|UFDC Home||myUFDC Home | Help|
This item has the following downloads:
EFFECT OF DIETARY n-3 FATTY ACID SOURCE ON PLASMA, RED BLOOD
CELL AND MILK COMPOSITION AND IMMUNE STATUS OF MARES AND
ELIZABETH LINDSAY STELZLENI
A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
Elizabeth Lindsay Stelzleni
This document is dedicated to my mother and step-father, Melanie and Jim Eisenhour, for
their unconditional love and support even when they had no idea what I was doing, and to
my husband Alex, who I could not have survived graduate school without.
First and foremost I want to thank my husband Alex for his never ending patience,
love and understanding through these first years of our marriage. His faith in me has
oftentimes exceeded the faith I have in myself, and without his constant encouragement I
could not have completed this work. I am so proud of him for completing his doctoral
degree this summer, while at the same time walking me through my first experiences of
graduate school. I am extremely lucky to have a husband who is also my best friend.
I owe great gratitude to Dr. Lori Warren, my committee chair. Her guidance and
wisdom have been invaluable to me, both inside the classroom and out. She has been
instrumental in my choices of future paths, and I thank her for this direction. I would
also like to thank Drs. Lokenga Badinga and Steeve Giguere, who served on my
committee and dedicated their time to improving my proj ect and thesis. Joel McQuagge
also deserves my appreciation for his encouragement, humor and friendship.
My sample analysis could not have been completed had it not been for the
supervision and instruction of Jan Kivipelto. Jan also receives my debt of gratitude for
being a shoulder to lean on and an open ear to talk to. I owe thanks as well to Steve
Vargas and the employees of the University of Florida Horse Research Center, especially
Cher Jackson. Their help, both in taking samples and organizing data, was crucial to this
proj ect. I would also like to thank the horses of the Horse Research Center and Horse
Teaching Unit. Without their cooperation and patience I could not have completed this
work. A special thank you goes to "Buster Buckley," who always made me smile.
I am fortunate enough to have friends throughout the department willing to offer
help and smiles during the past few years. Many thanks go to Sarah Dilling, Kelly
Spearman, Drew Cotton, Aimee Holton, Sarah White and Liz Greene for their assistance.
I also appreciate the support of the crew at the Horse Teaching Unit, especially Justin
Calahan and Kristin Detweiler.
Last, but most definitely not least, I would like to thank my grandmother, Lois
VanNatta; my "Baba," Dena Lovacheff; my sister- and brother-in-law, Jennifer and Todd
Schwent; and the rest of Alex' s and my family. I would like to lend a special thank you
to my in-laws, Lynne and Michael Stelzleni, for their unconditional love, support and
encouragement of Alex and me. Most importantly, I want to thank my mother, Melanie
Eisenhour, for showing me the kind of woman I want to be and my step-father, Jim
Eisenhour, for taking me in and loving me like his own daughter. They have offered me
nothing but undying love and support and have been my biggest fans. I am forever
indebted to them for all they have done.
TABLE OF CONTENTS
ACKNOWLEDGMENT S .............. .................... iv
LI ST OF T ABLE S ............ ..... .__ ...............x....
LI ST OF FIGURE S .............. .................... xvii
AB STRAC T ................ .............. xix
1 INTRODUCTION ................. ...............1.......... ......
2 REVIEW OF LITERATURE ................. ...............3.......... .....
Fatty Acid Structure, Digestion and Metabolism .............. ...............3.....
Fatty Acid Structure .............. ...............3.....
Fatty Acid Digestion............... ...............4
De novo Fatty Acid Synthesis .............. ...............7.....
Fatty Acid Degradation .............. ...............8.....
Polyunsaturated Fatty Acids ............... ... ............ ...............9......
n-6 and n-3 Polyunsaturated Fatty Acids .............. ....... ...............
Elongation of and Competition Between n-6 and n-3 Families .......................... 13
Eicosanoid Production and Function ................. ...............16................
The Immune System ................. ...............17........... ....
Acquired Immunity .............. ...............17....
Immunoglobulins .................. ...............17.................
Passive Immunity in the Foal .............. ...............20....
Failure of Passive Transfer............... ...............22
Innate Immunity .............. ...............24....
Inflam m ation ............... ..... ..... .. .. ... ..........2
Effects of Dietary PUFA Supplementation on Inflammation and Immune
Function .............. .... .. ... .. .............. ...............2
Blood and Tissue Responses to Experimental Feeding of n-3 PUFA .................26
Effects of PUFA supplementation on the acquisition of passive immunity
in the foal ................ ............ ... .......3
Effects of PUFA supplementation on the inflammatory response ...............3 3
Effects of PUFA supplementation on disease resistance and survival.........3 5
Characteristics of Mare Milk ................. ...............39........... ...
Mare Colostrmm .................. .... ...... ... ...... ........___..........3
Factors Affecting Mare Colostrmm IgG Content............... ...............39
Composition of Mare Milk ........._......... ....__ ........._.._ ...........4
Effect of Diet on Fat and Fatty Acid Composition of Milk .........._... ..............43
Fatty Acid Transfer across the Placenta .............. ...............46....
3 MATERIALS AND METHODS .............. ...............51....
Diets and Treatments .............. ...............52....
Bodyweights .............. .... .... ..............5
Blood Sample Collection and Processing .................._...... .........._. .......5
Colostrmm and Milk Collection and Processing ........._..._......_._ .........._.... 56
Fatty Acid Analysis .............. ...............57....
Intradermal Skin Test ................ .. ...............58.
Supplement and Feed Sample Analysis............... ...............59
Statistical Analysis............... ...............59
4 RE SULT S .............. ...............61....
Feed and Supplement Analysis............... ...............61
M are Fatty Acid Intake ............ ..... ._ ...............62...
Mare and Foal Bodyweight .............. ...............63....
Mare Plasma Fatty Acid Composition............... ..............6
Omega-6 Fatty Acids............... ...............63.
Omega-3 Fatty Acids...................... ..............6
Omega-6:Omega-3 Fatty Acid Ratios ................ ................................98
Mare Colostrmm and Milk Fatty Acid Composition............... ..............6
Foal Plasma Fatty Acid Composition ................. ...............66........... ...
Omega-6 Fatty Acids............... ...............66.
Omega-3 Fatty Acids...................... ..............6
Omega-6:Omega-3 Fatty Acid Ratios ................ ................................68
Fatty Acid Correlations................ ..................6
Fatty Acid Composition of Red Blood Cells ................. .............. ......... .....69
Mare Red Blood Cell Fatty Acids .............. ...............69....
Foal Red Blood Cell Fatty Acids............... ...............69.
Mare Serum, Colostrmm and Milk IgG............... ...............70..
Foal Serum IgG ........ ........._ .... ...... ... .......__ _.........7
Mare and Foal Responses to the Intradermal Skin Test ................. ............. .......71
Mare Response to PHA .............. ...............71....
Foal Response to PHA. .............. ... .............. ...............72......
Comparing Mare and Foal Responses to PHA ................. ................ ....._.72
5 DI SCUS SSION ................. ................. 105........ ....
Fatty Acid Composition of Feeds and Supplements ................. .......................105
Mare and Foal Bodyweight .............. ...............108....
Mare Plasma Fatty Acid Content ................. ...............109........... ...
Mare Milk Fatty Acid Content ................. ......... ...............111 ....
Foal Plasma Fatty Acid Content ................. ........... ......... ........ ....... 11
Mare and Foal Red Blood Cell Fatty Acid Content .........__.. .... ._.__............114
Effect of n-3 Supplementation on IgG. ....._._._ ....... .....__...........1
Mare and Foal Inflammatory Response .....__.....___ ........... .............1
6 IMPLICATIONS ............ ..... .__ ...............121...
A RAW DATA ............ ..... ._ ...............123...
Mare Expected and Actual Foaling Dates and Dates Started on Trial .....................123
Fatty Acid Composition of Monthly Pasture Samples ............_.. ........_........126
Mare Fatty Acid Intake ................. ...............127...............
M are Bodyweight .............. ...............128....
F oal B odywei ght............... ............. 13
M are Serum IgG .............. ...... .._ ...............134..
Mare Colostrmm and Milk IgG ............ .....__ ...............136
Foal Serum IgG. .............. ...... ...__ ......... .._.. ..........13
Fatty Acid Composition of Plasma from FISH Mares ............_.. ........_........142
Fatty Acid Composition of Plasma from FLAX Mares. ............_.. ........._.......147
Fatty Acid Composition of Plasma from CON Mares. ............_.. ........_.........152
Fatty Acid Composition of Colostrmm and Milk from FISH Mares ........................ 157
Fatty Acid Composition of Colostrum and Milk from FLAX Mares............._._... ....163
Fatty Acid Composition of Colostrum and Milk from CON Mares............._._... ......169
Fatty Acid Composition of Plasma from FISH Foals ........._.._....... ._._............175
Fatty Acid Composition of Plasma from FLAX Foals ............ ... ......_.........180
Fatty Acid Composition of Plasma from CON Foals ......___ ....... ...._..........185
Fatty Acid Composition of Red Blood Cells from FISH Mares ............. .............190
Fatty Acid Composition of Red Blood Cells from FLAX Mares ...........................195
Fatty Acid Composition of Red Blood Cells from CON Mares. ............. ..............200
Fatty Acid Composition of Red Blood Cells from FISH Foals ............... ..............205
Fatty Acid Composition of Red Blood Cells from FLAX Foals ............................210
Fatty Acid Composition of Red Blood Cells from CON Foals ............... .... ...........215
Response of Mares during an Intradermal Skin Test ................. ......._._. .........220
Response of Foals during an Intradermal Skin Test.................. ...............22
B PROCEDURE FOR IMMUNOGLOBULIN G ANALYSIS ........._..... ..............226
C PROCEDURE FOR FATTY ACID ANALYSIS .............. ...............228....
LIST OF REFERENCES ................. ...............232................
BIOGRAPHICAL SKETCH .............. ...............243....
LIST OF TABLES
2-1 Fatty acid composition of common feeds and fat supplements fed to horses .....12
2-2 Fatty acid composition of common forages fed to horses ................. ................13
2-1 Immunoglobulin concentrations of serum and milk in mature horses ................18
3-1 Nutrient composition of the grain mix concentrate and the milled flaxseed
and encapsulated fish oil supplements .............. ...............53....
3-2 Nutrient composition of the bahiagrass pasture (by month) and Coastal
bermudagrass hay .............. ...............54....
4-1 Fatty acid composition of the grain mix concentrate and the milled flaxseed
and encapsulated fish oil supplements .............. ...............73....
4-2 Fatty acid composition of winter and spring bahiagrass pasture and Coastal
bermudagrass hay .............. ...............74....
4-3 Mare average daily fatty acid intake from December-March............................7
4-4 Mare average daily fatty acid intake from April-June............... ...............7
4-5 M are bodyweights .............. ...............77....
4-6 Foal bodyweights............... ..............7
4-7 Overall effect of treatment on the fatty acid composition of mare plasma ........78
4-8 Omega-6 fatty acid content of mare plasma ................. ................ ........ .79
4-9 Omega-3 fatty acid content of mare plasma ................. ................. ........ 80
4-10 Omega-6:omega-3 fatty acid ratios in mare and foal plasma and mare milk......81
4-11 Overall effect of treatment on the total fat content of mare colostrmm and
m ilk ................. ...............82.................
4-12 Overall effect of treatment on the fatty acid composition of mare colostrmm
and m ilk ................ ...............8.. 2..............
4-13 Omega-6 fatty acid content of mare colostrum and milk ................. ................83
4-14 Omega-3 fatty acid content of mare colostrum and milk ................. ................84
4-15 Overall effect of treatment on the fatty acid composition of foal plasma..........85
.4-16 Omega-6 fatty acid content of foal plasma. ................ ............................86
4-17 Omega-3 fatty acid content of foal plasma. ................ ............................87
4-18 Correlations between mare milk and mare plasma fatty acid concentrations
and mare milk and foal plasma fatty acid concentrations .............. ..................88
4-19 Overall effect of treatment on the fatty acid content of mare red blood cells ....89
4-20 Linoleic acid content of mare red blood cells ................. ......... ...............90
4-22 Linoleic and alpha-linolenic acid contents of foal red blood cells ................... ...91
4-21 Overall treatment effect on the fatty acid composition of foal red blood cells ..92
4-23 Overall effect of treatment on mare serum and colostrmm IgG content at
foaling ................. ...............93.................
4-24 IgG content of mare milk ......._.........._.. ........___ ........._. ...._93
4-25 Correlations between IgG content of mare and foal serum, colostrum, and
m are age............... ...............94..
4-26 IgG content of foal serum ................. ...............95...............
4-27 Skin thickness of mares in response to an intradermal inj section of
phytohemagglutinin ........._._.._......_.. ...............95.....
4-28 Skin thickness of foals in response to an intradermal inj section of
phytohemagglutinin ........._.__........__. ...............96....
4-29 Skin response of mares and foals pooled across treatments to an intradermal
skin test using phytohemagglutinin as the stimulant............._ ........._._. .....97
A-1 FISH mare expected foaling dates, actual foaling dates and dates started on
trial ................ ............... 123........ .....
A-2 FLAX mare expected foaling dates, actual foaling dates and dates started on
trial ................ ............... 124........ .....
A-3 CON mare expected foaling dates, actual foaling dates and dates started on
trial ................ ............... 125........ .....
A-4 Fatty acid composition of bahiagrass pasture (by month) and Coastal
bermudagrass hay .............. ...............126....
A-5 Mare daily intake of forage, grain and supplement by month. ................... .......127
A-6 FISH mare bodyweights ................ ...............128...............
A-7 FLAX mare bodyweights .............. ...............129....
A-8 CON mare bodyweights .............. ...............130....
A-9 FISH foal bodyweights ................. ...............131...............
A-10 FLAX foal bodyweights ................. ...............132...............
A-11 CON foal bodyweights ................. ...............133...............
A-12 Serum IgG content of FISH mares at foaling ................. ........................134
A-13 Serum IgG content of FLAX mares at foaling ................. ........... ...........134
A-14 Serum IgG content of CON mares at foaling ................. ................. ......135
A-15 IgG content of colostrmm and milk from FISH mares............... ..................136
A-16 IgG content of colostrum and milk from FLAX mares ................. .................1 37
A-17 IgG content of colostrmm and milk from CON mares ............... ................138
A-18 IgG content of serum from FISH foals............... ...............139.
A-19 IgG content of serum from FLAX foals .....__ ................ ................ ...140
A-20 IgG content of serum from CON foals ....._____ .... ......... ..........__.....14
A-21 Fatty acid composition of FISH mare plasma at 28 d prior to expected foaling
date .............. ...............142....
A-23 Fatty acid composition of FISH mare plasma at 28 d post-foaling .................144
A-24 Fatty acid composition of FISH mare plasma at 56 d post-foaling .................145
A-26 Fatty acid composition of FLAX mare plasma at 28 d before expected foaling
date .............. ...............147....
A-27 Fatty acid composition of FLAX mare plasma at foaling ........._.._... ...............148
A-28 Fatty acid composition of FLAX mare plasma at 28 d post-foaling ...............149
A-29 Fatty acid composition of FLAX mare plasma at 56 d post-foaling ...............150
A-30 Fatty acid composition of FLAX mare plasma at 84 d post-foaling ...............15 1
A-31 Fatty acid composition of CON mare plasma at 28 d before expected foaling
date .............. ...............152....
A-32 Fatty acid composition of CON mare plasma at foaling ................. ................153
A-33 Fatty acid composition of CON mare plasma at 28 d post-foaling .................154
A-34 Fatty acid composition of CON mare plasma at 56 d post-foaling .................155
A-3 5 Fatty acid composition of CON mare plasma at 84 d post-foaling .................156
A-36 Fatty acid composition of FISH mare colostrum ................. ........__. ........157
A-37 Fatty acid composition of FISH mare milk at 36 h post-foaling .....................158
A-3 8 Fatty acid composition of FISH mare milk at 14 d post-foaling .....................159
A-39 Fatty acid composition of FISH mare milk at 28 d post-foaling .....................160
A-40 Fatty acid composition of FISH mare milk at 56 d post-foaling .....................161
A-41 Fatty acid composition of FISH mare milk at 84 d post-foaling .....................162
A-42 Fatty acid composition of FLAX mare colostrm ................. ............. .......163
A-43 Fatty acid composition of FLAX mare milk at 36 h post-foaling .............. .....164
A-44 Fatty acid composition of FLAX mare milk at 14 d post-foaling .............. .....165
A-45 Fatty acid composition of FLAX mare milk at 28 d post-foaling .............. .....166
A-46 Fatty acid composition of FLAX mare milk at 56 d post-foaling .............. .....167
A-47 Fatty acid composition of FLAX mare milk at 84 d post-foaling .............. .....168
A-48 Fatty acid composition of CON mare colostrm ................. ............ .........169
A-49 Fatty acid composition of CON mare milk at 36 h post-foaling .....................170
A-50 Fatty acid composition of CON mare milk at 14 d post-foaling .....................171
A-51 Fatty acid composition of CON mare milk at 28 d post-foaling .....................172
A-52 Fatty acid composition of CON mare milk at 56 d post-foaling .....................173
A-53 Fatty acid composition of CON mare milk at 84 d post-foaling .....................174
A-54 Fatty acid composition of FISH foal plasma at birth ................. ................. .175
A-55 Fatty acid composition of FISH foal plasma at 14 d of age .............. ..... ...........176
A-56 Fatty acid composition of FISH foal plasma at 28 d of age .............. ..... ...........177
A-57 Fatty acid composition of FISH foal plasma at 56 d of age .............. ..... ...........178
A-58 Fatty acid composition of FISH foal plasma at 84 d of age .............. .... ...........179
A-59 Fatty acid composition of FLAX foal plasma at birth ................. .........._.._.. .180
A-60 Fatty acid composition of FLAX foal plasma at 14 d of age ........._.._................181
A-61 Fatty acid composition of FLAX foal plasma at 28 d of age ........._.._................182
A-62 Fatty acid composition of FLAX foal plasma at 56 d of age ........._.._................183
A-63 Fatty acid composition of FLAX foal plasma at 84 d of age ........._.._................184
A-64 Fatty acid composition of CON foal plasma at birth ................. ................ ...185
A-65 Fatty acid composition of CON foal plasma at 14 d of age .............. ............... 186
A-66 Fatty acid composition of CON foal plasma at 28 d of age .............. ............... 187
A-67 Fatty acid composition of CON foal plasma at 56 d of age .............. ............... 188
A-68 Fatty acid composition of CON foal plasma at 84 d of age .............. ............... 189
A-69 Fatty acid composition of FISH mare red blood cells at 28 d before expected
foaling date ..... ._ ................ ...............190......
A-70 Fatty acid composition of FISH mare red blood cells at foaling............._.._.. ....191
A-71 Fatty acid composition of FISH mare red blood cells at 28 d post-foaling....... 192
A-72 Fatty acid composition of FISH mare red blood cells at 56 d post-foaling....... 193
A-73 Fatty acid composition of FISH mare red blood cells at 84 d post-foaling....... 194
A-74 Fatty acid composition of FLAX mare red blood cells at 28 d before expected
foaling date ................. ...............195......... ......
A-75 Fatty acid composition of FLAX mare red blood cells at foaling ................... ..196
A-76 Fatty acid composition of FLAX mare red blood cells at 28 d post-foaling ..... 197
A-77 Fatty acid composition of FLAX mare red blood cells at 56 d post-foaling ..... 198
A-78 Fatty acid composition of FLAX mare red blood cells at 84 d post-foaling ..... 199
A-79 Fatty acid composition of CON mare red blood cells at 28 d before expected
foaling date .............. ...............200....
A-80 Fatty acid composition of CON mare red blood cells at foaling ................... ....201
A-81 Fatty acid composition of CON mare red blood cells at 28 d post-foaling .......202
A-82 Fatty acid composition of CON mare red blood cells at 56 d post-foaling .......203
A-83 Fatty acid composition of CON mare red blood cells at 84 d post-foaling .......204
A-84 Fatty acid composition of FISH foal red blood cells at birth ................... .........205
A-85 Fatty acid composition of FISH foal red blood cells at 14 d of age ........._........206
A-86 Fatty acid composition of FISH foal red blood cells at 28 d of age ........._........207
A-87 Fatty acid composition of FISH foal red blood cells at 56 d of age ........._........208
A-88 Fatty acid composition of FISH foal red blood cells at 84 d of age ........._........209
A-89 Fatty acid composition of FLAX foal red blood cells at birth ................... .......210
A-90 Fatty acid composition of FLAX foal red blood cells at 14 d of age ................21 1
A-91 Fatty acid composition of FLAX foal red blood cells at 28 d of age ................212
A-92 Fatty acid composition of FLAX foal red blood cells at 56 d of age ................213
A-93 Fatty acid composition of FLAX foal red blood cells at 84 d of age ................214
A-94 Fatty acid composition of CON foal red blood cells at birth ............................215
A-95 Fatty acid composition of CON foal red blood cells at 14 d of age ..................216
A-96 Fatty acid composition of CON foal red blood cells at 28 d of age ..................217
A-97 Fatty acid composition of CON foal red blood cells at 56 d of age ..................218
A-98 Fatty acid composition of CON foal red blood cells at 84 d of age ..................219
A-99 Skin thickness values of FISH mares during an intradermal skin test ........._....220
A-100 Skin thickness values of FLAX mares during an intradermal skin test ............221
A-101 Skin thickness values of CON mares during an intradermal skin test .........._...222
A-102 Skin thickness values of FISH foals during an intradermal skin test ................223
A-103 Skin thickness values of FLAX foals during an intradermal skin test .............224
A-104 Skin thickness values of CON foals during an intradermal skin test ................225
LIST OF FIGURES
2-1 Essential fatty acid metabolism. ............. ...............14.....
4-1 Total omega-6 fatty acid content in mare plasma from 28 d pre-partum to 84
d post-foaling. ........._.__ ..... ._ ...............97....
4-2 Total omega-3 fatty acid content in mare plasma from 28 d pre-partum to 84
d post foaling. ............. ...............98.....
4-3 Total omega-6 fatty acid content of mare milk from foaling (dO) through 84 d
post-foaling. ........... ..... .._ ...............98....
4-4 Total omega-3 FA content of mares milk from foaling (dO) through 84 d post-
foaling ................. ...............99.................
4-5 Total omega-6 fatty acid content of foal plasma from birth (dO) through 84 d
of age. ............. ...............99.....
4-6 Total omega-3 fatty acid content of foal plasma from birth (dO) through 84 d
of age. ............. .....................100
4-7 Linoleic acid content of mare red blood cells from 28 d pre-partum to 84 d
post-foaling. ........... ..... .._ ...............100...
4-8 Linoleic acid content of foal red blood cells from birth (dO) to 84 d of age. ....101
4-9 Alpha-linolenic acid content of foal red blood cells from birth (dO) to 84 d of
4-10 Correlation between mare serum IgG concentration at foaling (dO) and foal
serum IgG concentration 36 h post-foaling. ................... ................0
4-11 Foal serum IgG concentration at birth and before nursing. .............. ..............102
4-12 Foal serum IgG content after colostrmm ingestion from 36 h to 84 d post-
foaling ................. ...............103................
4-13 Skin thickness of mares in response to an intradermal inj section of
phytohemagglutinin. ........... ..... .___ ...............103....
4-14 Skin thickness of foals in response to an intradermal inj section of
phytohemagglutinin. ........... ..... ._ __ ...............104....
4-15 Skin thickness of mares and foals in response to an intradermal inj section of
phytohemagglutinin. ........... ..... ._ __ ...............104....
Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science
EFFECT OF DIETARY n-3 FATTY ACID SOURCE ON PLASMA, RED BLOOD
CELL AND MILK COMPOSITION AND IMMUNE STATUS OF MARES AND
Elizabeth Lindsay Stelzleni
Chair: Lori K. Warren
Major Department: Animal Sciences
Supplementing the diets of horses with fat is a popular trend in today's equine
industry. However, little focus has been given to the effect of supplementing with
omega-3 fatty acids (FA) in the broodmare and her suckling foal. To study these effects,
36 Thoroughbred and Quarter Horse mares with an average bodyweight of 580.9 & 3.5 kg
(mean & SE) were randomly assigned to one of three treatment groups: 1) basal diet with
no supplementation (CON, n = 12); 2) basal diet plus milled flaxseed supplementation
(FLAX, n = 12); or 3) basal diet plus encapsulated fish oil supplementation (FISH, n =
12) from 28 days prior to expected foaling date until 84 days after foaling. The flaxseed
and fish oil supplements were fed to mares in amounts to provide 6 g total n-3 FA/100 kg
BW per day. The basal diet consisted of a commercial grain-based concentrate, Coastal
bermudagrass hay and bahiagrass pasture.
Blood samples were obtained from mares at 28 d pre-partum and milk and blood
samples were obtained from mares and foals at foaling, 36 h and 14, 28, 56 and 84 d
post-partum to determine FA and IgG content. On d 84, mares and foals received paired
intradermal inj sections of phytohemagluttinin (PHA) and skin thickness was determined
over a 48 h period as a measure of the inflammatory response. Bodyweights were
obtained from mares and foals at 14 d intervals throughout the trial.
Treatment had no effect on gestation length (P = 0.84), mare bodyweight (P = 0.80)
or foal bodyweight (P = 0.76). Mares fed FLAX had higher plasma alpha-linolenic acid
(ALA) (P=0.06) than mares fed FISH or CON mares. Mares fed FISH had higher plasma
eicosapentaenoic acid (EPA), docosahexanoic acid (DHA) and total n-3 (P=0.03) than
FLAX and CON mares. Across treatments, total milk n-3 increased (P=0.0005) and total
n-6 decreased (P=0.0001) from foaling to d 84. Milk from FLAX mares had higher ALA
(P=0.01) than milk from FISH and CON mares. Milk from FISH mares had higher EPA
and DHA and a lower n-6:n-3 ratio (P=0.007) than milk from FLAX and CON mares.
Foals suckling FLAX mares had higher plasma ALA (P=0.04) than foals suckling FISH
and CON mares. Foals suckling FISH mares had higher plasma EPA, DHA and total n-3
and a lower plasma n-6:n-3 ratio (P=0.002) than FLAX and CON foals. Treatment did
not affect colostrum, milk or foal serum IgG. Response to PHA inj section was greater
(P=0.0001) in mares compared to foals but similar between treatments. Although the
addition of n-3 FA to the mare's diet altered the FA content of mare milk and mare and
foal plasma, changes in total IgG and PHA intradermal responses were not detected..
Supplementing the diet with fat is a popular trend in the horse industry. Fat is
commonly fed to horses to improve the hair coat, improve body condition and increase
the energy density of the diet. However, most of the research that has examined fat
supplementation of the horse has been performed with little regard to the type of fatty
acids (FA) provided. In addition, most of this research has focused on mature
performance horses; relatively little information is available on fat supplementation of
mares and the effects on the suckling foal.
Corn oil, soybean oil, and rice bran are common sources of fat added to horse
rations; however, these feeds are high in omega-6 FA. High levels of n-6 FA have been
associated with more pronounced inflammatory responses in humans (Meydani et al.,
1993; Simopoulos, 1999); therefore, potential exists for such diets to also have negative
biological effects in the horse.
Based on the immunomodulatory effects of n-3 FA in humans and other animals
(Simopoulos, 1999; Anderson and Fritsche, 2002), there is interest in determining
whether n-3 FA supplementation can modify inflammatory and immune responses in
horses. In addition, the dietary source of n-3 FA may be important for eliciting the
desired health benefits. Flaxseed is an excellent source of alpha-linolenic acid (ALA),
whereas Hish oil is a good source of eicosapentaenoic acid (EPA) and docosahexaenoic
acid (DHA). Although both are rich in total n-3 FA, fish oil may be a more effective
means of providing biologically active n-3 FA than flax.
The immune status of foals is a vital concern for horse breeders, as suckling foals
are susceptible to many health problems including diarrhea and septicemia. These health
problems can cause significant veterinary expense, as well as endanger the life of the
foal. Previous research has shown that supplementation of broodmares. with linseed oil or
a mix of corn and linseed oil increases the n-3 content of her milk and the n-3 content of
her foal's blood (Duvaux-Ponter et al, 2004; Spearman et al., 2005). Therefore, it seems
possible to enhance the concentration of n-3 FA in the foal by manipulation of the mare's
The obj ectives of this research were to 1) examine the effect of dietary n-3
supplementation on the FA composition of mare milk and mare and foal plasma; 2)
examine the efficiency with which ground flaxseed or encapsulated fish oil augment the
presence of EPA and DHA in the mare and foal; 3) examine the effects of
supplementation with flaxseed vs. Eish oil on increasing colostrum, milk and foal plasma
IgG; and 4) examine the effects of feeding flaxseed vs. fish oil on the inflammatory
response in the mare and foal. We hypothesize that supplementing the mare with Hish oil
will increase the EPA and DHA concentrations in mare milk and mare and foal blood to a
higher extent than will flaxseed, will increase the IgG in mare colostrum and foal blood,
and will decrease the inflammatory response in mares and foals.
REVIEW OF LITERATURE
Fatty Acid Structure, Digestion and Metabolism
Fatty Acid Structure
Fatty acids (FA) consist of carbon (C), hydrogen (H) and oxygen (0) arranged in
a carbon chain with a carboxyl group (-COOH) at one end and a methyl group (-CH3) at
the other. FA are classified and named by their chain lengths and their degree of
unsaturation, or number of double bonds. Unsaturated FA can be monounsaturated (only
one double bond) or polyunsaturated (two or more double bonds), whereas saturated fatty
acids have no double bonds. Fatty acids are also classified as short, medium or long
chain, with short chain FA having less than 8 carbons, medium chain FA having 8 to 16
carbons and long chain FA having greater than 16 carbons. The numbering sequence of
the carbons in a fatty acid chain begins at the carboxyl end, with the carboxyl carbon
being C1. An older system used Greek letters to identify carbon atoms. In this system,
C2 (the first carbon after the carboxyl carbon) was the ot-carbon, C3 was the P-carbon
and so on, ending with the last carbon in the chain at the methyl end as the co-carbon
(Gurr et al., 2002).
Currently, the numbering system is the preferred method of naming individual FA.
In this system, the number of carbon atoms in the FA chain is given followed by a colon
and the number of double bonds. For example, stearic acid, a saturated FA of 18C, is
identified as C18:0. Linoleic acid, a polyunsaturated FA of 18C with two double bonds,
is identified as C18:2. While the current numbering system is preferred, the older system
is used to identify co-6 and co-3 fatty acids, where the last double bond in the fatty acid
chain is six and three carbons away from the co-carbon, respectively (Greene, 2006).
Newer research may substitute the ao with an n, but the meaning does not change.
The presence of double bonds in a fatty acid chain also allows for positional and
geometric isomerism. Positional isomerism refers to a different location of double bonds
in the carbon chain. Geometric isomerism refers to the orientation of the hydrogen
atoms around the carbon-carbon double bond. A cis configuration results when both
hydrogen atoms are on the same side of the bond, while a trans configurations results
when hydrogen atoms are on opposite sides of the bond. Most natural unsaturated FA are
in the cis configuration (Spallholz et al., 1999).
Fatty Acid Digestion
Dietary fats exist mostly as triglycerides (TG) which are made up of three FA
attached to a glycerol backbone (Mu and Hoy, 2004). The digestion of these TG begins
in the stomach by the action of gastric lipase released from the gastric mucosa. In
humans and rats, lingual lipase from the von Ebner glands, a group of serious glands on
the tongue, also aids in FA digestion in the stomach. This lipase is transferred with the
food bolus into the stomach where its activity begins (Mu and Hoy, 2004). Secretion of
the lingual lipase occurs continuously but is stimulated by dietary (high fat) and
mechanical (suckling) factors (Carey et al., 1983; Tso, 1989). Digestion by both lipases
produces free FA and diglycerides (Carey et al., 1983; Tso, 1989). The lingual lipase is
especially important in the newborn, as pancreatic lipase activity is not fully developed at
birth. In addition, the short and medium-chain TG present in milk fat are readily
hydrolyzed by the lingual lipase (Tso, 1989). Horse saliva, however, does not possess
this lingual lipase (Frape, 1998; Ellis and Hill, 2005). In fact, it is currently thought that
equine saliva does not contain any enzyme activity (Ellis and Hill, 2005). Therefore,
equine saliva is not as important in beginning digestion but is vital in providing feed
lubrication (Frape, 1998) and buffering of the feed-saliva mixture (Ellis and Hill, 2005).
While only 10-30% of dietary fat is hydrolyzed in the stomach, the maj ority of FA
digestion takes places in the small intestine, especially in the duodenum. In animals with
a gall bladder, the action of the food bolus entering the duodenum stimulates gall bladder
emptying, secretion of pancreatic lipase and the release of cholecystokinin (CCK). Bile
acids are also released from the gall bladder or directly from the liver to emulsify the fat
(Mu and Hoy, 2004). The horse, however, does not have a gall bladder, but this does not
seem to affect the digestion of fat (Cunha, 1991). In the horse, bile continuously drains
from the liver into the small intestine to facilitate the emulsion of fat (Frape, 1998).
Furthermore, the peristaltic and segmental contractions present in the intestine supply
mechanical energy to reduce the fat particle size and increase the interfacial area of the
fat droplets (Carey et al., 1983). The action of pancreatic lipase on a triglyceride
molecule releases two free FA and a 2-monoglyceride. These compounds, along with
biliary salts, form micelles that are absorbed into the intestinal mucosal cells by passive
diffusion (Doreau and Chilliard, 1997). In the horse, pancreatic lipase is secreted in high
amounts and increases as fat is added to the diet (Frank et al., 2004). Therefore, the horse
is able to digest high amounts of fat in the diet. Horses have been fed diets with 20% of
the DE provided by oil with good results and no negative effect on digestibility (Cunha,
Once the monoglycerides and free FA are absorbed into the intestinal cell, the long-
chain fatty acids (LCFA) must be transported to the endoplasmic reticulum, the maj or site
of absorbed lipid metabolism. One explanation for how the LCFA reach the endoplasmic
reticulum is by the action of fatty acid-binding protein (FABP). FABP is present in the
intestinal mucosa, liver, kidney, and adipose tissue and has no affinity for short or
medium-chain FA (Tso, 1985). It has been postulated that FABP may be responsible for
removing LCFA acids from their binding to the cytosolic side of the luminal membrane
and transferring them to the endoplasmic reticulum (Carlier et al., 1991). Unlike LCFA,
short and medium-chain FA are transferred directly from the intestinal cell into the portal
blood as free FA bound to albumin (Carlier et al., 1991).
Once inside the endoplasmic reticulum, LCFA and monoglycerides are recombined
into triglycerides by the monoglyceride pathway. The enzyme complex that makes up
this pathway is known as "triglyceride synthetase." This complex consists of three
enzymes: acyl-CoA synthetase, MG transacylase and diglyceride transacylase. The acyl-
CoA synthetase, in the presence of CoA, activates the LCFA to form acyl-CoA. The
acyl-CoA is then used for the reacylation of monoglyceride to diglycerides and finally to
triglycerides (Tso, 1985). The resulting triglycerides are then packaged with cholesterol
esters and phospholipids into chylomicrons, which are large lipoproteins that act as
carriers of dietary triglycerides. Chylomicron formation is activated by the addition of
apoproteins, which are proteins that play an important role in the formation and secretion
of chylomicrons by the enterocytes. Once chylomicrons are formed, they are released by
exocytosis into the lymphatic system where they can enter the blood stream via the
thoracic duct and be transported to the rest of the body (Carlier et al., 1991).
De novo Fatty Acid Synthesis
There are two primary sources of FA in the body: FA provided by the diet and FA
made by the animal via de novo synthesis (Lehner and Kuksis, 1996). The pathways for
de novo FA synthesis exist in the animal during the well-fed state and in monogastrics
occur primarily in the liver. Most of the carbon used for de novo FA formation is
supplied through the pyruvate pool and from the end product of glycolysis. There are
three substances needed for FA synthesis: acetyl CoA, malonyl CoA and NADPH. The
first step in the synthesis of FA is the formation of acetyl CoA from pyruvate in the
mitochondrial matrix by the action of pyruvate dehydrogenase. The acetyl CoA must
then be moved out of the mitochondria and into the cytosol where FA synthesis takes
place. Because the inner mitochondrial membrane is not permeable to acetyl CoA, the
acetyl CoA is combined with oxaloacetate to form citrate. Citrate is then translocated to
the cytosol where it is cleaved back to oxaloacetate and acetyl CoA by ATP:citrate lyase
(Gurr et al., 2002). This mechanism of moving acetyl CoA into the cytosol in the form of
citrate is called the citrate shuttle.
Once acetyl CoA reaches the cytosol, de novo FA synthesis begins. The first
reaction of this mechanism, which is also the rate limiting reaction, involves the
formation of malonyl CoA by the enzyme acetyl-CoA carboxylase (ACC) (Knowles,
1989). The malonyl CoA forms the source of the vast maj ority of the carbons of a FA
chain. The enzyme complex that synthesizes LCFA from acetyl and malonyl CoA is
fatty acid synthase (FAS). This enzyme complex has synthase, reductase and dehydrase
actions. The typical end product of animal FAS action is palmitic acid (C16:0) (Greene,
Once produced, palmitic acid can be elongated and desaturated. Type III synthases
(commonly called elongases) lengthen FA preformed in the animal 2C at a time. The
principal elongation reactions occur in the endoplasmic reticulum membranes and
involve acyl-CoA as a primer, malonyl-CoA as a donor of 2C units and NADPH as the
reducing coenzyme. This system is capable of producing FA chain with an excess of 20
carbons (Suneja et al., 1990). Desaturation, or the addition of double bonds, occurs
mainly by oxidative desaturation, a process by which a double bond is introduced directly
into the LCFA with 02 and NADH as cofactors (Scheuerbrandt and Bloch, 1962).
Mammalian desaturases are only able to introduce double bonds in the A9, A6 and A5
positions. Plant desaturases can introduce additional double bonds at the Al2 and Al5
positions, therefore creating n-6 and n-3 FA. All double bonds introduced by the process
of oxidative desaturation are in the cis configuration (Lehner and Kuksis, 1996).
Fatty Acid Degradation
The mobilization and oxidation of FA occur primarily during fasting, physical
exercise and stress in the animal in order to break down dietary or stored TG into FA to
provide energy. The mobilization of FA occurs via lipolysis in the adipose tissue, in
which FA are cleaved from their glycerol backbone mainly by hormone sensitive lipase
(HSL) and released into circulation (Johnson and Greenwood, 1998). The main forms of
FA oxidation are termed alpha (ot), beta (P) and omega (co), referring to which carbon on
the acyl chain is attacked. Of these three, P-oxidation is the most prevalent. In P-
oxidation, there is a stepwise removal of 2C units from the carboxyl end of the FA
The mitochondria and microbodies (peroxisomes and glyoxysomes) are capable of
performing P-oxidation. The process begins by converting the FA into fatty acyl-CoA as
soon as it enters the cytosol of the cell. The inner mitochondrial membrane, however, is
impermeable to fatty acyl-CoA. In order to move this molecule across the membrane, the
enzyme carnitine:palmitoyl transferase (CPT1), located on the outer mitochondrial
membrane, combines the fatty acyl-CoA with carnitine. The resulting acyl carnitine is
then transported across the membrane, crossing the inner membrane by a
carnitine: acylcarnitine translocase (Pande, 1975). Once the acyl carnitine is inside the
mitochondrial matrix, CPT2 transfers the acyl group from carnitine to CoA, therefore
reforming acyl-CoA as a substrate for further P-oxidation (Bieber, 1988).
The process of P-oxidation involves a repeated sequence of four reactions resulting
in the removal of 2C from the acyl chain. First, acyl-CoA dehydrogenase acts on the
acyl-CoA to form trans-3,3-enoyl-CoA. Enoyl hydratase then acts on the product of the
first reaction to form 3-hydroxy acyl-CoA. The third reaction is catalyzed by the enzyme
3-hydroxy acyl-CoA dyhyrogenase which works with NAD+ to form 3-oxoacyl-CoA.
The final reaction involves 3-oxoacyl-CoA thiolase which produces a shorter fatty acyl-
CoA and acetyl-CoA (Bieber, 1988). The resulting acyl-CoA is recycled back into P-
oxidation for the removal of additional 2 carbon units, while the acetyl-CoA can be used
in the TCA cycle to produce energy (Gurr et al., 2002).
Polyunsaturated Fatty Acids
n-6 and n-3 Polyunsaturated Fatty Acids
By definition, n-6 polyunsatured fatty acids (PUFA) have the last double bond in
the FA chain six carbons from the methyl (omega) end. The two most physiologically
import n-6 PUFA are linoleic acid (LA; C18:2) and arachidonic acid (AA; C20:4). Of
these, LA is considered a dietary essential fatty acid because it cannot be synthesized by
mammals. Sources of LA include vegetable oils such as corn, sunflower, peanut, and soy
oils (Carlier et al., 1991). Linoleic acid can be elongated and desaturated in the body to
produce AA in a mechanism that is discussed later in this chapter. Omega-6 PUFA, with
AA as the principal component, predominate in organs and tissues performing storage
functions (adipose tissue), chemical processing (liver), excretion (kidney) and mechanical
work (muscle) (Innis, 1991). In addition, plasma lipoproteins contain high amounts of
LA in triglycerides, cholesterol esters and phospholipids (Innis, 1992a). A very
important feature of n-6 PUFA is their effect on the body. In general, n-6 PUFA have
proinflammatory, prothrombotic and proaggregatory effects, characterized by increases in
blood viscosity, vasospasm, vasoconstriction and decreases in bleeding time
Omega-3 PUFA have the last double bond in their carbon chains three carbons
from the methyl end. The dietary essential PUFA from the n-3 family is ot-linolenic acid
(ALA; 18:3), but other physiologically important n-3 PUFA include eicosapentaenoic
acid (EPA; 20:5) and docosahexaenoic acid (DHA; 22:6) (Innis, 1992a). Using elongase
and desaturase actions similar to those in n-6 PUFA, ALA can be transformed into EPA,
which can be further transformed into DHA. Alpha-linolenic acid is found primarily in
the chloroplast of green leafy vegetables and in seeds of flax, linseed and walnuts. Fatty
fish and fish oils, however, are the main sources of EPA and DHA (Benatti et al., 2004).
The primary sites of n-3 PUFA accumulation in the body include the nervous tissue,
reproductive organs and retina membranes (Innis, 1991). Unlike the plasma
concentrations of LA, tissue and plasma triglyceride and cholesterol ester levels of ALA
are usually quite low (<1-2% FA) (Innis, 1992a). Polyunsaturated FA of the n-3 family
are known to have anti-inflammatory, antithrombotic, antiarrhythmic, hypolipidemic and
vasodilatory effects on the body (Simopoulos, 1999). To obtain optimal health, it is
important to have adequate dietary amounts of PUFA of both the n-6 and n-3 families,
but it may also be important to have a proper ratio between the two. A ratio of 4-5: 1 of
n-6:n-3 has been suggested as most beneficial for humans, but most investigation in this
area has been conducted in lab animals (Wiseman, 1997).
The efficiency at which the horse converts ALA to EPA and DHA is unknown. In
addition, while a recommendation for a beneficial n-6:n-3 ratio exists for humans, the
optimal ratio for horses is unknown. Most horse feeds today are high in n-6 FA, with the
horse' s maj or n-3 FA intake obtained from forages. The FA composition of common
grains and fat supplements fed to horses are presented in Table 2-1 and the FA
composition of common forages fed to horses are presented in Table 2-2.
Table 2-1. Fatty acid composition of common feeds and fat supplements fed to horses
SPresented as g fatty acid/100 g fat
2 COmmercial grain mix (Hallway Feeds, Lexington,
KY) containing barley, corn,
soybean meal, molasses, oats and supplemental pellet; 14.8% CP, 6.5% fat; from
O' Connor et al., 2004.
3 From Ellis and Hill, 2005.
4 From Chen et al., 2006.
SFrom Francois et al., 2003.
6 From Sierra et al., 2005.
SNA information not available.
Table 2-2. Fatty acid composition of common forages fed to horses
Fatty Fresh Fresh Perennial Bermudag:rass .ioh Ha4
acidly Bahiagrass2 Rye Grass3 Hay
C14:0 0.00 0.4 0.00 1.63
C16:0 22.56 14.6 39.14 NA
C18:0 4.28 1.2 6.72 NA
C18:1 3.00 1.7 7.05 NA
C18:2n-6 21.32 10.6 23.35 15.76
C18:3n-3 46.21 68.4 15.93 26.68
C20:4n-6 0.00 NA5 0.00 0.35
C20:5n-3 0.00 NA 0.00 0.36
C22:6n-3 0.00 NA 0.00 0.25
Presented as g fatty acid/100 g fat.
2 From the present study.
3 From Elgersma et al., 2003.
4 From O'Connor et al., 2004.
SNA information not available.
Elongation of and Competition Between n-6 and n-3 Families
As stated earlier, both LA and ALA can be elongated and desaturated to form their
longer chain derivatives (Figure 2-1). This conversion happens in the endoplasmic
reticulum (Benatti et al., 2004). The first step of the mechanism converting LA to AA is
catalyzed by A6-desaturase, the rate-limiting step of the pathway. This enzyme acts on
LA to insert a double bond between carbons 6 and 7. This product is then elongated by
the addition of two carbon units to form dihomo-y-linoleic acid (C20:3). Further
desaturation by A'-desaturase inserts a double bond between carbons 5 and 6, thereby
creating AA. Arachidonic acid can then be elongated to form adrenic acid (C22:4). The
enzyme A6-desaturase inserts a double bond between carbons 4 and 5 of adrenic acid to
form co6-docosapentaenoic acid (C22:5) (Innis, 1991).
(Vegetable fats and oils)
(animal fat) 1 ASdesaturation I
1 elongation \
C22:4n-6 C22:5n-3 (fish fat)
Figure 2-1. Essential fatty acid metabolism. Adapted from Innis, 1992a.
The conversion of ALA to its longer chain derivatives uses the same pathway and
enzymes as LA. The enzyme A6-desaturase acts first on ALA to form stearidonic acid
(C18:4). This acid is then elongated and desaturated by A'-desaturase to form EPA. To
form DHA, EPA is elongated to form co3-docosapentaenoic acid (DPA; C22:5), which is
then desaturated by A6-desaturase to form DHA (Innis, 1991). However, there is a
marked inefficiency of conversion of ALA to EPA, with only about 0.2% of plasma ALA
fated for synthesis of EPA in human blood (Pawlosky et al., 2001). There is 10-fold
greater rate of transfer, however, of EPA to DHA than there is from ALA to EPA,
showing that the initial desaturation/elongation to EPA is the most restrictive (Pawlosky
et al., 2001). The difficulty of conversion of ALA to DHA has been shown in rats, where
maternal rats were fed a diet high made in ALA by the addition of flaxseed oil (Bowen
and Clandinin, 2000). Maternal rats were started on the experimental diet on the day of
parturition and their pups were sacrificed at two weeks of age. Bowen and Clandinin
(2000) showed that supplementing maternal rats with a high ALA diet did not increase
the DHA content of the whole body, skin, epididymal fat pads or muscles in rat pups,
therefore suggesting that the conversion of ALA to DHA is inefficient in the rat.
However, the efficiency at which the horse converts ALA to EPA and DHA is unknown.
Therefore, providing animals with a dietary source of EPA and DHA (such as fish or fish
oil) may be a better way to ensure incorporation of these FA into the body than feeding a
source of ALA.
Because the n-6 and n-3 families use the same enzymes in the process of
desaturation to their longer chain derivatives, there is competition between them. The
maj or site of competition occurs at the site of A6-desaturase action, the rate limiting
reaction for PUFA desaturation. There is a strong preferential substrate affinity of the A6-
desaturase for n-3 PUFA, especially ALA over LA (Innis, 1991; Drevon, 1992).
Therefore, feeding animals a source of n-3 PUFA will often decrease the amount of n-6
PUFA processed in the body, as more of the A6-desaturase will act on the n-3 PUFA and
less on the n-6. Studies have shown, in both animals and humans, that providing a
dietary source of n-3 PUFA reduces the amount of AA found in the blood (Fritsche et al.,
1993; Sauerwald et al., 1996). This competition between n-3 and n-6 PUFA has been
established in sows assigned to diets containing 7% added fat where menhaden fish oil
was substituted for lard at 0, 3.5 and 7% of the total dietary fat (Fritsche et al., 1993).
Sows were fed from 107 days of gestation to 28 days of lactation. The substitution of
fish oil for lard at both 3.5 and 7% decreased serum levels of AA by approximately 50%
in sow serum (Fritsche et al., 1993). However, the opposite phenomenon has been
documented as well. Extensive research has shown that providing a diet rich in LA but
poor in ALA will result in the accumulation of AA and very little EPA and DHA
Eicosanoid Production and Function
Eicosanoids are a large family of oxygenated 20-carbon FA (Smith, 1989) that act
as local hormones to modulate the intensity and duration of inflammatory and immune
responses (Yaqoob, 2004). The family is made up of three groups: the prostanoids
(prostaglandins and thromboxanes) which are synthesized by cyclooxygenase (COX), the
leukotrienes which are synthesized by lipoxygenase (LOX) and the epoxides synthesized
by epoxygenase. Eicosanoids are produced from 20-carbon PUFA containing three to
five cis, methylene-interrupted double bonds. These PUFA include AA, a member of the
n-6 family, and EPA, a member of the n-3 family. Linoleic acid (18 carbons) and DHA
(22 carbons) can be converted to eicosanoid homologues, but these are not actual
eicosanoids and are thought to have limited biological function. Because AA is the most
abundant C20 pOlyunsaturate in mammalian systems, it is the maj or precursor of
eicosanoids (Smith, 1989). Macrophages and monocytes are important sources of
eicosanoids, as their membranes typically contain large amounts of AA (Yaqoob, 2004).
Arachidonic acid and EPA each produce eicosanoids of a different series.
Arachidonic acid is a substrate for the 2-series prostaglandins (PG), namely prostaglandin
E2 (PGE2) and prostaglandin F2 (PGF2) and the 4-series leukotrienes (LT), namely
leukotriene B4 (LTB4). Prostaglandin E2 and LTB4 have powerful proinflammatory
actions (James et al., 2000). Prostaglandin E2 induces fever and increases vascular
permeability, vasodilation, pain and edema. However, PGE2 alSo suppresses the
production of inflammatory cytokines TNF-ot, IL-1 and IL-6 by macrophages and T cells.
Leukotriene B4 inCreaSCS Vascular permeability and blood flow, is a chemotactic agent for
leukocytes, induces the release of neutrophil lysosomal enzymes, and enhances the
generation of reactive oxygen species. Leukotriene B4 alSo increases production of TNF-
ce, IL-1 and IL-6 by macrophages (Calder, 2001; Yaqoob, 2004). In contrast to AA, EPA
is a substrate for the 3-series PG, namely PGE3, and the 5-series LT, namely LTBS.
These eicosanoids have the same types of inflammatory effects as those generated from
AA, but they are far less biologically potent (Calder, 2001). Therefore, production of
eicosanoids from EPA could modulate the immune response.
The Immune System
The acquired immune system is capable of recognizing and selectively inhibiting
specific foreign antigens (Goldsby et al., 2003). T cells, B cells, antigen-presenting cells,
the maj or histocompatibility complex (MHC), and immunoglobulins all play important
roles in the acquired immune system. This system of immunity is classified as acquired
because the immune cells must be exposed to an antigen once to develop, or acquire,
memory for that antigen. A second exposure to the same antigen will trigger an enhanced
state of immune reactivity (Goldsby et al., 2003).
Along with playing an important role in acquired immunity, immunoglobulins are
also an important part of humoral immunity, or the type of immunity pertaining to
extracellular fluids including the plasma and lymph (Goldsby et al., 2003). Humoral
immunity is driven by B cells, which originate and mature in the bone marrow (Kuby,
1992). Immunoglobulins are a group of large glycoproteins found on B-cell membranes
or secreted by plasma cells. They are found most prevalently in blood serum but are also
present in mucosal tissues and external secretions such as milk. An antibody is an
immunoglobulin (Ig) that exhibits antigen-binding ability. Therefore, all antibodies are
Ig, but not all Ig are necessarily antibodies. The two terms, however, are often used
interchangeably. Antibodies have a wide range of functions, including targeting
infectious organisms, neutralization of toxins and removal of foreign antigens from body
circulation (Peakman and Vergani, 1997). Antibodies can serve as diagnostic tools for
clinical evaluations of immune diseases or disorders. For example, immunoglobulin G
(IgG) is measured in the serum of foals to determine if there has been a successful
transfer of maternal antibodies.
In horses, the maj or immunoglobulins are IgG, IgM, IgA and IgE (Nezlin, 1998).
Average concentrations of immunoglobulins in the serum of mature horses are presented
in Table 2-1.
Table 2-1. Immunoglobulin concentrations of serum and milk in mature horses
Sample IgG IgM IgA
Adult horse serum 1000-1500 100-200 60-350
Mare colostrum 1500-5000 100-3 50 500-1500
Mare milk 20-50 5-10 50-100
1From Tizard, 1996; presented as mg/dL.
IgG is synthesized and secreted from plasma cells found in the spleen, lymph nodes
and bone marrow (Tizard, 1996). IgG molecules have a long half-life (23-25 days) and
there is a continuous high-level stimulation for IgG production. As a result, the
concentration of IgG in blood and colostrum is higher than any other immunoglobulin
(Tizard, 1996). IgG is the smallest of the immunoglobulin classes, so it is therefore more
able to move through the body to travel to needed areas (Widmann and Itatani, 1998).
Four immunoglobulin subclasses have been described in horses: IgGa, IgGb, IgGc and
IgG(T) (Sheoran et al. 2000). IG(T) has also been divided into the subclasses IgG(Ta)
and IgG(Tb) (Tizard, 1996). These subclasses are distinguished from one another by
molecular structural differences and slight variations in biological function (Kuby, 1992).
Like IgG, IgM is also made and secreted from plasma cells in the spleen, lymph
nodes and bone marrow. It is found in the second highest concentration in serum,
following IgG (Tizard, 1996). IgM is the first immunoglobulin class produced by the
maturing B cell, and the first class synthesized by the neonate (Kuby, 1992). It is also the
first antibody produced in a primary response to an antigen (Widmann and Itatani, 1998;
Kuby, 1992). IgM is more efficient than other immunoglobulins in binding antigens
because of its larger molecular size (largest of the immunoglobulin classes) and its larger
number of binding sites. Because of its higher efficiency, IgM is also more able to
neutralize viral infectivity, cause agglutination and activate compliment than IgG
(Goldsby et al., 2003).
The immunoglobulin IgA is produced mainly by plasma cells in muscosa-
associated lymphoid tissues beneath surface epithelium (Widmann and Itatani, 1998).
While it is manufactured more than any other immunoglobulin class, serum concentration
of IgA is relatively low. This low concentration is due to the secretion of IgA in fluids
present on the epithelial surfaces of the alimentary, respiratory and reproductive tracts
and in such fluids as urine, saliva, tears and milk (Widmann and Itatani, 1998). Because
IgA is the maj or immunoglobulin in the external secretions of horses, it plays a vital role
in protecting the intestinal tract, respiratory tract, urogenital tract, mammary gland and
eyes against microbial invasion (Tizard, 1996).
Like IgM, IgE is produced predominantly by plasma cells located beneath body
surfaces. It is found in very low concentrations in the serum of healthy animals, partially
because the molecule is fairly unstable and has the shortest half-life of all the classes of
immunoglobulins (Tizard, 1996). IgE antibodies mediate immediate (type I)
hypersensitivity reactions that cause the symptoms of hay fever, asthma, hives and
anaphylactic shock (Kuby, 1992). IgE is also thought to be largely responsible for
immunity against parasitic worms (Tizard, 1996).
Passive Immunity in the Foal
Due to the mare's diffuse epitheliochorial placenta, there is no significant transfer
of immunoglobulins to the fetal circulation during pregnancy (Jeffcott, 1972, 1974a;
Erhard et al., 2001). Therefore, foals are born with a near absence of circulating
immunoglobulins and an easily compromised immune system. Although they are able to
produce their own antibodies soon after birth, foals will not produce levels approaching
those of the adult horse until 3-4 months of age (Jeffcott, 1974a). Foals receive the
needed antibodies via passive transfer from colostrum, or the mare's first milk. Prior to
birth, the mare's mammary gland is capable of selecting and concentrating a wide range
of serum Ig into the colostrum (Jeffcott, 1974a, 1975). When foals suckle this colostrmm
after birth, they take the antibody-rich fluid into their digestive tracts where the Ig can be
absorbed into the circulating blood.
For the transfer of passive immunity to be successful, the mare' s colostrum must
contain adequate amounts of the appropriate immunoglobulins, especially IgG (Rooke
and Bland, 2002). In addition, the immunoglobulins must be delivered intact to the site
of absorption and absorbed intact (Rooke and Bland, 2002). Because of the very low
level of proteolytic activity in the digestive tract of young foals, most immunoglobulins
are kept intact as they pass with the colostrum through the foal's stomach and small
intestine. Trypsin inhibitors found in colostrum further reduce the degradation of
immunoglobulins in the foals digestive tract (Krse, 1983; Tizard, 1996).
The immunoglobulins in colostrum are rapidly absorbed by non-specific
pinocytosis into the small intestine enterocytes. Maximum absorption occurs soon after
birth and declines thereafter, completely ceasing by 24 hours after birth (Raidal et al.,
2000). The foal's intestine shows selective permeability, with a greater affinity for IgG
and IgM (Tizard, 1996). Once inside the enterocyte, individual immunoglobulins merge
together to form one or more larger globules. These larger globules pass from the
enterocyte into the local lymphatics and later reach the systemic circulation (Jeffcott,
The critical event in the transfer of intact immunoglobulin to the foal's circulation
is cessation of transfer across the enterocyte basolateral membrane. For this reason, "gut
closure" is the term used to define the cessation of transfer of IgG to the foal's
circulation. Gut closure, usually reached around 24 hours of age (Rooke and Bland,
2002), is characterized by a replacement of the immature epithelial cells with more
mature cells that no longer engage in pinocytosis (Kruse, 1983).
IgG displays a unique behavior in foal serum. At birth, before the foal has suckled,
foal serum IgG may be as low as 30 mg/dL (Erhard et al., 2001). However, foal IgG rises
rapidly after colostrum is ingested. Peak IgG values in foal serum occur between 18 and
24 hours after birth (Jeffcott, 1974a) and have been reported to be as high as 2, 160 mg/dL
(McGuire and Crawford, 1973). The IgG values in foal plasma appear to stay at near
peak levels for at least the first two days after foaling (Jeffcott, 1974b; Duvaux-Ponter et
al., 2004). After this peak, passively derived IgG molecules will gradually decline until
they are completely absent by around 5 months of age (Jeffcott, 1974a). Foals may begin
to process their own IgG molecules as early as 2 weeks of age, but levels reaching those
of the adult horse are not seen until around 4 months of age (Jeffcott, 1974a). Erhard et
al. (2001) reported that 7 day old foals had a mean IgG value of 1000 ml/dL. This value
then decreased, indicating the elimination of maternal IgG, and reached the lowest level
of 790 mg/dL at 35 days of age. However, foal serum IgG increased to around 1 100
mg/dL at 42 days after birth, indicating that endogenous IgG production was increasing
in the foal (Erhard et al., 2001). Therefore, behavior of IgG in the foal seems to begin
with a very low presuckle value, experience a dramatic rise after colostrum ingestion,
undergo a steady decline as maternal IgG is eliminated and shows an increase as the foal
begins to produce its own IgG.
Failure of Passive Transfer
Failure of passive transfer (FPT) is defined as the failure of absorption of maternal
immunoglobulins by the neonatal foal, a condition that predisposes the foal to life-
threatening infections (Kohn et al., 1989). Failure of passive transfer is the most
commonly recognized immune deficiency in horses and may predispose affected foals to
septicemia, infective arthritis and pneumonia (Raidal, 1996; Raidal et al., 2000). There
are conflicting views in the literature as to what antibody levels actually constitute failure
of passive transfer. Liu (1980) and McGuire et al. (1977) defined failure of passive
transfer as less than 200 mg IgG/dL serum and partial failure as between 200 and 400
mg/dL. LeBlanc et al. (1986) suggested failure of passive transfer as levels below 400
mg/dL, while Raidal (1996) and Tyler-McGowan et al. (1997) noticed an increased
susceptibility of foals to disease when IgG levels dropped below 800 mg/dL. Today, it is
common to define failure of passive transfer as IgG levels below 400 mg/dL serum
(Tizard, 1996; Erhard et al., 2001) and partial failure as levels between 400 and 800
mg/dL. IgG levels above 800 mg/dL are considered necessary to provide optimal
immune function (Erhard et al., 2001). In order to prevent failure of passive transfer, the
minimal concentration of colostral immunoglobulin required has been estimated to be
between 1,000 and 3,000 mg IgG/dL colostrum (LeBlanc et al., 1986). The possibility of
failure of passive transfer cannot be evaluated until the foal is about 18 hours of age, as
antibody absorption is essentially complete at this time (Tizard, 1996). Therefore, the
standard industry practice of testing foal IgG levels at 12 hours after foaling may give
misleading results, as complete antibody absorption has not happened yet. This practice
may still be needed, however, in order to be able to administer plasma or colostrum to the
foal before the foal's ability of absorption is completed.
Possible causes of failure of passive transfer fall into three categories: production
failure, ingestion failure and absorption failure (Tizard, 1996). A failure of the mammary
gland to concentrate immunoglobulins from the blood into colostrum can occur in maiden
mares foaling for the first time. Premature lactation, however, is the most common
production failure cause of failure of passive transfer. In this case, initial colostrum may
be of adequate amount with adequate IgG concentration, but the mare will commence
lactation prior to parturition. This steady leak of colostrum may occur for several hours
or several days before birth and significantly reduces the amount of IgG available to the
foal (Jeffcott, 1974a).
While less common, inabilities of ingestion and absorption are additional failure of
passive transfer causes. Ingestion failures can arise from a mare not allowing her foal to
nurse, weak or deformed foals that take longer than normal to stand, or a delayed or
defective suckling reflex (Jeffcott, 1974a; Tizard, 1996). Foals usually overcome these
factors, but it may take longer than the 24-hour period of intestinal permeability to IgG.
Absorption failures can be linked to stress at the time of parturition. The adrenal
hormones play important roles in the onset of parturition and can influence changes in the
permeability of small intestine cells after birth (Jeffcott, 1972). In conditions of stress at
parturition, the mare or foal could produce abnormal amounts of corticosteroids which
would therefore have a detrimental affect on the foal's antibody absorption (Jeffcott,
Innate immunity is the first line of defense in Eighting an invading organism
(Goldsby et al., 2003). The skin, mucosal surfaces, macrophages and neutrophils all play
important roles in the innate immune system. The skin and mucosal surfaces act as
barriers against infection, and the macrophage and neutrophils act to phagocytize and kill
invading foreign cells. Inflammation is also an important part of innate immunity and
functions to draw immune cells to areas of injury or antigen attack. Because the innate
immune system is less specific than the acquired immune system, the innate system can
act quickly to begin an immune response (Goldsby et al., 2003).
By definition, inflammation is the response of tissue to the presence of
microorganisms or injury (Tizard, 1996). Inflammation is a vital protective mechanism
and the means by which defensive molecules and phagocytic cells gain access to the sites
of tissue microbial invasion or damage. Inflammation is classified according to its
severity and duration, with acute inflammation developing in less than an hour after
tissue damage and chronic inflammation occurring a much slower rate and being more
constant. There are five symptoms of acute inflammation: heat, redness, swelling, pain
and loss of function. These symptoms are a result of changes in the small blood vessels
in the damaged tissue (Tizard, 1996; Goldsby et al., 2003).
Immediately following microbial invasion or injury, blood flow to the effected area
greatly increases. This increase is due to a transient constriction of local arterioles and
dilation of all the small blood vessels in the area. The blood vessel permeability is also
increased, allowing fluid to move from the blood into the tissues where it causes edema
and swelling (Tizard, 1996). The changes in blood vessels allow an influx of
lymphocytes, neutrophils, monocytes and other immune cells into the area to participate
in clearance of the antigen (Kuby, 1992). Neutrophils are the first immune cells to arrive
in the inflamed tissues, followed by the slower moving monocytes. Once within the
inflamed tissues, these cells are attracted to sites of bacterial growth and tissue damage
and phagocytize and destroy any foreign material present. Monocytes will also remove
dead and dying tissue (Tizard, 1996).
Cytokines play an important role in the acute-phase inflammatory response. These
low-molecular-weight proteins secreted by macrophages exert a variety of effects on
lymphocytes and other immune cells to regulate the intensity and duration of an immune
response. The three cytokines that play the largest role in acute inflammation are tumor
necrosis factor-a (TNF-u), interleukin 1 (IL-1) and interleukin 6 (IL-6). All three
cytokines act locally on endothelial cells to induce coagulation and increase vascular
permeability (Kuby, 1992). TNF-u, IL-1 and IL-6 also act on the brain to induce fever
and suppress appetite and on skeletal muscle to drive protein catabolism and mobilize
available amino acids. In addition, these cytokines operate on liver cells to increase
protein synthesis and secretion of clotting factors, complement components and protease
inhibitors, all of which aid in the host defense (Tizard, 1996). All three cytokines
activate B and T cells, while IL-6 can also increase immunoglobulin synthesis (Goldsby
et al, 2003).
Effects of Dietary PUFA Supplementation on Inflammation and Immune Function
Blood and Tissue Responses to Experimental Feeding of n-3 PUFA
Numerous studies have investigated the levels of different n-3 PUFA resulting in
the blood when feeding ALA and DHA, either alone or in combination. In humans,
many studies have looked at providing adults with dietary fish oil, a good source of EPA
and DHA. Helland et al. (1998) supplemented pregnant women with cod liver oil for 14
days, between three and eight weeks post partum. The women were divided into four
groups: Group 1 served as the control and received no supplementation, Group 2
received 2.5 mL of cod liver oil/day, Group 3 received 5 mL of cod liver oil/day, and
Group 4 received 10 mL of cod liver oil/day. Helland et al. (1998) found that the
pregnant women in Groups 3 and 4 (receiving 5 and 10 mL of cod liver oil/day,
respectively) showed a decreased plasma LA content and an increased ALA and DHA
plasma content. When EPA and DHA intake were computed on a bodyweight basis, the
women receiving 5 mL of cod liver oil were consuming the equivalent of 14 mg EPA and
DHA/kg of bodyweight and the women receiving 10 mL of cod liver oil were consuming
the equivalent of 28 mg of EPA and DHA/kg bodyweight.
Henderson et al. (1992) also supplemented pregnant women with EPA and DHA,
but not in the form of fish oil. Henderson and coworkers supplemented pregnant women
with six capsules of Bio-EFA for a total supplement weight of six grams of
supplementation. This supplement provided women with 1080 mg EPA and 720 mg
DHA per day. Assuming an average bodyweight of 70 kg, this dosage provided 15.43
mg EPA/kg BW and 10.29 mg DHA/kg BW. Similar to Helland et al. (1998), however,
Henderson et al. (1992) also started their supplementation period after lactation had
already commenced, supplementing women between two and five weeks post partum for
a total of 21 days. The results of Henderson et al. (1992) showed that daily
supplementation of lactating women with 6 g of an EPA and DHA source increased the
women's red blood cell content of EPA, DPA and total n-3 PUFA. Although it was not
significant, there was also a trend toward red blood cell DHA increase. Infant red blood
cells were also affected by supplementation of the mother, as EPA and DPA
concentrations of infant red blood cells significantly increased after the supplementation
period. However, similar to maternal results, there was no significant change in infant
red blood cell DHA. Unfortunately, this study only used five women and their infants, so
it may have been hampered by a small sample size.
Further evidence suggests that infants breast fed from omnivorous mothers have a
higher DHA concentration in their red blood cells than do infants of vegan mothers
(Sanders and Reddy, 1992). In fact, the difference in infant red blood cell DHA was
quite large in this study, with infants feeding from vegan mothers having 1.9% of the
total FA found in their red blood cells as DHA and infants feeding from omnivorous
mothers having 6.2% of their total red blood cell FA as DHA. The content of DHA in
breast milk reported by Sanders and Reddy (1992) was also significantly lower in vegan
women when compared to omnivorous women (0. 14 and 0.37% of fat, respectively). In
addition, infants fed conventional formula (low in DHA) have consistently lower plasma
and red blood cell levels of DHA than infants fed breast milk, which is higher in DHA
(Innis, 1991; Innis, 1992b).
The ability to increase blood concentrations of n-3 PUFA by feeding sources of
these FA has also been documented in animals. Bauer et al. (1998) fed adult dogs either
ground flax or sunflower seeds for 84 days and showed that plasma ALA, EPA and DPA
were elevated when dogs were fed flaxseed, compared to when dogs were fed sunflower
seeds. Plasma DHA, however, remained unchanged in the flax fed dogs, showing the
difficulty in converting ALA to DHA. The flax fed dogs also showed a plasma reduction
in AA and 22:5 n-6, providing evidence of competition between n-3 and n-6 PUFA for
the A6-desaturase in dogs (Bauer et al., 1998). However, the exact amount of supplement
Bauer and coworkers added to the diets of the dogs was not stated, so it is therefore
difficult to compare levels of supplementation to other studies.
In horses, Hansen et al. (2002) examined the effects of ALA supplementation on
equine fatty acid status by feeding adult horses a diet consisting of 8% flaxseed oil for 18
weeks. Hansen and coworkers found that the flaxseed oil supplemented horses showed
an increased plasma ALA and EPA compared to horses that did not receive any fat
supplementation. On the other hand, there were no increases in DHA noticed. A
weakness of this study, however, is the low sample size of only 12 horses (6 horses in the
control group, 6 horses in the supplemented group). Duvaux-Ponter et al. (2004) also
tested the effects of ALA on horses, but used pregnant mares and young foals as subj ects.
In this study, 26 pregnant mares were divided into two groups. The first group acted as
the control and was fed extruded rapeseed (high in n-6 FA), while the second group was
fed extruded linseed (high in n-3 FA). The mares were supplemented 1.5 months prior to
foaling until one month after foaling. While mare blood was not tested, supplementation
with extruded linseed caused an increase in the ALA content of foal plasma from foaling
until 4 weeks post parturition, and this increase was greater than the increase seen in foals
nursing the mares given rapeseed (Duvaux-Ponter et al., 2004). However, the exact
amount of linseed provided to the mares is unclear, as it was not stated in the paper.
The effects of feeding sources of EPA and DHA have also been documented in
horses. Hall et al. (2004a) fed ten adult mares either menhaden fish oil or corn oil for a
period of 14 weeks. Mares fed the menhaden fish oil consumed 22.93 g EPA per day and
19.58 g DHA per day. These amounts equated to a daily intake of 4.6 g EPA/100 kg
bodyweight and 3.9 g DHA/100 kg bodyweight. As a result of this supplementation, Hall
et al. (2004a) noticed higher plasma ALA, EPA and DHA and lower plasma LA in the
mares fed fish oil compared to the mares fed corn oil. Brinsko et al. (2005) examined the
effects of feeding a DHA source to stallions to determine the effects of FA
supplementation on semen. In this study, eight stallions were used in a 2 x 2 crossover
design. Stallions were fed a grain mix top-dressed daily with either 250 g of a
commercial nutriceutical containing 30% n-3 FA (resulting in 75 g of n-3 FA) or a grain
mix with no supplementation. Stallions were fed their respective diets for 14 weeks,
separated by a 14 week wash out period, before treatments were switched for another 14
weeks of supplementation. The authors found that when stallions were supplemented
with n-3 FA, their semen had almost three times the levels of DHA/billion sperm
compared to when stallions were not supplemented. Even though the relative percentage
of DHA in semen fatty acids was not significantly different between the two groups,
treated stallions showed a 1.5-fold increase in their semen DHA:DPA ratio when they
received n-3 supplementation (Brinsko et al., 2005). Therefore, it seems that
supplementation of DHA in horses may have the ability to affect the fatty acid
composition of body constituents other than just the plasma.
Effects of PUFA supplementation on the acquisition of passive immunity in the foal
In order for IgG to be maximally absorbed into the foal's intestinal cells, the cell
membranes must be fluid enough to allow the molecules to navigate through the
membrane. Membrane bilayers tend to exist at the transition point between a fluid and
solid-like (gel) state. The phospholipid fatty acyl chains present in membranes are one of
the key chemical determinants of this balance. PUFA of the cis configuration tend to
increase the fluidity of the membrane (Murphy, 1990; Mills et al., 2005). The fatty acid
composition of membrane phospholipids is also easily changed by manipulation of
dietary fat (Murphy, 1990), which, in tumn, can influence membrane fluidity.
Brasitus et al. (1985) showed that adjusting the fatty acid composition of the diet in
rats changed the composition of their enterocyte membranes. To do so, rats were fed
either unsaturated or saturated triglycerides provided by comn oil or butter fat,
respectively, for 6 weeks. The supplementation of comn oil, which is rich in LA, caused
an enhanced fluidity of the membranes of several intestinal cells (Brasitus et al., 1985).
When humans were supplemented with dietary n-3 PUFA, the fluidity of their
erythrocyte membranes was substantially increased (Lund et al., 1999). In this study, 17
adults were supplemented with three 1 g capsules of fish oil per day for 42 days. Fluidity
of the red blood cell membrane was determined by measuring the lateral diffusion
coefficient of the fluorophore ODAF by fluorescence recovery after photobleaching. The
results of Lund et al. (1999) suggest that supplementation with fish oil increased the
lateral diffusion coefficient of ODAF, therefore increasing the membrane fluidity. This
increase in fluidity was seen at 21 days after supplementation and continued to rise until
the termination of the study at 42 days (Lund et al., 1999).
A correlation between membrane fluidity and permeability was shown in young
rats by Meddings and Theisen (1989). These researchers examined the changes that
naturally occur in the membrane of jejunal microvilli in rats as they aged from 9 to 25
days. Study results showed a decreasing lipid fluidity as the rats aged, assessed by a
steady-state fluorescence polarization technique. This decrease in membrane fluidity
correlated to a decrease in membrane permeability (Meddings and Theisen, 1989). This
correlation may suggest that if it is possible to enhance membrane fluidity by feeding n-3
FA, it may therefore be possible to augment the amount of IgG that travels both into the
mare's mammary gland and across the foal's intestine.
To determine the effects of feeding PUFA on the IgG content of mare colostrum,
Kruglik et al. (2005) fed mares either corn oil (rich in LA) or encapsulated fish oil (rich
in EPA and DHA) from 60 days before foaling to 21 days after foaling. The mares fed
encapsulated fish oil consumed 8.6 g of EPA and 10.4 g of DHA per day. The results of
Kruglik et al. (2005) showed a higher presuckle colostrum IgG content in the fish oil fed
mares, suggesting that supplementation with Hish oil may have improved the fluidity and
permeability of mammary epithelial cells. However, Kruglik et al. (2005) did not specify
the amount of colostrum collected, and it has been reported that IgG amounts can vary
between the first 250 and 500 mL of colostrum (Lavoie et al., 1989). Therefore, if the
volume of colostrum gathered was different between mares, the differences seen in IgG
may have been attributed to colostrum volume and not to treatment. Studies have also
been performed that suggest that n-6 PUFA may increase mare colostrum IgG. Hoffman
et al. (1998) fed mares a high fat diet (10 % fat) through gestation and lactation. The fat
in this diet was provided primarily by corn oil, which is high in LA. Colostrum IgG
levels were higher in the mares fed the high fat diet, even though the dietary fat was rich
in n-6 and not n-3 FA (Hoffman et al., 2004). However, the volumes of harvested
colostrum were not stated in this study. In addition, colostrum was collected between 6
and 12 hours after foaling. Because the IgG of colostrum can vary dramatically in the
first 12 hours after foaling (Pearson et al., 1984), colostrum IgG values obtained from
mares in this study may have shown differences that were due to time and not treatment.
Studies have also been executed to determine the effect of feeding PUFA on foal IgG.
Kruglik et al. (2005) showed that mares supplemented with Hish oil produced foals that
showed no differences in IgG levels when compared to foals born to corn oil fed dams,
even though the same study showed a higher colostrum IgG in the mares fed fish oil.
Kruglik et al. (2005) sampled foal plasma at 24 hours, so peak IgG content should have
been reached. Therefore, it is unclear as to why the higher IgG levels in fish oil fed mare
colostrum did not cause an increase in the plasma of foals born to these mares. Duvaux-
Ponter et al. (2004) also failed to show a difference in foal serum IgG when mares were
supplemented with a source of n-3 FA. In this study, mares fed extruded linseed did not
produce foals with a higher serum IgG than mares fed extruded rapeseed (Duvaux-Ponter
et al., 2004). Foal blood was again sampled at 24 hours after foaling in this study, so
peak foal IgG should have been recorded. However, mare colostrum IgG was not tested,
so it is unclear if ALA could increase colostrum IgG. More research needs to be done to
determine the capability of dietary PUFA on modifying mare mammary and foal
enterocyte membrane composition and fluidity on the enhancement of passive transfer of
Effects of PUFA supplementation on the inflammatory response
Numerous studies have shown anti-inflammatory effects of n-3 PUFA. Sadeghi et
al. (1999) fed groups of mice either a low fat diet (2.5% fat provided by corn oil) or diets
high fat diets providing 20% fat by coconut (rich in medium chain FA), olive (rich in
C18:1n-6), safflower (rich in LA) or fish oils. After 5 weeks on diet, mice were injected
with 1.0 mL of phosphate-buffered saline containing endotoxin from E. coli. In the mice
receiving the fish oil diet, lower plasma concentrations of the proinflammatory cytokines
TNF-u, IL-1 and IL-6 were seen after the inj section of endotoxin when compared to mice
fed olive or safflower oils. However, coconut oil fed rats also showed decreased amounts
of these cytokines (Sadeghi et al., 1999). Therefore, it is unclear if fish oil was
specifically responsible for the decreased proinflammatory cytokine production, or if this
difference was caused by a lack of n-6 PUFA. Unfortunately, the amount of feed offered
to the mice was not provided by the authors, so the amount of consumed FA could not be
calculated. Billiar et al. (1988) fed fish oil to rats for 6 weeks and observed a lower in
vitro production of IL-1 and TNF-a by macrophages. Unfortunately, the fish oil fed rats
in this study were compared to rats fed either corn or safflower oil, which are both high in
n-6 FA. Therefore, it is again unclear if fish oil reduces proinflammatory cytokine
production or if the feeding of n-6 FA increases cytokine production.
To test the inflammatory effects of PUFA in horses, Hall et al. (2004a) fed 10
adult mares 3.0% of their total diet (as-fed basis) with either corn oil or menhaden fish
oil. After 14 weeks of supplementation, the fish oil supplemented mares had neutrophils
with a 78-fold greater concentration of the lesser inflammatory LTBS when compared to
the neutrophils of mares fed corn oil (Hall et al., 2004a). In the same group of mares
during the same supplementation period, production of TNF-oc by bronchoalveolar lavage
fluid (BALF) cells was increased in both groups, but only the corn oil fed mares had an
increased production of inflammatory PGE2 their BALF cells (Hall et al., 2004b). When
mare BALF cells were stimulated with lipopolysaccharide, mares fed corn oil also
showed a higher production of inflammatory PGE2 (Hall et al., 2004b). Similar to the
studies discussed previously, however, Hall et al. (2004a, 2004b) compared horses fed
fish oil to horses fed corn oil and did not include a control group fed a diet without n-6 or
n-3 FA supplementation. Because the diets of both groups of horses were altered through
fat supplementation, it is unclear if the reported decreases in pro-inflammatory
eicosanoids seen in horses fed fish oil would have resulted if these horses had been
compared to a control group.
The delayed-type hypersensitivity (DTH) response has also been used to test the
anti-inflammatory effects of n-3 PUFA. Meydani et al. (1993) supplemental adult
humans with a low-fat, high-Hish diet (26% calories from fat, 1.23 g EPA and DHA
combined per day) or a low-fat, low-Hish diet (25% calories from fat, 0.27 g EPA and
DHA combined per day) for 24 weeks. This treatment period was compared to a
previous 6 week period where subj ects had been eating a current American diet of 3 5%
of calories from fat and 0.8% of calories from n-3 FA. A delayed-type hypersensitivity
(DTH) test was administered before and after the supplementation period using several
different antigens, including tetanus toxoid and Streptococcus (group C). Results of
Meydani et al. (1993) showed that the DTH response of adults consuming the low-fat,
high-fish diet was significantly less than the response of those consuming the low-fat,
low-fish diet, with diameter measurements of the DTH reactions of the low-fat, high-fish
diet participants being reduced by half.
Changing the n-6:n-3 ratio in dogs was also shown to effect the inflammatory
response (Wander et al., 1997). In this study, dogs were fed diets containing n-6:n-3
ratios of 31:1, 5.4: 1 or 1.4: 1 for 16 weeks, where the n-6 FA was provided by corn oil
and the n-3 FA was provided by fish oil. Dietary ratios were changed mostly by a
reduction in LA simultaneous to an increase in EPA and DHA. When the diameter of a
DTH response to keyhole limpet hemocyanin (KLH) was measured, dogs fed a n-6:n-3
ratio of 1.4 showed a much smaller reaction compared to the dogs fed ratios of 34: 1 and
5.4: 1 (Wander et al., 1997). Because the amount of n-6 FA in these diets decreased as
the n:6-n:3 ratios decreased (as opposed to holding the amount of n-6 FA stable and
increasing the amount of n-3), it is again unclear if differences seen in DTH response
were strictly caused by the increase in n-3 FA. It is quite plausible that these differences
may have been influenced by the decreasing n-6 FA. In contrast to dogs, the DTH
response of horses sensitized with KLH showed no differences between horses fed 3% of
the total diet (as-fed basis) either fish oil or corn oil (Hall et al., 2004b).
Effects of PUFA supplementation on disease resistance and survival
The maj ority of studies on inflammation and PUFA supplementation have shown
positive results with n-3 PUFA, particularly when n-3 supplementation reduces n-6 FA in
the diet. However, studies on the effect of PUFA on disease resistance show conflicting
results. When guinea pigs were fed diets high in either n-3 FA (1.4% and 0.9% fat
calories from EPA and DHA, respectively) or n-6 FA (15.4% fat calories from LA) for
13 weeks and infected with M~ tuberculosis, the guinea pigs fed a diet high in n-3 FA
showed a higher number of mycobacteria recovered from the spleen, the most
pronounced progression of the disease and a higher mean size of the tuberculin reaction
(Paul et al., 1997). The authors suggested that possible explanations for these results may
include the lower production of inflammatory mediators and the impairment in release of
lysosomal enzymes that kill mycobacteria (Paul et al., 1997). The study performed by
Paul et al. (1997) is possibly one of the best studies done to examine n-3 FA effects on
disease resistance, as animals consuming both fat supplemented diets were compared to a
no fat added control diet. In addition, the study utilized animals consuming the
experimental diets but that were not infected. These animals therefore acted as
uninfected controls within each diet. The benefits of a study design such as this is that a
direct comparison can be made between the n-3 FA supplemented group and the control
group, which in turn helps to determine disease effects are due strictly to the addition of
dietary n-3 FA.
Another well designed study compared disease responses of mice infected with
influenza (Byleveld et al., 1999). Challenging fish oil fed mice with influenza virus
produced a higher lung viral load, lower body weights and impaired production of IgG
and lung IgA when compared to mice fed beef tallow. Mice were fed fish oil or beef
tallow at 20% of dietary fat for 14 days, after which half of the mice from each treatment
were infected with influenza while the other half served as noninfected controls.
D'ambola et al. (1991) supplemented newborn rabbits with high (5 g/kg) or low (0.22
g/kg) doses of fish oil, safflower oil or saline for 7 days after birth. When the young
rabbits were supplemented with the higher levels of fat, both the fish and sunflower oil
supplemented rabbits had an impaired ability to clear Staphylococcus aureus when
compared to the saline control group (D'ambola et al., 1991). However, the low doses of
fish and safflower oil did not produce the same impaired ability to clear the bacteria. In
light of these results, the authors of this study concluded that high does of both n-3 and n-
6 FA can reduce the host's ability to kill S. aureus (D'ambola et al., 1991).
Positive results of supplementing with n-3 PUFA were shown when neonatal rat
pups were infected with group B streptococcus (Rayon et al., 1997). In this study,
researchers fed gestating rats a control diet (no fat added) or diets supplemented with
either comn or menhaden fish oil. Supplementation was begun on day 2 of gestation and
continued through lactation, but the amounts of diets and supplements fed were not
provided by the authors. Rat pups were then infected with the streptococcus bacteria at 7
days of age. The results of Rayon et al. (1997) showed that pups from mothers who had
been fed fish oil during gestation showed a significantly higher rate of survival (79%)
than those bomn to corn oil fed dams (49%), though this difference was not significant. In
this study, the lowered production of inflammatory mediators by fish fed rats when
compared to corn oil fed rats may have been responsible for the higher survival rates, as
group B streptococcal infections induce elevated levels of proinflammatory cytokines that
lead to septic shock (Rayon et al., 1997).
Feeding fish oil to weanling mice has been shown to prolong mice survival to a
murine retrovirus-induced immunodeficiency syndrome (MAIDS) that mimics human
AIDS (Femnandes, et al., 1992). Mice in this study were fed diets consisting of 5% corn
oil fed at an energy restriction of 40%, or diets fed ad libitum consisting of 5% comn oil,
20% corn oil or 20% menhaden fish oil. Mice were fed for 8 weeks before being inj ected
with the MAIDS plaque-forming units (Femnandes et al., 1992). Mice fed both the 5%
corn oil energy restricted and 20% fish oil diets showed significantly longer survival rates
than mice consuming the other diets. The authors explained the increase in survival rates
of these two groups as a result of a slowed the progression of the MAIDS disease
(Fernandes et al., 1992).
Thors et al. (2004) also showed positive immune effects on mice when feeding fish
oil. In this study, 120 female mice were fed a standardized, control diet for 6 weeks
before being divided into four groups and fed two different diets. The first two groups
were fed a diet enriched with fish oil at 10% of total diet weight, and the remaining two
groups were fed a diet enriched with corn oil at 10% of the total diet weight (Thors et al.,
2004). However, the amount of time these diets were fed was not clear. Mice were
intranasally inoculated with either Klebsiella or Streptococcus pneumoniae and the
inoculum was aspirated into the lungs. Survival rates of the mice fed a fish oil diet and
infected with Klebsiella pneumoniae were significantly higher than the rates seen in corn
oil fed mice infected with the same disease. However, survival rates of mice infected
with Streptococcus pneumoniae did not differ between the fish or corn oil fed mice
(Thors et al., 2004).
In general, the conflicting results of studies examining the effects of dietary PUFA
supplementation on disease resistance may be caused in part by study differences in the
type of animal used, type and amount of pathogen utilized, route of pathogen infection
and amount and duration of dietary PUFA supplementation. Because many of the above
studies did not clearly state this information, it is difficult to establish which differences
in disease response between studies could be attributed to PUFA treatment and which
could be attributed to differences in experimental design. However, Anderson and
Fritsche (2002) suggest that conflicting results may be rooted in the host' s ability to find
a proper balance between the necessary and excessive production of various
Characteristics of Mare Milk
Colostrum is the mare's first milk and is vital in transferring immunity to the
newborn foal. It has a much thicker, stickier consistency than milk and is often a pale to
deep yellow in color. Colostrum, produced in the mammary gland during the last
trimester of pregnancy, is only secreted for a very short time (Lavoie et al., 1989). By
24-96 hours after foaling, mammary secretions have completely transitioned from
colostrum to milk (Ullrey et al., 1966). Compositionally, colostrum is higher than milk in
fat content (Csap6, et al., 1995). The most important colostrum constituent, however, is
the immunoglobulins. Colostrum has high concentrations IgG but lower IgM and IgA
(Lavoie et al., 1989). Colostral IgG declines within the first 24 hours after birth. This
decline often corresponds to the change of a thick, pale yellow fluid to one of a thinner
consistency with a gray-white color (Pearson et al., 1984). Average colostral Ig
concentrations are shown in Table 2-1.
Factors Affecting Mare Colostrum IgG Content
Premature lactation, or "prelactation," is considered the most important cause of
failure of passive transfer in foals, as it is one of the main determinants of colostral IgG
levels (Jeffcott, 1974). Causes of premature lactation include placentitis and/or placental
separation, but the condition can occur without obvious placental pathology (Jeffcott,
1974b, 1975). Mares that experience prelactation for longer than 24 hours before foaling
tend to have lower colostral IgG concentrations than those who lactate normally (Koterba
et al., 1990). Morris et al. (1985) found that as the proportion of mares on a breeding
farm experiencing prelactation increased, so did the proportion of mares with low
colostral IgG concentrations. In addition, the proportion of foals with low serum IgG
concentration also increased.
Breed of mare may also affect colostral IgG concentration. Pearson et al. (1984)
found a significantly higher IgG concentration of more than 5,000 mg IgG/dL colostrmm
in Arabian mares when compared to Thoroughbred mares. Average time from birth until
colostrum IgG concentration declined to 1,000 mg/dL (the IgG concentration that cannot
prevent failure of passive transfer) was 19. 1 hours for the Arabians and only 8.9 hours for
the Thoroughbreds. LeBlanc et al. (1992) founder higher IgG colostral concentrations in
Thoroughbreds and Arabians when compared to Standardbreds. However, in another
study, LeBlanc et al. (1986) reported no differences between IgG colostral concentrations
in Thoroughbred, Quarter Horse, Arabian and Standardbred mares. The conflicting
results seen between these two studies should not have been due to different sampling
times or colostrum amounts taken, as both studies tested 10 mL of presuckle colostrum.
Therefore, the conflicting results may be explained by differences in body size and
weight between breeds. Larger breeds can sometimes produce larger volumes of
colostrum, and this large volume may lead to a dilution effect. However, the age of the
mare, number of lactations and herd management are factors that probably influence
colostral IgG concentration. Additionally, there is large individual variation in colostral
IgG content, making it difficult to attribute differences in colostral IgG as purely breed
oriented (Pearson et al., 1984). Further studies are needed to examine what, if any,
influence breed has on colostral IgG content.
Conflicting results exist in regard to the connection between a mare's age and her
colostrum quality. In a study involving Standardbred, Thoroughbred and Arabian mares,
mares between the ages of 3 and 10 years had the highest colostral IgG concentration and
FPT was most prevalent in foals born to dams of over 15 years (LeBlanc et al., 1992).
However, Morris et al. (1985) and Erhard et al. (2001) saw no significant effects of age
on IgG in mares of varying breeds. Both LeBlanc et al. (1992) and Erhard et al. (2001)
sampled colostrum before the foal had been allowed to suckle. However, Morris et al.
(1985) sampled colostrum during the first 2 hours after foaling. Since Morris et al.
(1985) sampled colostrum at a later time than the other two studies, any difference seen
in the colostrum of this study could have been attributed to time. However, time should
not have affected the values of LeBlanc et al. (1992) and Erhard et al. (2001). Therefore,
discrepancies in data reflecting the effect of mare age of colostrum IgG could be
explained by outside factors such as individual mare variation and management
Composition of Mare Milk
In mares kept without human influence, lactation lasts about one year, and drying
of the udder occurs several weeks to several days before the next foaling. There have
been, however, extreme cases noted of 2- or 3-year-old suckling foals (Feist and
McCullough, 1976). Today, the drying process is initiated by weaning foals at 4-6
months of age. Actual daily lactation yields of nursing mares are not well known, but are
estimated to be between 10 and 30 kg for light breed nursing mares (Doreau and Boulot,
1989). Peak lactation seems to occur at about two months postpartum (Bouwman and
van der Shee, 1978).
Compositionally, the fat content of mare milk is very low (Doreau and Boulot,
1989) but can be influenced by diet. Milk fat is also influenced by mare body condition
at foaling, with fat mares producing milk with a higher fat content than thin mares. The
increased lipid mobilization of fat mares may be explained this phenomenon (Doreau et
al., 1993). Crude protein in milk exists at between 1.7 and 3.0% (Doreau and Boulot,
1989) and decreases throughout lactation (Oftedal et al., 1983). Mare milk is different
from the milk of other species as it contains higher amounts of the amino acids cystine
and glycine (Doreau and Boulot, 1989). Milk carbohydrates are almost entirely made of
lactose, with very low levels of free glucose. Mare milk is also extremely low in ash,
with 0.7% as extreme (Doreau and Boulot, 1989). Milk is also different than colostrmm
in the amounts of immunoglobulins present. Levels of IgG, IgA and IgM all decrease as
colostrum transitions into milk and IgA becomes the predominant Ig present (Norcross,
1982). Average immunoglobulin concentrations in mare milk are shown in Table 2-1.
Peak colostrum IgG content is observed at foaling and rapidly declines during the
first 24 hours after foaling (Lavoie et al., 1989). In colostrum sampled within 2 hours
after foaling, mean IgG values were shown to be 16,583 mg/dL (Lavoie et al., 1989). At
4 hours post foaling, another study showed mean IgG values that were at lower levels of
5,450 mg/mL, and these values fell even further to 1,010 mg/dL by 9-12 hours after
foaling (Erhard et al., 2001). Colostrum IgG fell below 1,000 mg/dL by 13-16 hours post
foaling and continued to decrease until day 14 (Erhard et al., 2001). Duvaux-Ponter et al.
(2004) showed that these low milk IgG levels did not show any changes by 21 days after
foaling, suggesting that mare milk IgG levels stay at this low level for the duration of
Effect of Diet on Fat and Fatty Acid Composition of Milk
Milk fatty acids are either synthesized de novo by acetyl-CoA carboxylase and fatty
acid synthase or are supplied exogenously. The mammary epithelial cells of lactating
animals are highly active in triglyceride biosynthesis (Clegg et al., 2001). If the FA are
not synthesized in the mammary epithelial cells, they can enter the cells either from
albumin in the plasma or from hydrolysis of chylomicron triglycerides by lipoprotein
lipase. Once inside the cell, FA are bound to fatty acid binding protein in the cytoplasm
or activated with acetyl-coenzyme A (CoA) and used for triglyceride synthesis. The
endoplasmic reticulum synthesizes microlipid droplets that fuse to form cytoplasmic
droplets which move to the apical membrane where they are enveloped to form the milk
fat globule. This globule is then secreted in a membrane-bound form into the milk
(Neville and Picciano, 1997).
Mare milk contains relatively little fat, with triglycerides as the predominate lipid
class (Dils, 1986). Mare milk naturally contains very small quantities of stearic (C18:0)
and palmitoleic (C16:0) acids and high quantities of linolenic (C18:3n-3) and linoleic
(C18:2n-6) acids (Csap6 et al., 1995). The higher amounts of unsaturated FA are
explained by the fact that horses consume large amounts of forages rich in unsaturated
FA (Csap6 et al., 1995). Milk composition, however, may be changed by manipulating
the diet, with the largest effects seen in the fat content (Sutton and Morant, 1989). Mare
milk long-chain FA composition is strongly related to the FA composition of the diet, as
no microbial FA hydrogenation occurs before intestinal absorption in horses (Doreau et
al., 1992; Hoffman et al., 1998).
The ratio of forage to grain in the mare' s diet can effect her milk composition.
Generally, fat content decreases as the percentage of grain increases (Doureau and
Boulot, 1989). Doreau et al. (1992) fed nursing mares diets containing either 95% hay
and 5% grain or 50% hay and 50% concentrate. Milk fat concentrations were higher for
the mares fed the 95:5 forage:grain diet compared to the 50:50 forage:grain diet. The
mares eating mostly forage also had higher linolenic and lower linoleic acid milk
contents than those eating mostly grain (Doreau et al., 1992). This effect is
understandable considering the fact that forage is high in linolenic acid. However,
because exact amounts of hay and grain fed and the fat composition of the diet
ingredients was not given, it is difficult to determine accurate values for percent fat of
Studies in humans have also examined the effect of dietary fat on milk fat
composition. Henderson et al. (1992) found that supplementing pregnant women with 6
g of an EPA and DHA supplement for 21 days significantly increased EPA, DPA and
DHA and decreased total n-6 PUFA levels in breast milk when compared to pre-
supplementation levels. Helland et al. (1998) observed an increase in EPA and DHA in
breast milk when women were supplemented with 5 and 10 mL cod liver oil daily for 14
days compared to women receiving 5 mL of cod liver oil/day and those receiving no
supplementation. The changes in breast milk FA composition reported by Henderson et
al. (1992) and Helland et al. (1998) were noted as early as day two of supplementation.
Interestingly, daily supplementation of women with 20 g of flaxseed oil (approximately
10.7 g ALA/d) for 4 weeks increased the EPA and DPA breast milk content but failed to
produce an increase in DHA (Francois et al., 2003). The authors speculated that the
excess ALA supplied from flax oil may have competitively inhibited A6-desaturase from
converting DPA to DHA (Francois et al., 2003).
In dogs, feeding fat supplements with varying ratios between ALA and the sum of
EPA and DHA produced milk fat compositions highly correlated to the diet fed (Bauer et
al., 2004). Dogs were fed one of four diets containing 15% total fat as beef tallow and
varying amounts of linseed and menhaden fish oil to provide specific levels of ALA,
EPA and DHA. The diets were formulated as follows: the Lo/Lo diet contained 0.14%
ALA and 0.04% EPA and DHA, the Lo/Mod diet contained 0.29% ALA and 0.24% EPA
and DHA, the Lo/Hi diet contained 0.20% ALA and 0.66% EPA and DHA and the Hi/Lo
diet contained 6.82% ALA and 0.04% EPA and DHA (fatty acids are expressed as a
percentage of dry matter). Bitches fed the Hi/Lo diet had the highest milk ALA content,
while bitches fed the Lo/Hi diet had the highest EPA and DHA milk content. Milk
responses of EPA, DPA and DHA content were seen as a function of increasing dietary
n-3 PUFA content. There was no enrichment of DHA when the Hi/Lo diet was fed,
showing that ALA is inefficiently converted to DHA in the dog (Bauer et al., 2004).
Davidson et al. (1991) showed that mares fed a diet with 5% added fat produced
milk with a higher fat content than mares who were not supplemented with fat (2-3%
dietary fat). However, no differences in milk fat production were noted when mares were
fed a sugar and starch diet with 2.4% fat compared to a fat (corn oil) and fiber diet with
10.4% fat (Hoffman et al., 1998). Nonetheless, the FA composition of the milk mirrored
the FA supplied by the diet. The mares eating the high fat diet showed higher milk
concentrations of LA and lower concentrations of ALA, which can be explained, in part,
by the n-6 PUFA content of the corn oil (Hoffman et al., 1998). Spearman et al. (2005)
found that feeding gestating mares a mix of corn oil and linseed oil increased milk ALA
content when compared to mares fed corn oil. Duvaux-Ponter et al. (2004) observed
higher levels of ALA in mare milk when mares were supplemented with linseed oil.
Feeding mares 454 g of encapsulated fish oil per day increased EPA and DHA in the milk
but did not affect the ALA content when compared to mares fed corn oil (Kruglik et al.,
2005). Together, these studies show that the fat content and FA composition of the
mare's diet can influence milk composition.
Fatty Acid Transfer across the Placenta
The placenta is a pivotal organ in providing the developing fetus with essential
fatty acids. During the last trimester of pregnancy, fetal requirements for AA and DHA
are especially high due to rapid synthesis of brain tissue. To obtain these FA, the fetus
depends upon placental transfer, and thus on the FA status of the mother (Al et al., 2000).
Much of the research of placental FA transfer has been performed in humans, who
possess a discoid hemochorial placenta. In these studies, there has been considerable
evidence of transfer of ALA, EPA and DHA across the placenta (Innis, 2005). This
transfer is a multi-step process of FA uptake by fatty acid binding proteins and
intracellular translocation of the FA from the maternal to fetal environment. The fatty
acid binding proteins that facilitate this process favor the uptake of n-6 and n-3 PUFA
over non-essential FA (Innis, 2005).
Human placental preference for transfer of FA has been reported by one author to
be DHA>ALA>LA>AA (Haggarty et al., 1997), while others have speculated that DHA
and AA are preferred over all other FA (Campbell, 1996; Crawford, 2000).
Fetal plasma concentrations of AA and DHA are reported to be 300- to 400-fold higher
than maternal plasma levels while their LA and ALA levels are lower (Elias and Innis,
2001). However, human placenta does contain A6- and A'-desaturases (Innis, 2005), so
the higher concentration of AA and DHA in fetal circulation may be partially produced
by placental conversion of these FA from their 18 carbon precursors.
Human studies have shown that the maternal dietary intake of n-6 and n-3 PUFA
influences placental transfer of AA and DHA. Connor et al. (1996) supplemented
pregnant women with sardines and fish oil from the 26th to the 3 5th week of pregnancy in
amounts to provide 2.6 g of n-3 FA per day. When DHA blood levels of newborn infants
born to supplemented women were compared to those of newborn infants born to
unsupplemented women, newborn babies born to supplemented mothers had 35.2% more
DHA in red blood cells. Infants from supplemented women also showed a plasma DHA
content 45.5% higher than infants from unsupplemented mothers, concluding that
placental transfer of DHA in women is increased by maternal supplementation with DHA
(Connor et al., 1996). However, de Groot et al. (2004) reported that supplementing
pregnant women with ALA did not increase umbilical cord blood DHA, suggesting that
the placenta could not efficiently convert ALA to DHA. In this study, pregnant women
were supplemented daily with either 9.02 g LA and 2.82 g ALA (experimental group) or
10.94 g LA and 0.03 g ALA (control group) in the form of margarine. Supplementation
was provided from week 14 of pregnancy until delivery (de Groot et al., 2004). While
the umbilical venous plasma obtained from the subj ects at delivery showed no differences
in DHA content between groups, the experimental group did show an EPA concentration
twice that of the control group, suggesting that conversion of ALA to EPA in the human
placenta may be possible (de Groot et al., 2004).
In spite of its complex six-layered placenta, the transfer fatty acids from mare to
fetus is possible. Equine studies, while few in number, have shown a positive correlation
between maternal and umbilical vein plasma free FA levels (Stammers et al., 1991).
However, the same study also showed a difference in FA composition between maternal
and umbilical vein plasma. The phospholipids portion of the umbilical venous plasma
contained more longer chain derivatives of LA and ALA than was found in maternal
plasma, suggesting that these longer chain FA were of placenta origin, because maternal
plasma phospholipids in the horse contain very little longer chain PUFA (Stammers et al.,
The presence of A6 Of A5-desaturase, to the author' s knowledge, has not been
established in the equine placenta. However, many studies have produced results that
would imply these enzymes are present. In natural situations, long-chain PUFA
(particularly DHA) are virtually absent from maternal circulation and in very low
concentrations in other maternal lipid compartments. In spite of this occurrence, foal
plasma phospholipids are rich in long-chain PUFA which must therefore be provided to
the foal by placental formation and transfer (Stammers et al., 1987). Stammers et al.
(1988) showed that foals had higher plasma concentrations of AA, EPA and DHA than
did their dams. A 30 hour fast of the pregnant mares resulted in an even greater fetal
concentration of these fatty acids, resulting from the increased lipid mobilization in the
mares (Stammers et al., 1988). When Stammers et al. (1994) incubated equine placenta
in media enriched with LA, the lipid fractions released from the placenta consisted of
long-chain PUFA derivatives of LA such as C20:3n-6, C20:4n-6 and C22:6n-6. This
finding suggests that these PUFA would be seen in the umbilical plasma lipids rather than
the maternal plasma lipids. No studies exist in mares to test the ability of manipulation of
dietary fat to influence placental FA transfer, so much research needs to be done in this
Because many of the studies investigating the effects of feeding n-3 FA to the horse
have not utilized a true control group that received no fat supplementation, research is
needed to compare the effects of n-3 supplementation with no n-3 supplementation (i.e.,
unaltered diet). This is especially important considering the fact that high forage diets
contain significant quantities of n-3 FA, but the addition of grain to the diet shift the
proportion of FA in favor of n-6. Studies comparing n-3 FA supplementation to baseline
diets are needed to validate that the biological effects observed when feeding n-3 FA are
truly due to the increase in these FA, and not to a decrease in n-6 FA.
In addition, little research has addressed responses yielded by different n-3 FA
(e.g., ALA, EPA, DHA) to determine if differences in dietary FA source can influence
biological responses in the horse. In particular, little data exists that compares the effects
of different n-3 FA sources fed to the mare and the subsequent response of her nursing
foal. It is unclear if supplementing the mare during gestation with n-3 FA can affect the
IgG composition of her colostrum and milk and subsequently the IgG concentration in
her foal. Furthermore, it is unknown if increasing the gestating mare' s n-3 FA intake can
result in greater placental transfer of n-3 FA, therefore allowing the foal to be born with
an already elevated level of these FA.
Lastly, clear effects of supplementation with ALA or an EPA/DHA combination on
the inflammatory response in horses have yet to be elucidated. Therefore, in an attempt
to answer some of these questions, the obj ectives of this study were:
1. Examine the effect of dietary n-3 supplementation of mares on the FA composition
of mare milk and mare and foal plasma and red blood cells;
2. Examine the difference of efficiencies of ground flaxseed (ALA) and encapsulated
fish oil (EPA and DHA) in augmenting EPA and DHA in the mare and foal;
3. Determine if n-3 FA supplementation of the mare can increase the IgG content of
colostrum, milk and foal plasma.
4. Determine in supplementation with flaxseed or fish oil can alter the inflammatory
response in mares and foals.
MATERIALS AND METHODS
This trial used 36 pregnant Thoroughbred (n=24) and Quarter Horse (n=8) mares
and their subsequent foals. Mare age ranged from 4 to 20 years with a mean of 10.5 & 4.1
years (mean & SE). Mares were paired according to breed and stratified according to
expected foaling date before being assigned to three treatment groups. The order of
treatment assignment was determined by numbering three pennies, each penny
corresponding to a separate treatment, and placing them into a hat. Pennies were then
drawn at random to determine the order of treatment assignment. Treatment groups were
then balanced for mare age and parity.
For the duration of this trial, mares and foals were housed at the University of
Florida's Horse Research Center in Ocala, Florida. Pregnant mares were housed on
pasture until signs of foaling were evident. At this time, mares were moved into small
paddocks until foaling. All mares, with the exception of one, foaled outside. After
foaling, mares and foals were kept in a box stall for 24 hours and then turned out in a
small paddock for one week before being returned to pasture. A routine vaccination and
anthelmintic schedule was followed for all animals. This experiment was performed in
accordance with the regulations and approval of the Institutional Animal Care and Use
Committee of the University of Florida.
Diets and Treatments
The basal diet for all treatment groups consisted of a commercial grain-based
concentrate (Gest-O-Lac; Ocala Breeders Sales, Ocala, Florida) and pasture or hay. The
grain-based concentrate was offered at 1.0% BW in late gestation and 1.0-2.0% BW
during lactation in order to maintain bodyweight and a minimum body condition score of
5. The concentrate was formulated to meet or slightly exceed nutrient requirements for
late gestation and lactation based on NRC recommendations (NRC, 1989). From
December to March, mares were fed Coastal bermudagrass hay ad-libitum and had access
to dormant bahiagrass pasture. From April to June, mares only had access to bahiagrass
pasture. Trace mineralized salt blocks were available at all times. Foals were provided
with access to the same grain-based concentrate that was fed to mares via creep feeders
that were placed in the pasture.
Mares received one of three treatments: 1) basal diet with no supplementation
(CON, n = 12); 2) basal diet supplemented with milled flaxseed (Pizzey's Milling,
Manitoba, Canada; FLAX, n=12); or 3) basal diet supplemented with encapsulated fish
oil (United Feeds, Inc., Indiana; FISH, n = 12). Both FLAX and FISH were fed to mares
in amounts to provide 6 g total n-3 FA/100 kg BW per day. This level of
supplementation was chosen based on the studies of O'Connor et al. (2004) and Siciliano
et al. (2003) which demonstrated changes in plasma fatty acid composition when horses
were supplemented with similar levels of fish oil. Mares and foals were brought in from
pasture at 0700 and 1500 h each day, placed into box stalls and individually fed the grain
mix concentrate. Half of the daily allotment of flaxseed or fish oil supplement was hand
mixed into the grain provided in the morning feeding and the remaining half of the
supplements were mixed into the grain provided in the afternoon feeding. Foals had the
opportunity to share the mares' feed, but this depended upon the individual temperament
of each mare. Supplementation began 28 days before the expected foaling date and
continued until 84 days post-partum.
The nutrient composition of the grain-based concentrate and the flaxseed and
encapsulated fish oil supplements is presented in Table 3-1. The nutrient content of the
Coastal bermudagrass hay and bahiagrass pasture is presented in Table 3-2.
Table 3-1. Nutrient composition of the grain mix concentrate and the milled flaxseed and
encapsulated fish oil supplements
Nutrients Concentrate Flaxseed Fish Oil
DM, %2 92.7 91.5 91.2
DE, Mcal/kg3 3.41 3.0 3.7
CP, % 15.5 22.9 11.8
ADF, % 11.9 19.0 5.9
NDF, % 26.3 40.0 9.9
Fat, % 4.2 37.7 21.5
Ca, % 1.06 0.24 0.33
P, % 0.70 0.74 0.15
Zn, mg/kg 248 41 33
Cu, mg/kg 64 11 4
SValues are presented on a 100% DM basis (except DM).
2 DM, dry matter; DE, digestible energy; CP, crude protein; ADF, acid detergent fiber;
NDF, neutral detergent fiber.
3 Calculated using the equation: DE (Mcal/kg) = 4.07 0.055(%ADF) (NRC, 1989).
Table 3 -2. Nutrient composition of the bahiagrass pasture (by month) and Coastal
Jan. Feb. March April
64.1 65.0 44.1 30.5
1.9 2.0 1.9 2.2
10.7 11.7 11.8 16.0
41.4 40.2 43.5 35.4
68.5 64.5 67.7 54.7
2.1 2.5 2.4 3.1
0.61 0.68 0.64 0.72
0.24 0.25 0.25 0.36
38 36 40 32
6 6 8 8
Presented on a 100% DM basis (except DM).
2 DM, dry matter; DE, digestible energy; CP, crude protein; ADF, acid detergent fiber;
NDF, neutral detergent fiber.
3 Calculated using the equation: DE (Mcal/kg) = 4.22 0. 11(%ADF) + 0.0332(%CP) +
0.00112(%ADF)2 (NRC, 1989).
Mares were weighed at 28 and 14 d prior to expected foaling date (d-28, d-14), at
foaling (dO) and every 14 days thereafter. Foals were weighed at birth (dO) and every 14
days thereafter. A digital livestock scale with an accuracy of a 0.5 kg was used to obtain
Blood Sample Collection and Processing
Blood samples were collected from mares by jugular venipuncture at 28 and 14 d
prior to expected foaling, at foaling (dO), and at 28, 56 and 84 d after foaling for
acquisition of plasma, serum and/or red blood cells. Blood samples were collected from
foals via jugular venipuncture at birth before the foal was allowed to nurse (dO), 36 h
post-parturition, and 7, 28, 56 and 84 d post-foaling for acquisition of plasma, serum
and/or red blood cells. A square patch of hair was shaved over the foal's jugular vein to
allow for easier blood sampling. Precision Glide Vacutainer brand blood collection
needles (20G, 1 V/2 in. for mares; 20G, 1 in. for foals) were used to collect blood into
Beckton Dickinson Vacutainers containing sodium heparin, to facilitate harvesting of
plasma and red blood cells, or tubes containing no anticoagulant for harvesting of serum.
With the exception of samples obtained at birth or 36 h post-parturition, all blood
samples were collected between 0700 and 0900 h and prior to the mare's morning grain
feeding. After collection, blood samples were immediately placed on ice and transported
to the Animal Nutrition Laboratory for further processing.
In the laboratory, blood samples for obtaining serum were allowed to clot for 30
min to 1 h and then centrifuged at 5590 x g for 7 min to allow for separation of serum.
Serum was collected with plastic disposable pipets and aliquoted into polypropylene
cryogenic vials (2-3 vials, 0.5-1.0 mL each). Samples were frozen at -800C until further
analysis for IgG using a commercially available single radial immunodiffusion kit (SRID
Kit, VMRD, Inc., Pullman, WA). See Appendix B for a description of the IgG analysis.
Blood samples for obtaining plasma and red blood cells were first used to
determine the hematocrit (packed cell volume). Hematocrit values were determined in
duplicate using whole blood drawn into a microcapillary tube, centrifuged and read on a
microcapillary reader. After determination of hematocrit, each vacutainer was gently
rotated and 5.0 mL of whole blood was pipetted into a separate glass tube, labeled, and
centrifuged at 5590 x g for 15 minutes to separate the plasma and red blood cells. A
pipet was used to transfer 1.0 mL of plasma to each of four polypropylene cryogenic
vials. Samples were frozen on a slant at -200C to increase the surface area and ensure
more efficient freeze drying before being stored at -800C until further analysis of fatty
Once plasma had been removed, an aspirator was used to remove any additional
plasma and the thin layer of while blood cells lining the top of the red blood cells in each
tube. Two mL of cold saline was then added to each tube and the tubes were gently
mixed and centrifuged at 5590 x g for 7 min. After centrifuging, the supernatant was
aspirated off and an additional 2.0 mL of cold saline was added. The tubes were again
mixed and centrifuged at 5590 x g for 7 min. This procedure was repeated once more for
a total of three saline washes. After the supernatant of the final wash had been aspirated
off, exactly 2.0 mL of cold saline was added to the remaining red blood cells in each
tube. The tubes were mixed well before 2.0 mL of the red blood cell suspension was
transferred into labeled polypropylene cryogenic vials. These tubes were frozen at an
angle at -200C before being placed into storage at -800C until analyzed for fatty acid
Colostrum and Milk Collection and Processing
Colostrum was obtained from the mare within 1 h of birth and before the foal had
suckled (dO). Approximately 120 mL of colostrum was recovered into a pre-labeled, pre-
weighed plastic cup. The cup was covered with a lid and stored at 40C until transfer to
the Animal Nutrition Laboratory for processing.
Milk samples were obtained 36 h post-partum and between 0700 and 0900 h on 7,
14, 28, 56 and 84 d post-foaling for determination of fatty acid and IgG content. To
facilitate milk collection, foals were muzzled for approximately 30 min to allow the
mare's udder to fill. The entire udder was then milked out into a pre-labeled, pre-
weighed plastic cup. If the udder contained more milk than one cup could hold, the udder
was milked out into multiple cups whose content was then mixed in a larger container
and approximately 120 mL was transferred to the pre-labeled, pre-weighed sample cup.
The excess milk was discarded. After collection, milk samples were immediately placed
on ice and transported to the Animal Nutrition Laboratory for further processing.
In the laboratory, colostrum and milk samples were gently swirled to mix and
strained through four layers of cheesecloth to remove any dirt and debris in the sample.
The samples were then returned back to the original pre-weighed sample cups. After
straining, the sample was mixed again and approximately 1.0 mL was aliquoted into each
of three pre-labeled polypropylene cryogenic vials. These vials were then stored at -800C
until further analysis for IgG content. The remaining colostrum or milk sample was
weighed to determine a wet sample weight and then freeze dried. Freeze dried milk
samples were stored at -200C until used for the determination of fatty acid composition.
Fatty Acid Analysis
Fatty acids in plasma and red blood cells were extracted and methylated using the
procedure of Folch et al. (1957). Fatty acids were analyzed by gas chromatography (CP-
3800 Gas Chromatograph, Varian, Inc., Palo Alto, CA) using a WCOT fused silica
column (CP-SEL 88, lengthl00 m, internal diameter 0.25 mm, flow rate 5.0 mL/min,
Varian, Inc., Palo Alto, CA). The carrier gas was helium with a pressure of 29.5 psi (1
min), 35.4 psi (0.42 psi/min, total of 45 min) and 37.9 psi (0.17 psi/min, held for 50 min,
total of 110 min). The temperature program was 1200C for 1 min, increased to 1900C at
50C/min and held at 1900C for 30 min (total of 45 min), increased to 2200C at 20C/min
and held at 2200C for 50 min, giving a total run time of 110 min. Fatty acids were
identified by comparison of peak retention times for samples and reference standards
(Nu-Chek Prep, Inc., Elysian, MN). The FA identified included C8:0, C10:0, C12:0,
C14:0, C14:1, C16:0, C16:1, C17:0, C17:1, C18:0, C18:1n-9, C18:2n-6 (LA), C18:3n-3
(ALA), C20:0, C20:1, C20:2, C20:3, C20:4n-6 (AA), C20:5n-3 (EPA), C22:0, C22:5n-3,
C22:6n-3 (DHA) and C24:1. Nonadecanoic acid (C19:0) was added to the samples and
used as an internal standard to assess FA recovery. Total n-6 FA were defined as the sum
of C18:2 n-6 and C20:4n-6 while total n-3 FA were defined as the sum of C18:3n-3,
C20:5n-3, C22:5n-3 and C22:6n-3.
Intradermal Skin Test
To examine the effect of n-3 FA supplementation on the inflammatory response,
mares and foals were sensitized with phytohemagglutinin (PHA; Lectin from Pha~seobts
vulgaris, Sigma-Aldrich, Inc., St. Louis, Missouri) at 84 d post-partum. Twenty-Hyve
milligrams of PHA was reconstituted in 16.7 mL of phosphate buffered saline (PB S) to
give a Einal concentration of 150 Cpg/100 CLL. A 4 x 4 cm patch of hair was surgically
clipped on the midsection of both sides of the neck on mares and foals and inj ected
intradermally with 100 pIL of the PHA suspension. Precision Glide brand intradermal
inj section needles (26 G, 3/8 in.) were used to deliver the PHA. Needles were changed
between each inj section site on the right and left side of the neck. Skin thickness
measurements were obtained by pinching the skin between the thumb and forefinger and
measuring the skin fold thickness in mm with an electronic digital micrometer (Marathon
Watch Company, Ltd., Ontario, Canada). Measurements of each injection site were
obtained after clipping but before inj ecting (h 0) and at 2, 4, 6, 8, 12, 24 and 48 h after
inj section. Skin thickness measurements from the right and left sides of the neck were
averaged to give a single thickness measurement for each time point.
Supplement and Feed Sample Analysis
The same batch of milled flaxseed, encapsulated fish oil and Coastal bermudagrass
hay were available for the duration of the trial. However, the source of commercial grain
mix was replenished approximately every 2 wk due to storage limitations and the volume
of feed needed. Samples of the flaxseed, encapsulated fish oil and grain mix were
obtained at 4 wk intervals. These samples were then dried at 600C and stored at 200C for
later analysis. Samples of bahiagrass pasture were obtained at 4 wk intervals from four,
16 ha pastures. Pasture grass clippings were only obtained from areas where grazing was
evident. At each 4-wk collection, clippings from the four pastures were composite,
dried at 600C and stored at 200C for later analysis. Throughout the trial, each round bale
of Coastal bermudagrass hay that was offered to the mares was core sampled (5 cores per
bale), dried at 600C and stored at 200C. After the completion of the trial, all samples of
Coastal bermudagrass hay were composite into one sample for analysis. All feeds and
supplements were analyzed for fatty acid content using the method described above. In
addition, feeds were analyzed for DM, DE, CP, NDF, ADF, total fat, Ca, P, Mg, Zn and
Cu by wet chemistry (Dairy One Forage Analysis Lab, Ithaca, NY).
Four mares either delivered dead foals or their foals died shortly after birth. Only
pre-foaling data was used from these mares. One mare experienced a red bag during
foaling and her foal was subsequently given plasma, so IgG data from this mare and foal
were not included in the statistical analysis. One foal was euthanized at 30 d of age, so
only data taken up to that time point were used. The final number of mare and foal pairs
successfully completing this study was 11 CON, 11 FLAX and 9 FISH.
The MIXED procedure of SAS (V. 9.1, SAS Inst., Inc., Cary, NC) was used to
analyze fatty acid composition of colostrum, milk, plasma and all feeds, IgG content of
colostrum, milk, mare serum and foal serum, and mare and foal bodyweights. The
sources of variation included treatment, time and treatment x time interaction. Breed
effects were also tested for mare and foal bodyweights, mare serum IgG, mare colostrmm
and milk IgG and foal serum IgG. Sex effects were tested for foal serum IgG. In
addition, principle forage source (hay or pasture) was examined as a main effect for mare
plasma and red blood cell fatty acids. Fatty acids analyzed included linoleic (LA), co-
linolenic (ALA), arachidonic (AA), eicosapentaenoic (EPA) and docosahexaenoic (DHA)
acids as well as total n-3 and total n-6 fatty acids and the ratio of n-6:n-3 fatty acids. For
IgG and fatty acid analysis, horse nested within treatment was considered as a random
variable and used as an error term to test the effects of all sources of variation. Dunnett' s
test was used to separate means. The homogeneity of regression for skin thickness values
was evaluated using the GLM and MIXED procedures in SAS. Horse nested within
treatment was used as an error term.
Due to missing values and unbalanced treatment groups after foaling, all data are
expressed as least square means + SE unless otherwise stated. Values were considered
significant at p < 0.05 and trends were considered at p < 0.10.
Feed and Supplement Analysis
The fatty acid composition of the grain mix concentrate and the flaxseed and
encapsulated fish oil supplements is presented in Table 4-1. The fatty acid composition
of pasture forage was similar from December through March (Appendix A). Therefore,
the fatty acid composition of pasture samples collected in December, January, February
and March was averaged and presented as "winter" pasture (Table 4-2). Similarly, the
fatty acid composition of pasture forage was not different between the months of April,
May and June (Appendix A). Thus, the fatty acid composition of pasture samples from
these months was also averaged and presented as "spring" pasture (Table 4-2).
Spring pasture contained lower quantities of C18:0 (P = 0.01) and C18:1 (P =
0.004) and higher quantities of ALA (P = 0.03) and total n-3 FA (P = 0.03) than winter
pasture (Table 4-2). Hay contained higher concentrations of C16:0 (P = 0.0001), C18:0
(P = 0.003) and C18:1 (P = 0.0001) compared to winter and spring pasture forage (Table
4-2). In addition, hay contained lower concentrations of ALA (P = 0.0001) resulting in a
lower total n-3 FA content (P = 0.0001) and a higher n-6:n-3 FA ratio (P = 0.0001) in hay
compared to winter and spring pasture (Table 4-2). No differences were observed in LA,
AA, EPA, DHA or total n-6 FA content between hay and pasture forage. When principle
forage source (hay or pasture) was included in the model as a main effect, forage source
did not influence FA levels in any of the milk or blood samples examined in this study (P
= 0. 11).
Mare Fatty Acid Intake
Horses were housed on bahiagrass pastures throughout the trial. Because of winter
dormancy and reduced quantity of pasture forage, mares were offered unlimited access to
Coastal bermudagrass hay during the months of December, January, February and March.
During this four month period, hay was assumed to be the primary forage source.
Limited evidence of grazing in pastures and reasonable consumption of hay (based on the
number of round bales fed and expected DM intake) during this period support this
assumption. All hay was removed from pastures in the first week of April. Therefore,
pasture served as the primary forage source from April until the conclusion of the trial in
Average daily intake of long chain FA by mares on each treatment from December
to March (when Coastal bermudagrass hay was the primary forage source) and from
April to June (when bahiagrass pasture was the primary forage source) is shown in Tables
4-3 and Table 4-4, respectively.
From December to March (when hay was the primary forage source), FLAX mares
consumed 255% more n-3 FA and FISH mares consumed 257% more n-3 FA than CON
mares. From April to June (when pasture was the primary forage source), FLAX and
FISH mares consumed 138% and 137% more n-3 FA than CON mares, respectively.
This change in percentages reflects the higher n-3 FA content of spring bahiagrass
pasture compared to Coastal bermudagrass hay. From December to March, the total
diet provided a total n-3 FA intake of 4.3 g n-3 FA/100 kg BW in CON mares, 11.3 g n-3
FA/100 kg BW in FLAX mares and 11.3 g n-3 FA/100 kg BW in FISH mares. From
April to June, the total diet provided a total n-3 FA intake of 17.4 g n-3 FA/100 kg BW in
CON mares, 24.8 g n-3 FA/100 kg BW in FLAX mares and 24.3 g n-3 FA/100 kg BW in
FISH mares. Within the supplemented groups, the milled flaxseed and encapsulated fish
oil supplied 6.5 to 7.0 g total n-3 FA/100 kg BW, or approximately 65% of the total n-3
FA intake in the winter and 30% of the total n-3 FA intake in the spring.
For the duration of the trial, FLAX mares consumed higher ALA (P = 0.0001) than
FISH and CON mares while FISH mares consumed higher EPA (P = 0.0001) and DHA
(P = 0.0001) than FLAX and CON mares. There were no differences between the
consumption of total n-3 FA between FISH and FLAX mares (P = 0.94), but both
treatments consumed more total n-3 FA than CON mares (P = 0.0001). No treatment
effect was observed for total n-6 FA intake (P = 0.12).
Mare and Foal Bodyweight
Treatment had no effect on mare BW at any time during the trial (Table 4-5). CON
mares foaled 3 colts and 9 fillies, FLAX mares foaled 7 colts and 5 fillies and FISH
mares foaled 4 colts and 8 fillies. There were no differences in foal BW due to sex (P =
0.58), breed (P = 0.62) or treatment (P = 0.75). At birth, CON foals weighed 51.0 & 4.1
kg and gained 107.3 & 4.1 kg over the trial period. FLAX foals weighed 54.4 & 3.5 kg at
birth and gained 103.9 & 3.5 kg, and FISH foals weighed 56.3 & 3.8 kg at birth and
gained 107.6 & 3.9 kg over the trial period (Table 4-6). There was a significant effect of
time (P = 0.0001) on foal BW, which reflected an increase in BW as foals grew from
birth to 84 d of age.
Mare Plasma Fatty Acid Composition
Omega-6 Fatty Acids
The FA found in the highest concentration in mare plasma was LA, which made up
almost half of the total FA found in plasma (Table 4-7). An overall treatment effect was
noted for mare plasma LA, AA and total n-6 FA (Table 4-7). Before supplementation
began (d-28), plasma n-6 FA concentrations were similar between treatments (Table 4-8).
In response to supplementation, mares fed FISH had lower plasma LA (P = 0.05), greater
plasma AA (P = 0.03) and tended to have lower plasma total n-6 FA (P = 0. 10) than
FLAX or CON mares (Tables 4-7 and 4-8).
An overall effect of time was detected for plasma LA (P = 0.07), AA (P = 0.01) and
total n-6 FA (P = 0.08) and may have reflected the effects of both parturition and dietary
treatment (Table 4-8). Plasma LA declined from baseline (d-28) to foaling (dO) in CON
(P = 0.05) and FISH mares (P = 0.02). After foaling, plasma LA returned to pre-
treatment levels in CON mares but remained lower in FISH mares. Plasma AA increased
(P = 0.01) from baseline to foaling in FISH mares but did not change in CON mares.
Total plasma n-6 FA decreased from d-28 to dO in CON (P = 0.05) and FISH mares (P =
0.04, Figure 4-1). In contrast to the responses seen in FISH and CON mares, plasma LA,
AA and total n-6 FA did not change over the course of the trial in FLAX mares.
Omega-3 Fatty Acids
The n-3 FA found in the highest concentration in mare plasma was ALA, with
concentrations ranging from 2.99 to 3.65 g ALA/100 g fat (Table 4-7). Overall treatment
effects were detected for plasma ALA, EPA, DHA and total n-3 FA (Table 4-9). Before
supplementation, no differences in plasma n-3 concentrations were observed between
treatment (Table 4-9). In response to dietary treatment, FISH mares had higher plasma
EPA (P = 0.0001), higher plasma DHA (P = 0.0001) and higher plasma total n-3 FA (P =
0.01) than CON mares (Table 4-7). When mares were fed FLAX, plasma ALA tended to
be higher (P = 0.09) than that observed in the plasma of CON or FISH and total n-3 FA
plasma content was similar to both FISH and CON mares (Table 4-7).
Overall effects of time were detected in plasma ALA (P = 0.02), EPA (P = 0.0001),
DHA (P = 0.001) and total n-3 FA (P = 0.0005; Table 4-9). From d-28 to dO, plasma
ALA increased in FLAX mares (P = 0.05) but decreased in FISH mares (P = 0.02). After
foaling, the plasma ALA of FISH mares returned to baseline levels while the plasma
ALA of FLAX mares remained at an elevated level. Plasma ALA did not change over
time in CON mares. Plasma EPA increased (P = 0.002) through d+28 in mares fed FISH,
after which this FA stabilized at levels above CON and FLAX mares. Plasma DHA and
total n-3 increased (P = 0.001) in response to FISH, but remained unchanged in the
plasma of CON or FLAX mares for the duration of the trial (Table 4-9; Figure 4-2). In
contrast to the effects observed in FISH mares, plasma EPA, DHA and total n-3 FA did
not change in response to dietary treatment in FLAX or CON mares (Table 4-9; Figure 4-
Treatment did not affect the ratio of n-6:n-3 FA in mare plasma (P = 0.24; Table 4-
10). However, an overall effect of time was detected, as the ratio of plasma n-6:n-3 FA
decreased over the course of the trial in all treatments (P = 0.02; Table 4-10).
Mare Colostrum and Milk Fatty Acid Composition
Treatment had no effect on the total fat content of mare colostrum (P = 0.95) or
milk (P = 0.12, Table 4-11). As colostrum transitioned into milk, the total fat content
increased (P = 0.0003) for all treatments (Table 4-11). The FA found in the highest
concentrations in mare colostrum and milk were C16:0, C18:1 and LA (Table 4-12).
An overall effect of time was detected for all FA examined in mare milk (Tables 4-
13 and 4-14). All treatments experienced a decrease in total milk n-6 FA (P = 0.0001)
and an increase in total milk n-3 FA (P = 0.0001) as lactation progressed (Tables 4-13
and 4-14). As a result, the ratio of n-6:n-3 FA decreased in mare milk from foaling
through 84 d post-foaling (P = 0.0001; Table 4-10). From 36h to through 14 d post-
foaling, milk LA and total n-6 FA decreased in FLAX (P = 0.0005) and FISH mares (P =
0.0003), and then remained constant for the duration of the trial (Table 4-13). Milk from
mares fed FISH showed an increase in EPA (P = 0.0001) and DHA (P = 0.0001) content
from 36h to through 14 d post-foaling, and these levels remained steady until 84 d post-
foaling (Table 4-14).
Overall effects of treatment were not noted for any of the n-6 FA examined in
mare milk (Table 4-12, Figure 4-3). However, overall effects of treatment were observed
for milk ALA (P = 0.0001), EPA (P = 0.0001), DHA (P = 0.0001) and n-6:n-3 FA ratio
(P = 0.01; Tables 4-10 and 4-12). At foaling, the colostrum of FLAX mares contained
higher levels of ALA (P = 0.05) than CON and FISH mares (Table 4-14, Figure 4-4). As
lactation progressed, FLAX mares continued to have a higher ALA content in their milk
compared to FISH or CON mares (P = 0.01). FISH mares had a higher colostrum DHA
content than CON and FLAX mares (P = 0.03) and had higher EPA (P = 0.0001) and
DHA (P = 0.0001) concentrations in milk than CON or FLAX mares as lactation
progressed (Table 4-14, Figure 4-4). The colostrum of FLAX mares contained greater
total n-3 FA than the colostrum of CON mares (P = 0.05), but total n-3 FA was not
different than FISH mares. Over the course of lactation, the n-6:n-3 FA ratio tended to be
lower in the milk of FLAX mares (P = 0.09) when compared to the milk of CON and
FISH mares (Table 4-10).
Foal Plasma Fatty Acid Composition
Omega-6 Fatty Acids
Similar to the mare, the FA found in the highest concentration in foal plasma was
LA, making up roughly one third of the total FA found in plasma (Table 4-15). An
overall treatment effect was not observed for any of the n-6 FA examined in foal plasma
(Table 4-16, Figure 4-5). However, at foaling (dO), plasma AA concentrations were
highest in foals born to FLAX mares (P = 0.04, Table 4-16). At 14 d post-foaling, foals
suckling FISH mares showed a higher plasma AA concentration than foals suckling CON
mares (P = 0.04), but were similar to foals suckling FLAX mares (P = 0.30). No other
effects of treatment on n-6 FA were detected at any time point over the course of the trial.
An overall effect of time was detected in plasma LA (P = 0.0001), AA (P = 0.0001)
and total n-6 FA (P = 0.0001; Table 4-16, Figure 4-4). Plasma LA and total n-6 FA
increased (P = 0.0001) and plasma AA decreased (P = 0.0001) from foaling to 14 d post-
foaling in all treatments (Table 4-16). Plasma LA and total n-6 FA increased from 14 to
84 d of age in foals suckling CON (P = 0.005) and FISH mares (P = 0.008), but remained
stable in foals nursing FLAX mares. Plasma AA increased from 14 to 84 d of age in
Omega-3 Fatty Acids
The n-3 FA found in the highest concentration in foal plasma was ALA, with levels
ranging from 2.48 to 3.33 g ALA/100 g fat (Table 4-15). An overall effect of treatment
was observed in foal plasma ALA, EPA, DHA and total n-3 FA (Table 4-15, Figure 4-6).
At foaling, foals born to FISH mares tended to have a higher total n-3 FA plasma content
than foals born to CON or FLAX mares (P = 0.09; Table 4-17, Figure 4-6). Foals
suckling FLAX mares had higher plasma ALA (P = 0.04) than foals suckling CON mares
and foals nursing FISH mares had higher plasma EPA (P = 0.0001), DHA (P = 0.0001)
and total n-3 FA (P = 0.002) than foals nursing both CON and FLAX mares (Table 4-15
An overall effect of time was detected in all n-3 FA in foal plasma (P = 0.0001;
Table 4-17, Figure 4-6). From foaling to 84 d of age, the plasma ALA and total n-3 FA
content increased in foals, regardless of mare treatment (P = 0.0001). Plasma EPA
increased in FLAX foals from foaling to 28 d of age (P = 0.0001), but returned to foaling
levels by 56 d of age (Table 4-17). Plasma EPA increased from birth to 14 d of age in
FISH foals (P = 0.0001), but decreased from 28 to 56 d of age. However, the plasma
EPA concentration in FISH foals was still higher at 84 d of age than the concentrations at
foaling (P = 0.0005). From birth to 14 d of age, plasma DHA decreased in CON (P =
0.0001) and FLAX (P = 0.0001) foals and then remained steady until the end of the trial.
However, the DHA concentration in the plasma of FISH foals remained elevated through
56 d of age, declining slightly by 84 d (Table 4-17).
Omega-6:Omega-3 Fatty Acid Ratios
An overall effect of treatment on the n-6:n-3 ratio was observed in foal plasma
(Table 4-10). At birth, no difference in n-6: n-3 FA ratio was detected between
treatments. After suckling treated mares, FISH foals had a lower n-6:n-3 FA ratio (P =
0.002) than FLAX or CON foals (Table 4-10). An overall effect of time on the n-6:n-3
FA ratio was also detected in foal plasma (P = 0.001, Table 4-10). From birth to 14 d of
age, the plasma n-6:n-3 ratio increased in CON (P = 0.001) and FLAX (P = 0.01) foals.
The plasma n-6:n-3 FA ratio returned to levels seen at foaling by 28 d of age in FLAX
foals, while the plasma n-6:n-3 FA ratio of CON foals did not return to baseline levels
until 56 d of age. The plasma n-6:n-3 ratio in FISH foals remained steady over the course
of the trial (Table 4-10).
Fatty Acid Correlations
Positive correlations (P = 0.0001) between mare plasma and milk FA were found
for ALA, EPA, DHA and total n-3 FA, while a negative, but weak correlation between
mare plasma and mare milk was found for total n-6 FA (P = 0.003; Table 4-18). No
correlation was found between mare plasma LA and mare milk LA, while mare plasma
AA and mare milk AA tended to be negatively correlated (P = 0.10).
Mare milk ALA, EPA, DHA and total n-3 FA were positively correlated (P =
0.0001) with foal plasma ALA, EPA, DHA and total n-3 FA (Table 4-18). Mare milk
AA and total n-6 FA were negatively correlated to foal plasma AA (P = 0.0001) and total
n-6 FA (P = 0.03). No correlation between milk LA and foals plasma LA was detected.
Fatty Acid Composition of Red Blood Cells
Mare Red Blood Cell Fatty Acids
Fatty acids with chain lengths longer than 18 carbons were not detected in mare red
blood cells, and LA was the only n-6 FA observed (Table 4-19). An overall effect of
treatment was noted for LA (Table 4-19, Figure 4-7). While no differences in LA
concentration were found in mare red blood cells before supplementation, there was a
tendency (P = 0.10) for FISH mares to have a higher red blood cell LA content than both
CON and FLAX mares (P = 0.10, Table 4-19). An overall effect of time was not detected
for mare red blood cell n-6 FA content (Table 4-20, Figure 4-7). However, the red blood
cell LA content of FISH mares increased from pre-supplementation to foaling (P = 0.02;
Table 4-20). The LA content of CON and FLAX red blood cells did not fluctuate during
the study (Table 4-20).
Foal Red Blood Cell Fatty Acids
Linoleic acid was the only n-6 FA and ALA was the only n-3 FA found in foal red
blood cells; fatty acids with chain lengths longer than 18 carbons were not detected
(Table 4-21). An overall effect of treatment was detected for red blood cell LA, but not
ALA (Table 4-21). At foaling, no differences were observed in red blood cell LA content
(Table 4-22, Figure 4-8). In response to suckling supplemented mares, FISH foals had a
higher red blood cell LA content than both CON and FLAX foals (P = 0.04, Table 4-21,
Figure 4-8). Foals belonging to FLAX mares had higher (P = 0.03) ALA in red blood
cells at birth than foals belonging to CON mares, but had similar red blood cell ALA as
foals born to FISH mares. Treatment of the mare did not affect ALA content of foal red
blood cells at any other time point during the study (Table 4-22, Figure 4-9).
An overall effect of time was found in foal red blood cell LA (P = 0.03) but not
ALA (Table 4-22). The LA content of red blood cells increased in FISH foals from
foaling to 14 d of age (P = 0.01) and stayed elevated for the duration of the study. In
contrast, the LA in red blood cells of CON and FLAX foals did not change during the
trial (Table 4-22).
Mare Serum, Colostrum and Milk IgG
An overall effect of treatment was not detected in mare serum or colostrum IgG
concentrations (Table 4-23). Mare breed (P = 0.78) or age (P = 0.56) did not affect
serum IgG content. Similarly, colostrum IgG was not affected by mare breed (P = 0.67)
or age (P = 0.58).
Milk IgG was not affected by treatment (P = 0.65) or breed (P = 0.67, Table 4-24).
However, an overall effect of time on milk IgG was detected (P = 0.0001, Table 4-24).
Milk from all mares showed a decline in IgG from 36 h through 84 d post-foaling (Table
4-24). The overall decline in milk IgG concentration from 36 h to 84 d post-partum was
139.4 & 630.7 mg/dL for CON mares, 157.0 & 677.5 mg/dL for FLAX mares and 132.5 &
659.2 mg/dL for FISH mares.
Mare serum IgG at foaling was not correlated with mare age (Table 4-25).
Similarly, colostrum IgG was not correlated with mare age, although FLAX mares tended
to show an inverse relationship (r = -0.58, P = 0.06) between mare age and colostrum IgG
(Table 4-25). Mare serum IgG at foaling was not correlated to colostrum IgG, and
colostrum IgG was not correlated to foal serum IgG at 36 h post-foaling. However, a
weak correlation between mare serum IgG at foaling and foal serum IgG at 36 h post-
foaling was detected across treatments (r = 0.42, P = 0.02; Figure 4-10). Within
treatments, dO serum IgG from CON and FLAX mares showed no correlation with foal
36h serum IgG content, but serum IgG from FISH mares at dO was correlated to FISH
foals serum IgG at 36 h post-foaling (r = 0.63, P = 0.05, Table 4-25).
Foal Serum IgG
All foals had serum IgG concentrations that were very low at birth and reflected the
pre-suckle status of the foal (Figure 4-1 1). The IgG content of foal serum increased to,
and peaked at, 36 h post-foaling, indicating that passive transfer of IgG had taken place.
An overall effect of time was noted for foal serum IgG (P = 0.0001), as the IgG of all
foals steadily declined from 36 h to 84 d post-foaling (Table 4-25, Figure 4-12).
No overall effect of treatment was observed for foal serum IgG (Table 4-25).
However, foals suckling FISH mares tended to have lower serum IgG than foals nursing
FLAX mares at 36h (P = 0.09) and 7 d post-foaling (P = 0.10). In addition, FISH foals
tended to have lower IgG than CON foals at 28 d post-foaling (P = 0.10).
Mare and Foal Responses to the Intradermal Skin Test
Mare Response to PHA
An overall effect of treatment was not detected in the skin thickness of mares in
response to a paired intradermal skin test using PHA as the stimulant (P = 0.89; Table 4-
26). However, an overall effect of time was observed (P = 0.0001; Table 4-26). Before
inj section, no differences were observed in mare skin thickness (P = 0.56; Figure 4-13).
All mares experienced a significant increase in skin thickness from 0 to 2 h (P = 0.0001)
and from 2 to 4 h post-inj section (P = 0.0001; Table 4-26, Figure 4-13). Skin thickness
was greatest between 4 and 8 h post-injection and then decreased. At 48 h, skin thickness
was still elevated above that measured before PHA inj section at 0 h (P = 0.0001).
Foal Response to PHA
An overall effect of time (P = 0.0001) on skin thickness in foals in response to PHA
injection was detected, reflecting an inflammatory response (Figure 4-14). Foal skin
thickness increased (P = 0.0001) from 0 to 4 h, remained elevated through 8 h post-
injection and then declined through 48 h post-injection (P = 0.0001, Figure 4-14). At 48
h post-inj section, skin thickness had not yet declined to baseline thickness measured
before PHA inj section (P = 0.0001).
An overall effect of treatment on foal skin thickness was not detected (P = 0.58;
Table 4-27). However, CON foals peaked at 4 h (P = 0.0001), whereas skin thickness
remained elevated in FLAX and FISH foals through 6 h (P = 0.0001; Figure 4-14, Table
4-27). At 6 h post-inj section, FLAX foals had greater skin thickness than CON foals (P =
0.02), while the skin thickness of FISH foals was intermediate between FLAX and CON
Comparing Mare and Foal Responses to PHA
Across treatments, skin thickness in response to PHA inj section was different
between mares and foals (P = 0.0001; Table 4-28, Figure 4-15). Although thickness was
not different before inj section of PHA (Oh), mares exhibited a greater (P = 0.0001)
inflammatory response to intradermal PHA compared to foals (Table 4-28, Figure 4-15).
The skin thickness of neither the mares or the foals returned to pre-inj section values by 48
h post-inj section (P = 0.0001).
Table 4-1. Fatty acid composition of the grain mix concentrate and the milled flaxseed
and encapsulated fish oil supplements
Fatty acid Grain mix Flaxseed Fish Oil
C8:0 ND ND ND
C10:0 ND ND ND
C12:0 ND ND ND
C14:0 ND ND 8.62
C16:0 17.18 5.60 21.14
C16:1 0.21 ND 13.77
C17:0 ND ND ND
C17:1 ND ND ND
C18:0 2.20 2.77 3.79
C18:1 26.28 13.90 7.87
C18:2n-6 (LA) 49.63 16.31 7.23
C18:3n-3 (ALA) 3.72 61.20 2.35
C20:4n-6 (AA) ND ND 0.69
C20:5n-3 (EPA) ND ND 15.03
C22:5 n-3 (DPA) ND ND 2.11
C22:6n-3 (DHA) ND ND 12.54
Total n-62 49.63 16.31 7.92
Total n-33 3.72 61.20 32.03
n-6:n-3 13.34 0.27 0.25
SPresented as g fatty acid per 100 g fat; ND = not detected in the feed stuff
2 Calculated as C18:2 + C20:4.
3 Calculated as C18:3 + C20:5 + C22:5 + C22:6.
Table 4-2. Fatty acid composition of winter and spring bahiagrass pasture and Coastal
Fatty acid Winter2 Spring3 Hay
C8:0 ND ND ND
C10:0 ND ND ND
C12:0 ND ND ND
C14:0 ND ND ND
C16:0 22.07 & 0.69a 23.22 & 0.67a 39.30b
C16:1 ND ND ND
C17:0 0.93 & 0.05a 0.531 0.28a 0.00b
C17:1 ND ND ND
C18:0 4.97 & 0.38a 3.36 & 0.22b 6.65"
C18:1 4.21 & 0.60a 1.39 & 0.02b 7.080
C18:2n-6 (LA) 23.71 + 1.96 18.13 A 1.55 23.48
C18:3n-3 (ALA) 41.52 & 3.28a 52.47 & 1.76b 15.900
C20:4n-6 (AA) ND ND ND
C20:5n-3 (EPA) ND ND ND
C22:5 n-3 (DPA) ND ND ND
C22:6n-3 (DHA) ND ND ND
Total n-64 23.71 + 1.90 18.13 A 1.55 23.48
Total n-35 41.52 & 3.28a 52.47 & 1.76b 15.900
n-6:n-3 0.59 & 0.08a 0.35 & 0.04a 1.45b
SPresented as g fatty acid per 100 g fat; ND not detected in the forage.
2 Winter Mean of December, January, February and March.
3 Spring = Mean of April, May and June.
4 Calculated as C18:2 + C20:4.
SCalculated as C18:3 + C20:5 + C22:5 + C22:6.
a,b,c Values in the same row having different superscripts differ at P < 0.05.
Table 4-3. Mare average daily fatty acid intake from December-Marchl
SPresented as g fatty acid/d; ND
2 CON no supplement, FLAX =
=not detected in any of the feedstuffs.
supplemented with milled flaxseed, FISH:
supplemented with encapsulated fish oil.
3Hay intake estimated at 1.0% BW (DM basis).
4 Calculated as C18:2 + C20:4.
SCalculated as C18:3 + C20:5 + C22:5 + C22:6.
Table 4-4. Mare average daily fatty acid intake from April-Junel
Faty aidTretmet2 Grain Spig Supplement Total diet
SPresented as g fatty acid/d; ND
2 CON no supplement, FLAX =
=not detected in any of the feedstuffs.
supplemented with milled flaxseed, FISH:
supplemented with encapsulated fish oil.
3Pasture intake estimated at 1.0% BW (DM basis).
4 Calculated as C18:2 + C20:4.
SCalculated as C18:3 + C20:5 + C22:5 + C22:6.
Table 4-5. Mare bodyweightsl
SPresented in kg.
2 CON no supplement, FLAX = supplemented with milled flaxseed, FISH =
supplemented with encapsulated fish oil.
3 d-28 to d-14 d before expected foaling; dO = foaling; d+14 to d+84 = d post-foaling.
Table 4-6. Foal bodyweights1,2
Time4 CON FLAX FISH SEM
dO 51.0a 54.4a 56.3a 2.21
d+14 74.1b 75.5b 75.4b 2.20
d+28 91.90 95.40 95.0" 2.20
d+42 109.4d 116.0d 115.0d 2.25
d+56 125.1e 127.6e 131.5e 2.23
d+70 144.8f 142.4f 144.4f 2.34
d+84 158.3g 158.5g 163.9g 2.25
Presented in kg.
2 Effect of time (P = 0.0001), effect of treatment (P = 0.75), effect of treatment x time
3 CON no supplement, FLAX = supplemented with milled flaxseed, FISH =
supplemented with encapsulated fish oil.
4 dO foaling; d+14 to d+84 d post-foaling.
a~b~c~d~e~f~g Values in the same column having different sub scripts are different at P < 0.05.
Time3 CON FLAX
d-28 629.2 619.0
d-14 631.3 626.3
dO 554.9 544.2
d+14 553.0 551.7
d+28 556.0 550.7
d+42 560.9 552.9
d+56 561.7 548.2
d+70 556.6 542.8
d+84 556.6 552.4
Table 4-7. Overall effect of treatment on the fatty acid composition of mare plasma
Fatty Acid CON FLAX FISH SEM P-value
C8:0 ND ND ND NA NA
C10:0 ND ND ND NA NA
C12:0 ND ND ND NA NA
C14:0 ND ND ND NA NA
C16:0 16.15 16.01 16.48 0.25 0.42
C16:1 0.88 0.88 1.10 0.09 0.17
C17:0 0.70 0.73 0.73 0.04 0.78
C17:1 ND ND ND NA NA
C18:0 20.08 20.18 20.45 0.32 0.72
C18:1 10.28 9.78 9.70 0.28 0.29
C18:2n-6 (LA) 46.43 46.87 44.37 0.71 0.05
C18:3n-3 (ALA) 2.99 3.65 3.06 0.23 0.09
C20:4n-6 (AA) 1.50 1.33 1.82 0.13 0.03
C20:5n-3 (EPA) 0.02 0.02 0.56 0.06 0.0001
C22:6n-3 (DHA) 0.05 0.03 0.61 0.05 0.0001
Total n-63 48.64 48.92 47.04 0.64 0.10
Total n-34 3.03 3.69 4.22 0.26 0.01
n-6:n-3 16.33 16.21 13.32 1.35 0.24
SPresented as g fatty acid per 100 g fat, ND = not detected in plasma, NA = not
2 CON no supplement, FLAX = supplemented with milled flaxseed, FISH =
supplemented with encapsulated fish oil.
3 Calculated as C18:2 + C20:4.
4 CRIC111ated as C18:3 + C20:5 + C22:5 + C22:6.
Table 4-8. Omega-6 fatty acid content of mare plasma
dO d+28 d+56 d+84 SEM Treatment
CON 49.2Y 45.8z 49.5Y 48.9Y 49.7'
FLAX 48.4 47.6 49.1 49.5 50.0
FISH 49.2Y 45.7z 46.8Y~z 47.6Y~z 45.9z
Presented as g fatty acid per 100 g fat.
2 d-28 d before expected foaling; dO foaling; d+28 to d+84
3 Calculated as C18:2 + C20:4.
a~b Values in the same column for each fatty acid not sharing a common superscript are different at P < 0.05.
y~z Values in the same row not sharing a common superscript are different at P < 0.05.
Table 4-9. Omega-3 fatty acid content of mare plasma
dO d+28 d+56 d+84 SEM Treatment
SPresented as g fatty acid per 100 g fat.
2 d-28 d before expected foaling; dO foaling; d+28 to d+84 = d post-foaling.
3 Calculated as C18:3 + C20:5 + C22:5 + C22:6.
a~b Values in the same column for each fatty acid not sharing a common superscript are different at P < 0.05.
y~z Values in the same row not sharing a common superscript are different at P < 0.05.