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THESIS

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thesis entitled

THE USE OF DH42, A PROPIONIBACTERIUM FOR THE PREVENTION
OF LACTIC ACIDOSIS IN CATTLE

presented by

Ines Avilés

has been accepted towards fulfillment
of the requirements for

M.S. degree in Animal Science

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11m whom-9869.14

THE USE OF DH42, A PROPIONIBACTERIUM FOR THE PREVENTION OF
LACTIC ACIDOSIS IN CATTLE

By

Inés Avilés

A THESIS

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

MASTER IN SCIENCE
Department of Animal Science

1999

ABSTRACT

THE USE OF DH42, A PROPIONIBACTERIUM FOR THE PREVENTION OF
LACTIC ACIDOSIS IN CATTLE

By
Inés Avilés

Propionibacteria have the ability to produce propionic acid from lactic acid. This
characteristic was the basis for the hypothesis that if introduced into the rumen, lactic acid
levels would be lowered thus, preventing ruminal acidosis. The efl‘ects of adding
Propionibacterium acidipropionici DII42 to in-vitro ruminal fermentations at four
difl'erent inoculation rates and to rumen fistulated beef cattle receiving a high grain
finishing diet, via a rumen cannula and at inoculation rates higher than the in-vitro studies,
were examined. Propionibacterium acidipropionici DH42 added to in-vitro fermentations
resulted in no treatment effect (P > .05) on the levels of pH, lactic acid and volatile fatty
acids produced. Animal studies revealed a significant treatment effect (P < .05) on rumen
pH and blood pH. Blood and rumen lactic acid, total and individual volatile fatty acids,
and the average dry matter intake of animals were not affected (P > .05) by the addition of
strain DH42. Rumen and animal parameters in the treated and untreated groups
resembled those pertaining to cattle experiencing subacute acidosis. Microbiological
sampling of fermentors revealed growth of strain DH42 at O h only and absolutely no
growth was detected during the animal trials. In conclusion, P. acidipropionici DH42
does not appear to have the ability to survive in the rumen and prevent the accumulations

of lactic acid specifically when provided to cattle experiencing subacute ruminal acidosis.

To my Mother and my Father whom have shown an incredible amount of love throughout
the years and always told me to put forth my best effort despite the obstacles that exist in

life.

iii

ACKNOWLEDGEMENT

I would like to thank Dr. Melvin T. Yokoyama and Dr. Steven Rust for serving as
my co-advisors during my program and providing the necessary materials and equipment
to complete the research — thanks for the helpful suggestions, Dr. Yokoyama. I would
also like to express gratitude to my committee members Dr. Daniel Buskirk and Dr. Kent
Ames for their helpful advice.

My sincere thanks goes to Ms. Ruby Victoria Bato. She has been a wonderful
fiiend, mentor, and teacher during the course of this research. Thanks to her son, Robert,
for showing patience and understanding at such a young age. The fiiendship, the long
talks, the cooking, the challenges and surprises this experience has shown us has created a
lasting fiiendship that I will always treasure.

A warm thanks to Bob Burnett for great lab assistance and encouragement.

I would like to thank my husband, Paolo Spadoni for being patient, encouraging,
caring, and understanding. This work and writing would not have been possible without
your great attitude at home. “Grazie un milione arnore mio!”

Finally, I would like to thank the two most wonderful people in my life, my Mother
and Father. Words alone could not express the love and gratitude I feel as they have
supported me emotionally and financially throughout the many phases leading towards the
completion of the work and writing of this thesis. Thanks for believing in me “Marni and
Papi”.

Detours occur in life and we ask ourselves, “why?”. Then, one realizes why. God

knew what he was doing .....

iv

TABLE OF CONTENTS

LIST OF TABLES .................................................................................................. vii

REVIEW OF THE LITERATURE

LACTIC ACIDOSIS ................................................................................................. 1
General description ....................................................................................... 1
Chronic and acute acidosis ............................................................................ .2
Etiology of acidosis ...................................................................................... .3
Complications ................................................................................................ 4
The rumen microbial ecosystem and acidosis ................................................ .6
Lactic acid and VFA in ruminants .................................................................. 9

METHODS UTILIZED IN MANIPULATING RUB/[EN FERMENTATION .......... 13

Introduction .................................................................................................. 13
Ionophores and non-ionophores .................................................................... 14
Microbial feed additives ............................................................................... 21
THE PROPIONIBACTERIA ................................................................................... 27
General characteristics .................................................................................. 27
Metabolic pathways ...................................................................................... 29
Substrate preference, propionic acid production, pH, and temperature
requirements of propionibacteria .................................................................. 3O
Interactions between propionic and lactic acid producing bacteria ................. 32
Propionic acid and industrial uses ................................................................. 34
Bacteriocins of propionibacteria .................................................................... 36
Propionibacteria and probiotics ..................................................................... 39
Isolation and characterization of P. acidrpropionici DH42 ............................ 42
RESEARCH OBJECTIVES .................................................................................. 43
MATERIALS AND METHODS
In—Vitro Studies ....................................................................................................... 44
Preliminary studies on effectiveness of media ................................................ 44
Fermentor studies and experimental treatments ............................................. 44
Sampling schedule and analytical procedures ................................................ 45
HPLC analysis of fermentation end-products ................................................. 46
Statistical analysis ......................................................................................... 46
In-Vivo Studies ..................................................................................................... 47
Preliminary animal studies .......................................................................... 47
Animals, diet and experimental treatments .................................................. 48
Sampling schedule and analytical procedures .............................................. 49
Microbiological procedures and media ....................................................... 50

HPLC analysis of fermentation end-products and total

total blood lactic acid determination ............................................................ 50

Statistical analysis ....................................................................................... 51
RESULTS AND DISCUSSION
In-Vitro Studies ..................................................................................................... 53
In-Vivo Studies ...................................................................................................... 64
CONCLUSION .................................................................................................... 82
APPENDICES ...................................................................................................... 85
Appendix A

Figures of In-Vitro Studies ......................................................... 86
Appendix B

Figures of In-Vivo Studies .......................................................... 92
LITERATURE CITED ..................................................................... 101

Table 1

Table 2

Table 3

Table 4

Table 5

Table 6

Table 7

Table 8

Table 9

Table 10

Table 1 1

LIST OF TABLES

The effect of the addition of P. acidipropionici DH42
at different dose levels on rumen fluid pH
during in-vitro ferrnentations ................................................... 54

The effect of the addition of P. acidipropionici DH42
at different dose levels on the production of lactic acid
during in-vitro ferrnentations ................................................... 55

The effect of the addition of P. acidipropionici DH42
at different dose levels on the production of propionic acid
during in-vitro ferrnentations ................................................... 56

The effect of the addition of P. acidipropionici DH42
at different dose levels on the production of acetic acid
during in-vitro ferrnentations .................................................. 58

The effect of the addition of P. acidipropionici DH42
at different dose levels on the production of butyric acid
during in-vitro fermentations ................................................. 58

The effect of the addition of P. acidipropionici DH42
at different dose levels on the production of valeric acid
during in-vitro ferrnentations .................................................. 59

The effect of the addition of P. acidipropionici DH42
at different dose levels on the production of isobutyric acid
during in-vitro ferrnentations .................................................. 60

The effect of the addition of P. acidipropionici DH42 at difi‘erent
dose levels on the production of isovaleric acid during in-vitro
fermentat10ns60

The effect of the addition of P. acidipropionici DH42 at difl‘erent
dose levels on the production of total volatile fatty acids during
in-vitro ferrnentations ........................................................... 62

The recoverability of P. acidipropionici DH42 at 0 hours
from in-vitro ferrnentations with rumen fluid ............................... 63

The effect of P. acidipropionicr' DH42 (10ll cfii) on rumen pH
during experimentally induced acidosis in steers .................................. 64

vii

Table 12

Table 13

Table 14

Table 15

Table 16

Table 17

Table 18

Table 19

Table 20

Table 21

Table 22

The effect of P. acidipropionici DH42 (10ll cfu) on lactic acid
production during experimentally induced acidosis in steers ............ 66

The effect of P. acidipropionici DH42 (10” cfir) on
propionic acid production during experimentally
induced acidosis in steers ..................................................... 68

The effect of P. acidipropionici DH42 (10ll cfu) on
acetic acid production during experimentally
induced acidosis in steers ..................................................... 69

The effect of P. acidipropionici DH42 (10ll cfu) on
butyric acid production during experimentally
induced acidosis in steers ..................................................... 70

The effect of P. acidipropionici DH42 (10ll cfir) on
succinic acid production during experimentally
induced acidosis in steers ...................................................... 71

The effect of P. acidipropiom'ci DH42 (101l cfu) on
total volatile fatty acid production during experimentally
induced acidosis in steers ..................................................... 72

The effect of P. acidrpropionici DH42 (10ll cfir) on
blood pH during experimentally acidosis in steers ........................ 73

The effect of the addition of P. acidipropionici DH42 (10“cfu)
on total blood lactate during experimentally
induced acidosis in steers ........... ~ ........................................................ 74

The effect of the addition of P. acidrpropionici DH42 (10l lcfu)
on blood L(+) lactate during experimentally
induced acidosis in steers ................................................................... 75

The effect of the addition of P. acidipropionici DH42 (10"cfir)
on blood D(-) lactate dcuring experimentally
induced acidosis in steers ................................................................... 75

Table 20. The effect of the addition of 101 lcfil P. acidipropionici
DH42 on total blood lactate (mM),L+ lactate(mg/dl), D- lactate
(mg/d1), and blood pH of control and treated animals in the current
acidosis study as compared to results of published data in control
animals experiencing acidosis ............................................................. 72

Table 23

Table A-1

Table A-2

Table A-3

Table A-4

Table B-1

Table B-2

Table B-3

Table B-4

Table B-5

The effect of the addition of P. acidipropionici DH42
(10ll cfu) on the dry matter intake (DMI) of steers during
in vivo trials ....................................................................... 78

Data used for the analysis of volatile fatty acid production during
in-vitro fermentation studies with P. acidpropionici DH42 .................... 87

Data used for the analysis of recoverable numbers of
P. acidrpropionici DH42 during in-vitro fermentation studies ............... 89

A preliminary attempt at the recoverability of P. acidipropionici
DH42 from in-vitro anaerobic fermentors using rumen fluid from
a beef steer when plated on PBB with 1% erythritol and 1.5% agar ......... 91

A preliminary attempt at the recoverability of P. acidipropr’onici
DH42 from in-vitro fermentors using rumen fluid from a dairy cow
when plated on PBB with 1% erythritol and 1.5% agar .......................... 91

Data used for the analysis of fermentation end-products and pH for
in-vivo studies using P. acidrpropionici DH42 for the prevention
of experimentally induced acidosis in beef steers .................................... 93

Data used for the analysis of total plasma lactate, L(+) lactate, and

D(-) lactate concentrations and blood pH for in-vivo studies using

P. acidipropionici DH42 for the prevention of experimentally

induced acidosis in beef steers ............................................................... .95

The ammount of wheat necessary to provoke ruminal acidosis and

reduce ruminal pH below 5.0 when added as a wheat slurry (1 part

wheat:2 parts water) via rumen cannula to 4 steers during preliminary
studies .................................................................................................... 97

The amount of wheat used to induce experimental acidosis during

in-vivo trials when added as a slurry and prepared at 40 g/kg body
weight of steers ..................................................................................... .97

Data used for the analysis of Dry Matter Intake (DMI) for
steers during trial A and trial B in experimental acidosis ..................... 98

ix

REVIEW OF THE LITERATURE
LACTIC ACIDOSIS
MM!

Lactic acidosis in ruminants is a metabolic disorder often occurring in feedlot
cattle, as well as, in other ruminants when a large influx of readily fermentable
carbohydrates are suddenly consumed. It is characterized by a reduction in pH (4.0 to
5.0), increases in the concentrations of D (-) and L (+) lactic acids, and a disturbance in
the microbial population of the rumen. The population changes fiom a predominantly
gram (-) or lactate utilizing bacteria to a gram (+) or lactate producing organisms. In
addition, ruminal protozoa concentrations are reduced as the acid load increases (Owens
et al., 1998). The disorder can become systemic with changes in hematocrit levels (usually
hemoconcentration), and decreases in the bufl‘ering capacity of blood due to the surge of
lactate in the blood.

Energy intake and feed efliciencies are affected as decreased body weight and
variable intake patterns are observed. Diseases such as rumen ulcers, parakeratosis, liver
abscesses, and laminitis occur as a consequence of the acid accumulation in rumen. When
severe, the disorder is an overt illness characterized by anorexia, depression, dehydration,
recumbency, systemic acidosis, coma and death. Subclinical acidosis is less severe and
more dificult to detect and may be considered subliminal at times. Animals experiencing
subacute or subclinical acidosis often show no signs of an upset system. It is characterized
as a less overt and insidious illness therefore, causing great economic losses to the industry
in the forms of condemned livers, low weight gains, and wasted feed (Underwood, 1992;

Nocek, 1997; Nagaraja and Chengappa, 1998).

Acidosis can occur in any breed of cattle. Nevertheless, Brahman cattle have been
shown to be more susceptible to acidosis than the Angus breed and Holsteins appear to be
the most resistant to the disease (Elam, 1976). Weather changes, poor bunk management,
accidental access to concentrate feeds (Elam,l976), and muddy pens or feedlots
(Ritchie, 1984) are some of the other critical factors that have been identified as related to
acidosis.

Chronic and acute acidosis

There are two types of acidosis: chronic (subacute) and acute or clinical acidosis
(Blezinger and Mies, 1990). The pH levels used to diagnose acidosis of the rumen are 5.6
and 5.2 for chronic and acute acidosis, respectively. (Cooper and Klopfenstein, 1996
[cited by Owens et al, 1998]).

Chronic acidosis does not produce visibly ill animals but sometimes does result in
undetectable symptoms. It has been suggested (Britton and Stock, 1989) that variation in
feed intake might possibly be the only indicator of subacute acidosis. In contrast, others
have included a decrease in milk fat and milk yield (Allen and Beede, 1996) and a gray-
green pasty to watery consistency of feces (Dirksen, 1969) as other indicators of the
disease. Animals suffering from chronic or subacute acidosis usually are alert and do not
show a hesitation to move about (Dirksen,]969).

Acute or clinical acidosis is characterized as an illness following consumption of
readily fermented carbohydrates in amounts sumcient to reduce pH of ingesta. The
symptoms associated with this condition are total anorexia, increased water intake,
incoordination, loose feces or diarrhea, grinding of teeth, restlessness, and apathy. Heart

rate and rectal temperatures of cattle increase (100 to l40/min and 40 °C, respectively)

 

and respiratory rate increases to 40 to 60 breaths/minute (Dirksen, 1969; Howard,1981).
In addition, rumen motility also ceases (Slyter,1976; Elam,l976). As the disease
progresses, body temperatures fall to hypothermic levels (98 °F to 100 °F) and
hypoventilation, usually described as labored breathing, is evident with 10 to 20 breaths
per minute (Howard, 1981). If not treated within 12 to 24 h after the onset of acute
acidosis, recumbancy and coma can occur leading to eventual death (Johnson, 1991).
Etiology of acidosis

A variety of feedstuffs are available that can cause lactic acidosis. The
consumption of large amounts of grain, barley, corn , oats, and rye, or other sources of
feeds that are high in sugar including sugar beets with the beet tops attached, potatoes,
molasses, sugar cane, mangoes, and apples (Slyter,]97 6; Underwood, 1992) induce
acidosis. Other causative factors relating to occurrence of lactic acidosis is the industries
ever present need to incorporate economics into feed management. The desire to optimize
the rate of fermentation by making carbohydrates from feed more readily available allow
for several feed processing techniques that besides having a positive effect on nutritional
value, may actually have negative effects to ruminants. Those processing techniques
include steam flaking, rolling, cracking, crushing or grinding and in relation to forages,
their form and coarseness (Slyter, 1976; Dunlop,1972; Underwood, 1992). In a study
relating to the steam flaking of sorghum grain and among others, its effect on subacute
acidosis in feedlot steers, it was found that reducing flake density resulted in reduction of
rumen pH and a propensity towards acidosis (Reinhardt et al., 1997).

Lactic acidosis is more evident in those animals switched from a high forage diet

to a high concentrate diet. This was observed by Uhart and Carroll (1969), when they

2;)

changed ruminally fistulated steers on an all forage diet such as alfalfa, to a 90% grain
diet. This abrupt change produced all steers to go ofl‘ feed within 2 to 3 days. In contrast,
though, incrementing the daily available concentrate causes animals to go off feed, as well
as, the production of subtle symptoms. Lactic concentrations above 5 W are indicative
of acidosis and exceeding 40 mM indicate severe acidosis ( Owens et al., 1998).
Qomplicatiom

As a result of ruminal lactic acidosis, a sequel of other feedlot ailments occur that
bring about a decrease animal performance. Theses include rumenitis, liver abscesses,
laminitis, (Brent, 1976), grain bloat, and malabsorption (Britton and Stock, 1989).

The high acidity of lactic acid causes the rumen wall to become damaged. This
allows for the systemic invasion of bacteria into the portal blood system of the animal
causing deep ulcers in the rumen and abscessing of the liver thus, the terms rumenitis and
condemned livers. The bacteria responsible for the invasion and colonization of the rumen
wall is F usobacterium necrophorum (N ocek, 1997; Nagaraja and Chengappa, 1998).
Actinomyces pyogenes has also been implicated by others (Essig et al., 1988). This
colonization eventually causes entry of these bacteria into the blood stream and ruminal
wall abscesses. The bacteria present in the blood stream gain access to the liver by
filtration causing infection and abscesses. It has been suggested that 12 to 32 % of beef
cattle fed high or all concentrate diets may have abscessed livers (Brink et al., 1990).

Lactic acidosis has also been implicated in the incidence of laminitis in ruminants.
Laminitis can occur in acute and subclinical or chronic acidosis (Allen and Beede, 1996;
Nocek, 1997). Ruptured peripheral arterioles at the extremities caused by the release of

histamine and endotoxins from dying cells due to a decrease in rumen pH from ruminal

lactic acidosis, may be responsible for this condition as some theories state (Brent, 1976;
Mgassa et al., 1984). Signs of laminits seem to vary according to their severity. In severe
conditions, feet are warm to the touch, animals will be reluctant to stand, and pain will be
exhibited in all feet (Nocek, 1997). In chronic acidosis, animals will usually have
overgrown hooves; characterized by becoming more elongated, flattened, and broadened
(Nocek, 1997).

Another complication due to the onset of lactic acidosis includes a hardening of
rumen epithelium also known as parakeratosis (Nocek et al., 1984; Dirksen, 1989). This
usually occurs in mild, uncomplicated cases and causes a sloughing off of the epithelia on
the rumen wall during low pH related to a lactic acidosis episode. The epithelia heals and
forms blunt and clumped hardened papilla (Jensen and Mackey, 1979). This epithelial
hardening has shown to reduce volatile fatty acid absorption (Huntington and Britton,
1979). In a recent study (Krehbiel et al., 1995), absorption of acetate was found to be
13% lower for lambs experiencing acidosis than the control group, leading the workers to
conclude that acidosis affects the absorptionof volatile fatty acids.

Bloat is caused secondary to the lactic acidosis syndrome in ruminants. Gas is
normally produced in the rumen as a consequence of the fermentation process. Bloat is
characterized by the accumulation of this gas within the rumen in amounts suficient to
increase pressure in the rumen. This pressure causes distention of the rumen. In addition,
bacterial mucopolysaccharides such as slime can cause the ruminal contents to increase in
their viscosity contributing to a frothy appearance (Cheng et al., 1976). This froth or
foam prevents eructation adding to the distention of the rumen. Increased pressures in the

abdomen and thoracic areas of the animal cause blood from the viscera to be moved into

the peripheral blood system depositing more CD; into the plasma thus, contributing to an
already acidotic scenario (Blezinger and Mics, 1990).
The rumen microbial ecosystem and acidosis

The rumen is a highly complex and competitive microbial ecosystem. Population
densities of 1010 to 1011 have been found for bacteria, 105 to 107 for protozoa and smaller
amounts (< 104) of anaerobic fungi and facultatively anaerobic bacteria (Leedle, 1991;
McAllister and Cheng, 1996). It has an internal temperature of 37 °C and pH under normal
conditions of 6.8 to 7.0.

Rumen bacteria, in having the ability to ferment carbohydrates, are primarily
responsible for causing lactic acidosis in ruminants. Under a well balanced hay to
concentrate ratio, gram negative bacteria predominate (Dirksen, 1969). When the diet is
abruptly changed to all concentrate, a large influx of carbohydrate is suddenly available to
the rumen environment and gram positive bacteria dominate such as Streptococcus bovis
and the lactobacilli (Dirksen, 1969; Slyter, 1976; Goad et al., 1998). Streptococcus bovis
was first identified as playing an essential role in acidosis by Hungate in the 1950’s and
has been reviewed by others (Russell and Hino, 1985). Lactobacilli, have also been
implicated as growing in combination with the Streptococcus (Dirksen, 1969). Krogh
(1963), evaluated the rumen of cows and sheep suffering fi'om acute acidosis during
overfeeding and found a gram-positive flora predominating. The complete disappearance
of protozoa and cellulolytic bacteria has been observed as pH decreases from 5.5 to 5.0
with an eventual cessation of the bacterial population at a pH of 4.0. In a recent study
done by Goad et al. (1998), a comparison of two difl‘erent types of diets revealed that

amylolytic bacterial counts and lactobacilli counts were usually higher in grain adapted

 

steers after overfeeding with an all grain diet. In another study (Nagaraja et al., 1978),
data suggested that feeding grain to an animal adapted to forage diet caused the onset of
lactic acidosis, the proliferation of gram positive bacteria, the decrease in gram negative
bacteria, as well as, an increase in endotoxin. Endotoxin production was thought to be a
consequence of the lysis of gram negative bacteria and a subsequent decrease in their
numbers (Huber, 1976). However, in vitro incubations showed that there was no decrease
in the gram negative populations. Similar results were detected when after voluntary
engorgement of goats with a 90% concentrate diet, decreases in gram-negative bacteria
were not followed by increases in endotoxin concentrations (Suda, et al., 1997).

An overabundance of carbohydrate availability causes increases in the production
of acids by rumen bacteria (Slyter, 1976). The production of these end products could be
related to the growth of bacteria in the rumen (Dawson and Allison, 1988). When the
growth rate of Streptococcus bovis is low, end products such as acetate and propionate
are produced (Dawson and Allison, 1988). Nevertheless, when there is an abundance of a
carbohydrate source and the growth is high, then lactic acid is produced. The growth rate,
due to the excess availability of carbohydrates in the rumen environment, is not the only
mechanism by which some bacteria multiply to produce lactic acid. Low pH in the
environment has been found to trigger a shift in the metabolism of Streptococcus bovis
towards producing lactate by changing its regulation of lactate dehydrogenase (Russell
and Hino, 1985). This is known as the spiraling efl‘ect caused by Streptococcus bovis.
This action is responsible for the lowering of the ruminal pH (Dirksen, 1969). Because
they are sensitive to the acid they produce, they start to decrease in numbers and are

eventually overcome by the growth of more acid tolerant lactobacilli. Lactobacilli are gram

positive rods which can ferment sugars to lactic acid and produce optimum growth at a
pH of < 5.5 (Russel and Hino, 1985). Lactic acid produced by the lactobacilli is enough to
cause a final pH of 4.5 causing decreases in other microbial populations in the rumen with
an almost complete dominance of lactobacilli species in the rumen (Dawson and Allison,
1988). Nevertheless, the growth of the lactobacilli and the acid sensitivity of
Streptococcus bovis, does not appear to be associated only with a decrease in
Streptococcus bovis numbers. A recent study suggests the possibility of an antagonistic
mechanism in the lactobacilli which could also be afi‘ecting the populations of
Streptococcus bovis (Wells et al., 1997). In a study where there was a modest decline in
pH (< 5.6), Streptococcus bovis diminished in numbers and the lactobacilli increased. It
was suggested by the researchers that the pH alone could not account for the reduction in
Streptococcus bovis and that certain strains of lactobacilli could possess certain
bacteriocins, inhibiting their growth (Wells et al., 1997).

Gram negative bacteria including the lactate utilizers and protozoa decrease in
numbers and eventually die (Slyter, 1976) as the pH is affected by the heavy lactate
production from Streptococcus bovis and the lactobacilli. Normally, a feeding of diets
containing carbohydrates causes a temporary accumulation of lactic acid with eventual
conversion to acetic and propionic acid by rumen microorganisms. However, lactate
utilizing bacteria usually find difficulty in multiplying to efi‘ective numbers if there is a
sudden surge of lactic acid to the system (Mackie and Gilchrist, 1978, 1979).
Megasphaera elsdenii (considered by Slyter, 1976) as one of the most important lactate
utilizers), produces propionate and is usually able to compete with the demand of lactate

produced but only when the rumen is not overloaded with unreasonable amounts of

carbohydrates (Nocek, 1997). It utilizes 60 to 80 % of the lactate fermented in the rumen
(Counotte et al., 1981) and makes up 20% of the lactate utilizers in ruminants fed high
concentrate diets (Mackie et al., 1984). Megasphera elsdenii at 107 cfu/mL was able to
increase pH and reduced lactic acid concentrations (> 3 mM) in ruminal fermentation
studies in vitro involving other lactate utilizers (Hession and Kung, 1992). In another in
vitro study done by Kung and Hession (1995), two levels of Megasphera elsdenii were
tested on pH, lactate, and VFA concentrations. In relation to the lactic acid utilization, it
was found that the two levels tested (106 and 10s cfir/mL) were efl‘ective in utilizing it.
Other lactate utilizers found in the rumen include Selemonas ruminantium, Veillionella
alcalescens and Propionibactertum acnes (Slyter, 1976).

Yeasts and anaerobic fungi make up part of the microbial population of the rumen.
However, most of them are usually transient as they enter the rumen along with the feed.
Several of these organisms have been found to ferment carbohydrates such as the
anaerobic fungi Neocalimastixfrontalis, and Piromonas comum’s. The end-products
produced by these fungi are lactate and acetate (Orpin and Munn, 1986 [as cited by Orpin
and Joblin, 1988]). Yeast have also been found in the rumen and interestingly, they have
been found surviving at the low pH produced during acidosis (Dirksen, 1969).

Under normal conditions, lactate does not accumulate in the rumen at levels
exceeding 5 uM, but concentrations in the rumen of more that 40 mM indicate the onset of
severe acidosis (Owens et al, 1998). Bacteria in the rumen produce two forms of lactic
acid, the D(-) and the L(+) form. The L(+) form is identical to the lactate produce in

muscle tissue from glucose and the D(-) form is not. At a high pH (6.0), 20% of the total

lactate produce is attributed to the D-isomer (Dawson and Allison, 1988). At a pH lower
than 5.0 with lactate concentrations above 100 mM, the production of the D-isomer may
be as high as 50% of the lactate produced (Dawson and Allison, 1988). Lactate in the
rumen is produced by the catabolism of glucose via the Embden-Mayerhof-Parnas
pathway as proposed by Mountfort and Robertson (1978) after extensive studies with
pure cultures.

As mentioned previously, the enzyme responsible for the production of lactate in
Streptococcus bovis is lactate dehydrogenase. When growth rates in bacteria are low, the
concentrations of pyruvate within the cell are low and the enzyme is inactive. Pyruvate is
then converted into acetate or propionate via the Embden-Mayerhoff-Parnas pathway.
However, when energy is abundant and the growth rate of microorganisms intensifies,
pyruvate increases and lactate dehydrogenase shifts production towards lactate. D(+) and
L(-) lactate possess specific dehydrogenases. L(-) lactate is associated with L-lactate
dehydrogenase which can be found in the cytosol of animal tissues and D-2-hydroxy-acid
dehydrogenase which is associated with the D(+) lactate. Giesecke and Stangassinger
(1980), found that D-lactate must pass the mitochondrial membrane before it could be
oxidized by the D-2-hydroxy-acid dehydrogenase. The slow degradation of D-lactic acid
and the toxicity it produces during lactate accumulation, could be attributed to its
oxidation pathway (Dawson and Allison, 1988) and is responsible for the accumulation of
lactic acid in the rumen of cattle experiencing acidosis (Slyter, 1976).

Volatile fatty acid production in the rumen is attained primarily by microbial
fermentation. The primary volatile fatty acids produced in the rumen are acetic, propionic,

and butyric acid (Gottschalk, 1986; Hungate, 1988). As the foragezconcentrate ratio

10

 

decreases, the acetate:propionate ratio also decreases. For example, foragezconcentrate
ratios of 100:0, will give volatile fatty acid ratios of acetate, propionate, and butyrate of
71 : 16:7 .9. When the forage to concentrate ratio is 50:50, the molar ratios of acetate,
propionate and butyrate are 65.5: 18.4: 10.4. When there is more concentrate ofi'ered in
the diet (80:20), the volatile fatty acid ratios would be 53.62306: 10.6 (Annison and
Armstrong, 1970). In a study done by Goad et al. (1998) on hay adapted and grain
adapted steers, ruminal acetate proportions decreased over time in both groups and was
lower in the grain adapted steers at 60 h. The production of propionate increased with
time in both groups and was higher in the grain adapted steers at 72 h post-feeding. Goad
and Nagaraja (1988) have suggested that the volatile fatty acids may be responsible for the
low pH seen in subacute acidosis because lactate levels were low (< 5 mM) in contrast to
acute acidosis where lactate is seen in greater concentrations. Krehbiel et al. (1995),
studied the efl‘ects of increasing severity of acidosis on rates of absorption of ruminal
volatile fatty acids and found that the concentrations of ruminal volatile fatty acids in
lambs intraruminally dosed with glucose responded individually as the amounts of glucose
(0, 6, 12, and 18 g/kg) increased. Acetate concentrations decreased (63.3, 46.6, 37.5 and
37.7 mM), as glucose dose increased, propionate showed an increase of 21 .0 and 43.6 mM
at 0 and 6 g/kg, respectively with a decrease from 12 to 18 g/kg of glucose infused.
Butyrate tended to decrease (15.0, 8.5, 7.5 mM) when glucose was infused at 0, 6, and 12
g/kg and increased (14.0 mM) at 18 g/kg infused glucose. Amongst the parameters
studied by Patra and Swarup (1996) on studies with experimental acidosis in sheep, total
volatile fatty acid concentrations abruptly increased fiom 44.16 mmol - L'1 to 83.00

mmol - L’1 in 12 h after overfeeding. They attributed this to the rapid fermentation by the

starch degrading bacteria of the rumen (as discussed previously). Total volatile fatty acids
24 h after the engorgement, decreased to 57.60 mmol - L'l, leading them to deduce that the
decrease in total volatile fatty acids was caused by an increased absorption rate at a low
ruminal pH.

Before volatile fatty acids can be produced, carbohydrates must first pass through
the pyruvate intermediate. End product formation primarily depends on the type of
structural carbohydrate fermented and the type(s) of bacterial species involved in the
fermentation process.

Acetate production is formed by the pyruvate-formate lyase system. In this
system, formate and acetyl-coenzyme A (CoA) are formed as the intermediates. The
formate is converted to carbon dioxide and H2 by other bacteria. This is the main pathway
for acetate production in the rumen (Fahey and Berger, 198 8). Another pathway involved
in the production of acetate in the rumen utilizes pyruvate ferredoxin oxidoreductase. It
produces ferredoxin, which is the acceptor for some bacteria (Glass et al., 1977), carbon
dioxide and acetyl-CoA. In either pathway, the resulting end product is acetate and
bacteria such as Megasphera elsdenii and VeilIonella alcalescens have been observed
utilizing the ferredoxin-linked pyruvate oxidoreduction pathway (F ahey and Berger, 1988;
Russell and Wallace, 1988).

Propionate production is primarily via the dicarboxylic acid pathway. The
decarboxylation of succinate by Selenomonas ruminantium is the primary method of
propionate production in the rumen of cattle (Wolin and Miller, 1988). It must first
produce succinate as an intermediate. A carboxyl group is then removed from the

succinate by succinyl-coenzyme A ultimately leading to the production of propionate.

12

Nevertheless, there is an alternate pathway for the production of propionate. It has been
named the acrylate pathway and bacteria such as Megasphera elsdenii have been
implicated in utilizing it for the production of propionate (Paynter and Elsden, 1970). In
this pathway, succinate nor succinyl-CoA are used as intermediates. Instead, pyruvate is
converted to either form of lactate (L-, D-, or DL-) which is then converted to acrylyl-
CoA and reduced to propionyl-CoA (Gottschalk, 1986). One third of the total propionate
produced is via the acrylate pathway (Fahey and Berger, 1988). Nonetheless, the
production of propionate is of vital importance to the ruminant because of its involvement
in gluconeogenesis. Propionate is the only volatile fatty acid which makes contribution to
production of glucose in the ruminant animal (Fahey and Berger, 1988; Wolin and Miller,
1988). One study suggested that 27-54% of glucose is produced from propionate
(Lindsay, 1970).

Butyrate production is mainly via the reversal of the B-oxidation pathway ( Fahey
and Berger, 1988). In this pathway, acetacetyl-CoA, L(+)-B-hydroxybutyrly-COA, and
crotonyl-CoA are intermediates with butyryl-CoA being formed proceeded by butyryl
phosphate; thus giving rise to butyrate (Gottschalk, 1986).

METHODS UTILIZED IN MAN IPULATING RUMEN FERMENTATION
Introduction

The growing public concern over the use of antibiotics in the animal feed industry
to control rumen fermentation, diseases, and improve animal efficiency has lead
researchers to probe into the use of less artificial methods in animal feeds. Antibiotics
have been used as feed additives for farm animals since the second world war (Pusztai, et

al., 1990). It has been the repeated evidence of their beneficial effects on the grth and

13

health of livestock that has made them a popular choice in the industry. Nevertheless, the
use of antibiotics is still widely accepted in the industry but carries the risk of bacterial
strains becoming resistant and therefore afl’ecting future therapeutic uses in both man and
animal. As pressure by the public, media and the medical community increases, it would
be correct to state that in the 21" century, the world could be seeing the elimination of the
use of antibiotics in all livestock feed with an increase in the use of live feed supplements
such as rumen bacteria, fimgi and yeast.

In the next section, the various methods or additives used to modify rumen
fermentation activities and its microbial population along with the use of antibiotics will be
discussed.

Ionophores and non-ionophores

There are two types of antibiotics which are used in the livestock feed industry:
ionophore and non-ionophore antibiotics. Ionophore antibiotics are bacteriostatic and not
bactericidal (Nagaraja and Taylor, 1987). They are known for their ion bearing properties
(Bergen and Bates, 1984 [as cited by Wallace, 1992]), and it is their ability to permeate
the cell envelope of different species to different extents that is the basis of their selective
action (Wallace, 1992) and bacteriostatic activity. An example of an ionophore would be
monensin as a Na+ , K7 If antiporter (Wallace, 1992). Non-ionophore antibiotics do not
act on cation transport and difl‘er in their chemistry (ie. glycopeptide, tetracycline,
polypeptide), antibacterial spectrum, molecular weight and if they are absorbed from the
gut or not (Nagaraja, 1995). For example, avoparcin is a glycopeptide which inhibits cell
wall synthesis by inhibiting the incorporation of N-acetylglucosamine into the

peptidoglycans of the bacterial cell walls (Wallace, 1992; Jouany, 1994).

14

The efi‘ectiveness of using ionophores is in an increased efficiency of feed
conversion and an improvement in the rate of gain (Bauer et al., 1992; Herold et al., 1994;
DiConstanzo et al., 1997). These two effects are primarily attributed to the changes in
ruminal fermentation patterns as a consequence of the incorporation of ionophores
(Nagaraja, 1995) in feeds. According to Bergen and Bates (1984), amongst the three
major areas associated with ionophore feeding are: increased production of propionate and
decreased lactic acid production and froth formation in the rumen leading to decreased
ruminal upsets. The increased production of propionate by the use of ionophore feeding
may offer the animal considerable advantage over other end products such as acetate
because of its involvement in gluconeogenesis. There have been various studies where the
use of ionophores has lead to an increase in propionate production. Nagaraja et al. (1981)
induced acidosis in sheep treated with either lasalocid or monensin and found an increase
in propionate in treated animals after grain engorgement. In-vitro studies with salinomycin
was shown to enhance propionate production, but in-vivo studies in cattle using the same
ionophore, showed that propionate increased for 48 h then gradually declined (N agaraja et
al., 1985). Monensin in cattle has also shown to be beneficial in propionate production
(Burrin and Britton, 1986) and in-vitro fermentation studies of soluble carbohydrates
treated with monensin caused a 40% increase in molar proportions of propionate (Tung
and Kung, 1993). In contrast, studies with laidlomycin propionate reported no significant
influence on concentrations of volatile fatty acids including propionate (Herold, et al.,
1994; Bauer et al., 1995). It is thought that the increase in propionic acid is due to a
redirection in the utilization of hydrogen and a decrease in the production of methane

(Nagaraja, 1995; Jouany, 1994) in animals fed ionophores.

15

Ionophores have also been shown to reduce ruminal upsets. Because cereal grain
is one of the major components in feedlot diet, it has had a notoriety for causing situations
in which rapid fermentation of the feedstufl‘s has lead to the accumulation of organic acids
in the rumen. Lactic acidosis causes major economic loss to the ruminant livestock
industry. Ionophores are selective toward gram positive microbes and the major lactate
producing bacteria such as Streptococcu bovis and the Lactobacilli spp. , but do not affect
the lactate utilizers (Dennis et al., 1981). Nagaraja et al. (1985) have reported significant
reductions in lactate production and acidosis when ionophores have been administered
during a high carbohydrate diets. In a study done by Nagaraja et al. (1981), lasalocid and
monensin were administered into the diet and their effects on lactic acidosis in cattle were
observed. Antibiotic treated cattle showed higher rumen pH and lower L (+) and D (-)
lactate concentrations. Interestingly, administering the antibiotics 7 days before
experimentally inducing acidosis with corn (27.5 g/kg body weight) prevented acidosis
while administering antibiotics 2 days before the glucose induced acidosis was not enough
to prevent acidosis. The authors suggest this might be due to the differences in
carbohydrates used. Salinomycin was compared against the popular monensin and
lasalocid in preventing experimentally induced lactic acidosis (N agaraja et al., 1985). All
showed an increase in pH and a decrease in the lactate isomers, but salinomycin was
shown to be a least 3 times more powerful than monensin or lasalocid. Monensin alone
was studied to evaluate its response in cattle to subacute acidosis (Burrin and Britton,
1986). In this study, monensin was able to maintain a higher ruminal pH by reducing

concentrations of volatile fatty acids. Subacute acidosis is characterized as a less severe

l6

 

acid load with increases in volatile fatty acids and is thought to be the most common form
of acidosis in grain fed ruminants (Nagaraja, 1995).

Ionophore antibiotics are extremely effective against gram positive organisms and
impose no effect on gram negative bacteria. This was the outcome of an in-vitro study
conducted by Cheng and Wolin (1971). In this study the efi‘ect of monensin and lasalocid-
sodium on the growth of gram positive and gram negative bacteria was evaluated. The
results showed that Bacteroides ruminicola and Selenomonas ruminantium, two gram
negative rumen bacteria, were insensitive to either ionophore causing their proliferation.
The gram positive organisms utilized in this study, Ruminococcus albus, and R
flavefasciens, were inhibited and Butyrivibriofribisolvens, although possessing a gram
negative stain, but having gram positive cell wall structure, was inhibited as well. It was
also suggested by these workers that the selection of monensin and lasalocid-sodium for
gram negatives that produce succinate to propionate could lead to a beneficial propionate
formation in the rumen. Equally postulated, selection against the hydrogen and formate
producing gram positives used in this study, would lead to a decrease in the undesired and
wastefirl methane production in the rumen.

In essence, ionophore antibiotics inhibit gram-positive bacteria such as
Lactobacillus and Streptococcus and those that have a gram-positive cell wall structure
but stain gram-negative such as Butyrivibrio and Ruminococcus. Gram-negative bacteria
such as Megasphaera, Ruminobacter, Selenomonas, and Veillonella species, are usually
resistant to ionophores. The ruminal bacteria sensitive to ionophores produce lactate,

butyrate, formate or hydrogen and the bacteria which are resistant produce succinate or

17

 

propionate as the end products of fermentation (Cheng and Wolin, 1979; Nagaraja and
Taylor, 1989 [table 5; as cited by Nagaraja, 1995]).

Non-ionophore antibiotics are antibiotics which do not act on cation transport
across the cell membrane, as mentioned previously, yet their antibiotic activity is on the
inhibition of cell wall synthesis (Wallace, 1992). Such antibiotics include avoparcin,
virginiarnycin, thiopeptin, and tylosin. Nevertheless, in order for these antibiotics to be
accepted in livestock feed, they must not be absorbed from the gut or partially absorbed at
a low dosage; primarily because of the residues in milk and meat and because a withdrawal
time would not be required. The molecular weight of these non-ionophore antibiotics
have much to do with their absorption from the gut. For example, avoparcin, because of
its molecular weight in excess of 1000 (Table 13.8; Nagaraja, 1995), would not be
absorbed from the gut and thus, would not require a withdrawl period. In contrast,
virginiarnycin, because of a molecular weight of 525 (Table 13.8; Nagaraja, 1995), would
require a withdrawal period because of the antibiotic residues that would be detected in
either milk or meat products.

Non-ionophore antibiotics generally improve growth and enhance feed efficiency
and as in ionophore antibiotics, the response is mainly attributed to its effect on the rumen
flora. There have been many modes of action postulated (N agaraja, 1995) to account for
the grth effect that these antibiotics have on ruminants. One mode of action pertains to
the modification that non-ionophore antibiotics have on rumen fermentation. These act
directly on the microbial population to improve the fermentation efliciency. This is an
important factor when relating to digestive upsets in the rumen such as lactic acidosis from

the feeding of starch rich feeds. Muir and Barreto (1979) tested several antibiotics in their

18

 

ability to inhibit Streptococcus bovis. Amongst the many tested, thiopeptin was among
the most efiecfive antibiotics at inhibiting this microorganism. Thiopeptin is a sulfur—
containing peptide produced by Streptocmyces tateyamensis and has been shown to
possess strong activity against gram positive bacteria (Miyari et al., 1972). This prompted
Muir et al. (1980) to study the induced ruminal lactic acidosis in sheep fed wheat with or
without thiopeptin and related antibiotics. Their results indicated that thiopeptin given in a
single dose along with the wheat on the day of the challenge, prevented lactic acidosis
completely as it reduced rumen lactate concentrations by 80 to 90%. They also noticed a
normal rumen fermentation pattern during wheat supplementation with thiopeptin as an
increase in volatile fatty acids were noted. Penicillin was also tested for its eficiency in
preventing acidosis but was not found to be efi‘ective because of its broad spectrum
antimicrobial activity affecting even some gram negatives (Flulghum et al., 1968 [as cited
by Muir and Baretto, 1979]).

Virginiarnycin is a fermentation product of Streptomyces virginiae. It has been
shown to have antibiotic effects against gram positive bacteria (N agaraja et al., 1987). A
single drench of virginiamycin was evaluated in two trials for its ability to protect sheep
against lactic acidosis when fed wheat ad libitum (Thomiley et al., 1998). In this study, an
increase in pH was observed in sheep drenched with virginiamycin than in the control
sheep with an increase in weight and propionate concentrations were increased on the day
after drenching and exposure to the wheat while acetate and butyrate decreased. The
workers in this study concluded that sheep drenched with virginiamycin at 40 to 80 mg did
not become acidotic and that a single drench of the antibiotic was able to efl‘ectively

prevent lactic acidosis. Ruminal bacterial species were tested for their susceptibility and

19

 

resistance towards virginiamycin in studies done by Nagaraja (1987). In this study,
virginiamycin was inhibitory to the two major lactic acid producing bacteria, S. bovis and
Lactobacillus sp., responsible for lactic acidosis in ruminants (Slyter, 1976). The ability
for virginiamycin to inhibit lactic acid production in vitro was again confirmed in a study
done by Nagaraja et al. (1987). In this study, viginiarnycin inhibited lactic acid production
by 90 to 93% at the highest concentration (24.0 ug/mL), but decreased propionate
production when concentrations were < 6.0 ug/mL .

Avoparcin, produce by the yeast Streptomyces candidus, inhibits gram-positive
bacteria as it disrupts the peptidoglycan synthesis of the cell wall. It is presently not
approved in the United States, but may be used in Europe (N agaraja, 1997).

Nevertheless, in a study done by Nagaraja and Taylor (1987) avoparcin, was found to be
less inhibitory to S. bovis than the other non-ionophore compounds except tylosin. They
suggested that avoparcin might not be active against all gram-positive bacteria (Walton,
1978 [as cited by Nagaraja and Taylor, 1987]). Nagaraja et. al (1987) reported that out of
the non-ionophores tested, avoparcin increased the molar proportion of propionate with
increasing concentration (ug/mL) and was the least efl‘ective in reducing lactic acid
concentrations. This is in contrast to the study done by Nagaraja and Taylor (1987) which
reported a less inhibitory effect of avoparcin towards lactic acid producing bacteria.
Because avoparcin is also favored as a growth promotant, Dyer et al. (1980), evaluated
amongst the many parameters, growth rate and volatile fatty acid concentrations in cattle
fed a 77% barley diet. They reported cattle consuming less feed per unit gain (P < .05)
than control cattle and reported a trend in the increase of propionate with increasing levels

of avoparcin.

20

Tylosin is produced by Streptomycesfi'adiae (Berkman et al, 1961 [as cited by
Nagaraja, 1997]). Its antibiotics effects are on gram-positive and some gram-negative
bacteria (N agaraja and Taylor, 1987). Tylosin in combination with monensin is used in the
cattle industry for reducing the incidence of liver abscesses. The organism attributed to
the onset of liver abscesses is F usobacterium necrophorum. The organism is gram-
negative and tylosin has the ability to destroy some gram negative organisms as
mentioned. Tylosin showed some activity towards gram positive lactate producers such as
S. bovis and Lactobacillus sp. although not as strong as some other antibiotics tested
(Nagaraja and Taylor, 1987; Muir and Baretto, 1979). Tylosin showed an extreme
efi‘ectiveness in reducing lactic acid accumulation in vitro and showed a tendency to
increase the molar proportion of propionate. In contrast, tylosin at high concentrations
(>6.0 ug/mL) decreased the proportion of propionate. In studies with feedlot steers fed a
60% barley diet, Norton and Nicholson (1980) reported a decrease (P < .05) in liver
abscesses in steers fed tylosin, no beneficial efieas of the antibiotic on propionate
concentrations, and no major effect on gain or feed eficiency. This lead the researchers to
conclude no beneficial effects in using tylosin in the diets under their experimental
conditions. In contrast, the feeding of tylosin has been shown to increase weight gain and
feed efficiency in feedlot cattle (Potter et al., 1985).

Microbial feed additives

Much confusion has been associated with the use of the popular term “Probiotics”
in reference to bacteria, yeast and firngi (Martin and Nisbet, 1991). Nevertheless, in light
of the confirsion, the Food and Drug Administration in 1989, required manufactures to use

the term Direct Fed Microbials (DFM) and defined DMF’s as “a source of live (viable)

21

naturally occuring microorganisms”; this includes yeast, firngi, and bacteria (Miles and
Bootwalla, 1991 [as cited by Martin and Nisbet, 1991]).

Much work has been done with yeast and fungal cultures in benefiting the
microbial fermentation and eficiency in the rumen and has been reviewed extensively
(Williams and Newbold, 1990; Martin and Nisbet, 1991; Wallace, 1992; Kmet et al., 1993;
Jouany,1994; Newbold, 1995; Yoon and Stern, 1995; Girard, 1996; Chesson and Wallace,
1996). Yeast and fungi supplementation is primarily utilized in adult ruminants because of
their effects on feed intake and in preventing lactic acidosis (Chesson and Wallace, 1996).
However, the two main organism commonly used have been Saccharomyces cerevisiae
and Aspergillus oryzae (yeast and filamentous fiingus, respectively) (Chiquette, 1995).

Harrison et al. (1988) utilized a yeast culture supplement composed of
Saccharomyces cerevisiae to evaluate its effect on ruminal metabolism and digestibility in
dairy cattle. Amongst the parameters observed in this study, a low pH was reported along
with increases in the molar proportions of propionate. Anaerobic bacteria and cellulolytic
bacterial counts also showed an increase in the supplemented cows as opposed to the
controls. However, an in vitro study by Kung et al. (1997) revealed that supplementing
continuous fermetors with Saccharomyces cerevisiae on a 50:50 concentratezforage diet,
showed no effects on major fermentation acids (acetate and propionate) nor on the pH.

The in vitro effects of this yeast’s ability to influence lactate utilization of ruminal
bacteria have been studied. Saccharomyces cerevisiae (Levucell SC) was evaluated for its
efi’ect on lactate metabolism by S. bovis and Melsdem’ ( Chaucheyras et al., 1996). In
this study, the grth of the yeast and S. bovis in coculture reduced the lactate production

by the bacteria, suggesting less glucose availability for the bacteria. Saccharomyces

22

cerevisiae stimulated the utilization of L-lactate by M. elsdem'i with the effect dependent
on the concentration of yeast used. When S. bovis and M. eIsdem'i were grown in the
presence of S. cerevisiae, lactate reduction was improved. The fermentation products
produced by the bacteria were also increased in this study as a consequence of the yeast.
Researchers concluded S. cerevisiae to be efl‘ective in reducing lactic acid accumulation in
vitro.

It has been suggested that the mode of action of yeast in stimulating ruminal
fermentation is due to the removal of 0; from the ruminal environment which would
stimulate the growth of the strictly anaerobic bacteria (Rose, 1987 [as cited by Newbold et
al, 1996]). Newbold et al. (1996) investigated this theory and reported that S. cerevisiae
and a commercial preparation Yea-Sacc, increased the rate of oxygen disappearance fi'om
fermentors by between 46 and 89%.

The combination of S. cerevisiae and A. oryzae have been studied by some to
varying extents (Chiquette, 1995). When S. cerevisiae and A. oryzae were used alone or
in combination as a feed supplement for beef and dairy cattle, higher concentrations of
acetate, propionate and total volatile fatty acids were reported and a lower pH was
reported. In this study, bacterial counts were not affected nor their ability to colonize feed
particles. The addition of S. cerevisiae did not afi‘ect milk yield but the combination of
the yeast and fungi showed a positive effect.

Bacterial probiotics are primarily used in young ruminants principally during their
early days of life in order to reduce or prevent digestive upsets and promote the
development of normal rumen anaerobic flora (Kmet et al., 1993). The primary bacteria

used in feed for calves are microorganisms such as Lactobacillus, Enterococcus,

23

Bifidobacterium, and Bacillus spp. For example, in a study done by Bondai et al. (1986),
the incidence of diarrhea was reduced and a decreased count of fecal coliforrns in the
intestines were reported in studies by Gilliand et al.(l980) when Lactobacillus acidophilus
was used in calves. Beeman (1985), tested the effects of Lactobacillus acidophilus on
Holstein male calves with a history of diarrhea, and reported weight gains of 8 kg as
opposed to the 3.5 kg gain by control calves two weeks after the treatment with the
lactobacilli. Lactic acid bacteria have been suggested to have pH lowering efl‘ects and
alterations in the fermentation activities in the rumen of young ruminants (Kmet et al.,
1993; Chesson and Wallace,1996). For example, young Holstein calves benefited from a
yogurt containing L. acidophilus supplement as they showed rumination activities at 30
days after birth (Chesson and Wallace, 1996). Pollman (1985) evaluated the use of a
probiotic product containing L. acidophilus on the weight gain of calves convalescing
from neonatal diarrhea when treated with antibiotics. It was noted that on average, weight
was increased 17.7 lbs (8 kg) in treated animals as compared to 7.8 lbs (3.5 kg) in control
calves and coats were healthier in appearance. Lactic acid is used by the calf as a nutrient
and it promotes a decrease in the pH of the intestine making it a less desirable place for
pathogenic organisms such as E. coli to grow (Pollman, 1985).

Although mainly fungal additives are primarily used in adult ruminants (Kmet et
al., 1993; Yoon and Stern, 1995), recent attention has turned to the use of bacterial
organisms to enhance ruminal microbial fermentations and performance. Wiryawan and
Brooker (1995) were able to control acute lactic acidosis in sheep when an inoculum of
103 cfu of Selenomonas ruminantium preceded an acute grain feeding and the efi‘ect of

combining Selenomonas ruminantium and Megasphaera elsdem'i was shown to be even

24

more beneficial in combating the efl‘ects of an acidosis challenge. In this study, pH
remained high and rumen lactate levels were undetectable in sheep treated with
Selenomonas ruminantium. In sheep treated with both microorganisms, marked increases
in acetate, propionate and butyrate were observed. Huffman et. al. (1992) evaluated the
use of Lactobacillus acidophilus in steers experiencing subacute acidosis. A 108 cfu/d
inoculum reduced the amount of time that ruminal pH was below 6.0. In contrast,
Klopfenstein et a1 (1995), reported that Lactobacillus acidophilus had no influence on
subacute acidosis in cattle. The efl'ect of Megasphaera elsdenii in preventing lactate
accumulations in vitro was evaluated in a study done by Kung and Hession (1995). A
high and a low dose of M elsdenii revealed its ability to prevent an accumulation in lactic
acid and excessive drops in pH. The low dose consisting of 8.7 x 105 cfir, showed lactate
concentrations of 5 mM by 7 hours of fermentation and the high dose (8.7 x 106 cfir) held
steady at 2 mM. Propionate and acetate concentrations decreased after 4 h of
fermentation and butyrate increased. Additionally, the ability of M. elsdenii to withstand
an additional substrate challenge was evaluated as half of the fermentation media was
inoculated into a fresh substrate without the addition of M. elsdenii. Lactate
concentrations remained low and it was concluded that M. elsdem'i has the potential to
prevent the accumulation of lactic acid in high carbohydrate diets. In a study done by
Hession and Kung (1992), M. elsdem'i and Propionibacteria shermanii were evaluated for
their effects on rumen fermentation. In their preliminary studies, it was observed that high
(109 chL culture) and low (1010 cfu/mL culture) inocualtion rates of P. shermam'i
tended to decrease L-lactic acid concentrations when compare to untreated cultures, but

pH remained low. When P. shermam'i was inoculated at 108 cfu/mL culture pH remained

25

at 4.82, lactic acid levels rose to 30 mM, and higher levels of propionic and acetic acids
were reported. Inoculations of M elsdem'i at 106 cfir/mL culture showed and increase in
pH (5.4) five hours after treatment with lactic acid levels at 25 mM and after 7 h decreased
to less than 5 mM. M. elsdem'i at 107 chL culture held pH at 5.43 after 5 h of
treatment and lactic acid levels remained at 3 mM throughout the entire experiment. It
was concluded that P. shermam‘i would not be acceptable for treating acidosis because it
has a tendency to decrease pH due to the production of propionic and acetic acids. M.
elsdem'i would be a much better choice because of its production of weak acids that would
moderate the decrease in pH. M. elsdenii was also tested in vivo in a trial conducted by
Robinson et al. (1992). In this experiment, M. elsdenii was used to test its efl‘ectiveness in
preventing acute acidosis and in enhancing feed intake in cattle. M. elsdenit was
inoculated in rates of 1012 cfu/animal for three consecutive days before during and after the
acidosis producing feed was fed. It was found that feed intake was increased 24% more
than untreated animals, and workers found significant interactions between treatments and
days (P < 0.2) for ruminal pH, lactate, butyrate, valerate and total volatile fatty acids. It
was concluded that using .M. elsdem'i as an inoculant can accelerate the adaptation of
ruminants to high concentrate feeds.

Recently, attention has been brought to the possible manipulation of the rumen
microbial ecosystem with bacteriocins as another means of improving ruminant production
(Teather and Forster, 1998). Bacteriocins are proteinaceous “antibiotics” produced by
some bacteria (Teather and Forster, 1998) which have a common mode of action; to
interfere with energy transduction and membrane transport processes (Montville and

Bruno, 1994). Most of what is known about them comes fi'om gram (+) bacteria and they

26

are very resistant to inactivation unlike proteins; they are active at pHs as low as 2 and at
temperatures as high as 100 °C (Jack et al., 1995).

There has been limited work done in discovering rumen organisms capable of
producing bacteriocins. Iverson and Mills (1976) suggested that S. bovis could be a
probable producer of a bacteriocin. Staphylococcus and Enterococcus isolated from
calves were also shown to posses bacteriocin properties (Laukova et al., 1993; Laukova
and Marekova, 1993). Ruminococcus albus has also shown to produce a bacteriocin like
inhibitor (Odenyo et al., 1994). Nevertheless, these results suggest the presence of
bacteriocins in the rumen. After an extensive survey of 50 Butyrivibriofibrisolvens
isolated, 25 were found to have some sort of bacteriocin activity and one strain, AR] 0,
has been completely defined as having its bacteriocin like activity in a single peptide with a
molecular weight of approximately 4000 Da (Kalmokoff and Teather, 1996). It has been
named Butyrivibriocin AR10 and represents the first bacteriocin isolated from a rumen
anaerobe (Teather and Forster, 1998).

THE PROPIONIBACTERIA

WM

Propionibacteria belong to the recently proposed class Actinobacteria (Stackenbrandt et
al., 1997). They are generally described as pleomorphic rods with one end rounded and
the other tapered or pointed. The cells tend to arrange themselves in V or Y
configurations and are characteristically described as “Chinese characters”. They may be
arranged in pairs, short chains, or may occur singly.

Propionibacteria stain gram positive. They are non-motile, non-sporeforming,

catalase-positive, facultative anaerobes. Their catalase-positive reaction is contradictory

27

to what is generally observed in anaerobic bacteria; anaerobic bacteria are usually catalase-
negative, but the former has been shown to occur (Vorob’eva et al., 1968 [as cited by
Hettinga and Reinhold, 1972]). Cell colonies may appear white, gray, red, yellow, cream
or orange and the temperature range at which cells have been observed to grow rapidly is
between 30 to 37 °C (Cummins and Johnson, 1986).

Propionibacteria are divided into two groups according to their habitats: the
classical and the cutaneous strains. The classical strains include those bacteria found in
cheese and dairy products (Grappin et al., 1999). However, they have also been found in
silage ferrnentations and in fermenting vegetables (Babuchowski, et al., 1999; Merry and
Davies, 1999) and are known as Propionibacterium fieudenreichii, P. jensenii, P. thoenii,
and P. acidipropionici. The cutaneous strains include those bacteria that are found on
human skin. One acne bacillus, originally described as a C orynebacterium is an example
of this group and is referred to as the “acne group strains” or the “cutaneous
propionibacteria (Cummins and Johnson, 1986). These cutaneous bacteria are known as
Propionibacterium acnes, P. avidum, and P. granulosum.

There are distinctive characteristics amongst the morphology of these two types of
propionibacteria. The classical propionibacteria tend to exhibit a shorter and thicker rod
while P. acnes tends to show a longer and slender irregular rod (Cummins and Johnson,
1986). Nevertheless, there may be variability in the morphology from strain to strain
especially in the early log phase. For example, strains of P. acnes may be long and
irregularly slender (as discussed previously) in a young culture (early log phase), but in

older (post log phase) cultures, strains are more coccal (Cummins and Johnson, 1986).

28

MMlic pathways

The metabolism and grth of propionic acid bacteria has been reviewed
extensively by Hettinga and Reinbold (1972 a, b). Fermentation products of
propionibacteria include large amounts of propionic acid, acetic acids, and lesser amounts
of formic, succinic, lactic acid and carbon dioxide. The earliest study regarding the
products of the fermentation of propionic acid was conducted by Fitz in the l9“I century
(Wood, 1981). He proposed that 3 moles of lactate would yield 2 moles of propionate, 1
mole of acetate, 1 carbon dioxide, and 1 water. This stoichiometry has withstood the test
of time as Gottschalk (1986), proposed that many anaerobic bacteria ferment glucose to
propionate, acetate, and carbon dioxide and that a preferred substrate of propionate-
fonning bacteria is lactate; the fermentation products being as described by the Fitz
equation discussed above.

Piveteau (1999), in reviewing the metabolism of lactate and sugars by dairy
propionibacteria, discussed the production of propionate and the transcarboxylase cycles.
He suggested that the production of propionate involves reactions arranged in several
cycles. First, a transcarboxylation occurs in which the COOH group of methylmalonyl-
CoA reacts with pyruvate thus forming oxaloacetate and propionyl-CoA. The
oxaloacetate is then reduced to succinate via malate and firmarate. Succinate is then
converted to propionate via methyhnalonly-CoA intermediates such as succinyl-CoA and
propionyl-CoA. The cycle is completed when a carboxyl group is removed from
methylmalonyl-CoA and transferred to pyruvate to yield oxaloacetate. A second
transcarboxylation reaction can also be produced during propionate production, as

methylmalonyl-CoA is regenerated from succinyl-CoA and reacts with a new molecule of

29

pyruvate. According to Piveteau (1999), the recycling of CoA and the minimal carbon
dioxide fixation produced by the transcarboxylation reaction, are beneficial to the
production of propionate as there is a minimum loss of energy involved. This pathway has
also been called the succinate-propionate pathway as described by Gottschalk (1986). This
is the most widely used pathway by the propionibacteria.

There is another process unique to propionibacteria which involves the anaerobic
fermentation of erythritol. It was reviewed by Hettinga and Reinbold (1972) and briefly
mentioned by Wood (1981). Wood and Leaver (1953) proposed that the main products
fiom the fermentation of erythritol by Propionibacterium pentosaceum were propionic,

acetic, formic and succinic acids.

Substrate preference. propionic acid productioa, pH, and temperature rcguirementa of

 

p_ropionibactcria

Piveteau (1999) discussed the utilization of various substrates during propionic
acid fermentation. He also mentioned that more ATP was produced from sugars than
from lactate and that as a consequence, increased growth rates and cell yields were
obtained during the fermentation of lactose, glucose, and galactosc than fi'om lactate.
Nevertheless, when lactate is fermented, more propionate and acetate, arc produce per
mole of pyruvate than when any of the above mentioned sugars are used (Babuchowski et
al., 1993).

Piveteau et al. (1995) reported a preference towards lactate utilization in culture
when propionibacteria were incubated in a whey medium containing lactose and Marcoux
ct al. (1992) observed lactose utilization in a whey-based media, but only when the

majority of the lactate had been utilized. The growth P. acidipropionici on a mixture of

30

glucose and lactate was determined in studies by Perez-Chaia et al. (1994). It was found
that glucose was utilized first followed by a shift in the substrate utilization to lactate when
glucose levels were low. In contrast, Lee et a1. (1974) reported the opposite occurring as
lactate was utilized first by strains of P. shermanii . In a study done by Babuchowski et
al. (1993 ), various strains of propionibacteria were tested for their ability to produce
propionic acid on substrates such as maltose, lactose, lactate and starch. Based on the
results obtained from the tests on the substrates, it was found that Propionibacterium
acidipropionici produced the most propionic acid on lactate and that 14 g/L biomas in
140 h were produced when tested using a partially hydrolyzed com substrate in batch
fermentors. Lactate was the preferred substrate over lactose and glucose for propionic
acid production by P. acidipropionici using continuous, immobilized cell biorcactors in
studies done by Lewis and Yang (1992). Lactate fermentation yielded higher propionic
acid concentrations, lower cell yields and lower specific grth rate.

Rehberger and Glatz (1998) screened seventeen strains for acid production and
final pH on glucose, fructose, and maltose as the primary carbon sources.
Propionibacteria»: acidipropiom'ci was found to produce more acid. and reached a lower
final pH than any of the other strains. When tested for its ability to grow and survive at
low pH with lactic, hydrochloric or propionic acid, propionic acid was found to be the
most inhibitory. P. acidipropionici was not able to initiate grth and survive at lower
pH values. No growth could be detected for strains at a pH below 5.0. Nevertheless,
when started at neutral pH, the final pH reached values less than 4.4. Strains could grow
at neutral pH in the presence of increasing lactate and propionate concentrations (180 mM

and 150 mM, respectively). Hsu and Yang (1991) reported that P. acidipropionici

31

cultures are extremely pH—dependent. Quesada-Chanto ct al. (1994) reported that an
increase in the pH from 5.5 to 6.5 doubled the amount of propionic acid produced in a
continuous culture with P. acidipropionici and that the optimum pH value for the
production of propionic acid is between 6.5 and 6.8.

Quesada-Chanto et al. (1994) reported the optimum temperature for propionic
acid production to be 37 °C. They arrived at this conclusion after observing a biomass
increase and product concentration at temperatures ranging from 25 °C to 37 °C and a
decrease in cell concentration when P. acidipropionici was incubated at 40 °C or higher in
continuous culture.

According to Piveteau (1999), certain propionibacteria can utilize L-lactate
preferentially over D-lactate when mixed together independent of the pH, initial lactate
concentration and initial ratio of the two isomers. He explained the utilization of one
isomer over the other by the fact that L-lactate metabolism increases the intracellular
concentration of pyruvate thus inhibiting D-lactate dehydrogenase activity. Crow (1986)
reported a 75% decrease in D-lactatc dehydrogenase when tested against 20 mmol - L'l
pyruvate and 10 mmol - L'l lactate was used as the substrate. Nevertheless, when 40
mmol - L’1 pyruvate was used, it decreased both L-lactate dehydrogenase and D-lactate
dehydrogenase by 6 and 81, respectively. These tests were performed in P. fieudenreichii
subsp. shermanii, but similar trends have also been observed with P. freudenreichii subsp.
freudenreichii and P. acidipropionici (Piveteau, 1999).

Mions between propionic and lactic acid producing bacteria
Piveteau et al. (1995), studied the interactions between lactic acid and propionic

acid producing bacteria. In their studies, Lactobacillus helveticus and Streptococcus

32

thermophilus were able to stimulate the growth of P. fi'eudenreichtr' and P.
acidipropr'onici with the former being the most consistent. In this same study, L.
helveticus and S. thermophilus were chosen for a more detailed study of growth
stimulation. It was shown that the increase in grth coincided with an increase in the
conversion of lactate to propionate and acetate and that the lactate isomers behaved as
described by Crow (1986). In contrast, Pcrcz-Chaia et al. (1994), determined that P.
acidipropionici exhibited a slow growth rate when grown with Lactobacillus helveticus.
It was concluded the slow growth rate was due to the fast reduction of pH by
Lactobacillus as it has been known to cease the growth of P. acidipropionici (Hettinga
and Reinbold, 1972). For example, mixed-culturing P. acidipropiom'ci and P.
fieudenreichii with two strains of L. helveticus, caused an inhibition in growth ( Perez-
Chaia et al.,l995). However, a third strain of L. helveticus was able to stimulate growth
of P. fieudenreichii as it’s pH was shown to be the highest (> 4.5). Because this study
involved the use of different strains of lactobacilli and propionibacteria, it was determined
that the strains were cause in the behaviors and interactions observed between these
microorganisms. Lee et al. (1976) showed similar interactions between L. plantarum and
P. shermam'i. They reported an inhibition in the growth of P. shermanii at a pH of (4.9)
due to the high acid produced by L. plantarum when grown in a glucose-lactate mixture.
The preference of lactate over glucose by propionibacteria was evidenced in this study, as
there was a slight decrease in lactate concentrations at the beginning of the experiment;
although not enough to overcome its inhibition. These results are in contrast to studies by
Liu and Moon (1982) and Parker and Moon (1982), where P. shermam'i was able to use

the lactate produced by L. acidophilus at a high rate therefore, preventing the

33

accumulation of lactate in the medium; propionibacteria were therefore stimulated by
mixed culturing.
Propionic acid and industrial uses

Propionic acid has many commercial uses. The acid and its salts are used in
making plastics (ie. moulding plastics and textiles), fruit flavors (citronellyl propionate and
geranyl propionate), perfume bases, herbicides and butyl rubber (Boyaval and Corre,
1995). It is also used in animal feeds, grain preservation, antifiingal agents (calcium and
sodium salts) (Gu et al., 1999) and has therapeutic properties such as in the use of sodium
propionate in treating wound infections, arthritis, derrnatoses (Boyaval and Corre, 1995)
and in vitamin B12 production (Hettinga and Reinbold, 1972). In the food industry, it is
used to prevent the growth of mold and ropiness in breads, cheeses, meats, fruits,
vegetables, tobacco, grain and in preventing the blowing of canned frankfurter without
causing an alteration in flavor, dipping caps, containers and wrappers in propionic acid
solutions has also proved to be effective (Boyaval and Corre, 1995). Propionic acid, has
nevertheless, been accepted as a safe product for consumers as it has been summarized as
a Generally Recognized As Safe (GRAS) food and feed additive by the Food and Drug
Administration (FDA) (Boyaval and Corre, 1995).

The US. is the main producer of propionic acid with 50,000 tons produced in
1982 at a cost of $0.73 kg'1 and $1.00 kg'1 in 1992 ( Herrero, 1983 ;Boyaval and Corrc,
1995). Nevertheless, the production of propionic acid must overcome several barriers in
order to maximize its commercial efficiency and marketability. According to Gu et al.
(1999), these barriers are: increasing the rate at which propionic acid is produced as a

typical fermentation takes 3 days to reach only 20 g/L with a yield of less than 60%;

34

controlling end-product inhibition as it decreases the amount of propionic acid produced
during ferrnentations (Herrcro, 1983; Woskow and Glatz, 1991); and decreasing the costs
involved in the separation of the acid due to its low concentration (< 60 g/L propionic
acid) from acetic acid (a byproduct of the fermentation).

Several studies have been conducted in the ever increasing efl‘ort to maximize the
production of propionic acid. P. acidipropiom'ci was tested for its ability to produce
propionic acid while immobilized in calcium alginatc beads (Paik and Glatz, 1994). The
fermentation substrates were com steep liquor and a semi-defined laboratory medium in
batch, fed-batch, and continuous fermentation. The cell density of the beads was 9.8 x 109
cells/ g (wet weight). It was found that complete substrate consumption of glucose (in
laboratory media) and lactate (in corn steep liquor) and maximum acid production
occurred in 36 h in batch culture. This was much more than seen in free-cell fermentations.
Maximum propionic acid concentrations in fed-batch ferrnentations were 45 .6 g/L in corn
steep liquor and 57g/L in the scmidefincd medium. According to the authors, the
combination of rapid initiation of acid production with high accumulated levels of product
acid makes fed-batch fermentation with immobilized cells ideal for propionic acid
production. Continuous ferrnentations also fared well as high volumetric productivities of
propionic acid (0.96 g L'1 h '1) were reported; much more than the authors had observed
with free cells. Gu et a1 (1999) also used immobilized cell to increase propionic acid
productivity as well as extractive fermentation to reduce the inhibition of propionic acid
production by end-product. They reported the productivities of propionic and acetic acids
at 0.46 g/L‘1 h'1 and 0.12 g/L'1 h'l, respectively and an improved performance with

extractive fermentation at a concentration of 13 g/L'l. Schuppert et al. (1992), used a

three-electrode poised-potential system with cobalt sepulchrate as an artificial donor to
study its efi‘cct on the growth and production of propionate in P. acidipropionicr'. This
method allowed for the exclusive production of propionic acid without the production of
acetic acid fiom the acid whey substrate and in continuous culture, propionate was formed
as the only fermentation product up to a dilution rate of 0.04 h'l. The authors suggested
that even though productivities and growth yields were low, obtaining propionate as the
only fermentation product was advantageous as the separation of end-products produced
during propionate ferrnentations is difficult and costly.
Bacteriocins of propionibacteria

Bacteriocins are antimicrobial substances produced by certain bacterial species
(Lyon et al., 1993). They have also been described as protein-containing molecules that
exert a bactericidal action on susceptible bacteria (Tagg, et al., 1976). An excellent
review on bacteriocins is available and has been written by Barefoot and Nettles (1993).
Only two bacteriocins in the cutaneous propionibacteria classification have been identified
as belonging to P. acnes (Fujimura and Nakamura, 1978; Paul and Booth, 1988) and a
total of three in the classical propionibacteria produce by P. jensem‘i and P. thoem'i
(Grinstead and Barefoot, 1992; Lyon and Glatz, 1993; Ratnam et al., 1999).

Lyon and Glatz (1991) reported a bacteriocin produced by P. thoem'i P127. It was
shown to exhibit broad activity towards related species (P. thoem'i, P. jensem’i, and P.
acidipropionici) as well as possessed activity against gram-negative and gram positive
bacteria, molds and yeasts. When tested on psychrotrophic spoilage or pathogenic
organisms such as: Lysteria monocytogenes, Pseudomonasflourescens, V ibrio

parahaemolyticus, Yerstnia enterocolitica, and a C orynebactertum sp., strains were

36

inhibited when grown in skim milk fermented by P. thoenii P127. This was evidenced by
a loss in number of viable cells after 24 h at 10 °C. The uniqueness of this bacteriocin is in
its activity against gram-negative bacteria (Barefoot and Nettles, 1993). This bacteriocin,
named Propionicin PLG- 1 ,was isolated and purified in studies done by Lyon and Glatz
(1993). In these studies, isolation was made possible by allowing cultures to grow to a
late stationary phase then analyzing the supernatant for propionicin PLG-l. Maximum
production of propionicin from P. thoenii P127 was obtained at pH of 7.0 after anaerobic
incubation for 180 h in sodium lactate broth. At a pH of 6.0, 6.5 and 7 .5, and 8.0,
bacteriocin production declined. Purification with ammonium sulfate ion-exchange
chromatography and isoclectric focusing revealed a protein band with a molecular weight
of 10,000-Da.

Grinstead and Bearfoot (1992) reported on the first heat-stable bacteriocin
produced by the diary or classical propionibacteria. J ensenii G, is the bacteriocin
produced by P. jensenii P126. Its molecular size is 12,000-Da and has demonstrated heat-
stability at 100 °C for 15 minutes. This is in contrast to propionicin PLG-l, which is heat
liable at 85 °C (Lyon and Glatz, 1991). The above mentioned investigators found J ensenii
G to exhibit a narrow spectrum of activity towards related propionibacteria, lactococci
and lactobacilli. This bacteriocin showed a bactericidal rather than a bacteriolytic action
on P. acidipropionic as the addition crude jensenii G to 1.9 x 107 cells of the
propionibacteria strain caused in total cessation of growth but not cell death. Lower levels
of jensenii G (5 to 10 AU/mL) had no effect on grth of P. acidrpropionici P5. It was
also found that jensenii G has the ability to inhibit dairy lactobacilli. This was evident as

viable cell numbers of L. delbruecla'i subsp. lactis 4797 were reduced by 99%, indicating a

37

bactericidal action against the lactobacilli. The reason why jensenii G inhibits dairy
lactobacilli more that propionibacteria is not known. Although they have suggested the
availability of more receptors on the surface of L. delbrueckr'i subsp. lactis 4797 for
jensenii G than on P. acidipropionici P5.

After screening thirteen propionibacteria strains, eight were selected because of
their production of protease-sensitive, catalase—insensitive agents (bacteriocins) which
caused the inhibition of closely related species of propionibacteria, lactobacilli and
lactococcus (Ratnam et al., 1999). Zones of inhibition were observed in P. jensenii
B1264 and was found to produce the broadest inhibitory spectrum. It was chosen for
finther studies as the mode of action for this bacteriocin revealed inhibitory activity during
late stationary phase toward L. delbrueckii subsp. lactis ATCC 4797 producing a 90%
loss in viability within 60 minutes. The purification of the inhibiting substance was
reported to have an estimated molecular mass of 6 to 9 kDa. It was shown to hold its
stability when heated at 100 °C for 60 minutes which was much in contrast to jenseniin G
exhibiting stability at 100 °C for 15 minutes (Grinstead and Barefoot, 1993) and PLG-l
showing a heat stability of 585 °C (Lyon and Glatz, 1991). A wide range of tolerance
towards pH was reported ranging fi'om 3.0 - 12.0; a much wider range than reported for
PLG-l (3.0 - 9.0) (Lyon and Glatz, 1991) and a similarity towards jenseniin G at 3.0 -
12.0 (unpublished results; Ratnam et al., 1999). Dissirnilarities were observed from the
spectrum activities of propionicin PLG-l and jenseniin G. This antagonist inhibited P.
thoenii P127 and showed no action towards gram-negative bacteria (unlike propionicin

PLG-l ). In contrast to the results observed in jenseniin G activity (Grinstead and

38

Bearfoot, 1992), the antagonist inhibited the producer of jenseniin G, P. jensenii P126, as
well as L. acidophilus ATCC 4356 and L. casei ATCC 7469.
Propionibacteria and probiotics

Much of the work done on probiotics has been on lactic acid and bifidobacteria
(Mantere—Alhonen, 1995; Fuller, 1997) extending to the yeast and fungal organisms
utilized in cattle (Huber, 1997 ). Probiotics are bacteria that promote health towards
humans and animals (Mantere—Alhonen, 1995). Criteria have been established for the ideal
probiotic. These include the ability for the microorganisms to maintain viability during
processing and storage, demonstrate a resistance to adverse changes in pH, show
adherence to intestinal epithelial cells, and maintain viability and high cell concentration
during passage through the intestine (Mantere-Alhonen, 1995;Gibson, et al., 1997).

Propionibacteria have been evaluated for their effects on stimulating growth in
animals. Antipov and Subbotin (1980; as cited by Mantere-Alhonen, 1995) combined
propionibacteria with various lactobacilli and bifidobacteria species for ameliorating
disorders of the digestive tract in calves, piglets and chickens with positive results. In
these trials, the bacterial concentration used was 4.0 to 6.0 x 109 cfu/g. This is much in
agreement with Raibaud (1992) as he demonstrated that bacteria introduced into the gut
need to be at concentrations between 5 x108 to 1 x1011 in order to play a role in the
gastrointestinal ecosystem. Other work by Cema et al. (1991) showed a reduced daily
feed intake and increased daily weight gain in calves fed a preparation called Proma.
Proma is a preparation of L. plantarum, Enterococcus faecium, Lactococcu lactis
including the propionibacterium, P. freudenreichii. The concentration used in this study

was 2 x 108 cfu/g; again in agreement with Raibaud (1992). After withdrawal of a diet

39

supplemented with Proma, calves continued to show an increase in weight of 11.3 to 21.1
% faster than the control group. Tests with piglets fed different species of
propionibacteria were investigated and P. fieudenreichii ssp. shermanii was found to
exert the most effective probiotic properties (Mantere-Alhonen, 1982 [as cited by
Mantere-Alhonen, 1995). Mantere-Alhonen (1995) reports this as being the first study to
use pure propionibacteria in regards to its effect as a probiotic and a growth promoter. In
this study, 230 piglets were given 1 to 5 g/d per animal of P. fieudenreichii in their feed.
The concentration was 2 x 109 cfu/g. The treated groups had higher weight gains (9.2 to
14.5 %) and lower feed intakes (7.2 to 46.1 %) than control groups and reduction in
diarrhea was also observed. It was concluded that even though no colonization of the
propionibacteria was evident in the intestines of the piglets, the positive results obtained
were an indication of the benefits in utilizing propionibacteria as probiotic. As mentioned
before, an important characteristic for a probiotics is its tolerance to the various
interactions taking place in the gastro-intestinal tract of animals or humans. Mantere
Alhonen (1983), tested the survivability of P. fieudenreichii during an in-vitro gastric
digestion. In this study, the culture was treated with hydrochloric acid (0.15 N), rennet
(2.1%), and pepsin (1%) and incubated at 37.2 °C in a water bath supplemented with a
shaker to simulate the peristaltic movements of the gastro-intestinal tract. After 6 h of
incubation, colony counts revealed no effect on the viability of the microorganism as
counts remained steady at a log of 107 cfir/mL in spite of the decreasing pH fi'om 6.7 to a
final of 4.8.

Perez-Chaia et al. (1995) evaluated the effects of a dairy propionibacteria and it’s

establishment in the gut. The strain used in this study was a P. acidipropionici CRL 1198

40

isolated from Emrnental cheese. The approach towards determining the survival or
establishment in the gut was to feed mice a mixture of skim milk containing 10’
bacteria/mL for 7 days and determine the population of the surviving microorganism by
fecal sampling. Results revealed an increase of 1010 propionibacteria per gram and a week
after the cessation of the diet, the amounts of propionibacteria were reduced in the large
intestine but not in the small bowel. Perez-Chaia et al (1995) suggested the persistence
was due to adhesion factors in P. acidr'propionici. Mukai et al. (1994) suggested external
components on the cell wall of bacteria may contribute to the adhesion of the intestinal
mucosa. Perez-Chaia ct al. (1995) found that P. acidipropionici CRL 1198 possesses the
ability to reverse the hyperlipemic effect of a diet with a high fat content. Mice fed cream
supplemented with 108 propionibacteria /mL, showed lower serum lipid concentration than
the group fed the cream alone. These authors postulated that a lipid-lowering effect exists
in propionibacteria when given with milk.

Perez-Chaia et al. (1999) studied the potential probiotic properties of
propionibacteria in their ability to decrease fecal enzymes possessing the capacity to
convert pro-carcinogens into carcinogens. Red cooked meat, which induces carcinogenic
compounds (Mallett et al., 1983) was fed to mice which received skim milk supplemented
with 5 x 108 cell/day, skim milk or water for 14 d. The diet supplemented with P.
acidipropionic CRL 1198, was found to prevent the activity of B-glucuronidase caused by
the red meat diet on the 7th day of feeding when P. acidipropiom'c was found at 7 x 109
cfir per gram of feces. The lowest level of B-glucuronidase also coincided on the 7“I day
of feeding. Workers concluded that propionibacteria may play a role in preventing colon

cancer.

41

Is_olation and chagaterization of Propionibacterium acidipropionici DH42

Dawson et al. (1994) isolated a bacterium from ensiled, high-moisture corn that
produces propionic acid and acetic acid from glucose or lactate. It was described by
workers as a gram-positive, facultative anaerobe, non-sporeforrning, nonmotile,
pleiomorphic rod which formed clumps when grown in peptone-yeast extract-lactate
medium. The colonies were described as pin-point, lmrn in size, yellow in color, circular
and possessed a catalase reaction when grown aerobically for five days at 30 °C. This
bacterium produced propionate and acetate from lactate and glucose. Under anaerobic
conditions, the bacterium grows rapidly utilizing glucose in liquid media. Anaerobically,
the bacteria tends to grow slowly on solid media. Maximum growth peaked after 12 to 24
h on .5X MRS incubated at 30 °C and terminal pH after 48 h on purple base-glucose and
.5X MRS was 3.9 and 4.1

The bacterium was given the name Propionibacterium acidipropr'onici DH42 as
the phenotypic characteristics of this bacterium were much like that of the ATCC type
strain Propionibacterium acidipropionici. In a review of the literature by the authors, no

published studies were found regarding the isolation of a propionibacterium fiom ensiled

feeds.

42

RESEARCH OBJECTIVES
The goal of this research was to determine the ability of Propionibacterium

acidrpropionici DH42 in preventing the accumulation of lactic acid during in-vitro
ferrnentations and during experimentally induced acidosis in ruminally fistulated cattle.
SPECIFIC OBJECTIVES
1. To test the effectiveness of P. acidipropionici DH42 in reducing the lactic acid levels

produced by in-vitro ferrnentations with wheat.
2. To determine the probiotic efl‘ects of P. acidipropionici DH42 in its ability to prevent

reductions in pH and increases in lactic acid associated with lactic acidosis syndrome

during experimentally induced lactic acidosis in cattle.

43

MATERIALS AND METHODS

In-Vitro Studies

Preliminary studies on effectiveness of media. One and a halfpercent agar
(Difco, Detroit, MI) was added to a mixture of Purple Broth Base (BBL 11558
Cockeysville, MD) with 1% erythritol (Sigma Chemical Co. St. Louis, M0) were used as
the general media in studies assessing its use as a selective and differential media for the
enumeration of P. acidipropionici DH42 cells in pure culture, in a mixed culture (rumen
fluid) and compared against a control (rumen fluid without addition of P. acidipropionici
DH42). Rumen fluid utilized was fiom a dairy cow or a beef steer. One gram of freeze-
dried P. acidipropionici DH42 obtained from Laporte Biochem International, Milwakee,
WI, was used as the inoculant. Fermentors were incubated at 38 °C and sub-samples were
collected every 6 or 8 h for a total of 24 h. Cell recoverability was determined by the
spread plate technique with appropriate dilutions in duplicated plates. Plates were placed
in glass dessicators, gassed with C02 for one minute, and incubated anaerobically for 5
days at 38 °C. Yellow colonies were counted and the effectiveness of media was
determined as the number of colony forming units per 1 mL of sub-sample.

Fermentor studies and experimental treatments. Rumen fluid was collected
from a steer at the Beef Cattle Research and Teaching Center which was maintained on a
high concentrate diet, strained through four layers of cheese-cloth, and transported to the
laboratory. A total volume of 500 mL, consisting of 250 mL freshly strained rumen fluid
and 250 mL of McDougall’s buffer was added to each of five dual-port fermentors with
built in magnetic stir rods (Microcarrier Spinner Flask, Model #1965, Belco Glass, Inc.,

Vineland, NJ). Wheat, ground through a 1mm screen (Cyclotec, Tecator Inc., Hemdon,

44

VA) was added at 2% of the volume of each ferrnentor and served as the carbohydrate
source. Additions to the fermentors were 1 mL distilled water (Control), 1 mL of P.
acidipropionici DH42 at total amounts of: 109 cfu (F1), 108 cfu(F 2), 107 cfu(F 3), and
106 cfir (F4). P. acidipropionici DH42 was available in a commercial freeze-dried form
(Laporte Biochem International, Milwakee, WI). The cells in the packet were enumerated
for total viable numbers by using the spread plate technique before initiation of the
experiments. One gram was added to 500 mL peptone (Difco, Detroit, MI), serially
diluted, and plated in duplicates. Duplicated plates were incubated anaerobically at 38 C
for 5 days before counting colonies. Viability yielded 2.4 x 1011 cfir/g. The media used to
evaluate the viability of the freeze dried P. acidipropionici DH42 was Reinforced
Clostridial Medium (OXOID, Basingstoke, Hampshire, UK). Purple Base Broth (BBL
11558, Cockeysville, MD) supplemented with 1.5% agar (Difco, Detroit, MI) and 1%
erythritol (Sigma Chemical Co., St. Louis, MO) was used to enumerate the numbers of P.
actdipropionici DH42 during the fermentation studies. Incubation was as described
above.

Sampling schedule and analytical procedures. The fermentors were gassed for
one minute with C02, sealed tightly, and incubated at 38 °C on stir plates. The incubation
was for a total of two hours after the addition of the ground wheat to allow its
fermentation to proceed. The addition of DH42 was called time zero which initiated the
sampling time frames. Five mL of ferrnentor media was sampled every eight hours starting
with time zero, for a total sampling period of 48 h. Fermentors were re-gassed, rescaled
and re-incubated at each sampling. At the end of the first 24 h, 1010 cfu of DH42 were

added to all fermentors excluding the control. Sampling was continued as described for

45

the remainder of the 40 h. All samples were analyzed for pH, cell viability, and volatile
fatty acid determination. One mL aliquots were removed for viable cell counts as
described above and the remaining 4 mL were acidified with 1 mL of a 12 N solution of
sulfuric acid and frozen for later analysis of volatile fatty acids and lactic acid. The one
mL aliquots for microbiological analysis were plated in duplicates in their appropriate
dilutions. Plates were incubated anaerobically at 38 °C for 5 days before counting
colonies. Fermentation studies were conducted on two difierent days following the same
procedure as described above as to obtain duplicated results.

HPLC analysis of fermentation end-products. Lactic acid and volatile fatty
acids were quantified by ion-exchange-exclusion HPLC (BIORAD aminex HPX-87H,
Richmond, CA). The mobile phase consisted of .005 N H2304 at a flow rate of .6
mL/min. Column temperature was regulated by an external colunm heater (Waters
Millipore, Milford, MA) at 65 °C. Four mL of ferrnentor samples were centrifirged for 30
minutes at 26,000 x g and placed into 3 mL HPLC sample vials (National Scientific,
Atlanta, GA). Centrifuged samples were placed at 4 °C until analyzed. Fifteen mL of the
centrifuged samples were injected by an autoinjector (Waters WISP 712, Milford, MA)
and analytes were detected by refractive index (Waters 410 refractive index detector,
Milford, MA). Peak heights of individual volatile fatty acids were quantified by a
commercial HPLC software package (Turbochrom 3, PE Nelson, Cupertino, CA) and
compared to a mixed standard solution containing lactic acid and volatile fatty acids
(Supelco, Bellefonte, PA).

Statistical analysis. Data were analyzed using the mixed model analysis (Proc

Mixed) of SAS (1997), with the random effect of treatment nested within the experimental

46

replication. The least squares means were compared using the Tukey-Kramer procedure
for multiple comparisons. Fermentor pH, lactic acid, volatile fatty acids, total volatile
fatty acids, and cell recoverability were analyzed using a fixed classification model with
interactions among levels of main factors. The model was:
yijrt= llijk + Pj + TR ‘1' PTI‘jrt + eijk
where:
Yijk is the variable measure (ferrnentor pH, lactic acid, propionic acid, acetic acid,
butyric acid, valeric acid, isobutyric acid and isovaleric acid concentrations, total
volatile fatty acids, and cell recoverability) of fermentor (i) containing bacterial
dose (i) sampled after (k) hours;

Ujjk is the overall mean;

PJ- is the fixed effect of the sampling period (Six levels: 0, 8, 16, 24, 32, 40, and
hours);

Tk is the fixed efi‘ect of the treatment (Five levels: 109 cfir, 108 cfu, 107 cfu, 106
cfu,109 cfu);

PTJ-k is the two-way interaction of the sampling period and treatment ;

eijk is the random residual effect pertaining to every record.
In-Vivo Studies

Preliminary animal studies. Four rumen fistulated steers fed a high concentrate
diet composed of dry rolled corn and corn silage (80:20) were used to determine the
amount of wheat necessary to provoke experimental acidosis. The steers were maintained
a the Beef Cattle Research Teaching Center Metabolism Room. Four wheat slurry dose
levels were used and each dose corresponded to an individual animal. Ground wheat was
used as a slurry by adding water in the ratio of 1 part wheat: 2 parts warm water

(weight/volume) as follows: 10 g/kg BW, 20 g/kg BW, 30 g/kg BW and 40 g/kg BW.

47

The wheat slurry was prepared by hand mixing appropriate amount of wheat, as calculated
by the animal’s body weight, in large plastic tubs with warm tap water (30 to 35 °C) as
monitored using a hand held thermometer. Wheat slurry was then poured into the rumen
via the rumen fistula of each animal. Samples were collected every 2 h for 10 h and rumen
pH was recorded. Rumen contents were evacuated when ruminal pH approached 5 .0 as
to prevent irreversible acidosis. Evacuation of ruminal contents was performed by diluting
the rumen contents with warm water (30 to 35 °C) and then siphoning contents out with a
5 cm diameter plastic hose. This was repeated until rumen was empty. Animals were
reinoculated with 2 liters of rumen contents from a healthy donor steer and hay was
ofi‘ered. The following morning, animals were taken out of the Metabolism Room and
placed in large holding pens and allowed fi'ee roaming access.

Animals, diet and experimental treatments. Six rumen fistulated steers in a
two-period crossover design weighing 500 to 550 kg were provided a diet of dry rolled
corn and corn silage (80:20) prior to the trial. The steers were fed once a day in the
morning. Approval to perform the animal studies was granted by the All-University
Committee on Animal Use and Care (# 11/95-136-00). Three animals were treated with
1011 cfir as determined by plating for viability of the freeze-dried DH42 utilizing the same
method as in the in-vitro studies. Viability assessed before each trial. P. acidipropiom'ci
DH42 was administered via rumen fistula for 2 weeks before the initiation of the
experimental period and the other three were given no DH42 and served as the controls.
Propionibacteria in this study was available as a fi'eeze-dried commerical product from
Laporte Biochem International, Milwakee, WI. The treatment sequence involved 2

periods, and the interval between period 1 and period 2 was two weeks. Animals serving

48

as controls during the first period were dosed with DH42 in period 2 and vice versa.
Acidosis was induced by the intraruminal administration of a slurry of 1 part finely ground
wheat to 2 parts warm tap water forming a dose of 40 g/kg BW as determined by
preliminary animal studies. Animals were fasted 24 h prior to the feeding of the slurry.
Freeze-dried P. acidipropionict DH42 was reconstituted in 100 mL of warm tap water
(30 °C), swirled gently, and immediately poured into the rumen via the rumen fistula, 1 h
before the morning feeding. Cattle were not fed for 24 h before the carbohydrate overload,
but allowed fi'ee access to water. The last dose of DH42 was given with the last morning
feeding, approximately 24 h before the acidosis challenge.

The end of the test period was determined to be when rumen pH values fell below
5.0 at which time the rumen contents were totally evacuated, washed with warm water,
and re-inoculated with fresh rumen contents from a healthy donor steer receiving control
diet. Steers were then given hay and released from their pens the next morning, at which
time they resumed eating a dry corn and corn silage diet as described above.

Sampling schedule and analytical procedures. Rumen fluid and jugular blood
samples were obtained fi'om each steer before and at 2, 4, 6, 8, and 10 h (end of test
period as determined by rumen pH) after the initial carbohydrate dosing. Jugular catheters
were placed in all steers before the initiation of the experimental period and flushed
regularly with a sterile heparinized solution (200 U/mL heparin, .9 % NaCl, 1 % benzyl
alcohol). Rumen contents were hand mixed thoroughly and sampled at random in three
difi‘erent sites. Rumen fluid pH was recorded immediately after collection and strained
through 4 layers of cheesecloth. Two 30 mL aliquots samples of rumen fluid were

collected for enumeration of total numbers of P. acidipropionici DH42 and for the

49

analysis of lactic and volatile fatty acids. Samples collected for acid analysis were treated
with 1 mL 12 N sulfuric acid and were frozen for later analysis by HPLC. Jugular blood
for blood pH analysis and blood lactic acid concentrations were collected with a
heparinized syringe connected to flouride/oxalate vacutainers. Blood samples were
immediately placed on ice and analyzed within 30 minutes after collection for pH
determination using a blood gas analyzer (Stat 4 Profile, Nova Biomedicals). Sammes
were centrifirged and blood plasma was collected and stored flow for later analysis.
Blood plasma L(+) lactate was determined by an enzymatic method after being
deproteinized with 10 % trichloroacetic acid (Sigma Diagnostic Kit 826-B, Sigma
Chemical, St. Louis, MO).

Microbiological procedures and media. Ruminal fluid samples were collected at
the time frames as described above for the determination of the recoverable numbers of P.
acidipropionici DH42 during ruminal acidosis. Rumen fluid was plated on duplicate
spread plates with their appropriate dilutions. Plates were place in glass dessicators,
flushed with C02, and incubated anaerobically for 5 days at 38 °C. At the end of 5 days,
colonies were counted and the recoverable numbers of P. acidipropiom'ci DH42 were
determined. The medium used was as described above in the in-vitro studies in which
purple broth base (BBL 11558, Cockysville, 1WD), 1% erythritol (Sigma Chemical Co., St.
Louis, MO), and 1.5% agar (Difco, Detroit, MI) were used. Peptone broth (Difco,
Detroit, MI) was used for the serial dilutions. Samples were immediately transported to
the laboratory where they were analyzed within 30 minutes after collection.

HPLC analysis of fermentation end-products and total blood lactic acid

determination. Rumen lactic acid and VFA were analyzed utilizing the method described

50

in the in-vitro fermentation studies. Total blood lactate was analyzed using the same
method as described for the determination of lactic acid and volatile fatty acids in rumen
fluid with the exception that total plasma lactate was detected by reversing the polarity on
the refiactive index (Waters 410 refi'active index detector, Milford, MA). D (-) lactate was
obtained as the difference between the total blood lactate as analyzed by the HPLC
method and L(+) lactate as analyzed enzymatically.

Statistical analysis. Data were analyzed using the mixed model analysis procedure
(Proc Mixed) of SAS (1997) with animal used as a random efl‘ect. The least square means
were compared using the Tukey-Kramer procedure for multiple comparisons. A fixed
classification model with interactions among levels of main factors was applied to data of
rumen pH, blood pH, blood lactates, rumen lactic acid, rumen volatile fatty acids, rumen
total volatile fatty acids, and dry matter intake (lb/d). The model equation was:

y W = uiju+ T,~+ P I + Trk+ TPj1+ TrPtd + TrT kj+ eijkl
where:

y ijk] is the variable measured (rumen pH, blood pH, blood lactate, rumen lactic
acid, propionic acid, acetic acid, butyric acid and succinic acid, total rumen volatile fatty
acids, and dry matter intake) of animal (i) belonging to group (j) during trial (k) sampled
after (1) hours;

u t,“ is the overall mean;

T ,- is the fixed effect of treatment (two levels; control and treated);

P l is the fixed effect of the period of sampling (six levels: 0, 2, 4, 6, 8, 10 hours);

Tr k is the fixed effect of the trial (two levels: Trial A and Trial B);

TP j] is the two-way interaction of treatment and period of sampling;

51

TrPu is the two-way interaction of trial and period of sampling;
Terj is the two-way interaction of trial and treatment;

e ilk! is the random residual effect pertaining to every value.

52

RESULTS AND DISCUSSION

In-vitro Studies

In pure cultures, P. acidipropionici has the ability to utilize various carbon
sources, specifically, glucose and lactate (Lewis and Yang, 1992). However, P.
acidipropionici will use lactic acid preferentially when both glucose and lactic acid are
present (Lee et al., 1974). Because of these abilities, inoculation of P. acidipropionici
DH42 into simulated ruminal fermentations would be beneficial in preventing the
accumulation of lactate. In the present study, four treated and one control ferrnentor were
filled with rumen fluid from a steer adapted to a high-concentrate diet. F ermentors
included the following doses: Control (no DH42), Trt l (109 cfir), Trt 2 (108 cfu), Trt 3
(107 cfir) and Trt 4 (106 cfir). All the fermentors contained doses that were either at or
well below the numbers of naturally occurring lactate-utilizing bacteria (approx. 109)
found in the rumen of high-concentrate fed animals (Mackie and Gilchrist, 1979). It
should be noted that the low values for lactic and volatile fatty acids observed in the
controls at 0 h in the current study, are a direct consequence of missing data due to
human error (see Appendix A).

In this study, treatment had no effect (P > .05) on the pH of the rumen fluid
(Table 1) in our fermentors. A decrease (P > .05) in pH was observed from O to 8 h in all
fermentors and remained virtually constant with no further decreases throughout the
remainder of the sampling period. It is known that the feeding of carbohydrates causes
changes in the microbial population, its fermentation patterns, and in the decline of

rumen pH parallel to increases in lactic acid (Mackie and Gilchrist, 1978; Slyter and

’53

Table l. The effect of the addition of P. acidrpropionici DH42 at different dose levels on
rumen fluid pH during in-vitro ferrnentations.

Treatment Dose, cfu

 

Rumen Fluid, pH

Time 0 (control) 193 1_()8 1_()7 1_06 SEM
(h)
0 5.98 5.99 6.00 5.91 5.92 0.29
8 5.55 5.51 ~ 5.50 5.35 5.34
16 5.51 5.49 5.48 5.38 5.32
24 5.54 5.51 5.51 5.38 5.34
32 5.54 5.50 5.50 5.38 5.34
40 5.56 5.53 5.53 5.40 5.38

 

Means are averages of 2 replications

Rumsey, 1991). It can be suggested that because no significant reduction (P > .05) in
ruminal fluid pH was observed in any of the fermentors, the lactate-utilizing microbial
population of the rumen were possibly able to rapidly metabolize lactic acid as fast as it
was being produced. This could be a possibility because the ruminal fluid initially
collected for use in the current fermentation study was of a steer adapted to a high grain
diet. Mackie and Gilchrist (1978) have found elevated ruminal pH levels (pH > 5.78) in
sheep during a stepwise adaptation to a high concentrate diet and attributed it to the low
levels of lactic acid found in their experiments. However, fermentors treated with M.
elsdem'i, a lactate-utilizer, exhibited higher ruminal pH after stabilizing at 5.4 during the
entire study than the control cultures which decreased to 4.8 for the remainder of the 24 h
fermentation period (Kung and Hession, 1995).

Lactic acid was not produced in any of the fermentors as evidenced by HPLC
analysis or the quantities were too negligible (Table 2) to be detected. In addition,
treatment effect was not significant (P > .05). There was however, a significant increase
(P < .05) of lactic acid in the control culture at 16 h and then again at 40 h from O h and in

the fermentor treated with 109 cfir DH42 at 16 h.

54

Table 2. The effect of the addition of P. acidipropionicr’ DH42 at different dose levels on
the production of lactic acid during in-vitro ferrnentations.
Treatment Dose, cfir

 

 

Lactic acid, mM
T_im_e _(___)0 control 10_9 195 10_7 10_" M
(h)
0 0.00“ 0.00b 0.00 0.00 0.00 0.004
s 0.00" 0.00b 0.00 0.00 0.00
16 0.02‘ 0.018 0.00 0.00 0.00
24 0.00b 0.00b 0.00 0.00 0.00
32 0.00b 0.00" 0.00 0.00 0.00
40 0.02‘ 0.00" 0.00 0.00 0.00

 

’b Means within a column with unlike superscripts are significantly different (P < .05)
Means are averages of 2 replications

zSEM = .005 (due to missing data)

In fermentation studies by Kung and Hession (1995) with M. elsdem'i and by Wiriyawan
and Brooker (1995) with M. esldenii and Selenomonas ruminanttum, lactic acid was
always present in control cultures as a major end product of carbohydrate fermentation.
The presence of lactic acid was expected as it is well known that adding a source of
glucose or starch to ruminal contents under an anaerobic environment causes the
proliferation of amylolytic or lactate-producing bacteria (Slyter, 1976; Mackie and
Gilchrist, 1979). It is possible, however, that the microbial population of our rumen fluid
was not affected by the addition of our carbohydrate source (10 g wheat/500 mL rumen
fluid media) as the rumen fluid utilized in this study was of a steer adapted to a high-
concentrate diet or that the level of the substrate offered might of been too low to elicit an

effect. Low levels of lactic acid have been observed during step-wise adaptations to a

high-concentrate diet in sheep (Mackie and Gilchrist, 1979).

The addition of P. acidipropionici DH42 to our fermentors at different inoculation
rates caused no effect (P > .05) on the levels of propionic acid produced. A significant
increase (P < .05) in the production of propionic acid was only observed fi'om 0 to 8 h in
the control ferrnentor (18.46 to 31.78 mM) and remained slightly unchanged throughout
the end of the sampling period (Table 3). No significant increases were noted in the
DH42 treated fermentors as the trend past 8 h was much like that of the control
ferrnentor. In contrast to these results, Kung and Hession (1995) observed decreases (P <
.05) in propionic acid production after initial increases up to 4 h during in—vitro
fennentations with a ruminal lactate—utilizer, M. elsdem'i, regardless of dose (8.7 x 10’
cfu/mL or 8.7 x 106 cfu/mL) in treated cultures. Increases in propionic acid relative to the
trace amounts of lactic acid found in the current study could reflect the active metabolism
of lactic acid into propionic acid by lactate-utilizers.

Table 3. The effect of the addition of P. acidipropionici DH42 at different dose levels on

the production of propionic acid during in-vitro ferrnentations .
Treatment Dose, cfu

 

Propionic acid, mM

Lime #40 control _1_0_9 L(f 1_07 .126. M
(h)

0 18.46” 25.42 24.75 25.50 24.64 7.04
8 31.78‘ 31.15 29.50 30.21 29.35

16 31.60“ 30.83 29.92 30.98 30.36

24 29.053 30.23 29.46 30.18 31.29

32 28.42“ 29.48 29.68 30.98 32.01

40 30.60’ 30.34 30.01 31.34 30.62

 

in’Means within a column with unlike superscripts are significantly different (P < .05)
Means are averages of 2 replications
zSEM = 7.71 (due to missing data)

56

No treatment effect (P > .05) was observed on the production of acetic acid (Table
4) in the control and treated fermentors. However, a non-significant increase (P > .05) in
the production of acetic acid was noted from 0 to 8 h in all treated fermentors and
significant increase (P < .05) was noted in the control ferrnentor fi'om 0 to 8 h (43.22 to
81.76 mM). Again, levels remained constant throughout the end of the sampling period
(40 h) in all fermentors. Kung and Hession (1995) observed a different pattern. They
reported increases in acetic acid levels up to 4 h with decreases thereafter regardless of
dose.

Treatment had no effect on (P > .05) the levels of butyric acid (Table 5) produced
in this study. Within the treated fermentors, no significant differences (P > .05) were
observed over time. However, a significant increase (P < .05) from 0 to 8 h (8.59 to 20.08
mM) was observed in the control ferrnentor and is a direct consequence of missing data
as stated earlier. The main reason for this difference being that their choice of
microorganism for use in their fermentation studies, M. elsdem'i, is one of the main
producers of butyric acid in the rumen (Slyter, 1976; Ogimoto and Imai, 1981).

Treating our fermentors with different doses of P. acidrproptonici DH42,
produced no effect (P > .05) on the levels of valeric acid (Table 6). Within the treated
fermentors, significant increases (P < .05) in valeric acid were noted at 8 h. Past 16 h
production in this acid leveled off as amounts remained practically unchanged. A
significant increase (P < .05) in valeric acid was found at 16 h in the control ferrnentor
and remained unchanged past 16 h as compared with the treated ferrmentors. In Kung
and Hession’s (1995) study, valeric acid levels were increased (P < .05) ) in treated

cultures as opposed to the controls.

57

Table 4. The effect of the addition of P. acidipropionici DH42 at different dose levels on
the production of acetic acid during simulated in-vitro acidosis .
Treatment Dose, cfir

 

Acetic acid, mM
he 0 contr 1 11: 193 1.03 12‘ sen
(h)
0 38.571’: 60.24 58.83 57.73 53.90 11.60
8 81.76‘I 81.01 79.24 74.07 72.01
16 82.68“ 80.88 82.15 76.22 76.00
24 75.92' 80.20 81.01 74.83 75.77
32 ' 76.81“ 79.45 79.97 77.50 77.80
40 80. 58' 79 96 81 42 77. 67 74. 92

 

l’I’Means within a column with unlike superscripts are significantly different (P <05)

Means are averages of 2 replications
zSEM— - 15.72 (due to missing data)

Table 5. The effect of the addition of P. acidipropionr’ci DH42 at different dose levels on
the production of butyric acid during in-vitro ferrnentations .

Treatment Dose, cfu

 

Butyric acid,

mM .
Lime _(__.10 control 10’: m3 1_0_7 .116 SEM
(h)

0 8.59” 13.99 13.70 13.80 12.92 3.69
8 20.08' 19.99 19.04 17.92 17.44

16 20.62“ 20.60 20.19 18.66 18.41

24 19.05‘ 20.32 20.04 18.10 18.85

32 18.69a 19.88 20.15 18.71 19.25

40 19.88' 20.31 20.60 19.96 18.86

 

all’Means within a column with unlike superscripts are significantly different (P < .05)

Means are averages of 2 replications

zSEM = 4.55 (due to missing data)

58

Table 6. The effect of the addition of P. acidipropionici DH42 at different dose levels on
the production of valeric acid during in-vitro ferrnentations.

 

 

Treatment Dose, cfu
Valeric acid, mM
"Ti—n19 _l.___lO control 129 IQ“. L(Z 10_6 SM
00
0 2.01”: 183° 1.77° 1.76c 1.61‘1 0.80
8 288" 2.86” 2.76” 2.61b 251°
16 3.63“ 3.59“ 3.44“b 3.24“ 3.10”“
24 3.78“ 3.91“ 3.70“ 3.44“ 3.47“”
32 3.60“ 3.87“ 3.72“ 3.91“ 3.91“
40 4.01“ 4.04“ 3.88“ 3.79“ 3.83“

 

m Means within a column with unlike superscripts are significantly different (P < .05)
Means are averages of 2 replications
zSEM = 0.84 (due to missing data)
They suggested increases in valeric acid could be due to a substitution of propionyl-CoA
for acetyl-CoA during the synthesis of fatty acids which would ultimately cause
decreases in propionic acid levels (Marounek and Bentos, 1987).
Isobutyric (Table 7) and isovaleric (Table 8) acid levels were not affected (P >
.05) by the addition ofP. acidipropionici DH42. Fermentors dosed with 109 cfu and 108
cfu DH42, behaved much like the control ferrnentor as levels of isobutyric acid increased
significantly (P < .05) at 16 h and remained slightly unchanged throughout the remainder
of the experiment. Fermentors treated with 107 cfu and 10° cfu DH42 showed significant
increases (P < .05) in isobutyric acid at 8 h. Kung and Hession’s (1995) study reported a
similar pattern in the production of isobutyric acid as compared to the controls of the
current study, in which an initial concentration of approximately .78 mM as reported.
However, lower concentrations of this acid were reported at 8 h (approximately 1.0 mM)

as compared to the current study. Increases in isobutyric acids over time in fermentors

treated with the high (8.7 x 106 cfirlmL) and low (8.7 x 10’ cfu/mL) doses ofM. elsdenii

59

Table 7. The effect of the addition of P. acidipropionici DH42 at different dose levels on
the production of isobutyric acid during in-vitro ferrnentations.
Treatment Dose, cfu

 

 

Isobutyric acid, mM

Time 0 (control) 103 10_“ 1_07 11“ SEM
(h)

0 0.87“1 1.04“ 099° 0.97“ 091° .20
8 1.47“ 1.44““ 1.28“ 1.49“ 1.46“

16 1.95“ 1.91““ 1.86“ 1.74“ 1.74“

24 2.04“ 2.12“ 2.06“ 1.79“ 2.35“

32 2.13“ 2.23“ 2.76“ 1.94“ 2.06““

40 2.46“ 2.28“ 2.78“ 2.56“ 2.45“

 

d” Means within a column with unlike superscripts are significantly different (P < .05)
Means are averages of 2 replications
zSEM = 0.28 (due to missing data)

Table 8. The effect of the addition of P. acidipropionici DH42 at different dose levels on
the goduction of isovaleric acid during simulated in-vitro acidosis.
Treatment Dose, cfu

 

 

Isovaleric acid, mM

Time 0 control .129 mi 19: .126 SEM
(h)
0 1.48“1 2.44“ 2.34° 2.28“ 2.06° 1.30
8 4.53“ 4.40“ 4.43“ 3.99“ 3.83“
16 5.58““ 5.57“ 5.61““ 4.84““ 4.71““
24 5.60““ 5.85“ 5.97“ 5.08““ 5.25““
32 6.07“ 5.91“ 6.21“ 5.26“ 5.24“
40 6.00“ 6.10“ 6.45“ 5.73“ 5.53“

 

'b° Means within a column with unlike superscripts are significantly different (P < .05)
Means are averages of 2 replications

zSEM = 1.38 (due to missing data)

in Kung and Hession’s (1995) study are comparable to the fermentors treated with 109
and 10“ cfu DH42 in the current study. Isovaleric acid exhibited a significant increase
(P<.05) at 8 h in the control and treated fermentors. Past 8 b, no significant changes
(P>.05) were observed as production of this remained slightly unchanged for the
remainder of the 40 h fermentation period. Kung and Hession’s (1995) suggestion as to
the increases in branched-chain fatty acids was due to amino acid catabolism by the
microorganism used in their studies. In any event, branched-chain fatty acids are
essential to support the growth of many rumen bacteria (Yokoyama and Johnson, 1988)
and are produced primarily by the non- cellulolytic species. This could explain the type
of bacterial population present during the in-vitro ferrnentations in the current
experiment.

The addition of strain DH42 to our fermentors resulted in no significant
treatment effects (P > .05) on the production of total volatile fatty acids. However, within
treatments, significant increases (P < .05) were observed at 40, 24, and 32 h in fermentors
treated with 108, 107, and 106 cfu, respectively. The control ferrnentor showed a
significant increase (P < .05) at 8 h and remained unchanged throughout the remainder of
the experiment. The ferrnentor treated with 109 cfu DH42 exhibited increases in the
production of total volatile fatty acids at 8 h. However, this increase was not significant
(P > .05). The high concentrations of total volatile fatty acids observed in all fermentors
could be due to the fact that the in vitro fermentation system used to conduct the current
experiment was a batch system or a closed system. Therefore, an accumulation of

volatile fatty acids was occurring. This is very different to what occurs in the rumen

61

Table 9. The effect of the addition of P. acidipropionici DH42 at different dose levels on
the production of total volatile fatty acids acid during simulated in-vitro acidosis.
Treatment Dose, cfir

Total VFA,mM
m _1__)0 control 1_0_9 193 1_07 L0_6 am
0!)
0 62.24““ 104.98“ 102.40“ 102.06“ 96.08“ 14.18
8 142.52“ 140.86“ 136.25““ 13031““ 126.61““
16 146.08“ 143.39“ 143.18““ 135.69““ 134.34““
24 135.44“ 142.66“ 142.25““ 133.45“ 136.99”
32 135.74“ 140.84“ 142.53““ 138.31“ 140.27“
40 143.54“ 143.05“ 145.15“ 140.07“ 136.24“

 

'5 Means within a column with unlike superscripts are significantly different (P < .05)
Means are averages of 2 replications
zSEM = 20.05 (due to missing data)
where decreases in total volatile fatty acids are observed due to absorption through the
rumen epithelium at low pH (Annison and Armstrong, 1970).

Preliminary studies on the ability to selectively grow strain DH42 when added to
rumen fluid by using Purple Base Broth with 1% erythritol and 1.5% agar, suggested a
possibility for its use as a selective and differential media (Appendix A). Therefore, this
media was used for microbial analysis during the in-vitro fermentation studies with P.
acidipropionici DH42. Microbiological analysis (Table 10) on Purple Base Broth with
1.0% erythritol and 1.5 % agar revealed growth only at 0 h (sample taken immediately
after addition of wheat and dose of P. acidipropionici DH42) with mean recoverable
numbers in the following treatments as follows: Trt 1 (109 cfii) 5.59 log cfirlmL; Trt 2
(108 cfu) 6.89 log chL; Trt 3 (107cfir) 7.04 log cfu/mL; Trt 4 (106cfu) 0.00 log cfir/mL.
The table below includes the actual inoculation rate (cfu/mL) for each treatment. It is
evident from the ferrnentor treated with 109 cfir (or 4.8 x 106 cfu/mL) that strain DH42

decreased in numbers as only 5.59 log chL were recovered. However, the fermentors

treated with 108 cfu and 107 cfu (or 4.8 x 105 cfir/mL and 4.8 x10“ cfu/mL), DH42

62

numbers increased considerably. The ferrnentor treated with 106 cfu (or 4.8 x 103
chL) exhibited no growth. The increases in the numbers of strain DH42 recovered
(6.89 log chL and 7 .04 log cfu/mL), suggest active growth. It would have been
expected to observe higher recoverable counts at the highest treatment (109 Cfil) as this
dose of strain DH42 is equal to the number of lactate utilizers normally found in the
rumen. However, this discrepancy could also suggest a flaw in the methodology used to
sample for strain DH42 numbers. Eventhough Trt 1 (109cfir) contained P.
acidipropionici DH42 at levels equal to the naturally occurring lactateeutilizers found in
the rumen during the feeding of high concentrate diets (Mackie and Gilchrist, 1979), it
seems a factor was impairing its grth as it was not able to survive past 8 h in rumen
fluid. Fermentor pH should not have been an environmental condition affecting the
growth of strain DH42 as the minimum pH requirement for growth of propionibacteria is
5.0 (Hettinga and Reinbold. 1972). It is interesting however, to note that recoverability
at 0 h in some of the fermentors coincided with the slight increases observed in propionic
acid from 0 to 8 h relative to the control. This could indicate an attempt by strain DH42

Table 10. The recoverability of P. acidipropionici DH42 at 0 h from in-vitro
ferrnentations with rumen fluid“

Treatmentk Concentration/Fermentor“I Recovered DH42 SEM

 

(cfir) (cfir/mL) (Log cfir/mL)
Control (no DH42) 0.00“ 0.833
109 4.8x 106 5.59“
108 4.8 x 10’ 6.89“
107 4.8 x 10“ 7.04“
10“ 4.8 x 103 0.00“

 

‘1’ Means with unlike superscripts are significantly different (P < .05)

Colony counts are averages of duplicated plates

Means are averages of two studies

hNo recoverable numbers from 8 to 40 h

kJCalculated by dividing total DH42 counts (cfu) by volume of fennentor media (500 mL)

63

to initiate metabolic activity. No growth of the microorganism was observed past the
initial sampling suggesting P. acidipropionici DH42 was unable to survive.
In-vivo Studies

Intraruminal administration of 40 g/kg BW ground wheat slurry as determined by
preliminary animal dosing studies (Appendix B), was sufficient to produce a decrease (P
< .05) in pH (Table 11) from 0 to 6 h within the control animals (7.02 to 5.18) and from 0
to 6 h within the treated animals (6.49 to 5.51). The amount of wheat provided to each
animal via rumen fistula is shown in Appendix B. This amount is more than 10 times the
amount added to each ferrnentor during the in vitro experiments (10 g/500 mL in vitro;
approximately 107 g/SOOmL in vivo). The mean reduction in pH produced after the
administration of the ground wheat slurry in both groups falls within the range of 5.0 to
5.5 which has been primarily associated with subacute acidosis (Goad et al., 1998). An
increase in pH was observed at 8 and 10 h in the control and treated animals. However,

Table 11. The effect of P. acidipropionici DH42 (1011 cfu) on rumen pH during
experimentally induced acidosis in steers.

 

 

Treatment
Rumen Fluid, pH

Time Control DH42 SEM
(h)
0 7.02““ 7.40“ 0.22
2 5.70“1 ' 6.49“°
4 5.32“l 5.73“
6 5.18“ 5.51d
8 5.42““ 5.62d
10 5.64““ 6.07°

 

“Rah/leans lacking a common superscript are significantly different (P < .05)
Means are averages of 3 replications

‘SEM = 0.23 (due to missing data)

64

the increase was only significant (P < .05) in the treated animals at 10 h. An increase in
rumen pH could be attributed to the metabolism of lactic acid by lactate-utilizing rumen
bacteria.

Clinical signs of lactic acidosis such as diarrhea, dullness, and hyperventilation
(Underwood, 1992) were not observed in acidotic cattle which were alert and mobile
even though two steers had their rumens emptied due to rumen pH decreasing below 5 .0
(see Appendix B). The rumens of these animals were flooded with warm water and
contents were siphoned-out through a 5-cm (diameter) hose. The rumens of the two steers
were inoculated with rumen contents from a healthy donor steer and offered hay.

The inability to observe the characteristic clinical signs were possibly due to the fact that
some animals had their rumens emptied before the onset of severe systemic acidosis or
that the microbial population of those that were not emptied tolerated the carbohydrate
load as the cattle were on a high concentrate feed during the whole experiment. Mackie
and Gilchrist (1979) found that during a stepwise adaptation from a low to a high
concentrate diet, a surge in the number of amylolytic and lactate-utilizing bacteria
occurred and tended to balance each other, thus resulting in lower lactic acid
accumulation. Overall, a treatment effect (P < .05) was observed on ruminal pH. The
mean values for the control and treated animals were 5.68 and 6.13, respectively.

There were no significant differences (P > .05) in the control animals on the
production of ruminal lactic acid within time (Table 12). There was, however, a general
trend for an increase (P > .05) in lactic acid levels from 0 to 4 h in the control group.
Treated animals exhibited a significant increase (P < .05) in lactic acid production from 0

to 6 h. Increases in both groups corresponded to the decreases in rumen pH during the

65

Table 12. The effect of the addition of P. acidipropionici DH42 (10ll cfu) on ruminal
lactic acid concentration during experimentally induced acidosis in steers.

 

 

Treatment
Lactic acid, mM

Time Control DH42 SEM
(h) b b
0 0.00“ 0.00 18.09
2 0.55““ 10.72““
4 27.99““ 24.35““
6 10.59““ 37.52“
8 2.49““ 32.47““3
10 0.10“b 36.27“z

 

“Means lacking a common superscript are significantly different (P < .05)
Means are averages of 3 replications
zSEM = 18.93 (due to missing data)

same time frames. A decrease in rumen pH and increases in lactic acid levels are
associated with the proliferation of amylolytic and lactic acid producing bacteria due to
the availability of carbohydrates in the rumen (Dawson and Allison, 1988). However,
lactic acid levels decreased from 8 to 10 h in both groups (P > .05), and corresponded to
the increases in ruminal pH. This suggests that some metabolism of lactic acid was
occurring. Metabolism of lactic acid in ruminants is achieved by the orderly succession
of acid-sensitive and acid-tolerant genera of lactate-utilizing bacteria which have the
capacity to control ruminal fermentation thereby, preventing the accumulation of lactic
acid (Mackie and Gilchrist, 1979). Kung and Hession (1995) found decreasing lactic
acid levels in their control cultures, but only after peaking at 5 h; cultures treated with a
low dose (8.7 x 105 cfir/mL) of Megasphaera elsdenii B159, also exhibited the same
trend. However, the high dose (8.7 x 106 chL) of the microorganism was able to
control the accumulations of lactic acid. Lactic acid and pH data in the current
experiment do not agree with those of Wiryawan and Brooker (1995) in which

Selenomonas ruminantium, another lactate utilizing microorganism inoculated at 5.0 x

1010 cfir, was able to effectively reduce lactic acid levels in acutely grain-fed sheep. In
their studies, control animals exhibited a pH of 4.9 throughout the experiment as 120 mM
of lactic acid was produced in 8 h and no lactic acid accumulation was reported in treated
animals (pH above 6.2). In the current study, lower levels or no accumulation of lactic
acid were expected in treated animals because P. acidrpropionici DH42 has the ability to
metabolize lactic acid to other products such as propionic, acetic, and succinic acids
(Lewis and Yang, 1992). However, there was no treatment effect (P > .05) as treated
cultures behaved as the control cultures indicating strain DH42 was not preventing the
accumulation of lactic acid.

The production of propionic acid (Table 13) in the rumen showed a significant
increase (P < .05) at 2 h in the control animals and at 4 h in the treated animals from 0 h.
Afier 4 h, concentrations of propionic acid in both groups were maintained steady
towards the end of the trial. However, there was no treatment effect (P > .05) on the
levels of propionic acid produced. Control animals exhibited a mean concentration of
37.7 mM as opposed to the treated which showed a much lower mean production of
propionic acid (30.7 mM). This could be due to the lower levels of lactic acid found in
the control animals as opposed to the treated. Wiryawan and Brooker (1995) reported
increases of propionic acid from 0 to 16 h in treated animals (9.2-17.0 mM) as opposed to
their controls that showed a gradual decrease (8.8-0.7 mM) within the same time frame.
In contrast, in-vitro ferrnentations by Kung and Hession (1995) showed high productions
of propionic acid in control fermentors and lower production of propionic acid in treated
cultures from 0 to 9 h. Strain DH42 has the ability to produce propionic acid from lactic

acid (Dawson et al., 1994).

67

Table 13. The effect of P. acidipropionici DH42 (1011 cfir) on propionic acid production
during experimentally induced acidosis in steers.

 

 

 

Treatment
Propionic acid, mM
Time Control DH42 SEM

(h)

0 11.85f 10.34f 4.88
2 27.70““ 21.80“f

4 45.86““° 36.13““

6 47.79““ 42.27““°

8 51.55“ 40.39““:

10 41.63““° 31.48““

'MJMeans lacking a common superscript are significantly different (P < .05)

Means are averages of 3 replications
zSEM = 5.23 (due to ruissing data)

However, it is possible that the levels of propionic acid found could be attributed to other
propionic acid producing rumen microorganisms such as Veillonella alcalescens, P.
acnes, Selenomonas ruminantium, Anaerovibrio lipolytica, and M elsdenir' (Ogimoto and
Imai, 1981).

There was no treatment effect (P > .05) on the ruminal production of acetic acid
in the control and treated animals. Values for both groups (Table 14) ranged fi'om 27.11
to 53.43 and 24.94 to 50.11 mM, in the control and treated animals respectively over the
10-hour collection period. A significant increase (P < .05) in acetic acid levels was
observed at 8 h from 0 h in the control (27.11 to 66.22 mM) and treated (24.94 to 61.16
mM) groups with a slight tendency towards an decrease (P > .05) throughout the end of
the sampling period. In vivo studies by Wiryawan and Brooker (1995) on grain-engorged
sheep revealed an increase in acetic acid from O to 8 hours in treated animals as opposed
to control animals in which acetic acid exhibited a decrease at 8 hours. An increase in

acetic acid would have been expected in treated animals if indeed strain DH42 had

68

Table 14. The effect of P. acidipropionici DH42 (10ll cfu) on acetic acid production
during experimentally induced acidosis in steers.

 

 

Treatment
Acetic acid, mM

Time Control DH42 SEM
(h)
0 2711‘“ 24.94g 6.00
2 46.58““ 43.25“"
4 66.92“ 61.82““°“
6 66.50“ 66.47““
8 66.22“° 63.27“”:
10 53.43“°“° 52.04““

 

‘wg Means lacking a common superscript are significantly different (P < .05)

Means are averages of 3 replications

zSEM = 6.31 (due to missing data)

survived in the rumen. Acetic acid is produced in addition to propionic acid during the
fermentation of lactate and glucose by propionc acid bacteria (Wood, 1981).

Treating animals with P. acidipropionici DH42 had no effect (P > .05) on the
mean production of butyric acid during experimentally induced acidosis. However,
butyric acid levels (Table 15) increased significantly (P < .05) in control animals at 2 h
and in the treated animals at 4 h. Studies by Wiryawan and Brooker (1995) have shown
decreases in butyric acid in control animals and increases in this acid at 8 h in treated
animals. The current study does not agree with Wiryawan and Brooker’s (1995) study as
butyric acid increased significantly (P < .05) at 8 h from 2 h in the control animals and
treated animals showed increases (P > .05) at 8 h from 4 h. The presence of butyric acid
in the rumen of steers suffering from subacute acidosis have been found with increasing
amounts of propionic acid (Table 13) and decreasing amounts of acetic acid (Table 14)

(Goad and Nagaraja, 1988). Increasing levels of butyrate might be associated with M

elsdem’i or Butyrivibriofiibrisolvens, two ruminal butyrate producers.

69

Table 15. The effect of P. acidipropionici DH42 (lollcfir) on butyric acid production
during experimentally induced acidosis in steers.

 

Treatment
Butyric acid, mM

Time Control DH42 SEM
(h)
0 4.52° 4.65“ 2.01
2 7.69““1 7.64““
4 10.36““ 10.30“““
6 14.19““ 12.02““°
8 15.20“ 12.43““°‘
10 12.56““ 9.19““9

 

am Means lacking a common superscript are significantly different (P < .05)
Means are averages of 3 replications
‘SEM = 2.12 (due to missing data)

Bulyrivibriofibrisolvens has been shown to produce butyrate as a primary fermentation
end product and predominates in the rumen of sheep during adaptation to a high grain
diet (Mackie and Gilchrist, 1979). M. elsdenii, lactate-utilizer, is a major producer of
butyrate from lactate metabolism in the rumen (Goad et al., 1998). Butyric acid is an
important volatile fatty acid in ruminants because it synthesizes adipose and mammary
gland tissue (Fahey and Berger, 1988).

There was no treatment effect (P > .05) on the production of succinic acid (Table
16). Within time, control animals exhibited no time interaction. However, treated
animals showed a significant increase (P < .05) at 4 h from 0 h. At 10 h, a significant
decrease (P < .05) in succinic acid was noted from 4 h. Succinate is the main end-
product of many rumen bacteria (Yokoyama and Johnson, 1988). Lactate-utilizers such
as Selenomonas ruminanlium, Veillonela alcalescens, Anaerovibrio lipolytica and

Propionibacteria convert succinate into propionate. It is possible that the low levels of

70

Table 16. The effect of P. acidrpropionici DH42 (1011 cfu) on succinic acid production
during experimentally induced acidosis in steers.

 

Treatment
Succinic acid, mM

Time Control DH42 SEM
(h)
0 0.12“° 0.08“ 0.65
2 0.33“° 0.54““
4 1.07““° 2.49“
6 082““° 2.28“
8 0.38“° 1.90““
10 0.19“° 0.86““c

 

3'” Means lacking a common superscript are significantly different (P < .05.)

Means are averages of 3 replications

:SEM = 0.7] (due to missing data)

succinate observed in the current study reflect an active metabolism of this end-product
by lactate-utilizing species into propionate as evidenced by the increasing levels of
propionic acid (Table 13) in both control and treated animals.

.The treatment effect was not significant (P < .05) for the amounts of total volatile
fatty acids produced in both the control and treated animals. Total volatile fatty acids
increased significantly (P < .05) at 8 hours in the control group and at 6 hours in the
treated group. In Wiryawan and Booker’s (1995) studies, control animals showed
decreases in total volatile fatty acid while the treated group showed increases. When pH
is low, volatile fatty acids are absorbed from the rumen as the rumen epithelium is more
permeable to the undissociated form of the acids (Stevens, 1970). This explains the
decrease found in the total volatile fatty acids of control animals in the above mentioned
study as rumen pH levels were low (4.9). In the current study, rumen pH was never

below 5.0, suggesting that there was less absorption or slow absorption of volatile fatty

acids across the rumen wall.

71

Table 17. The effect of P. acidipropionici DH42 (10ll cfu) on total volatile fatty acid
production during experimentally induced acidosis in steers.

 

 

Treatment
Total VFA, mM

Time Control DH42 SEM
(h)
0 43.62“ 40.02° 9.74
2 82.31“ 73.24“
4 124.22“b 110.75““
6 129.33““ 123.06““
8 133.37“ 117.60““*
10 107.83“° 93.19““

 

m"“"TgiMeans lacking a common superscript are significantly different (P < .05)
Means are averages of 3 replications
stM = 10.46 (due to missing data)

Blood pH (Table 18) decreased significantly (P < .05) at 4 h from 2 h to 7.38 fi'om
7.43 in the control animals and was maintained steady towards the end of the experiment.
Treated animals showed a significant increase (P > .05) in blood pH levels at 2 h.
However, a significant decrease (P < .05) was observed at 4 h from 2 h and as in the
control animals, was maintained throughout the remainder of the experimental period. A
treatment effect was observed as the mean blood pH in the DH42 treated group was
higher than the controls (7.42 and 7.39 mM). From the blood pH levels associated with
this experiment, it would appear that animals were not suffering from an acute acidosis.
According to Howard (1981), a blood pH less than 7.2 is an indicator of a poor prognosis
along with a rumen pH of 4.5. The results in the current study are in agreement with
work done by Goad et al. (1998) who reported changes in systemic acid-base status to be
minimal during subacute acidosis. Their studies reported a blood pH of 7.41 8 at 0 h and
slightly higher levels at 12-h (7.415) than those reported in the control and treated

animals in the current study.

72

Table 18. The effect of P. acidrpropionici DH42 (10ll cfir) on blood pH during
experimentally inducted acidosis in steers.

 

 

Treatment
Blood pH

Time Control DH42 SEM
(h) bode bc
0 7.41 7.43 0.016
2 7.44“ 7.50“
4 7.38““ 7.40“°“°“
6 7.40“°“° 7.40“°“°
8 7.39““ 7.40“°“°
10 7.38° 7.39““

 

“we Means lacking a common superscript are significantly different (P < .05)
Means are averages of 3 replications
zSEM = 0.018 (due to missing data)

Treatment with DH42 had no significant (P > .05) effect on the levels of total
blood lactate (Table 19) produced in both the control and treated animals as means were
2.16 and 2.28, respectively. Over time, no significant differences (P > .05) in the
production of total blood lactate were observed. Total blood lactate levels are
comparable to those of Wiryawan and Brooker (1998) (Table 22). They reported
similarities in the total blood lactate levels for both control and treated animals. The data
reported for the control animals in the current study fall within the range of total blood
lactate levels (1.29 to 4.26 mM) reported in studies by Patra et al. (1996) (Table 22) at 0
and 12 h. In contrast, Goad et al. (1998) reported lower amounts of total blood lactate at
0 and 12 h (0.7 to 0.8 mM) in their studies with subacute acidosis in grain-adapted steers.
Animals in the current study were adjusted to a high-carbohydrate diet. It is possible that

the ground wheat offered after the 24 hour fast was sufficient to increase the amounts of

total lactate present in the blood much more than the method utilized in the Goad et al.

73

Table 19. The effect of the addition of P. acidipropiom’ci DH42 (1011 cfu) on total blood
lactate during experimentally induced acidosis in steers.

 

 

Treatment
Blood lactate, mM

Time Control DH42 SEM
(h)
0 2.04““1 2.17““ 0.22
2 2.17““ 2.19““
4 2.29““ 2.18““
6 2.44““ 2.67“
8 2.06““ 2.32““
10 1.94“ 2.16““

 

'5 Means lacking a common superscript are significantly different (P < .05)
Means are averages of 3 replications
:SEM = 0.25 (due to missing data)

(1998) study. Wheat is generally considered to be one of the worst grains to provoke the

development of ruminal acidosis (Elam, 1976). Dunlop and Hammond (1968) have

published guidelines for normal ruminant total blood lactate concentrations ranging from

0.5 to 2.0 mmol/L and Hyldgaard-Jensen and Simesen (1966; [as cited by Moller et al.,

1997]) have found peak total lactic acid levels in the blood of acutely acidotic dairy cattle

at 25 mmol/L. It is evident by the results obtained in the current study that neither

control nor treated animals were suffering fi'om acute acidosis. This leads to suggestion

that total blood lactate levels were not high enough to overwhelm the acid-base system of

the blood and the animals were therefore not systemically acidotic. Blood L(+) lactate

(Table 20) of control animals in the present study were slightly higher at 0 h (5.43

mg/dL) and lower at 6 h (4.96 mg/dL) than values reported in studies by Nagaraja et al.

(1985) (Table 22). Treated animals showed elevated blood L(+) lactate levels at 0 hours

(6.67 mg/dL) and a significant decrease (P < .05) by 6 hours (4.30 mg/dL). In general,

74

Table 20. The effect of the addition of P. acidipropionici DH42 (10ll cfir) on blood L (+)
lactate during experimentally induced acidosis in steers .

 

 

Treatment
L(+) lactate, mg/dL
Time Control DH42 SEM
(hours)

0 5.43“” 6.63““ 0.65
2 4.19“ 4.39“
4 5.09““ 5.30““
6 4.96““ 4.30“
8 3.98“ 5.78““
10 4.10“ 4.26““

 

‘1’ Means lacking a common superscript are significantly different (P < .05)
Means are averages of 3 replications

zSEM = 0.72 (due to missing data)

Table 21. The effect of the addition of P. acidipropionici DH42 (10ll cfu) on blood
D(-)Lactate during experimentally induced acidosis in steers.

 

 

Treatment
D(-) lactate, mg/dL

Time Control DH42 SEM
(h)
0 13.04“““ 12.27““ 2.24
2 15.40““ 15.36““
4 15.59““ 14.29““
6 17.02““ 19.75“
8 14.59““ 15.11““
10 13.43““ 14.24““

 

aI’Means lacking a common superscript are significantly different (P < .05)

Means are averages of 3 replications

zSEM = 2.48 (due to missing data)

levels of L(+) lactate were similar for both groups and no treatment effect (P > .05) was
observed between the control and treated animals.

D(-) lactate (Table 21) values are much higher than data published by Nagaraja et
al., (1985) (Table 22). Elevated D(-) lactate values in the current study could suggest
discrepancies in the method chosen to quantify this isomer. Blood D(-) lactic acid
concentrations have been shown to surpass those of L(+) lactic acid but only when
ruminal pH decreased below 4.5 in studies by Dougherty (1975). Nevertheless, treating
the animals with strain DH42 produced no significant effect (P > .05) on the levels of
blood D(-) lactate produced. However, there was a significant increase (P < .05) at 6 h to

19.75 mg/dL from 0 h in the treated group.

76

Table 22. The effect of the addition of 101 1017.! P. acidipropionici DH42 on total blood
lactate (mM), L+ lactate (mg/dL), D- lactate (mg/dL), and blood pH of control and
treated animals in the current acidosis study as compared to results of published data in

control animals experiencing acidosis.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Time Lactate Blood pH References
(11)
Current Published Current Published
study datau study data
Total
Control Treated
O 2.04 2.17 2.30 7.41 * Wiryawan and Brooker,
1995
1.29 7.41 7.42 Patra and Swarup, 1996
0.90 7 .41 7.40 Burrin and Britton, 1986
4 2.29 2.18 1.60 7.38 7.36 Burrin and Britton, 1986
8 2.06 2.32 1.70 7.39 * Wiryawan and Brooker,
1995
1.90 7.39 7.38 Burrin and Britton, 1986
L(fi lactate
0 5.43 6.63 4.40 7.41 7.41 Nagaraia et al., 1985
6 4.96 4.30 6.60 7.39 7.36 Nagaraja et al., 1985
D(-) lactate
0 13.04 12.27 0.10 7.41 7.41 Nagaraja et al., 1985
6 17.02 19.75 0.00 7.39 7.36 Nagaraia et al., 1985
*No blood pH reported

nPublished data of values reported in control animals

77

 

A trial effect (P < .05) was observed on the average dry matter intake (DMI) as
animals during trial A ate considerably more than animals in trial B (19.64 lb/d versus
17.42 lb/d). The decrease in the feed intake during trial B could suggest that the exposure
to the bout of subacute acidosis in trial A could have impacted feed intake. It is known
that reduce feed intake is a direct consequence of subacute acidosis (Huntington, 1988).
However, treating our animals with strain DH42 produced no significant effects (P > .05)
on the average DMI as control and treated animals ate an average of 18.33 lb/d and 18.73
lb/d, respectively. Cema et al. (1990) have reported a slightly lower feed intake of calves
when fed a microbial preparation which included a propionibacterium. The concentration
of the mixture including the propionibacterium was 2 x 108 cfu/g. In a study done with
swine, the feeding of propionibacterium caused a 7 .2 to 46.1 % decrease in the fodder
demand (Mantere-Alhonen, 1982 [as cited by Mantere-Alhonen, 1995]). Robinson et al.
(1992) intraruminally inoculated fistulated steers with Megasphaera elsdenii, a ruminal
microorganism, and found that on an average, feed intake was increased in the treated
group. The concentration used was 1012 cfu. The current study reports no significant

Table 23. The effect of the addition of P. acidipropionici DH42 (10ll cfu) on the average
dry matter intake (DMI) of steers during in vivo trials.

 

 

Treatment
DMI, lb/d
Period Control DH42 Average
A 19.46 19.84 19.65‘
B 17.21 17.62 17.42“
Average 18.34 18.73

 

“5 Means are significantly different (P < .05)
Means are averages of 3 replications
SEM = 0.7

78

treatment effect (P > .05) as the treated animals ate only 2.18% more than the control.

Microbial analysis of rumen contents revealed no presence of P. acidipropiom‘ci
DH42 during any of the time periods (0-10 hours) in control or treated animals. A
selective and differential media was used which included erythritol as a fermentation
substrate, bromoscresol purple as an acidic pH indicator dye and 1.5 % agar for plating.
Propionibacteria»: spp are unique in that they ferment erythritol to propionic and acetic
acids (Holdeman et al., 1977). Therefore, it was thought that this unique ability would
aide in the recovery and quantification of P. acidrpropionici DH42 fi'om rumen contents.
Dawson et al. (1994) was able to estimate numbers of propionic acid-producing bacteria
during studies with P. acidzpropionici DH42 by using purple base broth with 1 %
erythritol. However, their method was used as a most probable number dilution scheme.
Efforts in our lab to test this medium on in-vitro and in vivo fermentations have been met
with mixed results. In-vitro numbers of recoverable P. acidrpropionici DH42 were
usually higher than those recovered from steers on a high concentrate diet but not
subjected to experimental acidosis (Roman, 1999 personal communication). Inhibition of
growth in the ruminal environment due to acidic conditions should be excluded because
ruminal pH never decreased below 5.1 during both trials. According to Hettinga and
Reinbold (1972), a pH value lower than 5.0 has been proved inhibitory for the
development and grth of propionibacteria. The possibility of a bacteriophage(s) or
bacteriocin(s) inhibiting the growth of P. acidipropr'om’ci DH42 in the rumen should not
be excluded. It is possible that the rumen might possess microorganisms capable of
producing phages or bacteriocins that would inhibit closely related species.

Bacteriophages infecting dairy propionibacteria have been isolated from Swiss-type

79

cheeses (Gautier, 1999) and bacteriocins produced from two species of propionibacteria
have shown to inhibit P. acidipropionici strains (Grinstead and Barefoot, 1992; Lyon and
Glatz, 1993). The inability to recover P. acidipropionicr‘ DH42 from the ruminal contents
of acidotic steers could also be attributed to the fact that this microorganism is not a
normal inhabitant of the indigenous microflora of the ruminant as it was previously
isolated from silage. Fuller (1978) proposed that an effective colonization of the gut
would be more readily achieved if the organism being used as a bio-inoculant (probiotic)
originated from the gut itself. P. acidipropionici DH42 would have to effectively
compete for an ecological niche in the rumen for its survival. The predominating rumen
bacteria and indigenous lactate-utilizers might place the strain DH42 at a disadvantage as
its would be obliged to compete for two of the more important substrates (e. g. glucose
and lactate). Glucose and lactate preferences by strains of P. acidrpropionici have been
studied by several researchers (Perez-Chaia et al., 1994; Lee et al., 197 4). However, the
fact that no P. acidipropionici DH42 colonies were present on culture plates could be due
to the inability of this microorganism to reproduce itself in the complex ruminal
environment. Perez-Chaia et al. (1994) showed that P. acidipropionici exhibited low
population densities and low grth rates in mixed cultures than in pure cultures. A final
suggestion as to the inability of P. acidipropionici DH42 to survive in the rumen may be
due to the fact that this microorganism was administered as a conventional freeze-dried
preparation. Merry et al. (1995) have compared freeze-dried preparations of
Lactobacillus plantarum and fresh cultures of the same strain. Although their
measurements were not based on growth, they found that herbage treated with freshly

cultured inoculants shortened the lag times in terms of the decline in pH than in freeze-

80

dried treated herbages. F reeze-dried cells are usually rehydrated prior to use and could
require longer activation times after application (Merry et al., 1995). Therefore,
considering that P. acidipropionicr‘ DH42 was introduced into a foreign environment and
in a form that possibly lengthened its time to become metabolically active, the chances
for its survival could have been hampered by these factors.

The results suggest that even though cattle were exposed to subacute acidosis as
evidenced by a ruminal pH of no less than 5.0, P. acidipropionici DH42 was not able to
survive in the complex microbial population of the rumen. The production of volatile
fatty acids and lactic acid were probably a normal trend in the fermentation
characteristics of the indigenous ruminal microflora observed in cattle experiencing
subacute acidosis. Nagaraja et al. (1985) and Goad et al. (1998) both suggested that
reductions in pH could be associated with higher levels of volatile fatty acids than lactic

acid specifically when steers had a ruminal pH between 5.0 and 5.5.

81

CONCLUSION

Inoculating in-vitro rumen ferrnentations with difi‘erent doses of
Propionibacterium acidr'propionici DH42 had a slight effect on the pH, lactic acid, and
volatile fatty acids reported, but only at 0 hours. Microbiological analysis revealed the
inability of strain DH42 to survive in ruminal contents as only viable colonies were
detected at the first sampling and none thereafter. Thus it can only be speculated that P.
acidipropiom’ci DH42 produces no growth or shifts in the metabolic activities associated
with rumen fluid and that a factor or factors were responsible for these observations
reported.

Extrapolating the results of in-vitro studies to an in-vivo system may not
adequately reproduce the same conditiOns found in the rumen such as ruminal volume,
passage rate, and the actual microbial community present. Nevertheless, Owens and
Goetsch (1988) have suggested that in-vitro findings must always be tested in-vivo as
what normally is not efl‘ective in-vitro is efi‘ective in vivo. Hence, in order to test the
effectiveness of our in-vitro results, P. acidipropionici DH42 was inoculated into the
rumen at an even higher rate than in the in-vitro studies. Results showed that the treatment
had no efl‘ect in producing significant changes in the fermentation profiles of lactic and
volatile fatty acids, and in maintaining rumen pH near neutral between the control and
treated animals. The administration of the ground wheat slurry was enough to produce
subacute acidosis as evidenced by a decrease in ruminal pH to no less than 5.0.
Absolutely no grth was present at any of the sampling times corresponding to the
microbiological analysis during the in-vivo trials. Therefore, this leads to the suggestion

that the factor or factors responsible for the inability of P. acidipropionici DH42 to

82

 

 

that the factor or factors responsible for the inability of P. acidipropionici DH42 to
survive in-vitro were also evident when tested in a live and naturally functioning ruminal
environment.

There may be various suggestions as to what may be affecting the growth and
survival of P. acidipropionici DH42. The possibility of attack by bacteriocins could merit
some consideration as it has been know that some strains of P. acidipropionici produce
these and harm closely related species. Another consideration as to the inability of strain
DH42 to survive could be due to engulfinent by protozoa as they are known to engulf
non-rumen bacteria. Protozoa ingest ruminal bacteria as a source of protein as well as
compete for the same substrates utilized by the indigenous microflora, thus keeping
bacterial numbers in the rumen in check. It is possible however, that competition of the
microorganism in a batch fermentation system was hindered by the fact that P.
acidipropionici DH42 was added as a freeze-dried commercial preparation and by the fact
that propionibacteria are slow growers. Merry et al. (1995) have found that fi'esh cultures
of Lactobacillus plantarum were more effective in creating shorter lag time in relation to
the decline in pH needed for the stability of silage than the conventional freeze-dried
treated ones. According to Merry et al. (1995), because freeze-dried microbial
preparations need to be rehydrated prior to use, they probably require a longer activation
time after application to herbages. It would not be incorrect to speculate that our choice
of freeze-dried P. acidipropionici DH42 suffered these same conditions; and along with
the additive effect of being slow growers, found it impossible to compete with the already

established rumen microbial ecosystem.

83

Inoculating the rumen with lactate-utilizing microorganisms is not a novel idea and
many investigators have encountered positive results. However, it should be pointed out
that the microorganism utilized in those experiments are normal inhabitants of the rumen.
Thus, it would be expected for these bacteria to posses a more competitive edge in their
survival than a newly introduced species, as million of years of evolution have secured
their ecological niche within the rumen.

Further research or improvement on the existing methods utilized in the current
experiments warrant examination:

1. The use of the Tilly and Terry (1963) method for in-vitro batch ferrnentations as
it more closely accurately represents the natural aspects of ruminal fermentation than the
current method utilized.

2. Inoculating the rumen and in-vitro ferrnentations with fresh cultures of P.
acidipropionic DH42 and compare results to inoculations with freeze-dried commercial
preparations of the same microorganism.

3. Probe the rumen for the possibility of bacteriophages or bacteriocins exerting
specific negative effects towards Propionibacterium acidipropionici DH42.

4. Evaluate the effects of culturing Propionibacteria»: acidipropionici DH42 with

rumen protozoa and comparing the results to the survival rates of known

ruminal microorganisms cultured with protozoa in-vitro.

84

 

APPENDICES

85

 

APPENDIX A

Data Used For In-vitro Studies

86

 

Table A-1. Data used for the analysis of volatile fatty acid production during

in-vitro fermentation studies with P. acidipropiom‘ci DH42.

 

Reps Time trt prop lactic acetic butyric isobut valeric isovaleric Total pH
VFA
A 0 cont 13.12 0.00 43.22 6.38 0.86 1.16 2.52 67.24 6.63
A 0 trtl 14.80 0.00 48.50 7.73 1.06 1.34 2.83 76.26 6.64
A 0 trt2 13.65 0.00 45.45 7.02 0.94 1.24 2.59 70.89 6.64
A 0 trt3 14.55 0.00 48.04 7.47 1.02 1.33 2.82 75.22 6.62
A 0 trt4 13.17 0.00 42.03 6.37 0.90 1.11 2.47 66.06 6.67
A 8 cont 21.81 0.00 78.57 13.96 1.60 2.29 5.71 123.94 5.66
A 8 trtl 21.34 0.00 78.16 13.90 1.51 2.20 5.48 122.59 5.56
A 8 trt2 20.42 0.00 78.14 12.71 1.03 2.24 5.63 120.16 5.65
A 8 trt3 20.95 0.00 78.65 13.15 1.47 2.22 5.64 122.08 5.58
A 8 trt4 20.71 0.00 78.00 12.85 1.48 2.27 5.54 120.84 5.56
A 16 cont 25.64 0.00 89.50 17.57 1.97 2.75 6.68 144.13 5.57
A 16 trtl 25.07 0.00 87.40 17.87 1.88 2.63 6.65 141.50 5.52
A 16 trt2 24.52 0.00 90.89 16.79 1.90 ' 2.71 6.96 143.76 5.52
A 16 trt3 23.81 0.00 86.64 15.99 1.77 2.53 6.52 137.26 5.51
A 16 trt4 24.45 0.00 90.49 16.35 20.16 2.68 6.63 142.51 5.48
A 24 cont 23.51 0.00 81.91 16.45 1.88 2.62 6.32 132.69 5.61
A 24 trtl 25.51 0.00 89.29 18.34 1.94 2.72 6.73 144.54 5.53
A 24 trt2 24.72 0.00 91.10 17.35 2.01 2.73 7.09 145.00 5.54
A 24 trt3 23.83 0.00 86.49 16.50 1.86 2.60 6.76 138.03 5.53
A 24 trt4 24.78 0.00 89.85 16.98 1.99 2.76 6.79 143.15 5.50
A 32 cont 26.55 0.00 92.45 18.88 2.20 3.00 7.27 150.34 5.57
A 32 trtl 27.58 0.00 94.98 20.10 2.23 2.99 7.27 155.15 5.51
A 32 trt2 25.35 0.00 92.88 18.29 2.08 2.82 7.19 148.62 5.51
A 32 trt3 25.51 0.00 91.81 17.90 2.05 2.92 7.07 147.26 5.51
A 32 trt4 25.98 0.00 92.41 18.60 2.19 2.96 7.21 149.34 5.48
A 40 cont 26.18 0.00 92.08 18.61 2.25 2.91 7.10 149.12 5.60
A 40 trtl 26.77 0.00 92.64 19.74 2.16 2.92 7.13 151.37 5.54
A 40 trt2 26.79 0.00 96.59 19.45 2.41 3.03 7.73 156.00 5.55
A 40 trt3 26.30 0.00 95.56 18.78 2.21 2.94 7.45 153.25 5.53
A 40 trt4 25.83 0.00 92.90 18.86 2.16 3.05 7.30 150.11 5.50
B 0 cont . . . . . . . . 5.34
B 0 trtl 36.04 0.00 71.99 20.26 1.03 2.33 2.06 133.70 5.34
B O trt2 35.85 0.00 72.21 20.38 1.05 2.31 2.10 133.91 5.37
B 0 trt3 36.46 0.00 67.42 20.13 0.92 2.20 1.74 128.89 5.20
B 0 trt4 36.12 0.00 65.78 19.47 0.93 2.12 1.66 126.10 5.18
B 8 cont 41.75 0.00 84.95 26.21 1.34 3.48 3.36 161.10 5.45
B 8 trtl 40.96 0.00 83.86 26.08 1.37 3.52 3.32 159.12 5.47
B 8 trt2 38.58 0.00 80.35 25.37 1.53 3.29 3.23 152.34 5.35
B 8 trt3 39.48 0.00 69.50 22.70 1.51 3.01 2.35 138.54 5.14
B 8 trt4 37.99 0.00 66.03 22.03 1.45 2.75 2.12 132.37 5.12
B 16 cont 37.56 0.03 75.87 23.67 1.94 4.52 4.48 148.02 5.45

 

 

87

Table A-1 (cont’d).

 

 

Reps Time trt prop lactic acetic butyric isobut valeric Isovaleric Total pH
VFA
B 16 trtl 36.60 0.02 74.37 23.33 1.95 4.55 4.49 14528 5.47
B 16 trt2 35.32 0.00 73.42 23.60 1.83 4.17 4.26 142.50 5.45
B 16 trt3 38.16 0.00 65.81 21.34 1.71 3.95 3.16 13“? 5.20
B 16 trt4 36.28 0.00 61.51 20.48 1.58 3.53 2.79 123-17 5.16
B 24 cont 34.59 0.00 69.93 21.65 2.20 4.94 4.88 13-19 5.48
B 24 trtl 34.96 0.00 71.11 22.31 2.30 5.11 4.98 140-73 5.50
B 24 trt2 34.20 0.00 70.93 22.73 2.11 4.68 4.85 139-50 5.49
B 24 trt3 36.54 0.00 63.18 19.71 1.73 4.29 3.41 12335 5.23
B 24 trt4 37.81 0.00 61.70 20.73 2.71 4.18 3.71 130-34 5.19
B 32 cont 30.30 0.00 61.18 18.50 2.07 4.21 4.87 121-13 5.51
B 32 trtl 31.38 0.00 63.92 19.67 2.24 4.75 4.56 12353 5.50
B 32 trt2 34.02 0.00 67.07 22.02 3.45 4.63 5.24 133-43 5.50
B 32 trt3 36.46 0.00 63.19 19.53 1.83 4.90 3.45 129-33 5.26
B 32 trt4 38.04 0.00 63.19 19.90 1.94 4.87 3.27 131-21 5.21
B 40 cont 35.02 0.03 69.09 21.15 2.67 5.12 4.91 13736 5.52
B 40 trtl 33.92 0.00 67.29 20.88 2.40 5.16 5.08 134.73 5.53
B 40 trt2 33.23 0.00 66.25 21.76 3.16 4.74 5.17 13430 5.52
B 40 trt3 36.38 0.00 59.78 19.15 2.92 4.65 4.01 12639 5.27
B 40 trt4 35.42 0.00 56.94 18.87 2.74 4.62 3.77 12233 5.26

 

trt = treatment; cont = no DH42;

trt4=10“cfu

trt1=109cfu; trt 2 =10“ cfu; trt 3 =107 cfu;

88

 

 

Table A-2. Data used for the analysis of recoverable numbers of P. acidipropiom’ci
DH42 during in-vitro fermentation studies.

 

 

rep Time trt lg cfu/ml
A 0 cont 0.00
A 0 trtl 4.04
A 0 trt2 6. 18
A 0 trt3 6.30
A 0 trt4 0.00
A 8 cont 0.00
A 8 trtl 0.00
A 8 trt2 0.00
A 8 trt3 0.00
A 8 trt4 0.00
A 16 cont 0.00
A 16 trtl 0.00
A 16 trt2 0.00
A 16 trt3 0.00
A 16 trt4 0.00
A 24 cont 0.00
A 24 trtl 0.00
A 24 trt2 0.00
A 24 trt3 0.00
A 24 trt4 0.00
A 32 cont 0.00
A 32 trtl 0.00
A 32 trt2 0.00
A 32 trt3 0.00
A 32 trt4 0.00
A 40 cont 0.00
A 40 trtl 0.00
A 40 trt2 0.00
A 40 trt3 0.00
A 40 trt4 0.00
B 0 cont 0.00
B 0 trtl 7.15
B 0 trt2 7.60
B 0 trt3 7.78
B 0 trt4 0.00
B 8 cont 0.00
B 8 trtl 0.00
B 8 trt2 0.00
B 8 trt3 0.00
B 8 trt4 0.00
B 16 cont 0.00

 

89

 

Table A-2 (cont’d).

 

Rep tune trt log cfir/ml
B 16 trtl 0.00
B 16 trt2 0.00
B 16 trt3 0.00
B 16 trt4 0.00
B 24 cont 0.00
B 24 trtl 0.00
B 24 trt2 0.00
B 24 trt3 ' 0.00
B 24 trt4 0.00
B 32 cont 0.00
B 32 trtl 0.00
B 32 trt2 0.00
B 32 trt3 0.00
B 32 trt4 0.00
B 40 cont 0.00
B 40 trtl 0.00
B 40 trt2 0.00
B 40 trt3 0.00
B 40 trt4 0.00

 

Trt = treatment; cont = no DH42; trt l = 109 cfir/ml DH42; trt 2 = 103 cfu/ml DH42;
Trt 3 = 107 cfu/ml DH42; trt 4 10“ cfu/ml DH42

 

Table A-3. A preliminary attempt at the recoverability of
P. acidipropionici DH42 from in-vitro anaerobic fermentors using rumen
fluid from a beef steer when plated on PBB with 1% erythritol and 1.5%

agar.

 

 

Treatmentsk
log cfu/ml
Time Control RF. and DH42 DH42 (pure)

0!)

0 0.00 0.00 8.98

6 0.00 8.51 9.18

12 0.00 0.00 9.15

18 0.00 0.00 9.35

24 0.00 0.00 9.43

 

l“treatments consisted of one gram of freeze-dried DH42
’rumen fluid

Table A-4. A preliminary attempt at the recoverability of
P. acidipropionici DH42 from in-vitro anaerobic fermentors using rumen
fluid from a dairy cow when plated on PBB with 1% erythritol and 1.5%

agar.

 

 

Treatmentsk
log cfir/ml
Time Control RF‘ and DH42 DH42 (pure)

0‘)

0 0.00 0.00 9.23

8 0.00 0.00 9.19

16 0.00 8. 74 9. l 3

24 0.00 0.00 8.84

 

ktreatments consisted of one gram of freeze-dried DH42
'rumen fluid

91

APPENDIX B

Data Used for In-vivo Studies

92

Table B-1. Data used for the analysis of fermentation end-products and pH for in-vivo
studies using P. acidipropionici DH42 for the prevention of experimentally induced
acidosis in beef steers.

 

 

Trial trt animal time lactic prop acetic butyric succinic Total pH
VFA
A treated I 0 0.00 15.40 42.40 9.15 0.22 67.16 7.23
A treated 2 0 0.00 11.60 25.39 4.33 0.18 41.50 7.15
A treated 3 0 0.00 8.82 22.10 2.66 0.05 33.64 7.67
A control 4 0 0.00 7.65 21.29 2.85 0.22 32.02 6.91
A control 5 0 0.00 20.82 41.41 3.74 0.07 66.04 6.89
A control 6 0 0.00 5.12 13.81 2.57 0.00 21.50 7.55
A treated I 2 0.48 26.37 67.60 15.54 0.00 109.51 6.44
A treated 2 2 0.08 28.84 53.57 8.15 0.00 90.56 6.11
A treated 3 2 0.00 22.89 36.70 4.50 0.09 64.17 6.74
A control 4 2 1.48 35.68 56.83 6.80 1.12 100.43 4.90
A control 5 2 0.00 43.93 65.93 7.22 0.00 117.09 5.91
A control 6 2 1.87 16.57 30.21 6.40 0.07 53.25 6.83
A treated 1 4 10.38 33.53 79.05 23.15 0.00 135.72 5.77
A treated 2 4 5.94 395163.83 10.07 0.01 113.41 5.41
A treated 3 4 0.00 51.35 61.80 7.24 0.12 120.50 5.85
A control 4 4 44.41 44.07 62.24 9.82 0.26 116.39 4.38
A control 5 4 7.65 64.19 87.80 12.19 0.00 164.18 5.34
A control 6 4 0.57 322152.30 11.58 0.31 96.39 6.22
A treated I 6 44.77 33.66 81.12 25.73 0.00 140.51 5.72
A treated 2 6 3.45 41.49 62.72 11.52 0.00 115.74 5.43
A treated 3 6 0.86 70.42 77.67 10.03 0.32 158.44 5.51
A control 4 6 50.56 35.11 52.01 9.81 0.00 96.93 4.19
A control 5 6 2.97 70.12 91.85 15.09 0.05 177.10 5.28
A control 6 6 0.83 44.34 66.71 16.56 0.41 128.02 5.89
A treated I 8 114.99 26.22 75.06 16.53 1.10 118.92 5.71
A treated 2 8 0.00 42.87 62.07 12.38 0.00 117.32 5.60
A treated 3 8 0.00 63.50 69.26 8.65 0.00 141.41 5.61
A control 4 8 12.95 41.43 53.84 12.63 0.03 107.92 .
A control 5 8 1.18 77.63 92.94 18.62 0.00 189.18 5.34
A control 6 8 0.28 467665.91 16.36 0.93 129.96 5.96
A treated I 10 147.55 22.84 75.30 9.46 3.96 111.56 6.24
A treated 2 10 0.00 26.50 44.33 9.01 0.00 79.84 6.21
A treated 3 10 0.00 47.36 52.10 6.46 0.01 105.93 6.07
A control 4 10 0.00 29.72 42.36 10.39 0.10 82.57 .
A control 5 10 0.00 56.90 69.36 14.19 0.03 140.48 5.54
A control 6 10 0.28 46.76 65.86 16.36 0.92 129.90 6.69

 

 

93

Table B-1 (cont’d).

 

 

Trial trt animal time lactic prop acetic butyric succinic Total pH
VFA
B treated I 0 0.00 15.34 30.75 5.99 0.00 52.09 7.38
B treated 2 0 0.00 5.99 15.28 3.62 0.01 24.90 7.25
B treated 3 0 0.00 4.94 13.72 2.17 0.04 20.87 7.73
B control 4 0 0.00 11.16 22.61 4.92 0.16 38.84 6.80
B control 5 0 0.00 12.67 26.15 5.30 0.23 44.34 7.17
B control 6 0 0.00 13.72 37.43 7.78 0.08 59.01 6.80
B treated I 2 60.47 23.40 43.75 8.38 0.30 75.83 6.06
B treated 2 2 3.29 14.48 27.86 5.09 2.86 59.29 6.80
B treated 3 2 0.00 14.86 30.02 4.23 0.00 49.11 6.79
B control 4 2 0.00 30.88 45.47 7.54 0.15 84.03 5.88
B control 5 2 0.00 12.67 26.15 5.30 0.23 44.34 5.28
B control 6 2 0.00 26.48 54.94 12.90 0.42 94.74 5.43
B treated I 4 113.33 35.47 69.94 7.68 3.25 116.34 5.39
B treated 2 4 16.04 26.96 46.88 7.33 8.69 89.85 5.72
B treated 3 4 0.46 30.00 49.43 6.34 2.89 88.67 6.28
B control 4 4 1.28 47.66 60.00 8.64 2.61 118.90 5.38
B control 5 4 0.73 51.59 69.25 12.27 0.00 133.12 5.73
B control 6 4 113.33 35.47 69.94 7.68 3.25 116.34 4.90
B treated l 6 150.91 34.97 69.89 5.02 4.48 114.37 5.59
B treated 2 6 21.68 32.74 55.33 9.89 7.70 105.66 5.23
B treated 3 6 3.47 40.38 52.14 9.93 1.19 103.64 5.58
B control 4 6 5.61 46.64 55.88 8.77 4.50 115.79 5.07
B control 5 6 3.61 51.51 64.08 13.27 0.01 128.88 5.74
B control 6 6 0.00 39.07 68.52 21.67 0.00 129.25 4.92
B treated I 8 . . . . . . 5.94
B treated 2 8 11.14 34.41 56.42 11.53 6.92 109.28 5.31
B treated 3 8 0.06 41.94 50.64 11.28 0.25 104.10 5.55
B control 4 8 0.57 53.81 58.02 10.27 1.27 123.36 4.80
B control 5 8 0.00 49.29 60.92 11.86 0.00 122.06 5.82
B control 6 8 0.00 40.41 65.74 21.51 0.06 127.72 4.96
B treated I 10 . . . . . . 6.52
B treated 2 10 0.01 39.24 53.26 11.26 0.06 103.82 5.66
B treated 3 10 0.00 26.51 32.73 7.72 0.13 67.09 5.75
B control 4 10 0.32 38.78 44.05 8.22 0.07 91.13 4.45
B control 5 10 0.00 38.62 43.90 8.18 0.04 90.74 6.03
B control 6 10 0.00 39.03 55.05 18.07 0.00 112.14 5.07

 

94

 

Table B-2. Data used for the analysis of total plasma lactate, L(+) lactate, and D(-) lactate
concentrations and blood pH for in-vivo studies using P. acidr'propionici DH42 for the
prevention of experimentally induced acidosis in beef steers.

 

 

 

trial trt animal time blood pH total blood plasma L(+) plasma D(-)
lactate' lactate” lactatec
A treated I 0 7.43 1.73 4.44 11.16
A treated 2 0 7.44 2.05 3.26 15.15
A treated 3 0 7.44 2.42 7.70 14.08
A control 4 0 7.35 2.37 3.26 18.10
A control 5 0 7.46 1.39 5.87 6.67
A control 6 0 7.45 . . .
A treated I 2 7.57 2.43 3.39 18.45
A treated 2 2 7.62 2.18 4.05 15.57
A treated 3 2 7.50 2.33 2.87 18.07
A control 4 2 7.46 1.49 3 .26 10.12
A control 5 2 7.47 2.15 3.65 15.73
A control 6 2 7.40 2.61 3.78 19.68
A treated I 4 7.38 2.09 4.70 14.08
A treated 2 4 7.36 1.91 4.18 12.98
A treated 3 4 7.39 2.35 3.00 18.12
A control 4 4 7.28 2. 52 6. 53 16.16
A control 5 4 7.40 2.37 4.05 17.31
A control 6 4 7.40 2.39 2.74 18.80
A treated I 6 7.39 2.03 4.05 14.25
A treated 2 6 7.37 2.13 4.18 15.02
A treated 3 6 7.39 4.61 3.13 38.33
A control 4 6 7.26 2.31 6.66 14.16
A control 5 6 7.40 2.85 3.92 21.71
A control 6 6 7.39 2.36 3.78 17.46
A treated I 8 7.42 2.57 5.48 17.68
A treated 2 8 7.41 2.81 3.92 21.41
A treated 3 8 7.40 1.72 9.66 5.82
A control 4 8 7.28 2.56 3.52 19.52
A control 5 8 7.43 1.67 2.87 12.19
A control 6 8 7.39 1.59 3.65 10.69
A treated I 10 7.39 1.17 4.70 5.86
A treated 2 10 7.38 2.55 3.26 19.72
A treated 3 10 7.39 2.14 8.22 11.04
A control 4 10 7.26 2.42 4.70 17.08
A control 5 10 7.40 1.97 3 .00 14.70
A control 6 10 7.40 2.00 3 .00 15.00
B treated 1 0 7.45 2.58 7.18 13.10
B treated 2 0 7.44 1.96 9.27 8.37
B treated 3 0 7.43

 

95

Table B-2 (cont’d).

trial trt animal time blood pH total blood plasma L(+) plasma D(-)

 

lactate‘ lactate” lactate"

B control 4 0 7.40 2.58 4.31 18.91
B control 5 0 7.41 1.81 4.83 11.43
B control 6 0 7.44 2.20 10.05 9.75

B treated 1 2 7.45 1.99 4.44 13.44
B treated 2 2 7.44 1.65 4.44 10.38
B treated 3 2 7.45 2.61 7.18 16.28
B control 4 2 7.43 2.33 4.70 16.24
B control 5 2 7.41 2.02 5.87 12.31
B control 6 2 7.44 2.47 3.92 18.35
B treated I 4 . 2.02 7.18 11.00
B treated 2 4 7.43 2.75 5.61 19.11
B treated 3 4 7.45 1.96 7.18 10.46
B control 4 4 7.44 2.14 6.53 12.74
B control 5 4 7.41 1.90 6.00 11.10
B control 6 4 7.37 2.46 4.70 17.44
B treated I 6 7.41 2.51 8.09 14.53
B treated 2 6 7.44 2.81 1.57 23.75
B treated 3 6 7.45 1.94 4.83 12.63
B control 4 6 7.44 2.64 4.31 19.45
B control 5 6 7.44 2.39 5.74 15.74
B control 6 6 7.44 2.11 5.35 13.61
B treated I 8 7.41 2.38 5.22 16.20
B treated 2 8 7.42 2.69 5.22 19.02
B treated 3 8 7.39 1.75 5.22 10.56
B control 4 8 7.40 2.76 4.70 20.14
B control 5 8 7.42 3.03 4.96 22.34
B control 6 8 7.42 0.76 4.18 2.66

B treated I 10 7.39 3.01 5.74 21.38
B treated 2 10 7.42 2.07 4.70 13.90
B treated 3 10 7.41 2.06 4.96 13.58
B control 4 10 7.41 1.35 6.13 6.05

B control 5 10 7.41 2.32 4.05 16.83
B control 6 10 7.39 1.63 3.78 10.92

 

 

aconcentration expressed as mM; analyzed with HPLC
bconcentration expressed as mg/dl; analyzed with Sigma Kit 826-B
cconcentration expressed as mg/dl; difference between total and L(+) lactic acid

96

Table B-3. The amount of wheat necessary to provoke ruminal acidosis and reduce
ruminal pH below 5.0 when added as a wheat slurry (1 part wheat:2 parts water) via
rumen cannula to 4 steers during preliminary studies.

 

 

Treatments
pH
T_irn_e M 2_Osl|_<s 341822 m
(hours)
0 6.65 6.39 7.31 7.18
4 5.89 5.3 6.68 5.62
6 5.88 5.22 4.92 5.13
9 ** 4.73 * a:
10 6.83 * * *

 

‘data available or not available because of evacuation of rumen contents
"missing data

Table B-4. The amount of wheat used to induce experimental acidosis during
In-vivo trials when added as a slurry (1 part wheat:2 parts water) and prepared
at 40 g/kg body weight of steers.

 

 

Animal Body Weight’ Wheat

(k8) (kg)
A1 440.0 5.86
A2 442.0 5.89
A3 500.0 6.66
A4 483.0 6.44
A5 471.0 6.28
A6 533.0 7.11

 

‘weights of animals represent the average of 2 trials.

97

Table B-5. Data used for the analysis of Dry Matter Intake (DMI) for steers during

trial A and trial B of experimental acidosis.

 

 

Trial Animal Offered Whack Intake DMI Dry matter
A #331 35.80 0.00 35.80 26.30 73.45

42.26 7.10 35.16 25.83

39.04 6.00 33.04 24.27

35.80 7.70 28.10 20.64

32.60 0.90 31.70 23.28

39.04 5.20 33.84 24.86

39.04 8.10 30.94 22.73

35.80 9.60 26.20 19.24

32.60 0.00 32.60 23.94

32.60 7.80 24.80 18.22

30.40 0.00 30.40 22.33

35.80 5.40 30.40 22.33

Avg 22.83

#007 32.60 0.00 32.60 23.94
39.04 9.70 29.34 21.55

35.80 7.00 28.80 21.15

32.60 8.50 24.10 17.70

30.40 4.60 25.80 18.95

30.40 0.00 30.40 22.33

35.80 2.80 33.00 24.24

35.80 5.90 29.90 21.96

35.80 11.80 24.00 17.63

32.60 7.80 24.80 18.22

30.40 0.00 30.40 22.33

35.80 5.40 30.40 22.33

Avg 21 .03

#274 28.20 4.30 23.90 17.55
28.20 1.30 26.90 19.76

30.40 6.50 23.90 17.55

28.20 8.90 19.30 14.18

24.96 0.00 24.96 18.33

30.40 5.80 24.60 18.07

30.40 1 .40 29.00 21.30

28.20 0.00 28.20 20.71

32.60 8.00 24.60 18.07

30.40 4.20 26.20 19.24

30.40 8.40 22.00 16.16

Avg 18.27

#231 24.96 6.20 18.76 13.78
22.74 0.00 22.74 16.70

28.20 4.00 24.20 17.77

28.20 5.10 23.10 16.97

28.20 6.10 22.10 16.23

24.96 3.70 21.26 15.62

24.96 3.80 21.16 15.54

24.96 5.10 19.86 14.59

 

98

 

 

Table B-5 (cont’d).

 

 

 

Trial Animal Offered Whack Intake DMI Dry matter

24.96 5.10 19.86 14.59 73.45
24.96 2.06 22.90 16.82
22.74 1.90 20.84 15.31
24.96 9.00 15.96 11.72
Avg 15.55
#214 42.26 0.00 42.26 31.04
47.70 14.30 33.40 24.53
42.26 18.10 24.16 17.75
35.80 10.60 25.20 18.51
32.60 4.00 28.60 21.01
32.60 8.00 24.60 18.07
30.40 6.00 24.40 17.92
28.20 0.00 28.20 20.71
32.60 21.30 11.30 8.30
30.40 0.00 30.40 22.33
35.80 5.50 30.30 22.26
Avg 20.22

B #331 42.26 14.50 27.76 20.03 72.15
39.04 14.80 24.24 17.49
35.80 5.10 30.70 22.15
32.60 7.70 24.90 17.97
30.40 0.60 29.80 21 .50
35.80 3.70 32.10 23.16
35.80 8.20 27.60 19.91
32.60 4.20 28.40 20.49
32.60 2.10 30.50 22.01
32.60 0.20 32.40 23.38
Avg 20.81
#007 35.80 12.10 23.70 17.10
32.60 15.20 17.40 12.55
30.40 1.20 29.20 21.07
32.60 10.40 22.20 16.02
30.40 2.30 28.10 20.27
30.40 0.00 30.40 21 .93
32.60 2.40 30.20 21.79
32.60 3.60 29.00 20.92
32.60 3.50 29.10 21.00
32.60 5.50 27.10 19.55
Avg 19.22
#274 30.40 10.40 20.00 14.43
28.20 13.10 15.10 10.89
24.96 0.60 24.36 17.58
30.40 10.90 19.50 14.07
28.20 8.70 19.50 14.07
24.96 3.20 21 .76 15.70

99

Table B-5 (cont’d).

 

 

Trial Animal Offered Whack Intake DMI Dry matter
24.96 5.90 19.06 13.75 72.15
22.74 2.20 20.54 14.82
22.74 4.90 17.84 12.87
22.74 0.80 21.94 15.83
Avg 14.40
#231 24.96 1.10 23.86 17.21
28.20 12.00 16.20 11.69
24.96 5.20 19.76 14.26
22.74 5.20 17.54 12.66
19.52 0.80 18.72 13.51
24.96 2.00 22.96 16.57
24.96 6.10 18.86 13.61
22.74 6.00 16.74 12.08
19.52 1.90 17.62 12.71
22.74 2.40 20.34 14.68
Avg 13.90
#214 35.80 7.10 28.70 20.71
32.60 11.20 21.40 15.44
30.40 1 .20 29.20 21 .07
32.60 5.20 27.40 19.77
30.40 7.20 23.20 18.74
28.20 2.10 26.10 18.83
28.20 8.40 19.80 14.29
24.96 0.90 24.06 17.36
30.40 5.60 24.80 17.89
28.20 3.00 25.20 18.18
Avg 18.03

 

100

 

 

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