 g/kg of propionic acid

(Sigma-Aldrich, St. Louis, MO).

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Table 5-2. Fermentation indices and chemical composition and microbial counts (log cfu/g) of

corn silage that was inoculated with Escherichia coli O157:H7 (EC) alone or with

bacterial inoculants or propionic acid after 120 d of ensiling.

a–cMeans within a row with different superscripts differ (P < 0.05). 1Control = distilled water; ECCH = 1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen

Institute, Gainesville, FL); ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014 (Lallemand



(Sigma-Aldrich, St. Louis, MO). 2Lactic acid bacteria



pH 3.93 3.91 3.89 3.91 3.87 0.04 0.16

Lactate (% of DM) 2.99ab 3.10ab 3.50a 2.59b 3.06ab 0.20 0.01

Acetate (% of DM) 2.33b 2.43b 1.85b 3.39a 2.26b 0.30 0.01

Propionate (% of DM) 0.00b 0.00b 0.00b 0.00b 0.41a 0.05 0.01

Yeasts and molds 5.08a 5.30a 4.91a 3.43b 2.92b 0.48 0.01

LAB2 (log cfu/g) 7.06 7.41 7.79 7.89 7.46 0.51 0.52

DM (%) 30.3 30.2 30.6 30.8 31.0 0.44 0.30

NDF (% of DM) 42.5 42.3 43.3 42.5 43.3 1.89 0.97

ADF (% of DM) 28.4 28.0 28.6 28.7 28.6 1.33 0.98

CP (% of DM) 9.58 9.11 9.23 9.02 9.37 0.21 0.11

Ammonia-N (% of DM) 0.10ab 0.10ab 0.08b 0.12a 0.09ab 0.01 0.04

Ammonia-N (% of total N) 6.63ab 7.14ab 5.20b 8.41a 6.23ab 0.89 0.03

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Table 5-3. Bacterial community composition and diversity of whole-plant corn silage inoculated

with Escherichia coli O157:H7 (EC) alone or with bacterial inoculants or propionic

acid.

a–dMeans within a row with different superscripts differ (P < 0.05).

1Control = distilled water; ECCH = 1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen

Institute, Gainesville, FL); ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014 (Lallemand



(Sigma-Aldrich, St. Louis, MO). Values are averages of 4 replicates of each treatment.



Species diversity and richness

Shannon index 0.53a 0.41ab 0.30b 0.52a 0.42ab 0.06 0.02

Chao index 33.5 26.4 22.5 28.7 32.9 4.68 0.16

Phylum (%)

Firmicutes 97.8bc 98.4ab 99.0a 97.4c 98.7a 0.29 0.01

Proteobacteria 2.07ab 1.56bc 0.98c 2.63a 1.18c 0.28 0.01

Genus (%)

Lactobaccillus 94.9cd 96.5bc 98.3a 94.8d 97.9ab 0.54 0.01

Sphingobacterium 0.05 0.03 0.00 0.09 0.02 0.02 0.07

Weissella 1.57b 1.42b 0.72c 2.39a 0.79c 0.20 0.01

Acinetobacter 0.71b 0.63b 0.44b 1.16a 0.51b 0.12 0.01

Stenotrophomonas 0.22 0.20 0.17 0.36 0.15 0.07 0.06

Serratia 0.69 0.45 0.34 0.47 0.48 0.15 0.06

Lactobaccillus species (%)

L. diolivorans 94.3bc 95.3abc 96.8a 93.75c 95.5ab 0.55 0.01

L. plantarum 0.52dc 0.71bc 0.98ab 0.35d 1.08a 0.11 0.01

Uncultured 0.70 0.40 0.47 0.46 0.78 0.15 0.09

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Figure 5-1. Changes in the pH of whole-plant corn forage inoculated with 1 × 105 cfu/g of


and ensiled for different durations. Control = distilled water; ECCH = 1 × 105 cfu/g of

E. coli O157:H7 (Emerging Pathogen Institute, Gainesville, FL); ECLP = ECCH + 1

× 106 cfu/g of L. plantarum R2014 (Lallemand Animal Nutrition, Milwaukee, WI);

ECLB = ECCH + 1 × 106 cfu/g of L. buchneri R1102 (Lallemand Animal Nutrition,

Milwaukee, WI); ECA = ECCH + 2.2 g/kg of propionic acid (Sigma-Aldrich, St.

Louis, MO). Treatment × day SE and P-value = 0.02 and 0.001, respectively; an

asterisk (*) indicates that values differed at this ensiling duration (P < 0.05). Error

bars represent SE.

3.5

4

4.5

5

5.5

6

0 3 7 120

pH

Day of ensiling

Control

ECCH

ECLP

ECLB

ECA

*

/ /

140

Figure 5-2. Changes in Escherichia coli O157:H7 (EC) counts of whole-plant corn forage

inoculated with 1 × 105 cfu/g of Escherichia coli O157:H7 (EC) alone or with

bacterial inoculants or propionic acid and ensiled for different durations. Control =

distilled water; ECCH = 1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen

Institute, Gainesville, FL); ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014

(Lallemand Animal Nutrition, Milwaukee, WI); ECLB = ECCH + 1 × 106 cfu/g of L.

buchneri R1102 (Lallemand Animal Nutrition, Milwaukee, WI); ECA = ECCH + 2.2

g/kg of propionic acid (Sigma-Aldrich, St. Louis, MO). Treatment × day SE and P-

value = 0.02 and 0.001, respectively; an asterisk (*) indicates that values differed at

this ensiling duration (P < 0.05). Error bars represent SE.

0

1

2

3

4

5

6

0 3 7 120

EC

, lo

g 1

0cf

u/g

Day of ensiling

ECCH

ECLP

ECLB

ECA

*

/ /

141

Figure 5-3. Changes in lactate concentration of whole-plant corn forage inoculated with 1 × 105


propionic acid and ensiled for different durations. Control = distilled water; ECCH =

1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen Institute, Gainesville, FL);

ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014 (Lallemand Animal Nutrition,

Milwaukee, WI); ECLB = ECCH + 1 × 106 cfu/g of L. buchneri R1102 (Lallemand

Animal Nutrition, Milwaukee, WI); ECA = ECCH + 2.2 g/kg of propionic acid

(Sigma-Aldrich, St. Louis, MO). Treatment × day SE and P-value = 0.05 and 0.001,

respectively; an asterisk (*) indicates that counts differed at this ensiling duration (P

< 0.05). Error bars represent SE.

0

1

2

3

4

0 3 7 120

Lac

tate

, %

DM

Day of ensiling

Control

ECCH

ECLP

ECLB

ECA

*

*

/ /

142

Figure 5-4. Changes in acetate concentration of whole-plant corn forage inoculated with 1 × 105


propionic acid and ensiled for different durations. Control = distilled water; ECCH =

1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen Institute, Gainesville, FL);

ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014 (Lallemand Animal Nutrition,

Milwaukee, WI); ECLB = ECCH + 1 × 106 cfu/g of L. buchneri R1102 (Lallemand

Animal Nutrition, Milwaukee, WI); ECA = ECCH + 2.2 g/kg of propionic acid

(Sigma-Aldrich, St. Louis, MO). Treatment × day SE and P-value = 0.05 and 0.001,

respectively; an asterisk (*) indicates that counts differed at this ensiling duration (P

< 0.05). Error bars represent SE.

0

1

2

3

4

0 3 7 120

Ace

tate

, %

DM

Day of ensiling

Control

ECCH

ECLP

ECLB

ECA

*

/ /

143

Figure 5-5. Escherichia coli O157:H7 (EC) counts (log10 cfu/g) and pH of whole-plant corn

silage (d 128) re-inoculated with 5 × 105 cfu/g of EC 168 h after silo opening. Control

= distilled water; ECCH = 1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen

Institute, Gainesville, FL); ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014

(Lallemand Animal Nutrition, Milwaukee, WI); ECLB = ECCH + 1 × 106 cfu/g of L.

buchneri R1102 (Lallemand Animal Nutrition, Milwaukee, WI); ECA = ECCH + 2.2

g/kg of propionic acid (Sigma-Aldrich, St. Louis, MO). Treatment × day SE and P-

value = 0.05 and 0.001, respectively; an asterisk (*) indicates that counts differed at

this ensiling duration (P < 0.05). Error bars represent SE.

0

1

2

3

4

5

6

7


Treatment

pH EC

1.12b

4.24C

5.30a

6.13A

5.39a5.67B

0.00b

5.43B

3.96C

0.00b

a

b

144

Figure 5-6. The aerobic stability (h) of whole-plant corn forage inoculated with 1 × 105 cfu/g of


and ensiled for 120 d. Control = distilled water; ECCH = 1 × 105 cfu/g of E. coli

O157:H7 (Emerging Pathogen Institute, Gainesville, FL); ECLP = ECCH + 1 × 106

cfu/g of L. plantarum R2014 (Lallemand Animal Nutrition, Milwaukee, WI); ECLB

= ECCH + 1 × 106 cfu/g of L. buchneri R1102 (Lallemand Animal Nutrition,

Milwaukee, WI); ECA = ECCH + 2.2 g/kg of propionic acid (Sigma-Aldrich, St.

Louis, MO). Treatment × day SE and P-value = 0.05 and 0.001, respectively; an

asterisk (*) indicates that counts differed at this ensiling duration (P < 0.05). Error

bars represent SE.

40

60

80

100

120

140

160

180


Aer

obic

sta

bil

ity,

h

Treatment

168a 168a

76.0c

89.6bc92.8b

145

Figure 5-7. Unweighted UniFrac principal coordinate analysis (PCoA) plot of individual samples

in each treatment. Individual samples were represented with blue (Control), red

(ECCH), orange (ECLP), purple (ECLB), and yellow (ECA). Control = distilled

water; ECCH = 1 × 105 cfu/g of E. coli O157:H7 (Emerging Pathogen Institute,

Gainesville, FL); ECLP = ECCH + 1 × 106 cfu/g of L. plantarum R2014 (Lallemand

Animal Nutrition, Milwaukee, WI); ECLB = ECCH + 1 × 106 cfu/g of L. buchneri

R1102 (Lallemand Animal Nutrition, Milwaukee, WI); ECA = ECCH + 2.2 g/kg of

propionic acid (Sigma-Aldrich, St. Louis, MO). The PCoA plot indicates the

phylogenetic distance (variation) between samples using two principal coordinates

(PC1 and PC2). The percentage variation explained by each PC is indicated on each

axis.

PC

2 (

6.6

%)

PC1 (25.5%)


146

CHAPTER 6

EFFECTS OF THREE SEQUESTERING AGENTS ON MILK AFLATOXIN M1

CONCENTRATION AND THE PERFORMANCE AND IMMUNE STATUS OF DAIRY

COWS FED DIETS ARTIFICIALLY CONTAMINATED WITH AFLATOXIN B1

Background

Mycotoxins are a group of toxic compounds produced by fungi on growing forages or

stored feeds that can reduce the feed intake, growth, health, and reproductive performance of

livestock and humans. Aflatoxin B1 has been the most widely studied and problematic mycotoxin

in dairy cows because its metabolite, AFM1, can be transferred to milk (Diaz et al., 2004). Up to

166,000 cases of hepatocellular carcinoma per year in humans have been attributed to aflatoxins

(Liu and Wu, 2010). The United States FDA regulates the maximum permissible amount of

AFM1 in milk at 0.5 μg/kg and AFB1 in dairy feeds at 20 μg/kg because of the appreciable

consumption of milk by humans, particularly infants (FDA, 2000). The maximum milk AFM1

concentration allowed by the European Commission is 0.05 μg/kg (EFSA, 2004).

Various approaches have been employed to reduce the risk of aflatoxin ingestion by

livestock and the resultant aflatoxicoses. Ozonization and ammoniation can minimize the

production and effects of the toxins in corn and cottonseed meal (CAST, 2003), but these

approaches are costly and time-consuming (Kutz et al., 2009), so they are not typically used on

dairy farms. One of the more recent approaches is to use adsorbents that bind aflatoxin in the

gastrointestinal tract and reduce its bioavailability (Huwig et al., 2001, Phillips et al., 2002).

Clay-based aflatoxin-sequestering agents have shown the potential to bind aflatoxins and impair

their gastrointestinal absorption (Kabak et al., 2006), but high doses may reduce utilization of

vitamins and minerals (Chestnut et al., 1992). The outcomes of studies testing the effectiveness

of adsorbents at reducing the transfer of the toxins to milk have been inconsistent (Kabak et al.,

2006). Diaz et al. (2004) reported that 3 sodium bentonite products reduced the concentration of

147

AFM1 residues in milk by 50 to 65%. Another study (Stroud, 2006) compared 8 binding agents,

including 3 combinations of yeast and clay products and 5 clay products, and revealed that 1 of

the former and 3 of the latter reduced the transfer, secretion, or concentration of AFM1 in milk. A

recent study by Xiong et al. (2015) showed that a product based on yeast and sodium

montmorillonite reduced the transfer of aflatoxin to milk at a low dietary dose (20 μg/kg) but had

no effect at a high dose (40 μg/kg). We have shown that a high dose (1% of diet DM), but not a

low dose (0.2% of diet DM), of montmorillonite clay reduced the transmission of dietary AFM1

to the milk of dairy cows (Queiroz et al., 2012). There is clearly a need to continue to evaluate

the effects of aflatoxin-sequestering agents on milk AFM1 concentration, as well as on milk yield

and quality. The objective of this study was to examine whether dietary addition of 3

Saccharomyces cerevisiae fermentation product–based mycotoxin-sequestering agents would

reduce milk AFM1 concentration and improve the performance and health of dairy cattle that

were fed diets artificially contaminated with AFB1.


Cows, Treatments, and Design

Cows in this study were cared for according to the University of Florida Animal Research

Committee protocols, which were developed from guidelines outlined in the Guide for the Care

and Use of Agricultural Animals in Research and Teaching (FASS, 2010). Fifteen lactating

multiparous Holstein cows in mid lactation (150 to 200 DIM) were stratified by parity, DIM, and

milk production, and were randomly assigned to 1 of 5 treatments arranged in an incomplete

crossover design with four 28-d periods. At the end of each period, cows were switched to a

different treatment in a way that ensured no cow was on the same treatment in more than 1

period. Cows were housed in a freestall open-sided barn bedded with sand and equipped with

Calan gates (American Calan Inc., Northwood, NH) for individual feeding and with misters and

148

fans to minimize heat stress. The following dietary treatments were investigated: (1) a control

diet with no sequestering agent or mycotoxin added (C); (2) diet C fed with 1,725 μg of AFB1

(T); (3) diet T fed with 20 g/d of sequestering agent 1 (SEQ1); (4) diet T fed with 20 g/d of

sequestering agent 2 (SEQ2); (5) diet T fed with 20 g/d of sequestering agent 3 (SEQ3). The

sequestering agents were supplied by Diamond V Inc. (Cedar Rapids, IA). They were proprietary

formulations of an S. cerevisiae fermentation product containing a low (SEQ1) or high (SEQ2)

dose of a proprietary chlorophyll-based additive or a low dose of the additive combined with

sodium bentonite clay (SEQ3).

Diets were formulated to meet or exceed the nutrient requirements of cows producing at

least 30 kg of milk (NRC, 2001) using CPM-Dairy software version 3.0.10 (www.cpmdairy.net).

The ingredients and chemical composition of the basal diet are shown in Table 6-1. Cows were

fed 1,725 μg/head per day of AFB1 based on an average projected DMI of 23 kg/d, to supply a

daily dose of 75 μg/kg of AFB1 (in treatments T, SEQ1, SEQ2, SEQ3). Dietary AFB1 was

obtained from an Aspergillus parasiticus (NRRL-2999) culture at the University of Missouri

Diagnostic Laboratory (Columbia, MO). The AFB1 was mixed with ground corn and molasses,

weighed into gelatin capsules and dosed orally to cows once daily before the TMR was fed in the

morning; the sequestering agents (SEQ1, SEQ2, SEQ3) were top-dressed on the relevant TMR

from d 1 to 28 of each period. The first 20 d of each period were used to ensure adaptation to a

new diet and clearance of AFM1 residues from the milk from the preceding period. Mycotoxin-

contaminated corn was fed only during d 21 to 25 of each period in the relevant treatments, and

clearance of the toxin from the milk was monitored during d 26 to 28. Queiroz et al. (2012)

confirmed the suitability of the 5-d toxin-dosing and 3-d toxin-clearance periods for modeling

the effects of sequestering agents on ingested AFB1.

149

Experimental Measurements and Analytical Procedures

Cows were milked twice daily. Milk yield was recorded daily by the Afikim milking

system (AfiLab mini laboratory, Afimilk Ltd., Israel); milk samples were collected from both

1000 and 2200 h milkings on d 20 to 28 in each period and analyzed separately for milk

components. Concentrations of milk fat, milk protein, and SCC were analyzed by the Southeast

Milk dairy laboratory (Belleview, FL) using a Bentley 2000 near-infrared reflectance

spectrophotometer (Bentley Instruments Inc., Chaska, MN). Values for 3.5% FCM yield were

calculated based on the following equation: [(0.4324 × milk yield) + (16.218 × milk fat yield)]

(NRC, 2001). Milk AFM1 concentrations were measured on composited am and pm milk

samples on d 20 to 28 in each period and were quantified by the Southeast Milk dairy laboratory

using the radioimmunoassay test (CHARM II, Charm Sciences Inc., Malden, MA) described by

Diaz et al. (2004). Aflatoxin M1 excretion was calculated as the product of milk AFM1

concentration and milk yield. Transfer of the toxin to milk during the dosing period was

calculated as daily AFM1 excretion/ daily AFB1 intake. Transfer of the toxin during the

withdrawal period was calculated as daily AFM1 excretion/AFB1 intake on d 25. Aflatoxin M1

clearance rates were calculated by dividing the difference between the AFM1 concentration on d

26, 27, and 28 and that on d 25 by 24, 48, and 72 h, respectively.

The amount of feed offered at each feeding was recorded for each cow. Refused feed was

collected and weighed each morning before feeding and used to calculate intakes for the next

day. Diets were fed ad libitum (110% of the previous day’s DM intake). In each period, dietary

ingredients (corn silage, ryegrass triticale silage mixture, alfalfa orchardgrass hay mixture, and

concentrate mix) and orts were sampled (100 g) daily into labeled bags from d 20 to 28 and

stored in a refrigerator (4°C) for further analysis. Dietary samples were dried at 55°C for 48 h in

a forced-air oven, ground to pass the 1 mm screen of a Wiley mill (A. H. Thomas, Philadelphia,

150

PA), and composited across days. Feed samples were sent to the Dairy One forage testing

laboratory (Ithaca, NY) for DM, N, ash, NDF, ADF, and mineral analyses. Nitrogen was

determined by rapid combustion using a Macro Elemental N analyzer (AOAC International,

2000; Vario MAX CN, model ID 25.00–5003; Elementar, Hanau, Germany) and CP was

calculated by multiplying the N value by 6.25. Concentrations of NDF and ADF were measured

using the non-sequential method of Van Soest et al. (1991) in an Ankom A200 Fiber Analyzer

(Ankom Technologies, Macedon, NY). Heat-stable α-amylase was used in the NDF procedure,

and the results are expressed exclusive of residual ash. Minerals (Ca, P, Mg, K, Na, Fe, Zn, Cu,

Mn) were analyzed using an inductively coupled plasma radical spectrometer (ICAP, Thermo

Scientific Inc., Marietta, OH) after microwave digestion. The concentration of aflatoxins,

deoxynivalenol, zearalenone, and T-2 toxin in the feed ingredients was analyzed by HPLC at the

University of Missouri Veterinary Diagnostic Laboratory using the HPLC method described by

Kutz et al. (2009). On d 20 and 25 of each period, blood was collected before the morning

feeding from the coccygeal vessels into Vacutainer tubes (Becton Dickinson, Franklin Lakes,

NJ) containing sodium heparin anticoagulant, centrifuged at 2,500 × g for 20 min at 4°C and

stored at −20°C for further analysis of acute-phase proteins. Plasma haptoglobin concentrations

were determined by measuring haptoglobin/hemoglobin complexing based on differences in

peroxidase activity (Makimura and Suzuki, 1982). Plasma ceruloplasmin oxidase activity was

measured using the colorimetric procedure described by Demetriou et al. (1974). Acid-soluble

protein concentration was determined using the bicinchoninic acid kit for protein determination

(Sigma-Aldrich, St. Louis, MO) following plasma acid-soluble protein extraction (Nakajima et

al., 1982). Whole blood was analyzed for total and differential cell counts using an automated

ProCyte Dx hematology analyzer (Idexx Laboratories Inc., Westbrook, ME). The percentage of

151

neutrophils exhibiting phagocytosis and oxidative burst activities after Escherichia coli challenge

was measured according to procedures described by Martinez et al. (2012). Briefly, neutrophils

were loaded with dihydrorhodamine 123 (Sigma-Aldrich Co., St. Louis, MO) and then

cocultured with propidium iodide-labeled E. coli isolated from the milk of a cow with mastitis at

a bacteria:neutrophil ratio of 40:1. The samples were run within 2 h of preparation using a

Becton Dickinson Accuri C6 digital analyzer flow cytometer (Becton Dickinson). Neutrophils

were identified according to their size and granularity using forward and side scatter detectors,

and data were analyzed using Flowjo software (version 7.6.5., Tree Star Inc., Ashland, OR). The

parameters quantified were mean fluorescence intensity (MFI) for propidium iodide staining as a

proxy for the number of bacteria phagocytized per neutrophil, and MFI for dihydrorhodamine

123 staining as an indicator of the amount of reactive oxygen species generated per neutrophil.

Adhesion molecules that regulate the migration of leukocytes between blood vessel and

lymphoid organs, L-selectin (CD62L) and β2-integrin (CD18), were analyzed as described by

Silvestre et al. (2011). Interferon-γ, the proinflammatory cytokine produced by various activated

leukocyte populations, was measured as described by Garcia Orellana (2012) using the bovine

IFN-γ DuoSet ELISA Development Kit (R&D Systems, Inc., Minneapolis, MN).


The experiment had an incomplete crossover design with 12 experimental units per

treatment. Data were analyzed using the GLIMMIX procedure of SAS Institute Inc. (2012). The

model for analyzing outcomes measured repeatedly over time on the same experimental unit

(milk production and components; DMI; and aflatoxin concentration, transfer, and excretion)

included the effects of treatment, period, and day; interactions of treatment and day and day and

period; and the random effect of cow. Time was used in the repeated measures statement with

autoregressive order 1 [AR(1)] as the time-series covariance structure. We used pre-experimental

152

milk yields as covariates for milk yield data obtained during the predosing (d 16 to 20), toxin-

dosing (d 21 to 25), and withdrawal (d 26 to 28) periods. The model for analyzing measures that

were not collected serially on the same experimental unit (blood cell counts and immune

response data) included the effects of treatment and period and the random effect of cow. Blood

sample data from d 20 were used as covariates for analysis of data from samples collected on the

last day of toxin-dosing (d 25). We estimated denominator degrees of freedom using the

Kenward-Roger option in the MODEL statement for all data, and we corrected for potential

carryover effects of treatments between periods by including carryover coefficients in the model

(Littell et al., 2006; Jones and Kenward, 2015). We examined data for normality by examining

the distribution of residuals and transforming them as needed before statistical analysis. Post hoc

mean comparisons were performed using the Tukey-Kramer procedure for pairwise multiple

comparisons. Significance was declared at P ≤ 0.05, and trends toward significance were

declared at 0.05 < P ≤ 0.10.

Results and Discussion

Milk AFM1 Concentration

The amount of AFB1 dosed (75 µg/kg) exceeded the FDA action level of 20 µg/kg for

dairy cattle feeds and it was transferred into milk in sufficient quantities to raise the AFM1

concentration beyond the FDA action level of 0.5 µg/kg for milk. However, the 75 µg/kg dosed

represents a moderate natural level of AFB1 contamination in corn samples for livestock feeds,

which can be up to 158 µg/kg (Pietri et al., 2004), even though in other studies (Pietri et al.,

2007), most of the samples had concentrations less than 20 µg/kg. Doses used in mycotoxin

challenge studies usually range from 20 to 112 µg/kg (Diaz et al., 2004; Firmin et al., 2011;

Xiong et al., 2015).

153

The sequestering agent mixtures used in this study were tested to explore if combining

the aflatoxin-binding and immunomodulatory effects of Saccharomyces cerevisiae, and the

aflatoxin-binding effects of clay and chlorophyll-based products would result in synergistic

improvements in binding of aflatoxin, stimulation of the immune response, and enhancement of

the performance of dairy cows. The protective effect of Saccharomyces cerevisiae against

mycotoxins is due to adhesion of the toxin to the cell wall components (mannan and β-glucan) of

the yeast (Pfliegler et al., 2015). In addition, yeast-based products act as immune modulators

because β-glucan can stimulate non-specific and specific immunological responses, and thus

improve the performance of animals (Keller et al., 2015). When fed at levels above 1% of the

diet DM to dairy cows (Diaz et al., 2004; Queiroz et al., 2012), clays bind aflatoxins by

chemisorption via cation exchange (Phillips et al., 1995). Chlorophyll-based compounds

sequester aflatoxin and stimulate the immune response by trapping the toxin via the planar

ring structure of the chlorophyll (Arimoto et al., 1993) and stimulating increased activity of

gluthathione S-transferase (GST) enzyme, which prevents aflatoxin epoxide from forming an

adduct with DNA, thereby, facilitating the clearance of the toxin from the body (Hsu et al.,

2008). None of the treatments reduced the milk aflatoxin concentration, suggesting that the

ingredient amounts in the sequestering agent mixtures were inadequate for considerable toxin

binding. Nevertheless, amounts were sufficient to modulate the immune response. SEQ1 had

the greatest proportion of the S. cerevisiae fermentation product; its greater beneficial effects on

indices of the immune response highlights the immunomodulatory effects of the S. cerevisiaie

fermentation product. The presence of clay in SEQ3 and not in SEQ2 probably reduced

adverse effects of the toxin on the cows, such that they were able to mount a more pronounced

immune response. That SEQ2, which had the greatest proportion of the chlorophyll-based

154

additive, had the least beneficial effect on the immune response, suggests that the additive

may have been less effective at binding the aflatoxin or modulating the immune response than

the S. cerevisiae fermentation product or the clay.

We detected no AFM1 in the milk of the cows before they were dosed with AFB1 (data

not shown). During the toxin-dosing period, all cows except for those fed diet C, received 1725

µg of AFB1 daily and their DMI averaged 23.6 kg/d. Therefore, the average daily ingestion of

AFB1 was 73 µg/kg of DMI. No AFM1 was detected in the milk of cows fed diet C, whereas

cows fed diets T, SEQ1, SEQ2 and SEQ3 had mean milk AFM1 concentrations of 0.73, 0.74,

0.73 and 0.74 µg/kg, respectively (Table 6-2). Adding sequestering agents to diets containing the

toxin did not affect milk AFM1 concentration, excretion or transfer during the Toxin-dosing

period. Therefore, none of the sequestering agent treatments prevented milk AFM1

concentrations from rising beyond the FDA action level (0.50 µg/kg). This is consistent with

other studies on the AFB1-sequestering ability of yeast cell wall products that contain high levels

of AFB1 contamination. Milk AFM1 concentration was unaffected by adding 10 g/day of an

undisclosed yeast cell wall product or 125 g/d of a yeast cell wall-based product to dairy cow

diets containing 100 (Waltman et al., 2008) or 112 (Kutz et al., 2009) µg/kg of AFB1,

respectively. Likewise, Kissell et al. (2013) showed that yeast oligosaccharide-based additives

did not affect the milk AFM1 concentration of cows fed diets containing 80 µg/kg AFB1. In

addition, Firmin et al. (2011) reported that adding an undisclosed yeast cell wall product at 2

g/kg of the diet did not affect the concentration of AFM1 in the milk of dairy ewes fed a diet

containing 60 µg of AFB1/kg. However, yeast cell wall sequestering agents have effectively

sequestered AFB1 when the toxin concentration in the diet was lower (Diaz et al., 2004; Xiong et

al., 2015). Diaz et al. (2004) reported that adding a yeast cell wall-based product at 10 g/d to a

155

dairy cow diet contaminated with 55 µg/kg of AFB1 reduced the milk AFM1 concentration.

Certain combinations of yeast and clay-based products have also been less effective at high

contamination levels. Xiong et al. (2015) reported that adding a blend of yeast extract and yeast

culture, hydrated calcium sodium aluminosilicates, and other metabolites at 42.5 g/kg of DMI

reduced milk AFM1 concentration, secretion and transfer in diets contaminated with 20 µg/kg of

AFB1 but had no effect in diets contaminated with 40 µg/kg of AFB1. In general, the efficacy of

clay or yeast cell wall products alone or combinations of both depend on the dose of the

sequestering agent and the level of mycotoxin contamination in the diet.

Once AFB1 was withdrawn from the diet, milk AFM1 concentrations in cows fed the

sequestering agents decreased rapidly, such that they fell below the FDA action level within 24 h

(Table 6-2), whereas it took 48 h for milk AFM1 concentrations to decrease below the FDA

action level in cows fed diet T. In particular, milk AFM1 concentrations in cows fed SEQ1 and

SEQ3 were lower (P < 0.05) than those of cows fed T 24 h after withdrawing the toxin (Figure 6-

1). This was because the SEQ1 and SEQ3 diets tended to reduce (P ≤ 0.10) the transfer and

excretion of the toxin from d 25 into the milk on d 26. The SEQ2 diet tended to reduce (P =

0.06) the milk AFM1 concentration compared to that for cows fed diet T but had no effect on the

transfer and excretion of the toxin into the milk. Our previous study (Queiroz et al., 2012)

showed that during the Toxin-dosing period, a diet containing a montmorillonite clay adsorbent

(1% of the diet DM) reduced the milk AFM1 concentration below the FDA action level when

cows were fed a diet contaminated with 75 µg/kg DMI of AFB1. However, the adsorbent had no

effect when it was fed at 0.2% of the diet DM. The dose of clay fed in this current study (0.04%

of DMI) was lower than in our previous study, but when combined with the S. cerevisiae

156

fermentation product, it reduced the time required for a reduction in the milk AFM1

concentration to a safe level by 24 h after dietary withdrawal of the toxin.

By 48 h after withdrawing AFB1 from the diet, only trace concentrations of AFM1 were

detected in the milk in all treatments and no treatment effects (P > 0.10) on the concentration,

transfer, or excretion of the toxin were evident. This is consistent with other studies, which

reported that AFM1 concentrations in milk were reduced drastically within 2 to 3 d of

withdrawing AFB1 from the diet (Kutz et al., 2009; Queiroz et al., 2012; Xiong et al., 2015).

Clearance rates of milk AFM1 did not differ among cows fed the toxin during any of the days of

the Withdrawal period.

Animal Performance

During the predosing period, milk yield and DMI did not differ across treatments (26.7 ±

1.20, P = 0.64 and 22.6 ± 0.81, P = 0.30 kg/d, respectively; Table 6-3). Therefore, feeding the

sequestering agents during the adaptation or predosing periods did not affect the performance of

the cows. During the toxin-dosing period, DMI was not affected by dietary treatment (23.7 ±

0.85 kg/d, P = 0.16). Also, dietary treatments did not affect the change in milk yield from the

predosing to the Toxin-dosing periods. However, feeding diet T instead of C during the toxin-

dosing period tended to reduce milk yield (26.0 vs. 28.5 kg/d, P = 0.08) and 3.5% FCM yield

(27.1 vs. 29.8 kg/d, P = 0.09). This is consistent with the study of Queiroz et al. (2012) who

reported that feeding AFB1 at the rate used in this study tended to reduce the 3.5% FCM yield of

cows. A similar numerical trend was reported when 112 µg/kg of AFB1 was fed to dairy cows by

Kutz et al. (2009). However, studies have also reported no changes in milk yield due to feeding

117 µg/kg of DMI (Maki et al., 2016) or 80 µg of AFB1/kg of DMI (Kissell et al., 2013).

Feeding the SEQ3 diet instead of C reduced milk yield (24.8 vs. 28.5 kg/d, P = 0.01) and

3.5% FCM yield (25.9 vs. 29.8 kg/d, P = 0.01). Also, feeding the SEQ2 diet reduced 3.5% FCM

157

yield relative to feeding diet C (26.9 vs. 29.8 kg/d, P = 0.04). The reasons for these reductions

are not clear. In contrast, milk and FCM yields of cows fed the SEQ1 and C diets did not differ

(P > 0.10). Therefore, unlike the other sequestering agents, feeding the SEQ1 diet probably

prevented the adverse effect of the toxin on milk production by the cows. This may be due to the

numerically higher DMI for cows fed the SEQ1 diet relative to those fed SEQ2 and SEQ3. Feed

efficiency was unaffected (P > 0.10) by dietary treatments.

Compared with diet C, diet T tended to reduce the milk fat yield (P = 0.10) and the SEQ3

diet reduced milk fat yield (P = 0.01) and milk protein yield (P = 0.01; Table 6-3). These results

reflect the reduction in milk yields caused by feeding the T and SEQ3 diets because milk fat and

protein concentrations were not influenced by dietary treatment. The latter agrees with Smith et

al. (1994), who did not detect changes in milk protein and fat percentages due to feeding 200 μg

of AFB1/kg of diet DM to goats. In addition, Kutz et al. (2009) reported that milk protein and fat

percentages were unaffected when dairy cows were fed 112 μg of AFB1/kg of diet DM.

No dietary treatment effects on performance measures were evident during the

withdrawal period, except that the milk protein concentration of cows fed the SEQ1 diet was

greater (P = 0.04) than those of cows fed the SEQ2 diet, but none of the values differed from

those of cows fed diet C (Table 6-3).

Blood Parameters

Compared to diet C, feeding diet T reduced the red blood cell count (P = 0.05) and

hemoglobin concentration (P = 0.02; Table 6-4). However, the blood cell counts were within the

normal range for healthy adult dairy Holstein cows in mid-lactation (George et al., 2010). We

found no previous studies on the effect of dietary AFB1 on the blood cell counts of dairy cows,

but studies in poultry (Huff et al., 1986; Mohiuddin et al., 1986; Basmacioglu et al., 2005) and

rams (Dönmez et al., 2012) have shown that aflatoxin suppressed blood cell counts.

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Expression of CD62L and CD18 tended to be the highest in cows fed the SEQ1 (P =

0.10) and SEQ3 (P = 0.09) diets (Table 6-5). Blood neutrophils act as the first line of defense

against many pathogens once they have access to skin and mucosal surfaces, and migration of

neutrophils involves adhesion mediated by selectins (Smolen et al., 2000). After this initial

phase, the neutrophil is activated for adhesion of its integrins to the vascular endothelium

(Silvestre et al., 2011). The greater MFI of staining for CD62L and CD18 on neutrophils of cows

fed the SEQ1 and SEQ3 diets suggests a more prompt evasion of neutrophil cells from the

vasculature to the tissue. Dietary treatments did not affect the production of Interferon-γ and

percentage of cells exhibiting phagocytosis and oxidative burst activities (Table 6-5).

The production of proinflammatory cytokines is promoted by inflammasomes, which are

a specialized intracellular multiprotein complex (Schroder and Tschopp, 2010). These are innate

immune system receptors that trigger inflammation in response to infectious microbes and/or

toxins (Guo et al., 2015). Inflammasomes are activated by the lethal anthrax toxin (Boyden and

Dietrich, 2006), bacterial toxins such as Shiga toxin (Lee et al., 2015), and bacterial and viral

pathogens (Guo et al., 2015). To our knowledge, the role of inflammasomes in mycotoxin-

mediated inflammation is unknown. Future studies should examine the involvement of

inflammasomes in mycotoxin-mediated responses in dairy cows. Acute-phase proteins are blood

proteins that play major roles in systemic reaction to inflammation (Ceciliani et al., 2012), and

their increased secretion is an indication of inflammatory stress (Bertoni et al., 2008; Gonzalez et

al., 2008; Ceciliani et al., 2012). Haptoglobin and ceruloplasmin concentrations were not

affected by dietary treatments (Table 6-5). However, acid-soluble protein concentrations were

greater (P = 0.01) with diet T than with SEQ2, and tended to be greater with T than with SEQ1

(P = 0.08) and SEQ3 (P = 0.10). Increases of acid-soluble protein concentration in response to

159

inflammatory stress have been reported in goats (Gonzalez et al., 2008), sheep (Eckersall et al.,

2007), pigs (Eckersall et al., 1996), and cows (Nagahata et al., 1989). The result in this study

suggest that feeding the SEQ2 diet reduced the inflammatory response caused by diet T, and the

SEQ1 and SEQ3 diets tended to have a similar effect. Queiroz et al. (2012) fed the same dose (75

μg/kg) of AFB1 as that in the current study to dairy cows and reported that haptoglobin and β-

integrin concentrations were increased by the toxin. However, Xiong et al. (2015) reported that

adding 20 or 40 µg/kg of AFB1 to the diet of dairy cows had no effect on the plasma

concentrations of antibodies, IgG, IgM and IgA. The latter study may have been less successful

at detecting the effects of AFB1 on measures of the immune response because they fed lower

doses of AFB1 and examined different immune response indices. This study showed that the 3

sequestering agents altered the immune status of cows challenged with the toxin and reduced

toxin-induced inflammatory stress.

Conclusions

Feeding 75 μg/kg of AFB1 increased AFM1 concentration in the milk of lactating dairy

cows beyond the FDA action level of 0.5 μg/kg. Feeding the sequestering agents with the toxin

did not prevent the milk AFM1 concentration from exceeding the FDA action level. However,

within 24 h of withdrawing the toxin, diets that included the sequestering agents had reduced the

milk AFM1 concentration below the FDA action level and diet T had not. Feeding AFB1 tended

to reduce the yields of milk, 3.5% FCM, milk fat, and milk protein and reduced red blood cell

count and hemoglobin concentration. Feeding the SEQ1 diet prevented the reduction in milk and

3.5% FCM yield caused by the toxin and increased the MFI of CD62L and CD18 relative to

those for diets T and C. Feeding SEQ2 or SEQ3 did not prevent the adverse effect of the toxin on

milk yield or 3.5% FCM yield. However, the SEQ2 diet reduced the acid-soluble protein

concentration relative to that for cows fed diet T, and the SEQ3 diet increased the MFI of CD62L

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and CD18 relative to those for cows fed diets SEQ2, T, or C. These results imply that the

sequestering agents were not effective at preventing the toxin from being transferred to the milk,

but they did reduce toxin-induced inflammatory stress in cows. In addition, SEQ1 reduced the

adverse effects of the toxin on milk production. Future studies should validate the results of the

current study and elucidate the relative efficacy and synergistic effects of bentonite clays, S.

cerevisiae fermentation products, cell wall oligosaccharides, and chlorophyll-based additives at

alleviating the toxic effects of aflatoxin on dairy cows.

161

Table 6-1. Ingredient and chemical composition of the experimental diet.

Ingredient composition (% of DM) Amount

Corn silage 23.5

Alfalfa hay 6.9

Ryegrass/triticale mixture 8.9

Wet brewers grain 7.8

Corn grain 23.5

Dried citrus pulp 6.5

Soy hulls 5.9

Whole cottonseed 4.9

Soybean meal (47% CP) 5.9

Amino plus1 2.9

Mineral and vitamin mix2 3.3

Chemical composition

Crude protein, % 16.0

ADF, % 20.9

NDF, % 34.2

NEL, Mcal/kg 1.66

Calcium, % 0.86

Phosphorous, % 0.50

Magnesium, % 0.70

Potassium, % 1.31

Sodium, % 0.38

Iron, mg/kg 406

Zinc, mg/kg 110

Copper, mg/kg 29

Manganese, mg/kg 98

Aflatoxins, µg/kg ND3

Deoxynivalenol, mg/kg ND

T-2 toxin, mg/kg ND

Zearalenone, mg/kg ND 1Aminoplus (Ag Processing Inc., Omaha, NE) composition (% DM): Lysine 3.5, Methionine 0.75, Arginine 3.85,

Tryptophan 0.70, Histidine 1.47, Leucine 4.0, Isoleucine 2.43, Phenylalanine 2.64, Threonine 2.01, Valine 2.37,

Tyrosine 1.69, Choline 0.29. 2Vitamin mineral mixture (% as is): ProVAAL-Lys+Met 30 (Perdue Agribusiness,

Salisbury, MD), Sodium bicarbonate 28.5, DCAD Plus 13.0 (Soychlor), Calcium carbonate 9.0, Dicalcium

phosphate 7.0, Magnesium oxide 7.0, Salt (NaCl) 3.5, Zinpro Availa-4 1.2 (Zinpro Corporation Eden Prairie, Mn),

Sel-Plex1000 0.6 (Alltech, Nicholasville, KY), Rumensin90 0.22 (Elanco, Eli Lilly, Indianapolis, IN).

3Concentrations of respective toxins were below lower detection limits (5 μg/kg for aflatoxins, and 0.5 mg/kg for

deoxynivalenol, T-2 toxin, and zearalenone).

162

Table 6-2. Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents1 on the

concentration, transfer and excretion of aflatoxin M1 (AFM1) in the milk of dairy

cows during toxin-dosing and withdrawal periods.

Item

Treatment2 SEM P-value

T SEQ1 SEQ2 SEQ3

Toxin-dosing period (d 21-25)

Concentration (µg/kg) 0.73 0.74 0.73 0.74 0.01 0.74

Excretion (µg/d) 19.45 19.62 19.15 16.68 1.26 0.88

Transfer (%) 1.13 1.14 1.11 1.08 0.07 0.88

Withdrawal period: first day (d 26)

Concentration (µg/kg) 0.63a,x 0.43b,y 0.46ab,y 0.42b,y 0.06 0.01

Excretion (µg/d) 15.87x 11.14y 12.78xy 11.23y 1.99 0.09

Transfer (%) 0.92x 0.65y 0.74xy 0.65y 0.12 0.09

Clearance (µg/h) 0.01 0.01 0.01 0.01 0.00 0.85

Withdrawal period: second day (d 27)

Concentration (µg/kg) 0.06 0.04 0.05 0.03 0.03 0.76

Excretion (µg/d) 1.25 1.02 1.75 1.06 0.82 0.79

Transfer (%) 0.07 0.06 0.10 0.06 0.04 0.79

Clearance (µg/h) 0.01 0.01 0.01 0.01 0.00 0.88

Withdrawal period: third day (d 28)

Concentration (µg/kg) ND3 ND ND ND

Clearance (µg/h) 0.01 0.01 0.01 0.01 0.00 0.88

a-cMeans within a row with no common superscript differ (P ≤ 0.05). xyMeans within a row with no common superscript tend to differ (P ≤ 0.10). 1Proprietary Saccharomyces cerevisiae fermentation product-based sequestering agents

(Diamond V Inc. Cedar Rapids, IA) containing low (SEQ1) or high (SEQ2) doses of a

chlorophyll-based additive or a low dose of the additive and sodium bentonite clay (SEQ3). 2T = Control diet + AFB1; SEQ1 diet = T + SEQ1; SEQ2 diet = T + SEQ2; SEQ3 diet = T +

SEQ3. 3ND = Not detected

163

Table 6-3. Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents1 on the

performance of cows during Predosing (d 16 to 20), Toxin-dosing (d 21 to 25) and

Withdrawal (d 26 to 28) periods.

a-cMeans within a row with no common superscript differ (P ≤ 0.05). xyMeans within a row with no common superscript tend to differ (P ≤ 0.10). 1Proprietary Saccharomyces cerevisiae fermentation product-based sequestering agents



SEQ3. 3Calculated as mean milk yield during the toxin-dosing period– mean milk yield during the

predosing period.

Item Treatment2 SEM P-

value C T SEQ1 SEQ2 SEQ3

Predosing period

DMI (kg/d) 23.4 22.7 22.4 22.7 21.9 0.81 0.30

Milk yield (kg/d) 27.6 26.6 26.5 26.7 26.1 1.20 0.64

Toxin-dosing period

DMI (kg/d) 24.4 24.2 23.7 23.0 23.1 0.85 0.16

Milk yield (kg/d) 28.5a,x 26.0ab,y 26.7ab,xy 25.9ab,y 24.8b,y 1.34 0.01

Milk yield

change3 (kg/d) 0.8 -0.7 0.1 -0.7 -1.1 0.82 0.16

3.5% FCM

(kg/d) 29.8a,x 27.1ab,y 27.7ab,xy 26.9b,y 25.9b,y 1.52 0.01

Efficiency (3.5%

FCM/kg DMI) 1.21 1.12 1.19 1.25 1.17 0.07 0.37

Milk fat (%) 3.79 3.83 3.80 3.79 3.85 0.07 0.63

Milk fat (kg/d) 1.07a,x 0.99ab,y 1.00ab,xy 1.00ab,xy 0.95b,y 0.06 0.01

Milk protein (%) 3.23 3.24 3.24 3.26 3.24 0.02 0.64

Milk protein

(kg/d) 0.92a 0.84ab 0.85ab 0.86ab 0.79b 0.05 0.01

SCC

(×1000/mL) 221 169 209 283 237 59.6 0.34

Withdrawal period

DMI (kg/d) 24.4 23.5 24.4 24.0 23.2 0.92 0.64

Milk yield (kg/d) 26.2 23.5 25.5 23.9 23.1 1.63 0.21

3.5% FCM (kg) 27.3 24.5 26.4 24.7 24.2 1.69 0.17

Efficiency (3.5%

FCM/kg DMI) 1.03 1.00 1.02 0.96 0.99 0.07 0.89

Milk fat (%) 3.88 3.84 3.77 3.77 3.82 0.09 0.60

Milk fat (kg/d) 1.05 0.95 0.99 0.94 0.94 0.06 0.23

Milk protein (%) 3.23ab 3.26ab 3.30a 3.21b 3.22ab 0.03 0.04

Milk protein

(kg/d) 0.88 0.81 0.86 0.80 0.79 0.05 0.20

164

Table 6-4. Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents1 on blood

cell counts of dairy cows.

Item

Treatment2 SEM P-

value C T SEQ1 SEQ2 SEQ3

White blood cells

(103/µL) 15.22 14.99 14.85 15.14 14.65 0.52 0.94

Red blood cells

(106/µL) 6.08a 5.80b 5.87ab 5.93ab 5.85ab 0.07 0.05

Neutrophils (103/µL) 3.26 3.43 3.28 3.26 3.36 0.36 0.98

Monocytes (103/µL) 1.65 2.11 1.49 1.87 1.65 0.28 0.22

Eosinophils (103/µL) 0.37 0.38 0.40 0.40 0.40 0.03 0.93

Lymphocytes

(103/µL) 9.51 8.93 10.55 8.81 9.58 0.68 0.12

Basophils (103/µL) 0.11 0.05 0.02 0.02 0.00 0.03 0.12

Hemoglobin (g/dL) 9.97a 9.48b 9.65ab 9.62ab 9.57ab 0.14 0.02

Hematocrit (%) 29.85 28.59 28.85 29.06 28.78 0.52 0.16 a,bMeans within a row with no common superscript differ (P ≤ 0.05). 1Proprietary Saccharomyces cerevisiae fermentation product-based sequestering agents



SEQ3.

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Table 6-5. Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents1 on immune

response indices of dairy cows.

Item Treatment2 SEM P-

value

C T SEQ1 SEQ2 SEQ3

Granulocyte adhesion molecules

β2-integrin (CD18; MFI3) 918y 908y 1278x 956y 1312x 202 0.10

L-Selectin (CD62L; MFI) 2188y 2353y 3284x 2270y 3115x 516 0.09

Neutrophil function

Neutrophil phagocytosis (%

of cells)

27.72

26.55

28.27

27.69

28.45

2.28

0.94

Oxidative burst (% of cells) 2.94 2.08 3.76 3.06 1.61 0.78 0.11

Cytokine4

Interferon-γ (MFI) 4.79 6.56 6.75 4.55 6.16 1.52 0.46

Interferon-γ (% of cells) 3.27 3.08 3.48 3.90 2.43 0.82 0.46

Acute-phase proteins

Haptoglobin5 0.03 0.02 0.03 0.02 0.08 0.03 0.44

Ceruloplasmin (mg/100mL) 19.50 20.59 20.79 20.57 20.28 1.01 0.72

Acid-soluble protein

(mg/100 mL) 4.39ab,xy 5.10a,x 4.25ab,xy 3.69b,y 4.15ab,y 0.35 0.01

a,bMeans within a row with no common superscript differ (P ≤ 0.05). x,yMeans within a row with no common superscript tend to differ (P ≤ 0.10). 1Proprietary Saccharomyces cerevisiae fermentation product-based sequestering agents



SEQ3. 3MFI = mean fluorescence intensity. 4Fold increase of ionomycin and phytohemagglutinin-stimulated cells over the non-stimulated

cells. 5Arbitrary unit based on absorbance at 450 nm.

166

Figure 6-1. Effect of dosing aflatoxin B1 (AFB1) with or without sequestering agents on the

concentration of aflatoxin M1 (AFM1) in the milk of dairy cows.+Toxin-dosing period

= d 21 to 25, Withdrawal period = d 26 to 28. *AFM1 concentrations on the day

specified differ (P = 0.01). Error bars indicate standard errors. SEQ1, 2 and 3 are

proprietary Saccharomyces cerevisiae fermentation product-based sequestering

agents (Diamond V Inc., Cedar Rapids, IA) containing low (SEQ1) or high (SEQ2)

doses of a chlorophyll-based additive or a low dose of the additive and sodium

bentonite clay (SEQ3); T = Control diet + AFB1; SEQ1 diet = T + SEQ1; SEQ2 diet

= T + SEQ2; SEQ3 diet = T + SEQ3.

0.0

0.2

0.4

0.6

0.8

1.0

20 21 22 23 24 25 26 27 28

Afl

atox

in M

1 (

µg/k

g)

Day+

T SEQ1SEQ2 SEQ3FDA Action level

167

CHAPTER 7

GENERAL SUMMARY

The experiments presented in this dissertation had three main objectives. The first

objective was to evaluate the effects of treatment with either of two bacterial cultures or

propionic acid on the survival of EC during and after ensiling of corn and alfalfa. The second

objective was to determine the effects of inoculation with EC alone or either of two bacterial

cultures or propionic acid on the bacterial community composition of corn and alfalfa silages and

to examine if the abundance of known and unknown bacteria correlates with key silage

fermentation indices. The third objective was to examine if addition of Saccharomyces

cerevisiae to other aflatoxin-sequestering agents (clay or chlorophyll-based products), would

synergistically increase binding of aflatoxin in dairy cows, reduce adverse immune responses to

the toxin, and enhance the performance of dairy cows. In order to achieve these objectives, four

studies were performed.

The first study examined if adding L. plantarum or L. buchneri or propionic acid to

alfalfa silages contaminated with EC would inhibit the growth of the pathogen during or after

ensiling. Alfalfa forage was harvested at the early bloom stage, wilted to a DM concentration of

54%, chopped to 19-mm lengths and ensiled after treatment with one of the following: 1,

distilled water (Control); 2, 1 × 105 cfu/g of E. coli O157:H7 (EC); 3, EC and 1 × 106 cfu/g of

Lactobacillus plantarum (EC+LP); 4, EC and 1 × 106 cfu/g of Lactobacillus buchneri (EC+LB);

and 5, EC and 2.2 g/kg of propionic acid (EC+PA). Each treatment was ensiled in quadruplicate

in laboratory silos for 0, 3, 7, 16, and 100 d and analyzed for EC counts, pH, and organic acids.

In addition, samples from d 100 were analyzed for chemical composition, ammonia-N, counts of

yeasts and molds and aerobic stability. The pathogen was detected in all silages until d 7, but by

d 16, it was not detected in those treated with EC+LB and EC+LP, though it was still detected in

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EC and EC+PA silages. However, by d 100, the pathogen was not detected in any silage. The pH

dropped to 5.0 within 7 d for the EC+LP silage, followed by 16 d for the EC+LB silage and

beyond 16 d for the others. Nevertheless, all silages had attained a pH of or less than 5.0 by d

100. The rapid decrease in pH in EC+LP and EC+LB silages was because they had higher lactate

and acetate concentrations, respectively, relative to the other silages during the early

fermentation phase (d 3 to 16). Propionic acid was only detected in the EC+PA silage. Yeast

counts were lowest in EC+LB and EC+PA silages. Subsamples of all d 100 silages were re-

inoculated with 1 × 105 cfu/g of EC immediately after silo opening. When the pathogen was

subsequently enumerated after 168 h of aerobic exposure, it was not detected in silages treated

with EC+PA, EC+LB or EC+LP, which all had pH values less than 5.0. Whereas, the EC silage

had a pH value of 5.4 and 2.3 log cfu/g of the pathogen. Certain bacterial inoculants can hasten

the inhibition of EC during ensiling and like propionic acid, they can also prevent its growth on

silage contaminated with the pathogen after ensiling.

The second study applied the Illumina-based metagenomic sequencing technique to

examine the effects of silage additives on bacterial diversity and taxonomy of alfalfa silage. The

d 100 silage samples from Chapter 3 were analyzed for bacterial diversity and taxonomic

composition by Illumina MiSeq Sequencing. Compared to the Control, a less diverse bacterial

community was detected in the EC+LP and EC+LB silages. Overall, most sequences were

associated with phylum Firmicutes followed by Proteobacteria. Treatment with EC+LP and

EC+PA reduced the abundance of Firmicutes, whereas that of Proteobacteria increased compared

with the Control and EC+LB silages. Treatment with EC+LP resulted in the greatest abundance

of Lactobacillus (62.6%), increased abundance of Sphingomonas, Pantoea, Pseudomonas and

Erwinia and reduced abundance of Pediococcus, Weissella and Methylobacterium, relative to the

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Control. Abundance of Weissella and Methylobacterium were also reduced in EC+LB silage

relative to the Control. Application of propionic acid did not affect the abundance of

Lactobacillus, Weissella or Pediococcus. At the species level, EC+LB and EC+LP silages

contained the greatest abundance of L. buchneri and L. plantarum, respectively. Lactate

concentration correlated positively with abundance of Lactobacillus and negatively with

abundance of Pediococcus. Negative correlations were observed between ammonia-N

concentration and abundance of genera Sphingomonas, Pantoea, Pseudomonas and

Stenotrophomonas. Silage pH was negatively correlated with abundance of Lactobacillus,

Sphingomonas, Pantoea, Pseudomonas, Erwinia and Stenotrophomonas. The abundance of

bacterial species belonging to Rhodococcus, Pseudomonas, and Xanthomonadaceae, had positive

correlations with lactate and/or acetate, while unknown species belonging to Salana,

Frigoribacterium, Pseudorhodobacter, Pantoea, Stenotrophomonas, and Rhodococcus

had/negative correlations with ammonia-N. Future studies should aim to speciate, culture and

validate the functions of these unknown bacteria in order to better understand their roles in silage

fermentation.

The third study examined if addition of propionic acid or L. plantarum or L. buchneri

would inhibit the growth of EC on corn silage during ensiling, at silo opening, or after aerobic

exposure. The second objective was to examine how additives affect the bacterial community

composition in corn silage. Corn forage was harvested at approximately 35% dry matter,

chopped to a theoretical length of cut of 10 mm, and ensiled after treatment with one of the

following:1), Distilled water (Control); 2), 1 × 105 cfu/g of EC (ECCH); 3), EC and 1 × 106 cfu/g

of Lactobacillus plantarum (ECLP); 4), EC and 1 × 106 cfu/g of Lactobacillus buchneri (ECLB);

and 5), EC and 2.2 g/kg (fresh weight basis) of propionic acid, containing 99.5% of the acid

170

(ECA). Each treatment was ensiled in quadruplicate in laboratory silos for 0, 3, 7, and 120 d and

analyzed for EC, pH, and organic acids. Samples from d 0 and 120 were also analyzed for

chemical composition. Furthermore, samples from d 120 were analyzed for ammonia-N, yeasts

and molds, lactic acid bacteria (LAB), bacterial community composition and aerobic stability.

The pH of silages from all treatments decreased below 4 within 3 d of ensiling. Escherichia coli

O157:H7 was below the detection limit in all silages after 7 d of ensiling. Treatment with L.

buchneri or propionic acid resulted in fewer yeasts and greater aerobic stability compared with

the Control, ECCH, and ECLP silages. Compared to the Control, bacterial diversity was less in

the ECLP silage partly because it contained more Lactobacillus. The ECLB silage contained

greater abundance of Acinetobacter and Weissella than other silages. Subsamples of silages were

reinoculated with 5 × 105 cfu/g of EC either immediately after silo opening or after 168 h of

aerobic exposure, and EC were enumerated after 6 or 24 h, respectively. All silages reinoculated

with EC immediately after silo opening (120 h) had similar low pH values (< 4.0) and no EC was

detected in such silages. The ECCH and ECLP silages re-inoculated with EC after 168 h of

aerobic exposure had relatively high pH values (> 5.0) and EC counts (5.39 and 5.30 log cfu/g,

respectively) 24 h later. Whereas those treated with L. buchneri or propionic acid had lower pH

values (4.24 or 3.96, respectively) and lower EC counts (1.32 log cfu/g or none, respectively).

This study indicates that during ensiling, EC was eliminated from all silages once the pH

dropped below 4.0, whereas during aerobic exposure, the growth of EC was reduced or

prevented in silages that had been treated with L. buchneri or propionic acid at ensiling,

respectively.

The fourth study examined if adding three mycotoxin-sequestering agents to diets

contaminated with aflatoxin B1 (AFB1) would reduce milk aflatoxin M1 (AFM1) concentration

171

and enhance the performance and immune status of dairy cows. Fifteen lactating dairy cows

were used in an experiment with an incomplete crossover design with four 28-d periods.

Treatments included a Control diet (C), a toxin (T) diet (1725 µg of AFB1/hd/d; 75 µg/kg) and

diets containing the toxin and 20 g/hd/d of a proprietary mixture of Saccharomyces cerevisiae

fermentation product containing a low (SEQ1) or high (SEQ2) dose of a chlorophyll-based

additive, or a low dose of the chlorophyll-based additive with sodium bentonite clay (SEQ3).

Sequestering agents were top-dressed on the TMR daily in each period and the AFB1 was orally

dosed in gelatin capsules to cows before the TMR was fed on d 21 to 25. Milk was sampled

twice daily on d 20 to 28 and plasma was sampled on d 20 and 25. The AFM1 concentration of

the milk was not affected by the sequestering agents during the toxin-dosing period. However,

after withdrawal of AFB1 from the diets, the sequestering agents reduced the time required (24

vs. 48 h) for reduction of the milk AFM1 concentration below the Food and Drug Administration

(FDA) Action level of 0.5 µg/kg. Feeding T instead of C tended to reduce the milk and FCM

yields but feeding SEQ1 prevented these effects. Red blood cell count and hemoglobin

concentration were reduced by feeding diet T instead of C, but not by feeding diets containing

SEQ1, SEQ2 or SEQ3. Mean fluorescence intensity of antibody staining for two leukocyte

adhesion molecules, L-selectin (CD62L) and β-integrin (CD18), tended to be greatest when

SEQ1 and SEQ3 were fed. Plasma acid-soluble protein concentration was decreased when the

SEQ1, SEQ2 and SEQ3 diets were fed instead of diet T. Sequestering agents had no effect on

milk AFM1 concentration but they reduced the time required for reduction of milk AFM1

concentration to a safe level after withdrawal of AFB1. Only SEQ1 prevented adverse effects of

AFB1 on milk and FCM yields.

172

In addition to their well-established roles in improving silage fermentation, nutrient and

DM recovery, and aerobic stability of silage at feed-out, the series of studies demonstrates

conclusively that silage additives can be used to inhibit the growth of pathogens in feeds. The

results from these studies (Chapters 3 and 4) confirm that pathogenic E. coli can be found on

forages and show that L. plantarum or L. buchneri can increase the acidification of corn and

alfalfa silages and hasten inhibition of the pathogen during ensiling. In addition to improving the

aerobic stability of corn silage, L. buchneri and propionic acid also inhibited or prevented the

growth of EC when contamination occurred after silo opening and aerobic exposure. Therefore,

these silage additives can play a vital role in reducing cycling of EC on farms. This is vital for

reducing the frequency of EC outbreaks and the associated food-borne diseases that cost the US

approximately $271.4 million annually (USDA, 2014). Future research should examine the effect

of controlling the growth of EC in silage on the overall prevalence of EC on farms. If this

approach is successful, it would eliminate the high cost of vaccination to reduce EC shedding by

ruminants. Another aspect worthy of research is to examine if silage inoculation with L.

plantarum or L. buchneri can reduce fecal shedding of EC by ruminants. Future research should

also investigate the prevalence and risk factors associated with EC shedding in dairy cattle.

Better knowledge of these factors might guide effective mitigation strategies on farms to reduce

EC contamination and cycling.

Studies presented in Chapters 4 and 5 highlight the fact that several silage

microorganisms are unknown or uncultured and consequently their roles are unknown; however,

some unknown or yet to be cultured silage bacteria are strongly associated with fermentation

quality indices, such as lactate, acetate, and ammonia-N. Research is needed on functional

characterization of the complex microbial communities in silage and how they are influenced by

173

external factors like additives. Future research should apply function-based metagenomics to

identify and validate the roles of the unknown bacteria in silage fermentation. This may lead to

development of new microbial additives that can improve silage preservation and allow more

efficient utilization of forage protein by ruminants, thereby, reducing nitrogen excretion to the

environment. Future research should also characterize the silage metabolome to provide more

comprehensive information on chemical changes during ensiling and on how additives affect

silage quality. Future research should integrate the results of multi-omics silage characterization

approaches (metagenomics, metatranscriptomics, and metabolomics) to provide definitive

insights on the types and roles of micro-organisms present and to show how their metabolites are

involved in key signaling and metabolic pathways in silage.

Due to the ubiquitous occurrence of mycotoxins in grains and forages, frequent

contamination of livestock feeds with multiple mycotoxins, inconsistent effects of available

sequestering agents, and the global threat of mycotoxins to food and feed security, future

research should aim to develop more effective mycotoxin sequestering agents that can counteract

the additive and or synergistic effects of multiple mycotoxins. Studies on the ecology of specific

gut microbes that degrade mycotoxins, particularly aflatoxins, are also needed to reduce their

toxic effects and transfer to dairy products. To replace expensive chromatography methods,

hazardous radio immunoassay methods and laborious ELISA methods, future research also

should develop simple, rapid and accurate mycotoxin detection techniques to foster on-site or on-

farm detection of mycotoxins in feeds.

174

LIST OF REFERENCES

Abdul-Raouf, U. M., L. R. Beuchat, and M. S. Ammar. 1993. Survival and growth of

Escherichia coli O157:H7 on salad vegetables. Appl. Environ. Microbiol. 59:1999–2006.

Addah W., J. Baaha, E. K. Okineb, and T. A. McAllister. 2014. Silage Inoculants – Are they

worth the money? Adv. Dairy Tech. 26:193–206.

Ahlberg, S. H., V. Joutsjoki, and H. J. Korhonena 2015. Potential of lactic acid bacteria in

aflatoxin risk mitigation. Int. J. Food Microbiol. 207: 87-102

Alassane-Kpembi, I., G. Schatzmayr, I. Taranu, D. Marin, O. Puel, and I. P. Oswald. 2016.

Mycotoxins co-contamination: Methodological aspects and biological relevance of

combined toxicity studies. Crit. Rev. Food Sci. Nutr. http://dx.doi.org/10.1080/

10408398.2016.1140632.

Albrecht, K. A., and K. A. Beauchemin. 2003. Alfalfa and other perennial legume silage. Pages

633–665 in Silage Science and Technology. D. R. Buxton, R. E. Muck, and J. H.

Harrison, ed. American Society of Agronomy Inc., Crop Science Society of America Inc.,

Soil Science Society of America Inc. Publications, Madison, WI.

Alegakis A. K., A. M. Tsatsakis, M. I. Shtilman, D. L. Lysovenko, and I. G. Vlachonikolis.

1999. Deactivation of mycotoxins. I. An in vitro study of zearalenone adsorption on new

polymeric adsorbents. J. Environ. Sci. Health B. 34:633-644.

Allen, B., M. Kon, and Y. Bar-Yam. 2009. A new phylogenetic diversity measure generalizing

the Shannon index and its application to phyllostomid bats. Am. Nat. 174:236–243.

Alm, K., M. Dahlbom, M. Säynäjärvi, M. A. Andersson, M. S. Salkinoja-Salonen, M. C.

Andersson. 2002. Impaired semen quality of AI bulls fed with moldy hay: A case report.

Theriogenology 58:1497–1502.

Almuzara M. N., M. Vazquez, N. Tanaka, M. Turco, M. S. Ramirez, E. L. Lopez, F. Pasteran,

M. Rapoport, A. Procopio, and C. A. Vay. 2010. First case of human infection due to

Pseudomonas fulva, an environmental bacterium isolated from cerebrospinal fluid. J.

Clin. Microbiol. 48:660–664.

Amaral, R. C., L. G. Nussio, M. Zopollatto, R. S. Goulart, E. H. Cabezas Garcia, and S. G. de

Toledo Filho. 2010. Lower temperature profile and less aflatoxin on corn silage covered

by oxygen barrier plastic film. Adv. Anim. Biosciences 1:469-470.

Andreoletti, O., H. Budka, S. Buncic, P. Colin, J. Collins, A. De Koeiejer. 2008. Microbiological

risk assessment in feedingstuffs for food-producing animals, Scientific opinion of the

panel on biological hazards. The EFSA J. 720:1-84.

Anzai, Y., H. Kim, J. Y. Park, H. Wakabayashi, and H. Oyaizu. 2000. Phylogenetic affiliation of

the pseudomonads based on 16S rRNA sequences. Int. J. Syst. Evol. Microbiol. 50:1563–

89.

175

AOAC International. 2000. Official Methods of Analysis. 17th ed. AOAC Int., Arlington VA.

Arimoto, S., S. Fakuoka, C. Itome, H. Nakano, H. Rai, and H. Hayatsu. 1993. Binding of

polycyclic planar mutagens to chlorophyllin resulting in inhibition of the mutagenic

activity. Mutat. Res. 287:293-305.

Arriola, K. G., C. R. Staples, S. C. Kim, and A. T. Adesogan. 2011. Effect of applying inoculants

containing different types of bacteria on the performance of lactating dairy cows. J. Dairy

Sci. 94:3973-3979.

Auerbach, H., E. Oldenburg, and F. Weissbach. 1998. Incidence of Penicillium roqueforti and

roquefortine C in silages. J. Sci. Food Agric. 76:565–572.

Avery, S., L. Walters, and M. Hutchison. 2005. Fate of Escherichia coli O157 and detection of

stx phage during fermentation of maize, an animal feedstuff. Lett. Appl. Microbiol.

40:99–105.

Axelsson, L. 1998. Lactic acid bacteria: classification and physiology. Pages 1–72 in: Lactic acid

bacteria: microbiology and functional aspects, 2nd ed. S. Salminen and A. Von Wright,

Marcel Dekker Inc, New York.

Baath, H., O. Knabe, and P. Lepom. 1990. Occurrence of Fusarium species and their mycotoxins

in maize silage. Studies on the Fusarium infestation of maize silage plants. Arch. Anim.

Nutr. 40:397–405.

Bach, S. J., T. A. McAllister, D. M. Veira, V. P. J. Gannon, and R. A. Holley. 2003. Effects of a

Saccharomyces cerevisiae feed supplement on Escherichia coli O157:H7 in ruminal fluid

in vitro. Anim. Feed Sci. Technol. 104:179-189.

Bach, S. J., T. A. McAllister, J. Baah, L. J. Yanke, D. M. Veira, V. P. J. Gannon, and R. A.

Holley. 2002. Persistence of Escherichia coli O157:H7 in barley silage: Effect of a

bacterial inoculant. J. Appl. Microbiol. 93:288–294.

Baek, E., H. Kim, H. Choi, S. Yoon, and J. Kim. 2012. Antifungal activity of Leuconostoc

citreum and Weissella confuse in rice cakes. J. Microbiol. 50:842-848.

Bal-Krishna, K. C., A. Sathasivan, and M. P. Ginige. 2013. Microbial community changes with

decaying chloramine residuals in a lab-scale system. Water Res. 47:4666−4679.

Bao, W., Z. Mi, H. Xu, Y. Zheng, L. Y. Kwok, H. Zhang, and W. Zhang. 2016. Assessing

quality of Medicago sativa silage by monitoring bacterial composition with single

molecule, real-time sequencing technology and various physiological parameters. Sci.

Rep. 6:28358; doi: 10.1038/srep28358.

Bartolome, B., C. B. Faulds, M. Tuohy, G. P. Hazlewood, H. J. Gilbert, and G. Williamson.

1995. Influence of different xylanases on the activity of ferulic acid esterase of wheat

bran. Biotechnol. Appl. Biochem. 22:65–73.

176

Basmacioglu, H., H. Oguz, M. Ergul, R. Col, and Y. O Birdane. 2005. Effect of dietary esterified

glucomannan on performance, serum biochemistry and haematology in broilers exposed

to aflatoxin. Czech J. Anim. Sci. 50:31–39.

Bata, Á., and R. Lásztity. 1999. Detoxification of mycotoxin-contaminated food and feed by

microorganisms. Trends in Food Sci. Tech. 10:223-228.

Battacone, G., A. Nudda, M. Palomba, M. Pascale, P. Nicolussi, and G. Pulina. 2005. Transfer of

aflatoxin B1 from feed to milk and from milk to curd and whey in dairy sheep fed

artificially contaminated concentrates. J. Dairy Sci. 88:3063–3069.

Bauer, A. W., M. W. Kirby, J. C. Sherris, and M. Turck. 1966. Antibiotic susceptibility testing

by a standardized single disk method. Am. J. Clin. Pathol. 45:493–496.

Beck, T. 1972. The quantitative and qualitative composition of the lactic acid bacteria population

in silage. Landwirtschaftliche Forschung 27:55–63.

Berthier, F., and S. D. Elrhich. 1998. Rapid species identification within two groups of closely

related lactobacilli using PCR primers that target the 16S–23S rRNA spacer region.

FEMS Microbiol. Lett. 161:97–106.

Bertoni, G., E. Trevisi, X. Han, and M. Bionaz. 2008. Effects of Inflammatory conditions on

liver activity in the puerperium and consequences for performance in dairy cows. J. Dairy

Sci. 91:3300–3310.

Beuchat, L. R., and J. H. Ryu. 1997. Produce handling and processing practices. Emerg. Infect.

Dis. 3: 459–465.

Bhat, R., R. V. Rai, and A. A. Karim. 2010. Mycotoxins in food and feed: present status and

future concerns. Compr. Rev. Food Sci. Food Safety 9:57–81.

Binder, E. M., L. M. Tan, L. J. Chin, J. Handl, and J. Richard. 2007. Worldwide occurrence of

mycotoxins in commodities, feeds and feed ingredients. Anim. Feed Sci. Technol.

137:265–282.

Biomin 2015. Mycotoxin survey report. Accessed August 15, 2016.

https://info.biomin.net/acton/fs/blocks/showLandingPage/a/14109/p/p004e/t/page/fm/17.

Bjornsdottir, K., F. Breidit Jr., and R. F. McFeeters. 2006. Protective effect of organic acids on

survival of Escherichia coli O157:H7 in acidic environments. Appl. Environ. Microbiol.

72: 660–664.

Black, R. D., D. S. McVet, and F. W. Oehme. 1992. Immunotoxicity in the bovine animal. Vet.

Hum. Toxicol. 34:438–442.

Bottalico, A., 1998. Fusarium diseases of cereals: Species complex and related mycotoxin

profiles in Europe. J. Plant Pathol. 80:85–103.

https://info.biomin.net/acton/fs/blocks/showLandingPage/a/14109/p/p004e/t/page/fm/17

177

Boyden, E. D., and W. F. Dietrich. 2006. Nalp1b controls mouse macrophage susceptibility to

anthrax lethal toxin. Nat. Genet. 38:240-244.

Boysen, M. E., K. G. Jacobsson, and J. Schnurer. 2000. Molecular identification of species from

the Penicillium roqueforti group associated with spoiled animal feed. Appl. Environ.

Microb. 66:1523–1526.

Breidt, F., Jr., J. S. Hayes, and R. F. McFeeters. 2004. The independent effects of acetic acid and

pH on the survival of Escherichia coli O157:H7 in simulated acidified pickle products. J.

Food Prot. 67:12–18.

Brusetti, L., S. Borin, D. Mora, A. Rizzi, N. Raddadi, C. Sorlini, and D. Daffonchio. 2006.

Usefulness of length heterogeneity-PCR for monitoring lactic acid bacteria succession

during maize ensiling. FEMS Microbiol. Ecol. 56:154-164.

Buchanan, R. L., and S. G. Edelson. 1999. pH-dependent stationary-phase acid resistance

response of enterohemorrhagic Escherichia coli in the presence of various acidulants. J.

Food Prot. 62:211–218.

Buchanan, R. L., S. G. Edelson-Mammel, G. Boyd, and B. S. Marmer. 2004. Influence of

acidulant identity on the effects of pH and acid resistance on the radiation resistance of

Escherichia coli O157:H7. Food Microbiol. 21:51–57.

Bursian, S. J., R. J. Aulerich, J. K. Cameron, N. K. Ames, and B. A. Steficek. 1992. Efficacy of

hydrated sodium calcium aluminosilicate in reducing the toxicity of dietary zearalenone

to mink. J. Appl. Toxicol. 12:85–90.

Byrne, C. M., P. O. Kiely, D. J. Bolton, J. J. Sheridan, D. A. McDowell, and I. S. Blair. 2002.

Fate of Escherichia coli O157:H7 during silage fermentation. J. Food Prot. 65:1854–

1860.

Cai, Y., Y. Benno, M. Ogawa, S. Ohmomo, S. Kumai, and T. Nakase. 1998. Influence of

Lactobacillus spp. from an inoculant and of Weissella and Leuconostoc spp. from forage

crops on silage fermentation. Appl. Environ. Microbiol. 64:2982–2987.

Callaway, T. R., M. A. Car, T. S. Edrington, R. C. Anderson, and D. J. Nisbet. 2009. Escherichia

coli O157:H7, and cattle: A review after 10 years. Curr. Issues Mol. Biol. 11:67–79.

Callaway, T. R., T. S. Edrington, A. D. Brabban, R. C. Anderson, M. L. Rossman, M. J. Engler,

M. A. Carr, K. J. Genovese, J. E. Keen, M. L. Looper, E. M. Kutter, and D. J. Nisbet.

2008. Bacteriophage isolated from feedlot cattle can reduce Escherichia coli O157:H7

populations in ruminant gastrointestinal tracts. Foodborne Pathog. Dis. 5:183–191.

Callaway, T. R., T. S. Edrington, A. D. Brabban, J. E. Keen, R. C. Anderson, M. L. Rossman, M.

J. Engler, K. J. Genovese, B. L. Gwartney, J. O. Reagan, T. L. Poole, R. B. Harvey, E. M.

Kutter, and D. J. Nisbet. 2006. Fecal prevalence of Escherichia coli O157, Salmonella,

Listeria, and bacteriophage infecting E. coli O157:H7 in feedlot cattle in the southern

plains region of the United States. Foodborne Pathog. Dis. 3:234-244.

178

Callaway, T. R., and S. A. Martin. 2006. Use of competitive exclusion cultures and

oligosaccharides. In: Feedstuffs. Direct-fed Microbial, Enzyme and Forage Additive

Compendium, 8th Ed. (eds), Miller Publishing, Minnetonka, MN. pp. 34-39.

Campbell A. W., J. D. Thrasher, M. R. Gray, A. Vojdani. 2004. Mold and mycotoxins: effects on

the neurological and immune systems in humans. Adv. Appl. Microbiol. 55:375-406.

Canady, R. A., R. D. Coker, S. K. Rgan, R. Krska, T. Kuiper-Goodman, M. Olsen, J. J. Pestka,

S. Resnik, J. Schlatter. 2001. Deoxynivalenol. Safety Evaluation of Certain Mycotoxins

in Food. Fifty-Sixth Report of the Joint FAO/WHO Expert Committee on Food

Additives. International Programme on Chemical Safety, World Health Organization,

Geneva, pp. 420–555.

Caporaso J. G., J. Kuczynski, J. Stombaugh, K. Bittinger, F. D. Bushman, E. K. Costello, N.

Fierer, P. A. Gonzalez, J. K. Goodrich, J. I. Gordon, G. A. Huttley, S. T. Kelley, D.

Knights, J. E. Koenig, R. E. Ley, C. A. Lozupone, D. McDonald, B. D. Muegge, M.

Pirrung, J. Reeder, J. R. Sevinsky, P. J. Turnbaugh, W. A. Walters, J. Widmann, T.

Yatsunenko, J. Zaneveld, and R. Knight. 2010. QIIME allows analysis of high-

throughput community sequencing data. Nature Methods 7:335-336.

CAST, Council for Agricultural Science and Technology. 1989. Mycotoxins: economics and

health risks. Task Force Report No. 116. Ames, Iowa.

CAST, Council for Agricultural Science and Technology. 2003. Mycotoxins: risk in plant,

animal and human systems. Task Force Report No. 139. Ames, Iowa.

Cavallarin, L., E. Tabacco, S. Antoniazzi, and G. Borreani. 2011. Aflatoxin accumulation in

whole crop maize silage as a result of aerobic exposure. J. Sci. Food and Agric. 91:2419-

2425.

Ceciliani, F., J. J. Ceron, P. D. Eckersall, and H. Sauerwein. 2012. Acute phase proteins in

ruminants. J. Proteomics 75:4207–31.

Centers for Disease Control and Prevention, 2015. Escherichia coli. Accessed: 16 December

2016. http://www.cdc.gov/ecoli/general/.

Cernicchiaro, N., C. A. Cull, Z. D. Paddock, X. Shi, J. Bai, T. G. Nagaraja, D. G. Renter. 2013.

Prevalence of shiga toxin-producing Escherichia coli and associated virulence genes in

feces of commercial feedlot cattle. Foodborne Pathog. Dis. 10:835–841.

Charley, R. C., and L. Kung, Jr. 2005. Treatment of silage with Lactobacillus diolivorans.

Lallemand Animal Nutrition North America, assignee. US Pat. No. 20050281917.

Charmley, E., H. L. Trenholm, B. K. Thompson, D. Vudathala, J. W. G. Nicholson, D. B.

Prelusky, and L. L. Charmley. 1993. Influence of levels of deoxynivalenol in the diet of

dairy cows on feed intake, milk production, and its composition. J. Dairy Sci. 76: 3580-

3587.

http://www.cdc.gov/

http://www.cdc.gov/ecoli/general/

http://www.google.com/search?tbo=p&tbm=pts&hl=en&q=inassignee:%22Lallemand+Animal+Nutrition+North+America%22

http://www.google.com/patents/US20050281917?cl=en

179

Chase-Topping, M., D. Gally, C. Low, and M. Woolhouse. 2008. Super-shedding and the link

between human infection and livestock carriage with Escherichia coli O157. Nat. Rev.

6:904–912.

Cheeke, P. R., 1998. Mycotoxins associated with forages. In: Cheeke, P.R. (Ed.), Natural

Toxicants in Feeds, Forages, and Poisonous Plants. Interstate Publishers, Inc, Danville,

IL, pp. 243–274

Cheli, F., A. Campagnoli, and V. Dell’Orto. 2013. Fungal populations and mycotoxins in silages:

From occurrence to analysis. Anim. Feed Sci. Technol. 183:1–16.

Chen, Y., S. Sela, M. Gamburg, R. Pinto, and Z. G. Weinberg. 2005. Fate of Escherichia coli

during ensiling of wheat and corn. Appl. Environ. Microbiol. 71:5163–5170.

Chestnut, A. B., P. D. Anderson, M. A. Cochran, H. A. Fribourg, and K. D. Gwinn. 1992. Effects

of hydrated sodium calcium aluminosilicate on fescue toxicosis and mineral absorption.

J. Anim. Sci. 70:2838-2846.

Christensen, C. M., C. J. Mirocha, and R. A. Meronuck. 1977. Molds, Mycotoxins, and

Mycotoxicoses. Agricultural Experiment Station Miscellaneous Report 142. University of

Minnesota, St. Paul, MN.

Chu, F. S., 1974. A comparative study of the interaction of ochratoxins with bovine serum

albumin. Biochem. Pharmacol. 23:1105–1113

Chung, T. K., J. W. Erdman, and D. H. Baker. 1990. Hydrated sodium calcium aluminosilicate:

Effects on zinc, manganese, vitamin A, and riboflavin utilization. Poult. Sci. 69:1364–

1370.

Cobbold R. N., D. D. Hancock, D. H. Rice, J. Berg, R. Stilborn, C. J. Hovde, and T. E. Besser.

2007. Rectoanal junction colonization of feedlot cattle by Escherichia coli O157:H7 and

its association with supershedders and excretion dynamics. Appl. Environ. Microbiol.

73:1563–1568.

Cole, J. R., Q. Wang, J. A. Fish, B. Chai, D. M. McGarrell, Y. Sun, C. T. Brown, A. Porras-

Alfaro, C. R. Kuske, and J. M. Tiedje. 2014. Ribosomal Database Project: data and tools

for high throughput rRNA analysis. Nucl. Acids Res. 42:633-642.

Collins, M., and V. N. Owens. 2003. Preservation of forage as hay and silage. In Forages: an

introduction to grassland agriculture, pp. 443-471. R.F. Barnes, C. J. Nelson, K. J. Moore

and M. Collins, eds. Blackwell Publishing, Ames, IA.

Colvin, B. M., and L. R. Harrison. 1992. Fumonisin-induced pulmonary edema and hydrothorax

in swine. Mycopathologia 117:79–82.

Coppock, R. W., M. S. Mostrom, C. G. Sparling, B. Jacobsen, and S. C. Ross. 1990. Apparent

zearalenone intoxication in a dairy herd from feeding spoiled acid-treated corn. Vet.

Hum. Toxicol. 32:246-248.

180

Cote, L. M., A. M. Dahlem, T. Yoshizawa, S. P. Swanson, and W. B. Buck. 1986. Excretion of

deoxynivalenol and its metabolite in milk, urine, and faeces of lactating dairy cows. J.

Dairy Sci. 69:2416–2423.

Coulombe Jr, R. A. 1993. Biological Action of Mycotoxins. J. Dairy Sci. 76: 880-891.

Creepy, E. E. 2002. Update of survey, regulation and toxic effects of mycotoxins in Europe.

Toxicol. Lett. 127:19–28.

Croxen, M. A., R. J. Law, R. Scholz, K. M. Keeney, M. Wlodarska, and B. B. Finlay. 2013.

Recent advances in understanding enteric pathogenic Escherichia coli. Clin. Microbiol.

Rev. 26:822–880.

Cull, C. A., Z. D. Paddock, T. G. Nagaraja, N. M. Bello, A. H. Babcock, and D. G. Renter. 2012.

Efficacy of a vaccine and a direct-fed microbial against fecal shedding of Escherichia

coli O157:H7 in a randomized pen-level field trial of commercial feedlot cattle. Vaccine

30:6210– 6215.

Dänicke, S., K. Matthäus, P. Lebzien, H. Valenta, K. Stemme, K. H. Ueberschär, E. Razzazi–

Fazeli, J. Böhm, and G. Flachowsky. 2005. Effects of Fusarium toxin–contaminated

wheat grain on nutrient turnover, microbial protein synthesis and metabolism of

deoxynivalenol and zearalenone in the rumen of dairy cows. J. Anim. Physiol. Anim.

Nutr. 89:303–315.

Danner, H., M. Holzer, E. Mayrhuber, and R. Braun. 2003. Acetic acid increases stability of

silage under aerobic conditions. Appl. Environ. Microbiol. 69:562–567.

Davis, M. A., D. D. Hancock, D. H. Rice, D. R. Call, R. Digiacomo, M. Samadpour, and T. E.

Besser. 2003. Feedstuffs as a vehicle of cattle exposure to Escherichia coli O157:H7 and

Salmonella enterica. Vet. Microbiol. 95:199–210.

De Wolf, E., G. Kuldau, P. Lipps, G. Munkvold, P. Vincelli, C. Woloshuk, and D. Mills. 2005.

Subject: Mouldy grains, mycotoxins and feeding problems. Accessed November 5, 2016.

http://www.oardc.ohiostate.edu/ohiofieldcropdisease/Mycotoxins/myco- pagedefault.htm.

Dean-Nystrom, E. A., L. J. Gansheroff, M. Mills, and A. D. O’Brien. 2002. Vaccination of

pregnant dams with intimin (O157) protects suckling piglets from Escherichia coli

O157:H7 infection. Infect. Immun. 70:2414–2418.

Degnan P. H., and H. Ochman. 2012. Illumina-based analysis of microbial community diversity.

The ISME Journal 6:183–194.

Deiner, U. L., R. J. Cole, T. H Sanders, G. A. Payne, L. S. Lee and M. A. Klich. 1987.

Epidemiology of aflatoxin formation by Aspergillus flavus. Ann. Rev. Phytopathology

25:240-270.

Dellaglio, F., and S. Torriani. 1986. DNA–DNA homology, physiological characteristics and

distribution of lactic acid bacteria from maize silage. J. Appl. Bacteriol. 60:83–93.

http://www.oardc.ohiostate.edu/ohiofieldcropdisease/Mycotoxins/myco-%20pagedefault.htm

181

Demetriou, J. A., P. A. Drewes, and J. B. Gin. 1974. Ceruloplasmin. Pages 857–865 in Clinical

Chemistry. C. Cannon and W. W. Winkelman, eds. Harper and Row, Hagerstown, Md.

Desnoyers, M., S. Giger-Reverdin, G. Bertin, C. Duvaux-Ponter, and D. Sauvant. 2009. Meta-

analysis of the influence of Saccharomyces cerevisiae supplementation on ruminal

parameters and milk production of ruminants. J. Dairy Sci. 92:1620–1632.

Devegowda, G.M., V. L. N. Raju, and H.V.L.N. Swamy. 1998. Mycotoxins: Novel solutions for

their counteraction. Feedstuffs 70:12-16.

Diaz, G. J., and H. J. Boermans. 1994. Fumonisin toxicosis in domestic animals: A review. Vet.

Human Toxicol. 36:548-555.

Diaz, D. E., W. M. Hagler, J. T. Blackwelder, J. A. Eve, B. A. Hopkins, K. L. Anderson, F. T.

Jones, and L. W. Whitlow. 2004. Aflatoxin binders II: Reduction of aflatoxin M1 in milk

by sequestering agents of cows consuming aflatoxin in feed. Mycopathol. 157:233–241.

Diaz, D. E., B. A. Hopkins, L. M. Leonard, W. M. Hagler, Jr., and L. W. Whitlow. 2000. Effect

of fumonisin on lactating dairy cattle. J. Dairy Sci. 83:1171. (Abstr.)

Diaz, D., and J. Trujillo.2013. Disease risks posed by poorly fermented silages. Accessed

October 10, 2016.

https://www.cvmbs.colostate.edu/ilm/proinfo/wdn/2007/August%20WDN.pdf

Di Costanzo, A., L. Johnston, H. Windels, and M. Murphy. 1995. A review of the effects of

molds and mycotoxins in ruminants. Prof. Anim. Scientist 12:138-150.

Diekman, D. A., and M. L. Green. 1992. Mycotoxins and reproduction in domestic livestock. J.

Anim. Sci. 70: 1615- 1627.

Diez-Gonzalez, F., T. R. Callaway, M. G. Kizoulis, and J. B. Russell. 1998. Grain feeding and

the dissemination of acid-resistant Escherichia coli from cattle. Sci. 281:1666-1668.

Dolci, P., E. Tabacco, L. Cocolin, and G. Borreani. 2001. Microbial dynamics during aerobic

exposure of corn silage stored under oxygen barrier or polyethylene films. Appl. Environ.

Microbiol. 77:7499–7507.

Dollear, F. G., G. E. Mann, L. P. Codifer, Jr., J. K. Gardner, Jr., S. P. Koltun, and H. L. E. Vix.

1968. Elimination of aflatoxins from peanut meal. J. Am. Oil Chem. Soc. 45:862–865

Dönmez, N., H. H. Dönmez, E. Keskin, and I. Kisadere. 2012. Effects of aflatoxin on some

haematological parameters and protective effectiveness of esterified glucomannan in

Merino rams. Sci. World Journal. ID 342468:4.

Doronina, N. V., Y. A. Trotsenko, B. B. Kuznetsov, T. P. Tourova, and M. S. Salkinoja-Salonen.

2002. Methylobacterium suomiense sp. nov. and Methylobacterium lusitanum sp. nov.,

aerobic, pink-pigmented, facultatively methylotrophic bacteria. Int. J. Syst. Evol.

Microbiol. 52:773–776.

https://www.cvmbs.colostate.edu/ilm/proinfo/wdn/2007/August%20WDN.pdf

182

Drasar, B. S., and P. A. Barrow. 1985. Intestinal Microbiology. In: (eds), Amer. Soc. Microbiol.

Press, Washington, DC. pp. 19-40.

Driehuis, F. 2013. Silage and the safety and quality of dairy foods: A review. Agric. Food Sci.

22:16–34.

Driehuis, F., M. C. Spanjer, J. M. Scholten, and M. C. Giffel. 2008. Occurrence of mycotoxins in

feedstuffs of dairy cows and estimation of total dietary intakes. J. Dairy Sci. 91:4261–

4271.

Driehuis, F., S. J. W. H. Oude Elferink, and P. G. Van Wikselaar. 2001. Fermentation

characteristics and aerobic stability of grass silage inoculated with Lactobacillus

buchneri, with or without homofermentative lactic acid bacteria. Grass Forage Sci.

56:330–343.

Duniere, L., A. Gleizal, F. Chaucheyras-Durand, I. Chevallier, and D. Thevenot-Sergentet. 2011.

Fate of Escherichia coli O26 in corn silage experimentally contaminated at ensiling, at

opening or after aerobic exposure and protective effect of various bacterial inoculants.

Appl. Environ. Microbiol. 77:8696–8704.

Duniere, L., L. Jin, B. Smiley, M. Qi, W. Rutherford, Y. Wang, and T. McAllister. 2015. Impact

of adding Saccharomyces strains on fermentation, aerobic stability, nutritive value, and

select lactobacilli populations in corn silage. J. Anim. Sci. 93:2322–35.

Dunn, J. R., J. E. Keen, and R. A. Thompson. 2004. Prevalence of Shiga-toxigenic Escherichia

coli O157:H7 in adult dairy cattle. J. Am. Vet. Med. Assoc. 224:1151–1158.

Dutton M. F. 1988. Enzymes and aflatoxin biosynthesis. Microbiol. Rev. 52:274–295.

Eckard, S., F. E. Wettstein, H. R. Forrer, and S. Vogelgsang. 2011. Incidence of Fusarium

species and mycotoxins in maize silage. Toxins 3:949–967.

Eckersall, P. D., F. P. Lawson, L. Bence, M. M. Waterston, T. L. Lang, W. Donachie, and M. C.

Fontaine. 2007. Acute phase protein response in an experimental model of ovine caseous

lymphadenitis. BMC Vet. Res. 3:35.

Edgar, R. C. 2010. Search and clustering orders of magnitude faster than BLAST. Bioinformatics

26:2460–2461.

Edrington, T. S., A. B. Sarr, L. F. Kubena, R. B. Harvey, and T. D. Phillips. 1996. Hydrated

sodium calcium aluminosilicate (HSCAS), acidic HSCAS, and activated charcoal reduce

urinary excretion of aflatoxin M1 in turkey poults. Lack of effect by activated charcoal on

aflatoxicosis. Toxicol. Lett. 89:115-122.

Edrington, T. S., L. F. Kubena, R. B. Harvey, and G. E. Rottinghous. 1997. Influence of a

superactivated charcoal on the toxic effects of aflatoxin or T-2 toxin in growing broilers.

Poult. Sci. 76:1205-1211.

https://www.ncbi.nlm.nih.gov/pubmed/?term=Rutherford%20W%5BAuthor%5D&cauthor=true&cauthor_uid=26020328

https://www.ncbi.nlm.nih.gov/pubmed/?term=Wang%20Y%5BAuthor%5D&cauthor=true&cauthor_uid=26020328

https://www.ncbi.nlm.nih.gov/pubmed/?term=McAllister%20T%5BAuthor%5D&cauthor=true&cauthor_uid=26020328

183

Edwards, S. G. 2004. Influence of agricultural practices on Fusarium infection of cereals and

subsequent contamination of grain by trichothecenes mycotoxins. Toxicol. Lett. 153, 29–

35.

EFSA (European Food Safety Authority). 2004. Opinion of the scientific panel on contaminants

in the food chain on a request from the Commission related to aflatoxin B1 as undesirable

substance in animal feed. EFSA J. 39:1–27.

Egal, S., A. Hounsa, Y. Y. Gong, P. C. Turner, C. P. Wild, A. J. Hall, K. Hell, and K. F.

Cardwell. 2005. Dietary exposure to aflatoxin from maize and groundnut in young

children from Benin and Togo, West Africa. Int. J. Food Microbiol. 104:215– 224.

Eikmeyer, F. G., P. Köfinger, A. Poschenel, S. Jünemannd, M. Zakrzewski, S. Heinl, E.

Mayrhuber, R. Grabherr, A. Pühler, H. Schwabb, A. Schlüter, P. H. Degnan, and

H. Ochman. Illumina-based analysis of microbial community diversity. The ISME J.

1:12.

Elam, N. A., J. F. Gleghorn, J. D. Rivera, M. L. Galyean, P. J. Defoor, M. M. Brashears, and S.

M. Younts-Dahl. 2003. Effects of live cultures of Lactobacillus acidophilus (strains

NP45 and NP51) and Propionibacterium freudenreichii on performance, carcass, and

intestinal characteristics, and Escherichia coli strain O157 shedding of finishing beef

steers. J. Anim. Sci. 81:2686-2698.

Ercolini, D. 2004. PCR-DGGE fingerprinting: novel strategies for detection of microbes in food.

J. Microbiol. Methods 56:297–314.

Eugui, A. M., S. Almquist, C. D. Muller, and A. C. Allison. 1991. Lymphocyte-selective

cytostatic and immunosuppressive effects of mycophenolic acid in vitro: role of

deoxyguanoside nucleotide depletion. Scand. J. Immunol. 33: 161–173.

European Commission. 2006. Commission recommendation of 17 August 2006 on the presence

of deoxynivalenol, zearalenone, ochratoxin A, T–2 and HT–2 and fumonisins in products

intended for animal feeding. Off. J. Eur. Union 229:7–8.

Farrokh, C., K. Jordan, F. Auvray, K. Glass, H. Oppegaard, S. Raynaud, D. Thevenot, R.

Condron, K. De Reu, A. Govaris, K. Heggum, M. Heyndrickx, J. Hummerjohann, D.

Lindsay, S. Miszczycha, S. Moussiegt, K. Verstraete, and O. Cerf. 2013. Review of

Shiga-toxin-producing Escherichia coli (STEC) and their significance in dairy

production. Int. J. Food Microbiol. 162:190–212.

FASS. 2010. Guide for the care and use of agricultural animals in research and teaching. 3rd ed.

Fed Anim. Sci. Soc., Champaign, IL.

FDA, 2014. FDA Approval of Listeria-specific Bacteriophage Preparation on Ready-to-Eat

(RTE) Meat and Poultry Products. Accessed 25th January, 2017.

http://www.fda.gov/food/ingredientspackaginglabeling/ucm083572.htm.

http://www.fda.gov/food/ingredientspackaginglabeling/ucm083572.htm

184

FDA. 2011. Mycotoxin Regulatory Guidance: A Guide for Grain Elevators, Feed Manufacturers,

Grain Processors and Exporters. Accessed August 20, 2016. https://www.ngfa.org/wp-

content/uploads/NGFAComplianceGuide-FDARegulatoryGuidanceforMycotoxins8-

2011.pdf

Felis, G. E., and Dellaglio F. 2007. Taxonomy of lactobacilli and bifidobacteria. Curr. Issues

Intest. Microbiol. 8:44–61.

Felis, G. E., R. Dellaglio, L. Mizzi, and S. Torriani. 2001. Comparative sequence analysis of a

recA gene fragment brings new evidence for a change in the taxonomy of the

Lactobacillus casei group. Int. J. Syst. Evol. Microbiol. 51:2113–2117.

Fenlon, D. R., and J. Wilson. 2000. Growth of Escherichia coli O157 in poorly fermented

laboratory silage: A possible environmental dimension in the epidemiology of E. coli

O157. Lett. Appl. Microbiol. 30:118–121.

Filya, I. 2003. The effect of Lactobacillus buchneri, with or without homofermentative lactic

acid bacteria, on the fermentation, aerobic stability and ruminal degradability of wheat,

sorghum and maize silages. J. Appl. Microbiol. 95:1080–1086.

Filya, I., R. E. Muck, and F. E. Contreras-Govea. 2007. Inoculant effects on alfalfa silage:

Fermentation profile products and nutritive value. J. Dairy Sci. 90:5108–5114.

Fink–Gremmels, J. 2008. Mycotoxins in cattle feeds and carry–over to dairy milk: A review.

Food Addit. Contam. 25:172–180.

Firmin, S., D. P. Morgavi, A. Yeannikouris, and H. Boudra. 2011. Effectiveness of modified

yeast cell wall extracts to reduce aflatoxin B1 absorption in dairy ewes. J. Dairy Sci.

94:5611–5619.

Fitzgerald, C. M., P. Camejo, J. Z. Oshlag, and D. R. Noguera. 2015. Ammonia-oxidizing

microbial communities in reactors with efficient nitrification at low-dissolved oxygen.

Water Res. 70:38–51.

Fox, J. T., D. U. Thomson, J. S. Drouillard, A. B. Thornton, D. T. Burkhardt, D. A. Emery, and

T. G. Nagaraja. 2009. Efficacy of Escherichia coli O157:H7 siderophore receptor/porin

proteins-based vaccine in feedlot cattle naturally shedding E. coli O157. Foodborne

Pathog. Dis. 6:893–899.

Freimund, S., M. Sauter, and P. Rys. 2003. Efficient adsorption of the mycotoxins zearalenone a

nd T2 toxin on a modified yeast glucan. J. Environ. Sci. Health 38:243255.

Frenzen, P. D., A. Drake, and F. J. Angulo. 2005. Economic cost of illness due to Escherichia

coli O157 infections in the United States. J. Food Prot. 68:2623–2630.

Frey, K. G., J. E. Herrera-Galeano, C. L. Redden, T. V. Luu, S. L. Servetas, A. J. Mateczun, V.

P. Mokashi, and K. A. Bishop-Lilly. 2014. Comparison of three next-generation

185

sequencing platforms for metagenomic sequencing and identification of pathogens in

blood, BMC Genomics 15:96-110.

Fruhauf, S., H. Schwartz, F. Ottner, R. Krska, and E. Vekirua. 2012. Yeast cell based feed

additives: Studies on aflatoxin B1 and zearalenone. Food Addit. Contam. Part A. Chem.

Anal. Control Expo. Risk. Assess. 29:217–231.

Fuhs, G. W, and M. Chen. 1975. Microbiological basis of phosphate removal in the activated

sludge process for treatment wastewater. Microbial Ecol. 2:119–138.

Fuller, R. 1989. A review: Probiotics in man and animals. J. Appl. Bacteriol. 66:365–378.

Fung, F., and R. F. Clark. 2004. Health effects of mycotoxins: a toxicological overview. Toxicol.

Clin. Toxicol. 42:217-234

Fusco, V., G. M. Quero, G. S. Cho, J. Kabisch, D. Meske, H. Neve, H. Bockelmann, and C. M.

A. P, Franz. 2015. The genus Weissella: Taxonomy, Ecology and Biotechnological

Potential. Front. Microbiol.6:155.

Gallo, A., T. Bertuzzi, G. Giuberti, M. Moschini, S. Bruschi, C. Cerioli, and F. Masoero. 2015.

New assessment based on the use of principal factor analysis to investigate corn silage

quality from nutritional traits, fermentation end products and mycotoxins. J. Sci. Food

Agric. 96:437-48.

Galvano, F., A. Pietri, T. Bertuzzi, G. Fusconi, M. Galvano, A. Piva, and G. Piva. 1996.

Reduction of carry over of aflatoxin from cow feed to milk by addition of activated

carbons. J. Food Prot. 59:551-554.

Garcia Orellana, M. 2012. Effect of supplementing essential fatty acids to prepartum Holstein

cows and preweaned calves on calf performance, metabolism, immunity, health and

hepatic gene expression. Ph.D. Dissertation. University of Florida.

Garon, D., E. Richard, L. Sage, V. Bouchart, D. Pottier, and P. Lebailly. 2006. Microflora and

multimycotoxin detection in corn silage: experimental study. Journal of Agriculture and

Food Chemistry 54:3479– 3484.

Garrett, W. N., H. Heitman, and A. N. Booth. 1968. Aflatoxin toxicity in beef cattle. Proc. Soc.

Exp. Biol. Med. 127:188–190.

Garvie, E. I. 1986. Genus Pediococcus. Pages 1075–1079 in Bergey's Manual of Systematic

Bacteriology. Vol. 2, 9th ed. Williams and Wilkins, Baltimore, MD.

Gaspar P., F. A. Al-Bayati, P. W. Andrew, A. R. Neves, and H. Yesilkaya. 2014. Lactate

dehydrogenase is the key enzyme for pneumococcal pyruvate metabolism and

pneumococcal survival in blood. Infect. Immun. 82:5099–5109.

186

Gelderblom, W. C. A., M. E. Cawood, S. D. Snyman, R. Vleggaar, and W. F. O. Marasas. 1993.

Structure-activity relationships of fumonisins in short-term carcinogenesis and

cytotoxicity studies. Food Chem. Toxicol. 31:407–414.

Gentry, P. A., M. L. Ross, and P. K. C. Chan. 1984. Effect of T-2 toxin on bovine hematological

and serum enzyme parameters. Vet. Hum. Toxicol. 26:24–28.

George J. W., J. Snipes, and V. M. Lane. 2010. Comparison of bovine hematology reference

intervals from 1957 to 2006. Vet. Clin. Pathol. 39:138–148.

Getachew, G., E. J. DePeters, W. Pittroff, D. H. Putnam, and A. M. Dandekar. 2006. Does

protein in alfalfa need protection from rumen microbes? Prof. Anim. Scientist 22:3.

Gloor, G. B., R. Hummelen, J. M. Macklaim, R. J. Dickson, A. D. Fernandes, R. MacPhee, and

G. Reid. 2010. Microbiome profiling by Illumina sequencing of combinatorial sequence-

tagged PCR products. PLoS One 5: e15406.

Gollop, N., V. Zakin, and Z. G. Weinberg. 2005. Antibacterial activity of lactic acid bacteria

included in inoculants for silage and in silages treated with these inoculants. J. Appl.


Gonzalez-Pereyra, M. L., V. A. Alonso, R. Sager, M. B. Morlaco, C. E. MAgnoli, A. L.

Astoreca, C. A. R. Rosa, S. M. Chiacchiera, M. Dalcero, and L. R. Cavaglieri. 2008.

Fungi and selected mycotoxins from pre- and post-fermented corn silage. J. Appl.

Microbiol. 104:1034-1041.

Gonzalez, F. H. D., F. Tecles, S. Martinez-Sublela, A. Tvarijonaviciute, L. Soler, and J. J. Ceron.

2008. Acute phase protein response in goats. J. Vet. Diagn. Invest. 20:580–584.

Goodson, M., and R. J. Rowbury. 1989. Habituation to normally lethal acidity by prior growth of

Escherichia coli at a sub-lethal acid pH value. Lett. Appl. Microbiol. 8:77–79.

Gorbacheva, I. V., and N. A. Rodinova. 1977. Studies on xylan degrading enzymes. II. Action

pattern of endo-1,4-xylanase from Aspergillus niger strain 14 on xylan and

xylooligosaccharides. Biochim. Biophys. Acta. 484:94–102.

Gourama, H., and L. B. Bullerman. 1995. Inhibition of growth and aflatoxin production of

Aspergillus flavus by Lactobacillus species. J. Food Prot. 58, 1249–1256.

Gowda N. K. S., H. V. L. N. Swamy, and P. Mahajan. 2013. Recent advances for control,

counteraction and ame lioration of potential aflatoxins in animal feeds. In: Aflatoxins:

Recent Advances and Future Prospects. Ed. Mehdi Razzaghi Abyaneh. InTech, pp. 129–

140.

Grada, A., and K. Weinbrecht. 2013. Next-generation sequencing: methodology and application.

J. Invest. Dermatol. 133:8-11.

187

Graf, K., A. Ulrich, C. Idler, and M. Klocke. 2016. Bacterial community dynamics during

ensiling of perennial ryegrass at two compaction levels monitored by terminal restriction

fragment length polymorphism. J. Appl. Microbiol. 120:1471-1491.

Gregory, J. L., L. Wang, and E. A. Johnson. 1995. Acid adaptation of Escherichia coli O157:H7

increases survival in acidic foods. Appl. Environ. Microbiol. 61:3752–3755.

Gregory, N. G., L. H. Jacobson, T. A. Nagle, R. W. Muirhead and G. J. Leroux. 2000. Effect of

preslaughter feeding system on weight loss, gut bacteria, and the physicochemical

properties of digesta in cattle. New Zealand J. Agric. Res. 43:351-361.

Grenier, B., and P. I. Oswald. 2012. Mycotoxin co-contamination of food and feed: Meta-

analysis of publications describing toxicological interactions. World Mycotoxin J. 4:285–

313.

Griffin, P. M. 1995. Escherichia coli O157:H7 and other enterohemorrhagic Escherichia coli.

Pages 739–761 in Infections of the Gastrointestinal Tract. M. J. Blaser, P. D. Smith, J. I.

Ravdin, H. B. Greenberg, and R. L. Guerrant, ed. Raven Press Ltd., New York.

Grim, R. E. 1968. Clay Mineralogy. McGraw-Hill, Inc., New York, 596 pages.

Gumprecht, L. A., A. Marcucci, R. F. Weigel, R. F. Vesonder, R. T. Riley, J. L. Showker, V. R.

Beasley, and W. M. Haschek. 1995. Effects of intravenous fumonisin B1 in rabbits:

Nephrotoxicity and sphingolipid alterations. Natural Toxins 3:395–403.

Guo, H., J. B. Callaway, and J. P. Ting. 2015. Inflammasomes: mechanism of action, role in disease,

and therapeutics. Nature Med. 21:677–687.

Guthrie, L. D., and D. M. Bedell. 1979. Effects of aflatoxin in corn on production and

reproduction in dairy cattle. Proc. Annu. Mtg. US Animal Health Assoc. 83:202.

Haggblom, P. 1990. Isolation of roquefortine C from feed grain. Appl. Environ. Microbiol.

56:2924–2926.

Hall, N. 2007. Advanced sequencing technologies and their wider impact in microbiology. J.

Exp. Biol. 210:1518-1525.

Hancock, D., T. Besser, J. Lejeune, M. Davis, and D. Rice. 2001. The control of VTEC in the

animal reservoir. Int. J. Food Microbiol. 66:71–78.

Hanshew, A. S., C. J. Mason, K. F. Raffa, and C. R. Currie. 2013. Minimization of chloroplast

contamination in 16S rRNA gene pyrosequencing of insect herbivore bacterial

communities. J. Microbiol. Methods 95:149-155.

Henderson, N. 1993. Silage additives. Anim. Feed Sci. Tech. 45:35-56.

Hodges, K., and G. Ravinder. 2010. Infectious diarrhea – cellular and molecular mechanisms.

Gut Microbes 1:4-21.

http://www.nature.com/nm/journal/v21/n7/full/nm.3893.html#auth-1



188

Hohler, D., K. H. Sudekum, S. Wolffram, A. A. Frolich, and R. R. Marquart. 1999. Metabolism

and excretion of ochratoxin A fed to sheep. J. Anim. Sci. 77:1217:1223.

Holzer, M., E. Mayrhuber, H. Danner, and R. Braun. 2003. The role of Lactobacillus buchneri in

forage preservation. Trends Biotechnol. 21:282–287.

Hoogenboom, L. A., J. Tulliez, J. P. Gautier, R. D. Coker, J. P. Malcion, M. J. Nagler, and T. H.

Polman. 2001. Absorption distribution and excretion of aflatoxin-derived ammoniation

products in lactating cows. Food Addi. Cont. 18:47– 58.

Hsu, C., Y. Chenc, P. Chao, C. Chena, L. Hsiehe, and S. Hu. 2008. Naturally occurring

chlorophyll derivatives inhibit aflatoxin B1-DNA adduct formation in hepatoma cells.

Mutat. Res. 657:98–104.

Hsu, I. C., C. B. Smalley, F. M. Strong, and W. E. Ribelin. 1972. Identification of T-2 toxin in

moldy corn associated with a lethal toxicosis in dairy cattle. Appl. Microbiol. 24:684-

690.

Hu, W., R. J. Schmidt, E. E. McDonell, C. M. Klingerman, and L. Kung, Jr. 2009. The effect of

Lactobacillus buchneri 40788 or Lactobacillus plantarum MTD-1 on the fermentation

and aerobic stability of corn silages ensiled at two dry matter contents. J. Dairy Sci.

92:3907–3914.

Huff, W. E., R. B. Kubena, R. H. Harvey, D. E. Corrier, and H. H. Mollenhaur. 1986.

Progression of aflatoxicosis in broiler chickens. Poult. Sci. 65:1891–1899.

Huff, W. E., L. F. Kubena, R. B. Harvey, and T. D. Phillips. 1992. Efficacy of hydrated sodium

calcium aluminosilicate to reduce the individual and combined toxicity of

aflatoxin and ochratoxin A. Poult. Sci. 71:6469.

Huisden, C. M., A. T. Adesogan, S. C. Kim, and T. Ososanya. 2009. Effect of applying molasses

or inoculants containing homofermentative or heterofermentative bacteria at two rates on

the fermentation and aerobic stability of corn silage. J. Dairy Sci. 92:690–697.

Hult, K., Teiling, A., and Gatenback, S., 1976. Degradation of ochratoxin A by the ruminant.

Appl. Environ. Microbiol. 32:443–444.

Huwig, A., S. Freimund, O. Kappeli, and H. Dutler. 2001. Mycotoxin detoxification of animal

feed by different adsorbents. Toxicol. Lett. 122:179–188.

IARC. 2002. Some Traditional Herbal Medicines, Some Mycotoxins, Naphthalene and Styrene.

IARC monographs on the evaluation of carcinogenic risks to humans Volume 82. Lyon:

IARC Scientific Publication, pp. 9-13.

Ibáñez-Vea, M., E. González-Peñas, E. Lizarraga, and A. López de Cerain. 2012. Co-occurrence

of aflatoxins, ochratoxin A and zearalenone in barley from a northern region of Spain.

Food Chem. 132:35–42.

189

Iglesias, C., A. Bach, M. Devant, C. Adelantado, and M. A. Calvo. 2005. The effect of

Lactobacillus buchneri inoculation on corn silages conservation. Pages 611–613 in Proc.

XI Jornadas sobre Producción Animal. Gobierno de Aragón, Servicio de Investigación

Agroalimentaria, Zaragoza, Spain.

Ingle, D. J., M. Tauschek, D. J. Edwards, D.M. Hocking, D. J. Pickard, and K. I. Azzopardi.

2016. Evolution of a typical enteropathogenic E. coli by repeated acquisition of LEE

pathogenicity island variants. Nat. Microbiol. 1:15010.

Jack, R. W., J. R. Tagg, and B. Ray. 1995. Bacteriocins of gram-positive bacteria. Microbiol.

Rev. 59:171–200.

Jackson, C. R., E. E. Roden, and P. F. Churchill. 2000. Denaturing gradient gel electrophoresis

can fail to separate 16S rDNA fragments with multiple base differences. Mol. Biol.

Today 1:49–51.

Jafari, A., M. M. Aslani, and S. Bouzari. 2013. Escherichia coli: a brief review of diarrheagenic

pathotypes and their role in diarrheal diseases in Iran. Iran J. Microbiol. 4:102-117.

Jafarian-Dehkordi, A., and N. Pourradi. 2013. Aflatoxin M1 contamination of human breast milk

in Isfahan, Iran. Adv. Biomedical Res. 2:86.

Jassim, S. A., and R. G. Limoges. 2014. Natural solution to antibiotic resistance: bacteriophages

the living drugs. World J. Microbiol. Biotechnol. 30:2153–2170.

Jean Pierre, J. 2007. Methods for preventing, decontaminating and minimizing the toxicity of

mycotoxins in feeds. Anim. Feed Sci. Tech. 137:342-362.

Jiang, Y., I. M. Ogunade, S. Qi, T. J. Hackmann, C. R. Staples, and A. T. Adesogan. 2017.

Effects of the dose and viability of Saccharomyces cerevisiae. 1. Diversity of ruminal

microbes as analyzed by Illumina MiSeq sequencing and quantitative PCR. J. Dairy

Sci. 100:325-342.

Johnson, L. M., J. H. Harrison, D. Davidson, W. C. Mahanna, K. Shinners, and D. Linder. 2002.

Corn silage management: Effects of maturity, inoculation, and mechanical processing on

pack density and aerobic stability. J. Dairy Sci. 85:434–444.

Johnson, M., E. Emmoth, A. C. Salomonsson, and A. Albihn. 2005. Potential risks when

spreading anaerobic digestion residues on grass silage crops: survival of bacteria, moulds

and viruses. Grass Forage Sci. 60: 175-185.

Jones B. A., R. D. Hatfield, and R. E. Muck. 1992. Effects of fermentation and bacterial

inoculation on lucerne cell walls. J. Sci. Food Agriculture 60:147–155.

Jones, B., and M. G. Kenward. 2015. Design and Analysis of Cross-Over Trials, Third Edition.

CRC Press, Taylor Francis Group. Boca Raton, FL.

190

Jones, R. K. 1981. Effect of nitrogen fertilizer, planting date, and harvest date on aflatoxin

production in corn inoculated withAspergillus flavus. Plant Disease 65:741–744.

Jouany, J. P. 2001. The impact of mycotoxins on performance and health of dairy cattle. In:

Alltech’s 17th Annual Symposium. Proceedings, pages 191–222.

Jouany, J. P. 2007. Methods for preventing, decontaminating and minimizing the toxicity of

mycotoxins in feeds. In: Morgavi, D.P., Riley, R.T. (Eds.), Fusarium and their toxins:

Mycology, occurrence, toxicity, control and economic impact. Anim. Feed Sci. Technol.

137:342–362.

Jouany, J. P., A. Yiannikouris, and G. Bertin. 2009. Risk assessment of mycotoxins in ruminants

and ruminant products. In: Papachristou, T.G., Parissi, Z.M., Ben Salem, H. and Morand-

Fehr, P. (eds.) Nutritional and foraging ecology of sheep and goats. CIHEAM/

FAO/NAGREF, Zaragoza, Spain, Options Méditerranéennes: Série A. Séminaires

Méditerranéens; n. 85, pp. 205-224.

Jubert, C., J. Mata, G. Bench, R. Dashwood, C. Pereira, and W. Tracewell. 2009. Effects of

chlorophyll and chlorophyllin on low-dose aflatoxin B1 pharmacokinetics in human

volunteers. Cancer Prevention Res. 2:1015–1022.

Kabak, B., A. D. W. Dobson, and I. Var. 2006. Strategies to prevent mycotoxin contamination of

food and animal feed: A review. Cr. Rev. Food Sci. 46:593–619.

Kaiser, A. G. 1999. Silage Additives. Department of Primar Industries, Wagga Wagga

Agricultural Institute, Wagga Wagga, NSW.

Kaiser, A. G., and M. J. Evans. 1997. Forage conservation on Australian dairy farms. Animal

Industry Rep. 3. NSW Agriculture, Wagga Wagga, Australia.

Kämpfer, P., and S. P. Glaeser. 2012. Prokaryotic taxonomy in the sequencing era–the

polyphasic approach revisited. Environ. Microbiol. 14:291-317.

Kang, T. W., A. T. Adesogan, S. C. Kim, and S. S. Lee. 2009. Effects of an esterase-producing

inoculant on fermentation, aerobic stability, and neutral detergent fiber digestibility of

corn silage. J. Dairy Sci. 92:732–738.

Kaper, J. B., J. P. Nataro, and H. L. Mobley. 2004. Pathogenic Escherichia coli. Nat. Rev.


Kateryna, K., and M. Rai. 2016. Antibiotic resistance: mechanisms and new antimicrobial

approaches. Academic Press.

Keen, J. E., and R. O. Elder. 2002. Isolation of Shiga-toxigenic Escherichia coli O157 from hide

surfaces and the oral cavity of finished beef feedlot cattle. J. Amer. Vet. Med. Assoc.

220:756-763

191

Keen, J. E., G. A. Uhlich, and R. O. Elder. 1999. Effects of hay- and grain-based diets on fecal

shedding in naturally acquired enterohemorrhagic E. coli (EHEC) O157 in beef feedlot

cattle. Procs. 80th Conf. Research Workers in Animal Diseases. Chicago, IL. (Abstr.).

Kegl, T., and A. Vanyi. 1991. T-2 fusariotoxicosis in a cattle stock. Magyar Allatorvosok Lapja

46:467-471.

Keller, L. A. M., M. L. González Pereyra, K. M. Keller, V. A. Alonso, A. A. Oliveira, T. X.

Almeida, T. S. Barbosa, L. M. T. Nunes, L. R. Cavaglieri, and C. A. R. Rosa. 2013.

Fungal and mycotoxins contamination in corn silage: Monitoring risk before and after

fermentation. J. Stored Prod. Res. 52:42–47.

Keller, L., L. Abrunhosa, K. Keller, C. A. Rosa, L. Cavaglieri, and A. Venâncio. 2015.

Zearalenone and Its derivatives, α-Zearalenol and β-Zearalenol, decontamination

by Saccharomyces cerevisiae strains isolated from bovine forage. Toxins. 7:3297-

3308.

Kellerman, T. S., W. F. O. Marasas, P. G. Thiel, W. C. A. Gelderblom, M. E. Cawood, and J. A.

W. Coetzer. 1990. Leukoencephalomalacia in two horses induced by oral dosing of

fumonisin B1. Onderstepoort J. Vet. Res. 57:269–275.

Kennedy, D. G., S. A. Hewitt, J.D. McEvoy, J. W. Currie, A. Cannavan, W. J. Blanchflower, and

C. T. Elliot. 1998. Zeralenol is formed from Fusarium spp. toxins in cattle in vivo. Food

Addit. Contam. 15:393–400.

Kenney, N. M., E. S. Vanzant, D. L. Harmon, and K. R. McLeod. 2015. Direct-fed microbials

containing lactate-producing bacteria influence ruminal fermentation but not lactate

utilization in steers fed a high-concentrate diet. J. Anim. Sci. 93:2336–2348.

Kiessling K. H., H. Pettersson, K. Sandholm, and M. Olsen. 1984. Metabolism of aflatoxin,

ochratoxin, zearalenone, and trichothecenes by intact rumen fluid, rumen protozoa, and

rumen bacteria. Appl. Environ. Microbiol. 47:1070–1073.

Kim, E. K., C. M. Maragos, and D. F. Kendra. 2004. Liquid chromatographic determination of

fumonisins B1, B2, and B3 in corn silage. J. Agric. Food Chem. 52:196–200.

Kirk, J. L., L. A. Beaudette, M. Hart, P. Moutoglis, J. N. Klironomos, H. Lee, and J. T. Trevors.

2004. Methods of studying soil microbial diversity. J. Microbiol. Methods. 58:169–188.

Kissell, L., S. Davison, B. A. Hopkins, G. W. Smith, and L. W. Whitlow. 2013. Effects of

experimental feed additives on aflatoxin in milk of dairy cows fed aflatoxin-contaminated

diets. J. Anim. Physiol. Anim. Nutr. 97:694–700.

Kizilsimsek, M., R. J. Schmidt, and L. Kung Jr. 2007. Effects of a mixture of lactic acid bacteria

applied as a freeze-died or fresh culture on the fermentation of alfalfa silage. J. Dairy Sci.

90:5698–5705.

192

Kleinschmit, D. H., and L. Kung, Jr. 2006a. A meta-analysis of the effects of Lactobacillus

buchneri on the fermentation and aerobic stability of corn, grass and small grain silages.

J. Dairy Sci. 89:4005–4013.

Kleinschmit, D. H., and Kung, Jr., L. 2006b. The effects of Lactobacillus buchneri 40788

and Pediococcus pentosaceus R1094 on the fermentation of corn silage. J. Dairy

Sci. 89:3999–4004.

Kleinschmit, D. H., R. J. Schmidt, and L. Kung, Jr. 2005. The effects of various antifungal

additives on the fermentation and aerobic stability of corn silage. J. Dairy Sci. 88:2130–

2139.

Klieve, A. V., and T. Bauchop. 1988. Morphological diversity of ruminal bacteriophages from

sheep and cattle. Appl. Environ. Microbiol. 54:1637–1641.

Klindworth A., E. Pruesse, T. Schweer, J. Peplies, C. Quast, M. Horn, and F. O. Glockner. 2013.

Evaluation of general 16S ribosomal RNA gene PCR primers for classical and next-

generation sequencing-based diversity studies. Nucleic Acids Res. 41:1.

Knief, C., N. Delmotte, S. Chaffron, M. Stark, G. Innerebner, R. Wassmann, C. von Mering, and

J. A. Vorholt. 2012. Metaproteogenomic analysis of microbial communities in the

phyllosphere and rhizosphere of rice. ISME J. 6:1378 –1390.

Köhl, J., J. Postma, P. Nicot, M. Ruocco and B. Blum, 2011. Stepwise screening of

microorganisms for commercial use in biological control of plant-pathogenic fungi and

bacteria. Biol. Control 57:1‒12.

Konowalchuk J., J. I. Speirs, and S. Stavric. 1977. Vero response to a cytotoxin of Escherichia

coli. Infect. Immun. 18:775–779.

Korosteleva, S. N., T. K. Smith, and H. J. Boermans. 2009. Effects of feed naturally

contaminated with Fusarium mycotoxins on metabolism and immunity of dairy cows. J

Dairy Sci 92:1585-1593.

Kraut-Cohen, J., V. Tripathi, Y. Chen, J. Gatica, V. Volchinski, S. Sela, Z. Weinberg, and E.

Cytryn. 2016. Temporal and spatial assessment of microbial communities in commercial

silages from bunker silos. Appl. Microbiol. Biotechnol. 100:6827-6835.

Krehbiel, C. R., S. R. Rust, G. Zhang, and S. E. Gilliland. 2003. Bacterial direct-fed microbials

in ruminant diets: performance response and mode of action. J. Anim. Sci. 81:120–132.

Kristensen, N. B., K. H. Sloth, O. Højberg, N. H. Spliid, C. Jensen, and R. Thøgersen. 2010.

Effects of microbial inoculants on corn silage fermentation, microbial contents, aerobic

stability, and milk production under field conditions. J. Dairy Sci. 93:3764–3774.

Krooneman, J., F. Faber, A. C. Alderkamp, S. J. H. W. Oude Elferink, F. Driehuis, I.

Cleenwerck, J. Swings, J. C. Gottschal, and M. Vancanneyt. 2002. Lactobacillus

193

diolivorans sp. Nov., a 1,2-propanediol-degrading bacterium isolated from aerobically

stable maize silage. Int. J. Syst. Evol. Microbiol. 52:639–646.

Kudva, I. T., P. G. Hatfield, and C. J. Hovde. 1997. Characterization of Escherichia coli

O157:H7 and other shiga toxin-producing E. coli serotypes isolated from sheep. J. Clin.

Microbiol. 35:892-899.

Kuilman, M. E. M., R. F. M. Maas, D. J. Judah, and J. Gremmels. 1998. Bovine hepatic

metabolism of aflatoxin B1. J. Agric. Food. Chem. 46:2707-2713.

Kung L., Jr. 2010. Aerobic stability of silage. In: Proceedings of California Alfalfa and Forage

Symposium and Crop/Cereal Conference, Visalia, CA, USA. Accessed July 1, 2016.

http:// alfalfa.ucdavis.edu/+symposium/proceedings/2010/10- 89.pdf.

Kung L., Jr. 2010. Understanding the biology of silage preservation to maximize quality and

protect the environment. Pages 41-54 in Proc. Alfalfa & Forage Symposium, Visalia, CA.

Kung, L., Jr. 1998. A review on silage additives and enzymes. Pages 121–135 in Proc. 59th

Minneapolis Nutrition Conference, Minneapolis, MN.

Kung, L., Jr., D. B. Grieve, J. W. Thomas, and J. T. Huber. 1984. Added ammonia or microbial

inoculant for fermentation and nitrogenous compounds of alfalfa ensiled at various

percentages of dry matter. J. Dairy Sci. 67:299–306.

Kung, L., Jr., and R. Shaver. 2001. Interpretation and use of silage fermentation analysis reports.

Focus on Forage 3:1–5.

Kung, L., Jr., J. R. Robinson, N. K. Ranjit, J. H. Chen, C. M. Golt, and J. D. Pesek. 2000.

Microbial populations, fermentation end-products, and aerobic stability of corn silage

treated with ammonia or a propionic acid-based preservative. J. Dairy Sci. 83:1479–1486.

Kung, L., Jr., L. D. Satter, B. A. Jones, K. W. Genin, A. L. Sudoma, G. L. Enders, and H. S.

Kim. 1987. Microbial inoculation of low moisture alfalfa silage. J. Dairy Sci. 70:2069–

2077.

Kung, L., Jr., A. C. Sheperd, A. M. Smagala, K. M. Endres, C. A. Bessett, N. K. Ranjit, and J. L.

Glancey. 1998. The effect of preservatives based on propionic acid on the fermentation

and aerobic stability of corn silage and a total mixed ration. J. Dairy Sci. 81:1322–1330.

Kung, L., Jr., C. C. Taylor, M. P. Lynch, and J. M. Neylon. 2003. The effect of treating alfalfa

with Lactobacillus buchneri 40788 on silage fermentation, aerobic stability, and nutritive

value for lactating dairy cows. J. Dairy Sci. 86:336–343.

Kurtzman, C. P., B. W. Horn, and C. W. Hesseltine. 1987. Aspergillus nomius, a new aflatoxin-

producing species related to Aspergillus flavus and Aspergillus tamarii. Antonie van

Leeuwenhoek 53:147-158.

194

Kutz, R. E., J. D. Sampson, L. B. Pompeu, D. R. Ledoux, J. N. Spain, M. Vazquez-Anon, and G.

E. Rottinghaus. 2009. Efficacy of Solis, NovasilPlus, and MTB-100 to reduce aflatoxin

M1 levels in milk of early to mid-lactation dairy cows fed aflatoxin B1. J. Dairy Sci.

92:3959–3963.

Lahti, E., O. Ruoho, L. Rantala, M. L. Hanninen, and T. Honkanen-Buzalski. 2003. Longitudinal

study of Escherichia coli O157 in a cattle finishing unit. Appl. Environ. Microbiol.

69:554–561.

Lambert, R. J., and M. Stratford. 1999. Weak-acid preservatives: modelling microbial inhibition

and response. J. Appl. Microbiology 86:157–164.

Lee, M. S., H. Kwon, E. Y. Lee, D. J. Kim, J. H. Park, V. L. Tesh, T. K. Oh, M. H. Kim.

2015. Shiga toxins activate the NLRP3 inflammasome pathway to promote both

production of the proinflammatory cytokine interleukin-1β and apoptotic cell

death. Infect. Immun. 84:172–186.

LeJeune, J. T., and A. N. Wetzel. 2007. Preharvest control of Escherichia coli O157 in cattle. J.

Anim. Sci. 85:73–80.

Lema, M., L. Williams, and D. R. Rao. 2001. Reduction of fecal shedding of enterohemorrhagic

Escherichia coli O157:H7 in lambs by feeding microbial feed supplement. Small Rum.

Res. 39:31-39.

Lepom, P., O. Knabe, and H. Baath. 1990. Occurrence of Fusarium strains and their mycotoxins

in corn silage. 7. Formation of deoxynivalenol in a silage corn plot artificially inoculated

with Fusarium culmorum and the effect of silaging on the stability of the DON formed.

Arch. Tierernahr. 40:1005-1012.

Li, W., L. Fu, B. Niu, S. Wu, and J. Wooley. 2012. Ultrafast clustering algorithms for

metagenomic sequence analysis. Brief Bioinform. 13:656.

Li, Y., and N. Nishino. 2011. Effects of inoculation of Lactobacillus rhamnosus and

Lactobacillus buchneri on fermentation, aerobic stability and microbial communities in

whole crop corn silage. J. Jpn. Grassl. Sci. 57:184–191.

Lim, J. Y., J. Li, H. Sheng, T. E. Besser, K. Potter, and C. J. Hovde. 2007. Escherichia coli

O157:H7 colonization at the rectoanal junction of long-duration culture-positive cattle.

Appl. Environ. Microbiol. 73:1380-1382.

Lin, C., K. K. Bolsen, B. E. Brent, and D. Y. C. Fung. 1992. Epiphytic lactic acid bacteria

succession during the pre-ensiling and ensiling periods of alfalfa and maize. J. Appl.

Bact. 73:373–387.

Littell, R. C., G. A. Milliken, W. W. Stroup, R. D. Wolfinger, and O. Schabenberber. 2006. SAS

for Mixed Models. Second edition. SAS Inst., Inc., Cary, NC.

195

Liu, Y., and F. Wu. 2010. Global burden of aflatoxin-induced hepatocellular carcinoma: a risk

assessment. Environ. Health Perspect. 118:818–824.

Liu, Y., K. Liu, X. Yu, B. Li, and B. Cao. 2014. Identification and control of a Pseudomonas spp

(P. fulva and P. putida) bloodstream infection outbreak in a teaching hospital in Beijing,

China. Int. J. Infect. Dis. 23:105–108.

Loman N. J., C. Constantinidou, J. Z. Chan, M. Halachev, M. Sergeant, C. W. Penn, E. R.

Robinson, and M. J. Pallen. 2012. High-throughput bacterial genome sequencing: an

embarrassment of choice, a world of opportunity. Nat. Rev. Microbiol. 10:599-606.

Magoc, T., and S. L. Salzberg. 2011. FLASH: Fast Length Adjustment of Short Reads to

Improve Genome Assemblies. Bioinform. 27:2957–2963.

Maki C. R., A. D. Thomas, S. E. Elmore, A. A. Romoser, R. B. Harvey, H. A. Ramirez-Ramirez,

and T. D. Phillips. 2016. Effects of calcium montmorillonite clay and aflatoxin exposure

on dry matter intake, milk production, and milk composition. J. Dairy Sci. 99:1039–1046.

Makimura, S., and N. Suzuki. 1982. Quantitative determination of bovine serum haptoglobin and

its elevation in some inflammatory diseases. Jpn. J. Vet. Sci. 44:15–21.

Malekinejad, H., R. Maas-Bakker, and J. Fink-Gremmels. 2006. Species differences in the

hepatic biotransformation of zearalenone. The Vet. Journal 172:96–102.

Mansfield, M. A., A. D. Jones, and G. A. Kuldau, 2008. Contamination of fresh and ensiled

maize by multiple Penicillium mycotoxins. Phytopathology 98:330–336.

Mansfield, M. A., D. D. Archibald, A. D. Jones, and G. A. Kuldau. 2007. Relationship of

spinganine analog mycotoxin contamination in maize silage to seasonal weather

conditions and to agronomic and ensiling practices. Phytopathology 97:504-511.

Mansfield, M. A., E. D. De Wolf, and G. A. Kuldau. 2005. Relationships Between Weather

Conditions, Agronomic Practices, and Fermentation Characteristics with Deoxynivalenol

Content in Fresh and Ensiled Maize. Plant Dis. 89:1151–1157.

Marasas, W. F. O., 1996. Fumonisins: History, world-wide occurrence and impact. Pp. 1–17. In

L. S. Jackson, J. W. DeVries, and L. B. Bullerman (Eds.). Fumonisins in Food. Plenum

Press, New York.

March, S. B., and S. Ratnam. 1989. Latex agglutination test for detection of Escherichia coli

serotype O157. J. Clin. Microbiol. 27:1675–7.

Marczuk, J., K. Obremski, K. Lutnicki, M. Gajecka, and M. Gajecki. 2012. Zearalenone and

deoxynivalenol mycotoxicosis in dairy cattle herds. Pol. J. of Vet. Sci. 15: 365-372.

Mari, L. J., R. J. Schmidt, L. G. Nussio, C. M. Hallada, and L. Kung. 2009. Short

communication: An evaluation of the effectiveness of Lactobacillus buchneri 40788 to

196

alter fermentation and improve the aerobic stability of corn silage in farm silos. J. Dairy

Sci. 92:1174-1176.

Martinez, N., F. S. Lima, R. S. Bisinotto, L. F. Greco, E. S. Ribeiro, F. Maunsell, K. N. Galvão,

C. A. Risco, and J. E. P. Santos. 2012. Evaluation of peripartal calcium status, energetic

profile, and neutrophil function in dairy cows at low or high risk of developing uterine

disease. J. Dairy Sci. 95:7158–7172.

Masimango, N., J. Remacle, and J. Ramaut. 1979. Elimination, par des argiles gonflantes de

L’aflatoxine B1 des milieus contamines. Ann. Nutr. Alim. 33:137–147.

Mathur, S., P. D. Constable, R. M. Eppley, A. L. Waggoner, M. E. Tumbleson, and W. M.

Haschek. 2001. Fumonisin B1 is hepatotoxic and nephrotoxic in milk–fed calves.

Toxicol. Sci. 60:385–396.

Matthews L., J. C. Low, D. L. Gally, M. C. Pearce, D. J. Mellor, J. A. P. Heesterbeek, M.

Chase-Topping, S. W. Naylor, D. J. Shaw, S. W. J. Reid, G. J. Gunn, M. E. J.

Woolhouse. 2006a. Heterogenous shedding of Escherichia coli O157 in cattle and its

implications for control. Proc Natl. Acad. Sci. USA 103: 547–552.

Matthews L., I. J. McKendrick, H. Ternent, G. J. Gunn, B. Synge, and M. E. J. Woolhouse.

2006b. Super-shedding cattle and the transmission dynamics of Escherichia coli O157.

Epidemiol. Infect. 134:131–142.

Matthews, L., R. Reeve, D. L. Gally, J. C. Low, M. E. Woolhouse, S. P. McAteer, M. E.

Locking, M. E. Chase-Topping, D. T. Haydon, L. J. Allison, M. F. Hanson, G. J. Gunn,

and S. W. Reid. 2013. Predicting the public health benefit of vaccinating cattle against

Escherichia coli O157. Proc. Natl. Acad. Sci. U.S.A. 110:16265–16270.

McAllister, T. A., K. A. Beauchemin, A. Y. Alazzeh, J. Baah, R. M. Teather, and K. Stanford.

2011. Review: The use of direct fed microbials to mitigate pathogens and enhance

production in cattle. Can. J. Anim. Sci. 91:193–211.

McDonald P., A. R. Henderson, and S. J. E. Heron. 1991. The Biochemistry of Silage. 2nd

edition. Chalcombe Publications, Marlow, Bucks, UK.

McEniry, J., P. O’Kiely, N. J. W. Clipson, P. D. Forristal, and E. M. Doyle. 2008. Bacterial

community dynamics during ensilage of wilted grass. J. of Appl. Microbiol. 105:359-371.

McGarvey, J. A., R. B. Franco, J. D. Palumbo, R. Hnasko, L. Stanker, and F. M. Mitloehner.

2013. Bacterial population dynamics during the ensiling of Medicago sativa (alfalfa) and

subsequent exposure to air. J. Appl. Microbiol. 114:1661–1670.

McKenzie, K. S. 1997. Degradation and detoxification of common chemical contaminants of

food and water using ozone generated by electrolysis. [Dissertation]. College Station,

TX: Texas A & M University. 200 p.

197

Mead, P. S., L. Slutsker, V. Dietz, L. F. McCaig, J. S. Bresee, C. Shapiro, P. M. Griffin, and R.

V. Tauxe. 1999. Food-related illness and death in the United States. Emerg. Infect. Dis.

5:607–625.

Milicevic, D. R., M. Skrinjar, and T. Baltic. 2010. Real and perceived risks for mycotoxin

contamination in foods and feeds: Challenges for food safety control. Toxins 2:572–592.

Miller, J. D., and Trenholm, H. L., 1994. Mycotoxins In Grain: Compounds Other Than

Aflatoxin, 1st Ed. Eagan Press, 3340 Pilot Knob Rd., St. Paul, MN pp. 302-303.

Mills, D. K., J. A. Entry, and P. M. Gillevet. 2007. Assessing microbial community diversity

using amplicon length heterogeneity polymerase chain reaction. Soil Sci. Soc. Am. J.

71:572–578.

Min, B. R., W. E. Pinchak, R. C. Anderson, and T. R. Callaway. 2007. Effect of tannins on the in

vitro growth of Escherichia coli O157:H7 and in vivo growth of generic Escherichia coli

excreted from steers. J. Food Prot. 70:543-550.

Mir, R. A., T. A. Weppelmann, M. Kang, T. M. Bliss, N. DiLorenzo, G. C. Lamb, S. Ahn, and

K. C. Jeong. 2015. Association between animal age and the prevalence of Shiga toxin-

producing Escherichia coli in a cohort of beef cattle. Vet. Microbiol. 175:325–331.

Mohiuddin, S. M., M. V. Reddy, M. M. Reddy, and K. Ramakrishna. 1986. Studies on

phagocytic activity and haematological changes in aflatoxicosis in poultry. Indian Vet. J.

63:442–445.

Mohr, A. I., I. Lorentz, B. Baum, M. Hewicker-Trautwein, M. Pfaffl, A. Dzidic, H. H. D. Meyer,

J. Bauer, and K. Meyer. 2007. Influence of oral application of mycophenolic acid on the

clinical health status of sheep. J. Vet. Med. A. 54:76-81.

Moon, N. J. 1983. Inhibition of the growth of acid tolerant yeasts by acetate, lactate and

propionate and their synergistic mixtures. J. Appl. Bacteriol. 55:454–460.

Moschini, M., A. Gallo, G. Piva, and F. Masoeo. 2008. The effects of rumen fluid on in vitro

aflatoxin binding capacity of different sequestering agents and in vivo release of

sequestered toxin. Anim. Feed Sci. Technol. 147:292–309.

Muck R. E., and B. J. Holmes. 2000. Factors affecting bunker silo densities. Appl. Eng. in Agric.

16:613-619.

Muck, R. E. 1990. Dry matter level effects on alfalfa silage quality II. Fermentation products and

starch hydrolysis. Trans. ASAE. 33:373–381.

Muck, R. E. 2008. Improving alfalfa silage quality with inoculants and silo management. Pages

137–146 in Proc. Cornell Nutr. Conf. East Syracuse, NY. Cornell University, Ithaca, NY.

Muck, R. E. 2010. Silage microbiology and its control through additives. R. Bras. Zootec.

39:183–191.

198

Muck, R. E. 2013. Recent advances in silage microbiology. Agric. Food Sci. 22:3–15.

Muck, R. E., 1998. Preparing high quality alfalfa silage. pp 1-11 in Int. Seminar of Conservation

of High Quality Flora, Los Angeles, Chile.

Muck, R. E., and J. T. Dickerson. 1988. Storage temperature effects on proteolysis in alfalfa

silage. Trans. ASAE 31:1005–1009

Muck, R. E., and L. Kung, Jr. 1997. Effects of silage additives on ensiling. In Silage: Field to

Feedbunk, NRAES-99, pp. 187-199. Ithaca, NY: Northeast Regional Agric. Engng.

Service.

Muhlbach, P. 1998. Potential for silage production in Brazil. Pages 135–144 in Proc.

Biotechnology in the Feed Industry Symposium. T. P. Lyons and K. A. Jacques (ed.),

Nottingham University Press, Nottingham, UK.

Munkvold, G. P., 2014. Crop management practices to minimize the risk of mycotoxins

contamination in temperate-zone maize. Mycotoxin Reduction in Grain Chains, 59-75.

Munns, K. D., L. B. Selinger, K. Stanford, L. Guan, T. R. Callaway, and T. A. McAllister. 2015.

Perspectives on super-shedding of Escherichia coli O157: H7 by cattle. Foodborne

Pathog. Dis. 12:89-103.

Muyzer, G., E. C. D. Waal, and A. G. Uitterlinden. 1993. Profiling of complex microbial

populations by denaturing gradient gel electrophoresis analysis of polymerase chain

reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol. 59:695–700.

Nagahata, H., K. Taguchi, and H. Noda. 1989. Preliminary studies on the acid soluble

glycoproteins in serum and their diagnostic value for acute inflammatory disease in

cattle. Vet. Res. Comm. 13: 257–263.

Nagel, S., and G. A. Broderick. 1992. Effect of formic acid or formaldehyde treatment of alfalfa

silage on nutrient utilization by dairy cows. J. Dairy Sci. 75:140–154.

Nakajima T., S. Watanabe, Y. Sato, T. Kameya, T. Hirota, and Y. Shimosato. 1982. An

immunoperoxidase study of S-100 protein distribution in normal and neoplastic tissues.

Am. J. Surg. Pathol. 6:715–727.

Nataro, J. P., and J. B. Kaper. 1998. Diarrheagenic Escherichia coli. Clin. Microbiol. Rev.

11:142-201.

Nataro, J. P., J. P. Kaper, R. Robins-Browne, V. Prado, P. Vial, and M. M. Levine (1987).

Patterns of adherence of diarrheagenic Escherichia coli to HEp-2 cells. Pediatr. Infect.

Dis. J. 6:829–831.

National Toxicology Program. 1999. Toxicology and carcinogenesis studies of fumonisin B1

(CAS NO. 116355-83-0) in F344/ N Rats and B6C3F1 Mice. NTP Technical Report 496.

199

National Institutes of Health Publication No. 99-3955, Research Triangle Park, North

Carolina.

Ndagano D., T. Lamoureux, C. Dortu, S. Vandermoten, and P. Thonart. 2011. Antifungal

activity of 2 lactic acid bacteria of the Weissella genus isolated from food. J. Food

Sci. 76:305–311.

Nelson, K. E., C. Weinel, I. T. Paulsen, R. J. Dodson, H. Hilbert, V. A. Martins dos Santos, D. E.

Fouts, S. R. Gill, M. Pop, M. Holmes, L. Brinkac, M. Beanan, R. T. DeBoy, S.

Daugherty, J. Kolonay, R. Madupu, W. Nelson, O. White, J. Peterson, H. Khouri, I.

Hance, P. C. Lee, E. Holtzapple, D. Scanlan, K. Tran, A. Moazzez, T. Utterback, M.

Rizzo, K. Lee, D. Kosack, D. Moestl, H. Wedler, J. Lauber, D. Stjepandic, J. Hoheisel,

M. Straetz, S. Heim, C. Kiewitz, J. A. Eisen, K. N. Timmis, A. Dusterhoft, B. Tummler,

and C. M. Fraser. 2002. Complete genome sequence and comparative analysis of the

metabolically versatile Pseudomonas putida KT2440. Environ. Microbiol. 4:799–808.

Nevarez, L., V. Vasseur, A. Le Madec, M. A. Le Bras, L. Coroller, and I. Leguérine, and G.

Barbier. 2009. Physiological traits of Penicillium glabrum strain LCP 08.5568, a

filamentous fungus isolated from bottled aromatised mineral water. Int. J. Food

Microbiol. 130:166-171.

Ni, K., T. T. Minh, T. T. M. Tu, T. Tsuruta, H. Pang, and N. Nishino. 2016. Comparative

microbiota assessment of wilted Italian ryegrass, whole crop corn, and wilted alfalfa

silage using denaturing gradient gel electrophoresis and next-generation sequencing.

Appl. Microbiol. Biotechnol. 101:1-10.

Niu Y. D., R. P. Johnson, Y. Xu, T. A. McAllister, R. Sharma, M. Louie, K. Stanford. 2009.

Host range and lytic capability of four bacteriophages against bovine and clinical human

isolates of Shiga toxin-producing Escherichia coli O157:H7. J. Appl. Microbiol.

107:646–656.

Nkosi, B.D., R. Meeske, H. J. Van der Merwe, and I. B. Groenewald. 2010. Effects of

homofermentative and heterofermentative bacterial silage inoculants on potato hash

silage fermentation and digestibility in rams. Anim. Feed Sci. Technol. 157:195–200.

Noel, R. J., and L. G. Hambleton. 1976. Collaborative study of a semi-automated method for

determination of crude protein in animal feeds. J. Assoc. Off. Anal. Chem. 59:134–140.

NRC. 2001. Nutrient Requirements of Dairy Cattle. 7th rev. ed. Natl. Acad. Press, Washington,

DC.

Nsereko, V. L., B. K. Smiley, W. M. Rutherford, A. Spielbauer, K. J. Forrester, G. H. Hettinger,

E. K. Harman, and B. R. Harman. 2008. Influence of inoculating forage with lactic acid

bacterial strains that produce ferulate esterase on ensilage and ruminal degradation of

fiber. Anim. Feed Sci. Technol. 145:122–135.

O’Brien, M., K. F. Nielsen, P. O’Kiely, P. D. Forristal, H. T. Fuller, and J. C. Frisvad. 2006.

Mycotoxins and other secondary metabolites produced in vitro by Penicillium paneum

200

Frisvad and Penicillium roqueforti Thom isolated from baled grass silage in Ireland. J.

Agric. Food Chem. 54:9268–9276.

Obrig, T. G., C. B. Louise, C. A. Lingwood, B. Boyd, L. Barley-Maloney, and T. O. Daniel.

1993. Endothelial heterogeneity in Shiga toxin receptors and responses. J. Biol.

Chem. 268:15484-15488.

Ogiy, S., Y. Chen, R. Pasvolsky, Z.G. Weinberg, and M. Moshe. 2015. High Resolution Melt

analysis to confirm the establishment of Lactobacillus plantarum and Enterococcus

faecium from silage inoculants during ensiling of wheat. Grassland Sci. 62: 29-36.

Ohya, T., M. Akiba, and H. Ito. 2001. Use of a trial probiotic product in calves experimentally

infected with Escherichia coli O157. Japan Agric. Res. Quart. 35:189-194.

Oldenburg, E., and F. Eiiner. 2005. Fusarium mycotoxins in forage maize—Detection and

evaluation. Mycotoxin Res. 21:105–107.

Oliveira A. S, Z. G. Weinberg, I. M. Ogunade, A. A. P. Cervantes, K. G. Arriola, Y. Jiang, D. H.

Kim, X. Li, M. C. M. Gonçalves, D. Vyas, A. T. Adesogan. 2017. Meta-analysis of

effects of homofermentative and facultative heterofermentative lactic acid bacteria

inoculation on silage fermentation and aerobic stability and the performance of dairy

cows. J. Dairy Sci. (In press). https://doi.org/10.3168/jds.2016-11815.

Omisakin F., M. MacRae, I. D. Ogden, and N. J. C. Strachan. 2003. Concentration and

prevalence of Escherichia coli O157 in cattle feces at slaughter. Appl. Environ.

Microbiol. 69:2444–2447.

Osweiler, G. D., M. E. Kehrli, J. R. Stabel, J. R. Thurston, P. F. Ross, and T. M. Wilson. 1993.

Effects of fumonisin–contaminated corn screenings on growth and health of feeder

calves. J. Anim. Sci. 71:459–466.

Oude Elferink, S. J. W. H., J. Krooneman, J. C. Gottschal, S. F. Spoelstra, F. Faber, and F.

Driehuis. 2001. Anaerobic conversion of lactic acid to acetic acid and 1, 2-propanediol by

Lactobacillus buchneri. Appl. Environ. Microbiol. 67:125–132.

Pahlow, G., C. Rammer, D. Slottner, and M. Tuori. 2001. Ensiling of legumes. Pages 27-30 in

Proc. of Legume silages for animal production. R. J. Wilkins, C. Paul, ed. LEGSIL,

Braunschweig.

Pahlow, G., R. E. Muck, F. Driehuis, S. J. W. H. Oude Elferink, and S. F. Spoelstra. 2003.

Microbiology of ensiling. Pages 31–93 in Silage Science and Technology. D. R. Buxton,

R. E. Muck, and J. H. Harrison, ed. American Society of Agronomy, Inc., Crop Science

Society of America, Inc., Soil Science Society of America, Inc. Publications, Madison,

WI.

Pang, H., G. Qin, Z. Tan, Z. Li, Y. Wang, and Y. Cai. 2011. Natural populations of lactic acid

bacteria associated with silage fermentation as determined by phenotype, 16S ribosomal

RNA and recA gene analysis. Syst. Appl. Microbiol. 34:235–241.

201

Pantaya, D., D. P. Morgavi, M. Silberberg, F. Chaucheyras-Durand, C. Martin, K. G. Wiryawan,

and H. Boudra. 2016. Bioavailability of aflatoxin B1 and ochratoxin A, but not fumonisin

B1 or deoxynivalenol, is increased in starch-induced low ruminal pH in nonlactating dairy

cows. J. Dairy Sci. 99:9759-9767.

Park, D. L., L. S. Lee, and S. A. Kolton. 1984. Distribution of ammonia-related aflatoxin

reaction products in cottonseed meal. J. Am. Oil Chem. Soc. 61:1071–1074.

Park, S. H., S. I. Lee, and S. C. Ricke. 2016. Microbial populations in naked neck chicken ceca

raised on pasture flock fed with commercial yeast cell wall prebiotics via an Illumina

MiSeq platform. PLOS One 1:1–14.

Parvin, S., C. Wang, Y. Li, and N. Nishino. 2010. Effects of inoculation with lactic acid bacteria

on the bacterial communities of Italian ryegrass, whole crop maize, guinea grass and

rhodes grass silages. Anim. Feed Sci. Technol. 160:160–166.

Patterson, R., and L. G. Young. 1993. Efficacy of hydrated sodium calcium aluminosilicate,

screening and dilution in reducing the effects of mold contaminated corn in pigs.

Canadian J. Anim. Sci. 73:616–624.

Pedroso, A., A. T. Adesogan, O. C. M. Queiroz, and S. Williams. 2010. Control of Escherichia

coli O157:H7 in corn silage with or without various inoculants: efficacy and mode of

action. J. Dairy Sci. 93:1098–1104.

Pestka, J. J. 2007. Deoxynivalenol: Toxicity, mechanisms and animal health risks. Anim. Feed

Sci. Technol. 137:283–298.

Pfliegler, W. P., T. Pusztahelyi, and I. Pócsi. 2015. Mycotoxins– Prevention and

decontamination by yeasts. J. Basic Microbiol. 55:805–818.

Phillips, T. D. 1999. Dietary clay in chemoprevention of aflatoxin-induced disease. Toxicol. Sci.

(Suppl.) 52:118-126.

Phillips, T. D., A. B. Sarr, and P. G. Grant. 1995. Selective chemisorption and detoxification of

aflatoxins by phyllosilicate Clay. Nat. Toxins 3:204-213.

Phillips, T. D., B. A. Clement, and D. L. Park. 1994. Approaches to reduction of aflatoxin in

foods and feeds. Pp. 383–406. In L. D. Eaton and J. D. Groopman (Eds.). The Toxicology

of Aflatoxins: Human Health, Veterinary Agricultural Significance. Academic Press,

New York.

Phillips, T. D., L. F. Kubena, R. B. Harvey, D. R. Taylor, and N. D. Heidelbaugh. 1987.

Mycotoxin hazards in agriculture: New approach to control. J. Am. Vet. Med. Assoc.

190:1617-1631.

Phillips, T. D., L. F. Kubena, R. B. Harvey, D. R. Taylor, and N. D. Heidelbaugh. 1988.

Hydrated sodium calcium aluminosilicate: A high affinity sorbent for aflatoxin. Poult.

Sci. 67:253-260.

202

Phillips, T. D., S. L. Lemke, and P. G. Grant. 2002. Characterization of clay-based

enterosorbents for the prevention of aflatoxicosis. Adv. Exp. Med. Biol. 504:157–171.

Pietri, A., and G. Piva. 2007. Aflatoxins in foods. Italian J. Public Health. 4:32-38.

Pietri, A., T. Bertuzzi, L. Pallaroni, and G. Piva. 2004. Occurrence of mycotoxins and ergosterol

in maize harvested over 5 years in Northern Italy. Food Addit. Contam. Part A Chem.

Anal. Control Expo. Risk Assess. 21:479–487.

Pitt, R. E., 1986. Dry matter losses due to oxygen infiltration into silos. J. of Agric. Engng.

Res. 35:193–205.

Pittet, A., 1998. Natural occurrence of mycotoxins in foods and feeds- an update review. Rev.

Med. Vet. 149:479–492.

Polley, H. W., B. J. Wilsey, and J. D. Derner. 2007. Dominant species constrain effects of

species diversity on temporal variability in biomass production of tallgrass prairie. Oikos

116:2044–2052.

Potter A. A., S. Klashinsky, Y. Li, E. Frey, H. Townsend, D. Rogan, G. Erickson, S. Hinkley, T.

Klopfenstein, R. A. Moxley, D. R. Smith, and B. B. Finlay. 2004. Decreased shedding of

Escherichia coli O157:H7 by cattle following vaccination with type III secreted proteins

Vaccine 22:362–369.

Puntenney, S. B., Y. Wang, and N. E. Forsberg. 2003. Mycotic infections in livestock: recent

insights and studies on etiology, diagnostics and prevention of hemorrhagic bowel

syndrome. pp.49–62 In: Southwest Animal Nutrition and Management Conference

Proceedings.

Quail, M. A., M. Smith, P. Coupland, T. D. Otto, S. R. Harris, T. R. Connor, A. Bertoni, H. P.

Swerdlow, and Y. Gu. 2012. A tale of three next generation sequencing platforms:

comparison of Ion Torrent, Pacific Biosciences and Illumina MiSeq sequencers. BMC

Genomics 13:341-354.

Queiroz, O. C. M., J. H. Han, C. R. Staples, and A. T. Adesogan. 2012. Effect of adding a

mycotoxin-sequestering agent on milk aflatoxin M1 concentration and the performance

and immune response of dairy cattle fed an aflatoxin B1-contaminated diet. J. Dairy Sci.

95:5901–5908.

Queiroz, O. C. M., K. G. Arriola, J. L. P. Daniel, and A. T. Adesogan. 2013. Effects of 8

chemical and bacterial additives on the quality of corn silage. J. Dairy Sci. 96:5836–

5843.

Raju, M. V. L. N., and G. Devegowda. 2000. Influence of Esterified-Glucomannan on

performance and organ morphology, serum biochemistry and haematology in broiler

exposed to individual and combined mycotoxicosis (aflatoxin, ochratoxin and T-2 toxin).

Br. Poult. Sci. 41:640–650.

203

Ranjit, N. K., C. C. Taylor, and L. Kung Jr. 2002. Effect of Lactobacillus buchneri 40788 on the

fermentation, aerobic stability and nutritive value of maize silage. Grass Forage Sci.

57:73–81.

Rasby, R., R. Stock, B. Anderson, and N. Schneider. 1988. Nitrates in livestock feeding.

Nebraska Cooperative Extension Service, G74-170.

Rastogi, G., and R. K. Sani. 2016. Molecular techniques to assess microbial community

structure, function, and dynamics in the environment. In ahmad et al. (eds.), microbes and

microbial technology: Agricultural and Environmental Applications, DOI 10.1007/978-1-

4419-7931-5_2.

Raters, M., and R. Matissek. 2008. Thermal stability of aflatoxin B1 and ochratoxin A.

Mycotoxin Research 24:130-134.

Rawel, H. M., and S. Rohn. 2010. Nature of hydroxycinnamate–protein interactions. Phytochem

Rev. 9:93–109.

Rayner, E. T., S. P. Koltun, and F. G. Dollear. 1977. Solvent extraction of aflatoxins from

contaminated agricultural products. J. Am. Oil Chem. Soc. 54:242–244.

Raza, K., and S. Ahmad. 2016. Principle, analysis, application and challenges of next-generation

sequencing: a review. arXiv:1606.05254.

Razzaq, S. 2006. Hemolytic uremic syndrome: an emerging health risk. Am. Fam. Physician

74:991–996.

Reed, K. F. M., and D. D. Moore. 2009. A preliminary survey of zearalenone and other

mycotoxins in Australian silage and pasture. Anim. Prod. Sci. 49: 696–703.

Reinstein, S., J. T. Fox, X. Shi, and T. G. Nagaraja. 2007. Prevalence of Escherichia coli

O157:H7 in gallbladders of beef cattle. Appl. Environ. Microbiol. 73:1002-1004.

Richard, E., N. Heutte, V. Bouchart, and D. Garon. 2009. Evaluation of fungal contamination

and mycotoxin production in maize silage. Anim. Feed Sci. Technol. 148:309-320.

Riley, L. W., R. S. Remis, S. D. Helgerson, H. B. McGee, J. G. Wells, B. R. Davis, R. J. Hebert,

E. S. Olcott, L. M. Johnson, N. T. Hargrett, P. A. Blake, and M. L. Cohen. 1983.

Hemorrhagic colitis associated with a rare Escherichia coli serotype. N. Engl. J. Med.

308:681–685.

Robins-Browne, R. M., K. E. Holt, D. J. Ingle, D. M. Hocking, J. Yang, and M. Tauschek. 2016.

Are Escherichia coli pathotypes still relevant in the era of whole-genome

sequencing? Frontiers in Cellular Infect. Microbiol. 6.

Rodrigues, I., and K. Naehrer. 2012. A three-year survey on the worldwide occurrence of

mycotoxins in feedstuffs and feed. Toxins 4:663-675.

204

Ronaghi, M., S. Karamohamed, B. Pettersson, M. Uhlen, and P. Nyren. 1996. Real-time DNA

sequencing using detection of pyrophosphate release. Anal. Biochem. 242:84–89.

Rossi, F., and Dellaglio, F., 2007. Quality of silages from Italian farms as attested by number and

identity of microbial indicators. J. Appl. Microbiol. 103:707–1715.

Rotter, B. A., D. B. Prelusky, and J. J. Pestka.1996. Toxicology of deoxynivalenol (vomitoxin).

J. Toxicol. Environ. Health 48:1–34.

Russell, J. B., F. Diez-Gonzalez, and G. N. Jarvis. 2000. Potential effects of cattle diets on the

transmission of pathogenic Escherichia coli to humans. Microbes Infect. 2:45–53.

Saeger, S. D., L. Sibanda, and C. V. Peteghem. 2003. Analysis of zearalenone and alpha-

zearalenol in animal feed using high-performance liquid chromatography. Analytica

Chemica Acta. 478: 137-143.

Saito M., and K. Okubo. 1974. Trichothecene toxins of Fusarium species. In: Mycotoxins (Ed.

by I.F.H. Purchase), p. 263. Elsevier Scientific Publishing Co., Amsterdam.

Samuel, J. E., L. P. Perera, S. Ward, A. D. O'brien, V. Ginsburg, and H. C. Krivan. 1990.

Comparison of the glycolipid receptor specificities of Shiga-like toxin type II and Shiga-

like toxin type II variants. Infect. and immun. 58:611-618.

Sanger, F., S. Nicklen, and A. R. Coulson. 1977. DNA sequencing with chain-terminating

inhibitors. PNAS. 74:5463-5467.

Santos, A. O., C. L. S. Avila, J. C. Pinto, B. F. Carvalho, D. R. Dias, and R. F. Schwan. 2015.

Fermentative profile and bacterial diversity of corn silages inoculated with new tropical

lactic acid bacteria. J. of Appl. Microbiol. 120:266-279.

Sargeant J. M., M. R. Amezcua, A. Rajic, and L. Waddell. 2007. Pre-harvest interventions to

reduce the shedding of E. coli O157 in the feces of weaned domestic ruminants: a

systematic review. Zoonoses Public Health 54:260–77.

Sarr, A. B., B. A. Clement, and T. D. Phillips. 1990. Effects of molecular structure on the

chemisorption of aflatoxin B1 and related compounds by hydrated sodium calcium

aluminosilicate. Toxicologist 10:163-167.

Sarr, A B., B. A Clement, and T. D. Phillips. 1991. Molecular mechanism of aflatoxin B1

chemisorption by hydrated calcium aluminosilicate. Toxicologist 11:97-110.

SAS Institute Inc. 2012. What’s New in SAS 9, Inc., Cary, NC.

Satoh H., W. D. Ramey, F. A. Koch, W. K. Oldham, T. Mino, and T. Matsuo. 1996. Anaerobic

substrate uptake by the enhanced biological phosphorus removal activated sludge treating

real sewage. Water Sci. Tech. 34:8-15.

205

Scallan E., R. M. Hoekstra, F. J. Angulo, R. V. Tauxe, M. A. Widdowson, and S. L. Roy

2011. Foodborne illness acquired in the United States—major pathogens. Emerg. Infect.

Dis. 17:7–15.

Schatzmayr, G., F. Zehner, M. Taubel, D. Schatzmayr, A. Klimitsch, A. P. Loibner, and E. M.

Binder. 2006. Microbiologicals for deactivating mycotoxins. Mol. Nutr. Food Res.

50:543-551.

Schell, T. C., M. D. Lindemann, E. T. Kornegay, and D. J. Blodgett. 1993. Effects of feeding

aflatoxincontaminated diets with and without clay to weanling and growing pigs on

performance, liverfunction, and mineral metabolism. J. Anim. Sci. 71:1209-1218.

Schmidt, R. J., M. G. Emara, and L. Kung Jr. 2008. The use of a quantitative real-time

polymerase chain reaction assay for identification and enumeration of Lactobacillus

buchneri in silages. J. Appl. Microbiol. 105:920–929.

Schmidt, R. J., and L. Kung, Jr. 2010. The effects of Lactobacillus buchneri with or without a

homolactic bacterium on the fermentation and aerobic stability of corn silages made at

different locations. J. Dairy Sci. 93:1616–1624.

Schneweis, I., K. Meyer, S. Hormansdorfer, and J. Bauer. 2000. Mycophenolic acid in silage.

Appl. Environ. Microbiol. 66:3639– 3641.

Schollenberger, M., H. M. Müller, M. Rüfle, S. Suchy, S. Plank, and W. Drochner. 2006. Natural

occurrence of 16 Fusarium toxins in grains and feedstuffs of plant origin from Germany.

Mycopathologia 161:43–52.

Schroder, K., and J. Tschopp. 2010. The inflammasomes. Cell 140:821–832.

Scott, P. M., T. Delgado, D. B. Prelusky, H. L. Trenholm, and J. D. Miller. 1994. Determination

of fumonisin in milk. J. Environ. Sci. Health. 29:989-998.

Scudamore, K. A., and C. T. Livesey 1998. Occurrence and significance of mycotoxins in forage

crops and silage: A review. J. Sci. Food Agric. 77:1–17.

Seeling, K., S. Danicke, K. H. Ueberschar, P. Lebzien, and G. Flachowsky. 2005. On the effects

of Fusarium toxin–contaminated wheat and the feed intake level on the metabolism and

carryover of zearalenone in dairy cows. Food Addit. Contam. 22:847–855.

Seok, Y., H. Shin, Y. Lee, I. Cho, S. Na, D. Yong, S. H. Jeong, and K. Lee. 2010. First report of

bloodstream infection caused by Pseudomonas fulva. J. Clin. Microbiol. 48:2656 –2657

Shrestha, R., S. H. Lee, J. H. Hur, and C. K. Lim. 2005. The effects of temperature, pH and

bactericides on the growth of Erwinia pyrifoliae and Erwinia amylovora. Plant Pathol. J.

21:127–131.

Siegler, R. L. 2003. Post-diarrheal Shiga toxin–mediated hemolytic uremic syndrome. JAMA

290:1379–1381.

206

Silvestre, F. T., T. S. Carvalho, P. C. Crawford, J. E. Santos, C. R. Staples, T. Jenkins, and W.

W. Thatcher. 2011. Effects of differential supplementation of fatty acids during the

peripartum and breeding periods of Holstein cows: II. Neutrophil fatty acids and function,

and acute phase proteins. J. Dairy Sci. 94:2285–2301.

Siwela, A. H., M. Siwela, G. Matindi, S. Dube, and N. Nziramasanga. 2005. Decontamination of

aflatoxin-contaminated maize by dehulling. J. Sci. Food Agric. 85:2535-2538.

Smit, E., P. Leeflang, and K. Wernars. 1997. Detection of shifts in microbial community

structure and diversity in soil caused by copper contamination using amplified ribosomal

DNA restriction analysis. FEMS Microbiol. Ecol. 23:249–261.

Smith, D. R., R. A. Moxley, S. L. Clowser, J. D. Folmer, S. Hinkley, G. E. Erickson and T. J.

Klopfenstein. 2005. Use of rope devices to describe and explain the feedlot ecology of

Escherichia coli O157:H7 by time and place. Foodborne Path. Dis. 2:50–60.

Smith, E. E., T. D. Phillips, J. A. Ellis, R. B. Harvey, L. F. Kubena, J. Thompson, and G.

Newton. 1994. Dietary hydrated sodium calcium aluminosilicate reduction of aflatoxin

M1 residue in dairy goat milk and effects on milk production and components. J. Anim.

Sci. 72:677–682.

Smolen, J. E., T. K. Petersen, C. Koch, S. J. O’Keefe, W. A. Hanlon, S. Seo, D. Pearson, M. C.

Fossett, and S. I. Simon. 2000. L-Selectin signaling of neutrophil adhesion and

degranulation involves p38 mitogen-activated protein kinase. J. Biol. Chem. 275:15876–

15884.

Snedeker, K. G., M. Campbell, and J. M. Sargeant. 2011. A systematic review of vaccinations to

reduce the shedding of Escherichia coli O157 in the feces of domestic ruminants.

Zoonoses Public Health 59:126–38.

Soriano, J. M., L. González, and A. I. Catalá 2005. Mechanism of action of sphingolipids and

their metabolites in the toxicity of fumonisin B1. Prog. Lipid Res. 44:345–356.

Squire, R. A. 1981. Ranking animal carcinogens: a proposed regulatory approach. Science 214:

877–880.

Stanford K., C. A. Agopsowicz, and T. A. McAllister. 2012. Genetic diversity and antimicrobial

resistance among isolates of Escherichia coli O157:H7 from feces and hides of super-

shedders and low-shedding pen-mates in two commercial beef feedlots. BMC Vet. Res.

8:178.

Stanford, K., D. Croy, S. J. Bach, G. L. Wallins, H. Zahiroddini, and T. A. McAllister. 2005.

Ecology of Escherichia coli O157: H7 in commercial dairies in southern Alberta. J. Dairy

Sci. 88:4441-4451.

Stanton, T. L., and D. Schutz. 2000. Effect of switching from high grain to hay five days prior to

slaughter on finishing cattle performance. Colorado State University Research Report.

Colorado State University, Fort Collins, CO.

207

Stephens T. P., T. A. McAllister, and K. Stanford. 2009. Perineal swabs reveal effect of super

shedders on the transmission of Escherichia coli O157:H7 in commercial feedlots. J.

Anim. Sci. 87:4151–4160.

Stevenson D. M., R. E. Muck, K. J. Shinners, and P. J. Weimer. 2006. Use of real time PCR to

determine population profiles of individual species of lactic acid bacteria in alfalfa silage

and stored corn stover. Appl. Microbiol. Biotechnol. 71:329–338.

Storm, I. M. L. D., N. B. Kristensen, B. M. L. Raun, J. Smedsgaard, and U. Thrane. 2010.

Dynamics in the microbiology of maize silage during whole-season storage. J. Appl.

Microbiol. 109:1017–1026.

Storm, I. M. L. D., R. R. Rasmussen, and P. H. Rasmussen. 2014. Occurrence of pre- and post-

harvest mycotoxins and other secondary metabolites in Danish maize silage. Toxins

6:2256–2269.

Storm, I. M., J. L. Sørensen, R. R. Rasmussen, K. F. Nielsen, and U. Thrane. 2008. Mycotoxins

in silage. Stewart Posthar. Rev. 4.

Stroud, J. S. 2006. The effect of feed additives on aflatoxin in milk of dairy cows fed aflatoxin

contaminated diets. M.S. Thesis. North Carolina State Univ., Raleigh.

Sulzberger, S. A., S. Melnichenko, and F. C. Cardoso. 2017. Effects of clay after an aflatoxin

challenge on aflatoxin clearance, milk production, and metabolism of Holstein cows. J.

Dairy Sci. 100:1-14 (In press).

Sumarah, M. W., J. D. Miller, and B. A. Blackwell. 2005. Isolation and metabolite production by

Penicillium roqueforti, P. paneum and P. crustosum isolated in Canada. Mycopathologia.

159:571-577.

Swanson, S. P., J. Nicoletti, H. D. Rood, W. B. Buck, and L. M. Coe. 1987. Metabolism of three

trichothecene mycotoxins, T-2 toxin, diacetoxyscirpenol and deoxynivalenol, by bovine

rumen microorganisms. J. Chromatogr. 414:335.

Sweeney, M. J., and A. D. W. Dobson. 1998. Mycotoxin production by Aspergillus, Fusarium

and Penicillium species. Int. J. Food Microbiol. 43:141–158.

Tabacco, E., S. Piano, L. Cavallarin, T. F. Bernades, and G. Borreani. 2009. Clostridial spore

formation during aerobic deterioration of maize and sorghum silages as influenced by

Lactobacillus buchneri and Lactobacillus plantarum inoculants. J. Appl. Microbiol.

107:1632–1641.

Tanaka, T., S. Yamamoto, A. Hasegawa, N. Aoki, J. R. Besling, Y. Sugiura, and Y. Ueno. 1990.

A survey of the natural occurrence of Fusarium mycotoxins, deoxynivalenol, nivalenol

and zearalenone, in cereals harvested in the Netherlands. Mycopathologia 110:19–22.

208

Tao, L., X. S. Guo, H. Zhou, D. J. Undersander, and A. Nandety. 2012. Characteristics of

proteolytic activities of endo- and exopeptidases in alfalfa herbage and their implications

for proteolysis in silage. J. Dairy Sci. 95:4591-4595.

Taylor, C. C., N. J. Ranjit, J. A. Mills, J. M. Neylon, and L. Kung, Jr. 2002. The effect of treating

whole-plant barley with Lactobacillus buchneri 40788 on silage fermentation, aerobic

stability, and nutritive value for dairy cows. J. Dairy Sci. 85:1793–1800.

Tchaptchet, S., and J. Hansen. 2011. The Yin and Yang of host-commensal mutualism. Gut

Microbes 2:347–352.

Teller, R. S., R. J. Schmidt, L. W. Whitlow, and L. Kung, Jr., 2012. Effect of physical damage to

ears of corn before harvest and treatment with various additives on the concentration of

mycotoxins, silage fermentation, and aerobic stability of corn silage. J. Dairy Sci.

95:1428-1436.

Temmerman, R., I. Sheirlinck, G. Huys, and J. Swings. 2003. Culture-independent analysis of

probiotic products by denaturing gradient gel electrophoresis. Appl. Environ. Microbiol.

69:220–226.

Thies, J. E. 2007. Soil microbial community analysis using terminal restriction fragment length

polymorphisms. Soil Sci. Soc. Am. J. 71:579–591.

Thomson, D. U., G. H. Loneragan, A. B. Thornton, K. F. Lechtenberg, D. A. Emery, and D. T.

Burkhardt. 2009. Use of a siderophore receptor and porin proteins-based vaccine to

control the burden of Escherichia coli O157:H7 in feedlot cattle. Foodborne Pathog. Dis

6:871–877.

Thorpe, C. M. 2004. Shiga toxin-producing Escherichia coli infection. Clin. Infect. Dis. 38:

1298–1303.

Torriani, S., G. E. Felis, and F. Dellaglio 2001. Differentiation of Lactobacillus plantarum, L.

pentosus, and L. paraplantarum by recA gene sequence analysis and multiplex PCR

assay with recA gene-derived primers. Appl. Environ. Microbiol. 67:3450– 3454.

Trenholm, H. L., B. K. Thompson, K. E. Hartin, R. Greenhalgh, and A. J. McAllister. 1985.

Ingestion of vomitoxin (deoxynivalenol)-contaminated wheat by non-lactating dairy

cows. J. Dairy Sci. 68:1000–1005.

Trenholm, H. L., R. M. Hamilton, D. W. Friend, B. K. Thompson, and K. E. Hartin. 1984.

Feeding trials with vomitoxin (deoxynivalenol)-contaminated wheat: Effects on pigs,

poultry, and dairy cattle. J. Am. Vet. Med. Assoc. 185:527–531.

Tsitsigiannis, D. I., M. Dimakopoulou, P. P. Antoniou, and E. C. Tjamos. 2012. Biological

control strategies of mycotoxigenic fungi and associated mycotoxins in Mediterranean

basin crops. Phytopathol. Mediterr. 51:158–174.

209

Tsitsigiannis, D. I., M. Georgiadou, S. Agoritsis, G. Zakynthinos, T. H. Varzakas, S. E. Tjamos,

P. P. Antoniou, M. Dimakopoulou, G. Karnavas, E. Paplomatas, S. Gianniotis, and E. C.

Tjamos, 2010. Ecology, epidemiology and control of Aspergillus spp. in pistachio

orchards in Greece. Petria 20:95‒96.

Tulayakul P., S. Sakuda, K. S. Dong, and S. Kumagai. 2005. Comparative activities of

Glutathione-S-transferase and dialdehyde reductase toward aflatoxin B1 in livers of

experimental and farm animals. Toxicon. 46:204–209.

Upadhaya, S. D., M. A. Park, and J. K. Ha. 2010. Mycotoxins and their biotransformation in the

rumen: a review. J. Anim. Sci. 23:1250-1260.

USDA. 1998. Pathogen reduction/ HACCP regulatory requirements. Accessed February 8, 2017.

https://www.fsis.usda.gov/wps/portal/fsis/topics/regulatory-compliance/haccp/pr-and-

haccp-guidance-documents/pathogen-reduction-haccp-guidance.

USDA. 2014. Cost estimates of foodborne illnesses. Accessed February 8, 2017.

https://www.ers.usda.gov/data-products/cost-estimates-of-foodborne

illnesses.aspx#.VDW27r4mUfy.

USDA. 2016. Crop Production 2015 Summary. Accessed February 8, 2017

https://www.usda.gov/nass/PUBS/TODAYRPT/cropan16.pdf.

USDA-APHIS. 2001. Escherichia coli O157 in United States feedlots. Center for Epidemiology

and Animal Health. Fort Collins, CO.

USDA-ERS. 2001. ERS estimates foodborne disease costs at $6.9 billion per year. Economic

Research Service- United States Department of Agriculture. Accessed August 27, 2016.

http://www.ers.usda.gov/publications/aer741/aer741.pdf.

Van Soest, P. J., J. B. Robertson, and B. A. Lewis. 1991. Methods for dietary fiber, neutral

detergent fiber and non-starch polysaccharides in relation to animal nutrition. J. Dairy

Sci. 74:3583–3597.

Varel, V. H., and H. G. Jung. 1986. Influence of forage phenolics on ruminal fibrolytic bacteria

and in vitro fiber degradation. Appl. Environ. Microbiol. 52:275–280.

Velde, B., and A. Meunier. 2008. Fundamentals of clay mineral crystal structure and

physicochemical properties, in: The origin of clay minerals in soils and weathered rocks,

1st ed., Springer, Berlin, pp. 3–74.

Veldman, A., G. J. Borggreve, E. J. Mulders, and D. Van de Lagemaat. 1992. Occurrence of the

mycotoxins ochratoxin A, zearalenone and deoxynivalenol in feed components. Food

Add. Contaminants 9:647-655.

Viazis, S., and F. Diez-Gonzalez. 2011. Enterohemorrhagic Escherichia coli: the twentieth

century’s emerging foodborne pathogen: a review. Adv. Agronom. 111:1-50.

https://www.fsis.usda.gov/wps/portal/fsis/topics/regulatory-compliance/haccp/pr-and-haccp-guidance-documents/pathogen-reduction-haccp-guidance

https://www.fsis.usda.gov/wps/portal/fsis/topics/regulatory-compliance/haccp/pr-and-haccp-guidance-documents/pathogen-reduction-haccp-guidance

https://www.ers.usda.gov/data-products/cost-estimates-of-foodborne%20illnesses.aspx#.VDW27r4mUfy

https://www.ers.usda.gov/data-products/cost-estimates-of-foodborne%20illnesses.aspx#.VDW27r4mUfy

https://www.usda.gov/nass/PUBS/TODAYRPT/cropan16.pdf

210

Vigier, B., L. M. Reid, K. A. Seifert, D. W. Stewart, and R. I. Hamilton. 1997. Distribution and

prediction of Fusarium species associated with maize ear rot in Ontario. Can. J. Plant

Pathol. 19:60–65.

Vissers, M. M. M., F. Driehuis, M. C. Te Giffel, P. De Jong, and J. M. G. Lankveld. 2006.

Improving farm management by modeling the contamination of farm tank milk with

butyric acid bacteria. J. Dairy Sci. 89:850–858.

Voss, K. A., W. J. Chamberlain, C. W. Bacon, R. A. Herbert, D. B. Walters, and W. P. Norred.

1995. Subchronic feeding study of the mycotoxin fumonisin B1 in B6C3F1 mice and

Fischer 344 rats. Fund Appl. Toxicol. 24:102–110.

Walczak, J. J., and S. P. Xu. 2011. Manure as a source of antibiotic-resistant Escherichia coli

and Enterococci: A Case Study of a Wisconsin, USA Family Dairy Farm. Water Air Soil

Poll. 219:579–589.

Wallace, R. J., M. L. Falconer, and P. K. Bhargava. 1989. Toxicity of volatile fatty acids at

rumen pH prevents enrichment of Escherichia coli by sorbitol in rumen contents. Current

Microbiol. 19:277-281.

Waltman, L., S. Davidson, B. A. Hopkins, G. W. Smith, and L. W. Whitlow. 2008. Changes in

milk aflatoxin concentrations in response to investigational sequestering agents added to

aflatoxin- contaminated diets fed to lactating Holstein cows. J. Dairy Sci. 91(E–Suppl.

1):268. (Abstr.).

Wang, X., S. Haruta, P. Wang, M. Ishii, Y. Igarashi, and Z. Cui. 2006. Diversity of a stable

enrichment culture which is useful for silage inoculant and its succession in alfalfa silage.

FEMS Microbiol. Ecol. 57:106–115.

Ward, R. T. 2008. Fermentation analysis of silage: use and interpretation. Accessed July 01,

2016 http://www.foragelab.com/media/fermentation-silage-nfmp-oct-2008.

Weaver, G. A., H. J. Kurtz, J. C. Behrens, T. S. Robison, B. E. Seguin, F. Y. Bates, and C. J.

Mirocha 1986. Effect of zearalenone on the fertility of virgin dairy heifers. Am. J. Vet.

Res. 47:1395–1397.

Weinberg Z. G., G. Ashbell, Y. Chen, and A. Azrieli. 1995. The effect of propionic acid bacterial

inoculant applied at ensiling on the aerobic stability of wheat and sorghum silages. J. Ind.

Microbiol. Biotechnol. 15:493–497.

Weinberg, Z. G., and R. E. Muck. 1996. New trends in development and use of inoculants for

silage. FEMS Microbiol. Rev. 19:53–68.

Weinberg, Z. G., G. Ashbell, Y. Chen, A. Azrieli, G. Szakacs, and I. Filya. 2002. Ensiling

whole-crop wheat and corn in large containers with Lactobacillus plantarum and

Lactobacillus buchneri. J. Ind. Microbiol. Biotechnol. 28:7–11.

211

Weinberg, Z. G., G. Ashbell, Y. Chen, M. Gamburg, and S. Sela. 2004. The effect of sewage

irrigation on safety and hygiene of forage crops and silage. Anim. Feed Sci. Technol.

116:271–280.

Wells, J. E., E. D. Berry, and V. H. Varel. 2005. Effects of common forage phenolic acids on

Escherichia coli O157:H7 viability in bovine feces. Appl. Environ. Microbiol. 71:7974-

7979.

Wells, J. E., E. D. Berry, and V.H. Varel. 2005. Effects of common forage phenolic acids on

Escherichia coli O157:H7 viability in bovine feces. Appl. Environ. Microbiol. 71:7974-

7979.

Whiter, A. G., and L. Kung, Jr. 2001. The effect of a dry or liquid application of Lactobacillus

plantarum MTD1 on the fermentation of alfalfa silage. J. Dairy Sci. 84:2195–2202.

Whitlow L. W., and W. M. Hagler. 2005. Mycotoxins in dairy cattle: occurrence, toxicity,

prevention and treatment. Proc. Southwest Nutr. Conf. 124–138.

Whitlow, L. W. 1993. Mycotoxin contamination of silages: A potential cause of poor production

and health in dairy herds silage production from seed to animal. Proceedings of the

NRAES-93. National Silage Production Conference, Syracuse, NY, pp. 220-231.

Whitlow, L. W. 2005. Moulds and mycotoxins in feedstuffs - Prevention and treatment.

Proceedings of the 16th Annual Florida Ruminant Nutrition Symposium, Gainesville,

Florida:123-142.

Whitlow, L. W. 2006. Evaluation of mycotoxin binders, p. 132–143. In N. G. Zimmermann

(ed.), Proceedings of the 4th Mid-Atlantic Nutrition Conference. Maryland Feed Industry

Council, Department of Animal and Avian Sciences, University of Maryland, College

Park.

Whitlow, L. W., R. L. Nebel, and W. M. Hagler. 1994. The association of deoxynivalenol in

grain with milk production loss in dairy cows. Pp. 131-139. In: G. C. Llewellyn, W. V.

Dashek and C. E. O’Rear. (Eds.) "Biodeterioration Research 4". Plenum Press, New

York.

Wicklow, D. T. 1983. Taxonomic features and ecological significance of sclerotia. In: U. L.

Deiner, R. L. Asquith and J. W. Dickens. (Eds) "Aflatoxin and Aspergillus flavus in

Corn". Southern Coop. Serv. Bull. 279. Craftmaster, Opelika, AL.

Wilkinson, J. M., and D. R. Davies. 2013. The aerobic stability of silage: key findings and recent

developments. Grass Forage Sci. 68:1–19.

Winfield, M. D., and E. A. Groisman. 2003. Role of non-host environments in the lifestyles of

Salmonella and Escherichia coli. Appl. Environ. Microbiol. 69:3687-3694.

Woolford, M. K. 1975. Microbial screening of food preservatives, cold sterilants, and specific

antimicrobial agents as potential silage additives. J. Sci. Food Agric. 26:229–237.

212

World Health Organization. 2000. Fumonisin B1 (Environmental Health Criteria 219), Geneva,

153 pp.

Wu, J. J., R. P. Du, M. Gao, Y. Q. Sui, L. Xiu, and X. Wang. 2014. Naturally occurring lactic

acid bacteria isolated from tomato pomace silage. Asian-Australasian J. Anim. Sci.

27:648–657.

Wu, Q., A. Jezkova, Z. Yuan, L. Pavlikova, and V. Dohnal. 2009. Biological degradation of

aflatoxins. Drug Metab. Rev. 41:1–7.

Xiong, J. L., Y. M. Wang, T. D. Nennich, Y. Li, and J. X. Liu. 2015. Transfer of dietary

aflatoxin B1 to milk aflatoxin M1 and effect of inclusion of adsorbent in the diet of dairy

cows. J. Dairy Sci. 98:2545–2554.

Xu, X., L. V. Madden, and S. G. Edwards. 2014. Modeling the effects of environmental

conditions on HT2 and T2 toxin accumulation in field oat grains. Phytopathology 104:57-

66.

Yang, L. 2010. Effects of feed types on Ochratoxin A biodegradation by Korean native goats.

master’s thesis, Seoul National University, Korea.

Yiannikouris, A., and J. P. Jouany. 2002. Mycotoxins in feeds and their fate in animals: a review.

Anim. Res. 51:81–99.

Yitbarek, M. B., and B. Tamir. 2014. Silage additives: a review. Open J. Appl. Sci. 4:258-274.

Younts-Dahl, S. M., M. L. Galyean, G. H. Loneragan, N. A. Elam, and M. M. Brashears. 2004.

Dietary supplementation with Lactobacillus- and Propionibacterium-based direct fed

microbials and prevalence of Escherichia coli O157 in beef feedlot cattle and on hides at

harvest. J. Food Prot. 67:889-893.

Yu, P., J. J. McKinnon, and D. A. Christensen. 2005. Improving the nutritional value of oat hulls

for ruminant animals with pretreatment of a multienzyme cocktail: In vitro studies. J.

Anim. Sci. 83:1133-1141.

Zadik, P. M., P. A. Chapman, and C. A. Siddons. 1993. Use of tellurite for the selection of

verocytotoxigenic Escherichia coli O157. J. Med. Microbiol. 39:155–158.

Zain, M. E. 2011. Impact of mycotoxins on humans and animals. J. Saudi Chem. Soc. 15:129–

144.

Zhang. C., M. J. Brandt, C. Schwab, and M. G. Gänzle. 2010. Propionic acid production by co-

fermentation of Lactobacillus buchneri and Lactobacillus diolivorans in sourdough. Food


Zhao, T., M. P. Doyle, J. Shere, and L. Garber. 1995. Prevalence of enterohemorrhagic

Escherichia coli O157:H7 in a survey of dairy herds. Appl. Environ. Microbiol. 61:1290–

1293.

213

Zinedine, A., J. M. Soriano, J. C. Molto, and J. Manes. 2007. Review on the toxicity, occurrence,

metabolism, detoxification, regulations and intake of zearalenone: An oestrogenic

mycotoxin. Food Chem. Toxicol. 45:1–18.

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BIOGRAPHICAL SKETCH

Ibukun Ogunade was born in Lagos, Nigeria. He attended the University of Agriculture,

Abeokuta, Nigeria where he earned his Bachelor of Science and Master of Science degrees in

2009 and 2011, respectively. His master’s thesis focused on reducing fecal shedding of

Salmonella in laying hens with the use of probiotics and prebiotics. Immediately after obtaining

his Master of Science degree in Animal Nutrition, he was employed as a Lecturer at the same

university from 2011 to 2013 where he taught several undergraduate animal nutrition classes. In

June 2013, Ibukun became a Doctor of Philosophy candidate in the Department of Animal

Sciences at the University of Florida under Dr. Adegbola Adesogan’s supervision, and he

received his Ph.D in the spring of 2017. During his doctoral research program, Ibukun performed

several applied dairy nutrition research studies investigating forage quality and conservation,

feed microbiology and safety, and dietary mycotoxin sequestration strategies. He also

collaborated with other researchers within and outside the University on several studies that

focused on using fibrolytic enzymes and novel proteins to improve forage utilization, evaluating

the use of alternative feed ingredients, and developing alternatives to antibiotics for mitigating

greenhouse gas production by ruminant livestock, and improving their productive efficiency.

Ibukun’s long-term plan is to become a faculty member at a university in the US.

Documents

EFFECTS OF MYCOTOXIN ADSORBENTS ON ...ufdcimages.uflib.ufl.edu/UF/E0/05/08/73/00001/OGUNADE_I.pdfEFFECTS OF MYCOTOXIN ADSORBENTS ON AFLATOXINS AND OF MICROBIAL ADDITIVES ON Escherichia