AN ABSTRACT OF THE THESIS OF
Karen M. von Deneen for the degree of Master of Science in Animal Science presented on July 10, 2002 Title: Plasma and Fecal Progestins during Placentation in the Mare.
Abstract approved:
Donald W. Holtan
Aims of this research were to find the time period and specific progestins reflective of the transition from ovarian source to placental source during early gestation in mares. An attempt was made to describe an accurate profile of total and individual progestins in plasma and feces from 64 to 150 days of gestation. There was also an attempt to determine if plasma and fecal progestins increased around 90 days of gestation as previously reported in this laboratory for urinary hormones, estrone sulfate (E1S) and pregnanediol glucuronide (PdG). Blood and fecal samples were collected and ovaries were examined via ultrasound every other day from days 64 to 150 of gestation. Plasma and fecal samples were analyzed using gas chromatography/mass spectrometry (GC/MS) with steroid derivatization. Eight progestins in plasma and ten progestins in feces were identified and quantified. In plasma, 20a-hydroxy-5a-pregnan-3-one (20a-5a) was found in highest concentration, while 5u-pregnane-3,2O-diol (f3-diol) was highest in feces. Progestins averaged approximately 100 times higher in feces than in plasma, while
-diol was about 800 times higher. There was a linear, parallel increase over time
(P<zO.05, ANOVA) for total progestins. There was a high, positive correlation between plasma and feces for most progestins, the exception being progesterone (P4), which was negatively correlated because over time, concentrations in plasma decreased while increasing in feces. Log ratios of feces to plasma indicated some progestins increased faster in plasma than feces, and vice-versa, while some remained constant. Spline regression (Gauss-Newton, SAS) analysis of the four main progestins, 3-hydroxy-5apregnan-3-one (3f-5a), 20a-5a, 5a-pregnane-3,20a-diol (1k-diol), and 3f3-diol,

indicated a difference in the rate of increase over time, occurring at about 112.7 days of gestation in plasma and 109 days of pregnancy in feces, which was nearly 20 days later than that reported for urine. Although the time of change in the rate of increase was highly variable among mares and specific progestins, for most, a clear change was evident by 115 days of gestation. This likely represents a functional feto-placental unit, fully capable of pregnancy maintenance.

Plasma and Fecal Progestins during Placentation in the Mare by
Karen M. von Deneen
A THESIS
Submitted to
Oregon State University in partial fulfillment of the requirements for the degree of
Master of Science
Completed July 10, 2002
Commencement June 2003

Master of Science thesis of Karen M. von Deneen presented on July 10, 2002
APPROVED:
Major Professor, representing Animal Science
Chair of Department of Animal Sciences
Dean of Graduate School
I understand that my thesis will become part of the permanent collection of Oregon State
University libraries. My signature below authorizes release of my thesis to any reader upon request.
Karen M. von Deneen, Author

TABLE OF CONTENTS
CHAPTER 1: INTRODUCTION
Definition of the Problem
Statement of Purpose
CHAPTER 2: LITERATURE REVIEW
Overview
Ovarian Anatomy and Function
Description of Progestins and Progestagens in the Mare
Description of Estrogens in the Mare
Endometrial Cups in the Mare
Description of Equine Chorionic Gonadotropin and Other
Hormones in the Mare
Steroidogenesis and Endocrinology in the Pregnant Mare
Embryological Development of the Equine Fetus
Placenta and Mechanisms of Placentation in the Mare
Structure and Function of Fetal Gonads in the Horse
Fetal Loss in the Mare
Exogenous Hormone Treatment in the Mare
Ovariectomy (OVX) and Hysterectomy (HX) in the Mare
CHAPTER 3: EXPERIMENTAL DESIGN
Objectives
Materials
Sample Collection
Ultrasound Procedures
Sample Preparation
Sample Analysis
Statistical Analysis
Page
1
1
3
3
5
7
8
10
12
16
18
23
25
27
30
33
34
35
32
32
32
33

TABLE OF CONTENTS (CONTINUED)
CHAPTER 4: RESULTS
Group Mean Results
Individual Mare Results
CHAPTER
5:
DISCUSSION
Overview
Hormone Identification and Quantification
Group Mean Results
Individual Mare Results
Future Considerations
LiTERATURE CITED
47
47
49
53
53
55
Page
36
43

UST OF FIGURES
Figure Page
2
3
4
5
6
Mean concentration and pooled standard error (SE) of total progestins
(log) in plasma (bottom line, SE = 0.057) and feces (top line,
SE = 0.069) from 64 to 150 days of gestation in mares (n = 4).
38
Mean log concentration and pooled standard error (SE) in plasma
(decreasing, SE = 0.103) and fecal (increasing, SE = 0.113) progesterone (P4) from 64 to 150 days of gestation in mares (n = 4).
40
Mean concentration and pooled standard errors (SE) of plasma
33-hydroxy-5a-pregnan-20-one (313-5a, SE = 179.753), 20a-hydroxy-
5a-pregnan-3-one (20a-5a, SE = 49.919), 5a-pregnane-33,203-dio1
(3-diol, SE = 944.158), and 5a-pregnane-33,20a-dio1 (13a-diol,
SE = 424.952) from 64 to 150 days of gestation in mares (n = 4). Spline regression (SR) indicates mean day (112.7) of increase for all progestins listed.
41
Mean concentration and pooled standard errors (SE) of fecal
33-hydroxy-5a-pregnan-20-one (3f-5u, SE = 1.405),
20a-hydroxy-5a-pregnan-3-one (20a-5a, SE = 2.655), 5a-pregnane-
31,20f3-dio1 (3-diol, SE = 1.463), and 5a-pregnane-33,20a-diol
(f3a-diol, SE 2.124) from 64 to 150 days of gestation in mares (n = 4).
Spline regression (SR) indicates mean day (109) of increase for 313-5a and
20a-5a only; 3f3-dio1 and 13a-diol did not show a defined change.
42
Mean ratio of feces to plasma (log) in 3j3-hydroxy-5a-pregnan-20.-one
(A, 3 -5u), 20a-hydroxy-5a-pregnan-3-one (B, 20a-5a), and
5a-pregnane-33,20u-dio1 (C, I3ct-diol) from 64 to 150 days of gestation in mares (n = 4).
44
Plasma (A) and fecal (B) 313-hydroxy-5a-pregnan-20-one (33-5a) in individual mares from 64 to 150 days of gestation.
45

UST OF TABLES
Table
2
3
4
Mean concentration of progestins in plasma (ng/ml) and feces
(ng/g) of mares (n =4) sampled from 64 to 150 days of gestation.
ANOVA table for the log concentration of total progestins in plasma and feces from 64 to 150 days of gestation in mares (n = 4).
Summary table of individual plasma and fecal progestins as compared to total progestins using ANOVA analyses.
Spline regression ranges for plasma and fecal
3-hydroxy-5u-pregnan-20-one (3f3-5a), 20a-hydroxy-5apregnan-3-one (20a-5a), 5a-pregnane-33,20l3-diol (1313-diol), and 5a-pregnane-3f3,20a-diol (3a-diol) in individual mares (n =4) from 64 to 150 days of gestation.
39
39
Page
37
46

LIST OF ABBREVIATIONS
Systematic Name
5a-pregnane-3,20-dione
4-pregnene-3,20-dione, progesterone
3a-hydroxy-5a-pregnan-3-one
20a-hydroxy-5a-pregnan-3-one
2O-hydroxy-5a-pregnan-3-one
5a-pregnane-3a,20a-diol
5c-pregnane-3a,2O-dio1
5a-pregnane-3,2Oa-dio1
5a-preane-3,2O-dio1
3-hydroxy-5-pregnen-2O-one
5-preene-3,2O3-dio1
5-pregnane-3,2O-dione
Estrone sulfate
Pregnanediol glucuronide
Estradiol 17
Prostaglandin F2ct
Dehydroepiandrosterone
Abbreviated Name
5a-DHP
P4
3a-5a
3-5a
2Oct-5a
2O-5u act-diol c4-dio1 a-dio1 f3-dio1
P5
P5-
513-DHP
El S
PdG
E17
PGF2a
DHEA

PLASMA AND FECAL PROGESTINS DURING PLACENTATION IN THE MARE
According to Evans et al. (1990), the mare is regarded as having a low reproductive efficiency because many mares that have successful performance records are kept as broodmares without consideration to existing reproductive problems.
Reproduction in the mare is a complex process, unique in many ways, including the anatomical site of ovulation, follicular growth patterns, long estrous periods, differentiation of fertilized and nonfertilized ova, and equine chorionic gonadotropin
(eCG) secretion in pregnant mares.
Progestin production in the mare is critical for the maintenance of pregnancy. For example, progesterone (P4) plays an essential role in placentation, embryo development, and placental functions. Profiling progestins, such as P4, in mid-gestation could be indicative of fetal well being and a successful pregnancy to term. Determining the time period when certain progestins increase rapidly might reflect pregnancy maintenance by the placenta. This information may allow for the development of diagnostic tests to measure the concentration of those particular progestins. Their determination may preclude exogenous progestin therapy. Exogenous progestin therapy is administered to ensure P4 levels are above 2 ng/ml (an adequate level for pregnancy maintenance).
Detecting progestins in both plasma and feces could be useful in reproductive studies of feral horses. For instance, if the same progestins correlate in concentration in both sources, only one sample would be necessary to obtain valid results.
Aims of this research were to determine the time period and specific progestins reflective of the transition from ovarian source to the placental source of progestins in

mare plasma and feces using a spline regression; to describe an accurate profile of total and individual progestins found in plasma and fecal samples taken every other day from
2
64 to 150 days of gestation; to correlate individual plasma to fecal progestins over time; and to see if certain plasma and fecal progestins increased around 90 days of gestation similarly to urinary hormones, estrone sulfate (E1S) and pregnanediol glucuronide (PdG)
(Riad and Holtan, 1993).
In one study by Heistermann et al. (1996), excretion patterns of estrogens and progestins in feces were highly correlated with those in urine throughout gestation in bonobos, but fecal steroids showed a lesser increase. Measuring E1S and PdG in urine was used to assess ovarian cycles since they reflected the changes in corresponding circulatory hormones in primates. That study found fecal P4 and PdG were significantly correlated with urinary PdG concentrations; therefore, measuring either steroid in feces could reliably monitor corpora lutea (CL) function.
On the contrary, the study by Sist et al. (1987) showed there was no correlation between plasma and fecal values of El S in the mare. Despite these findings, our studies will measure plasma and fecal progestin concentrations, and compare them to urinary
E1S and PdG levels.

3
Overview
Reproductive physiology and endocrinology of mares is similar to other domestic animals in some aspects, but quite different in other ways. A complete review of reproduction and endocrinology in the mare can be found in the 1992 edition of
Reproductive Biology in the Mare by O.J. Ginther. A complete review of reproduction and endocrinology of other domestic animals is located in Reproduction in Farm Animals by E.S.E. Hafez (1993). These features, along with exogenous hormone treatment in the mare, will be discussed.
Ovaries are considered to be unique endocrine and exocrine glands, and essential organs in reproduction and pregnancy. The inside of ovaries is composed of connective tissue, fofficles, and CL that are in varying stages of development or regression during the breeding season or pregnancy. Follicles not confined to a particular area or outer layer, as in other animals, are distributed throughout the ovary. Follicles are classified as primordial, growing, or Graafian. Primordial follicles consist of an oocyte surrounded by the zona pellucida. As one or more primordial follicles develop, they become surrounded by several layers of granulosa cells and are referred to as growing follicles. These types of follicles continue to develop into Graafian or dominant follicles (Evans et aL, 1990).
Future dominant follicles have an early size advantage over future subordinate follicles
(Gastal et al., 1997).
As follicles increase in size and develop, they migrate to the ovulation fossa.
Rupturing of these follicles is spontaneous with the follicular wall collapsing into the follicular cavity, along with hemorrhage from the thecal capillaries. Granulosa cells

undergo luteinization, and theca intema cells invade granulosa walls (Evans et al.,
1990), resulting in a structure known as the primary CL.
Recent research suggests that in the pregnant mare, the primary CL becomes the sole source of P4 maintaining pregnancy for the first 70 to 90 days (Discafani et al.,
1995; Albrecht et al., 1997). On the contrary, Allen (1980) stated that most primary CL activity declines 12 to 14 days post-ovulation. Large secondary follicles develop between 28 to 51 days of gestation, mature, ovulate, and form secondary CL that produce maximal levels of P4 around 55 to 80 days of gestation (Squires and Ginther, 1975;
Knowles et al., 1993). The mean concentration of P4 at that time is from 14.9 to 31.9
ng/ml (Schwab et al., 1990). Martin et al. (1989) also found that plasma P4 concentrations correlate to total luteal mass.
Formations of accessory CL in pregnant mares are maximal in the second and third month of gestation (Evans et al., 1990). Martin et al. (1989) noted that two-thirds of secondary CL form from luteinization of unruptured follicles.
In pregnant mares, follicular development begins to wane in early gestation.
Knowles et al. (1994) found no follicles greater than 29 mm in intact mares after 100 days of gestation, and no follicles greater than 19 mm at 148 days of pregnancy. All secondary CL were progressively smaller from 100 to 160 days of gestation (the mean number of CL was 11 on day 100 of gestation, and 14.6 between 100 to 160 days of pregnancy). Mean day of peak accessory CL identification was around 121.2 days of gestation, while by 160 days of gestation, the mean number of secondary CL identified was 10. Most accessory CL generally regress around 140 to 210 days of gestation
(Knowles et al., 1993). In addition, Daels et al. (1998) stated that secondary CL do not develop in all pregnant mares, and their development in time and numbers are extremely variable, indicating they may not be important in the outcome of pregnancy.
After each secondary ovulation, plasma progestagen concentrations increase and as many as 10 to 15 additional luteal structures are present by day 120 to 150 of gestation before degeneration occurs. At this stage, the placenta is well developed and maintains the function of progestagen production. However, a slight decrease in serum progestagen concentrations may occur by day 90 of pregnancy due to degeneration of secondary CL before the placenta has completely taken over hormone production (van Niekerk and van

Niekerk, 1 998a). In addition, increased estrogen secretion coinciding with eCG secretion is consistently associated with the presence of active CL and vice-versa between days 35 to 50 of pregnancy (Daels et al., 1991 a).
According to Discafam et al. (1995), the number of accessory CL was not dependent on the day of gestation. Holtan et al. (1979) noted there was no apparent relationship between the side of placentation, locations, and numbers of accessory CL. It was impossible to differentiate between primary and secondary CL, since most accessory
CL regressed by day 200 of pregnancy. However, CL in mares were functional long after
5 they were necessary to maintain pregnancy. This could be due to individual variations in follicular dynamics, eCG concentrations in the blood, and differences in secretion of other reproductive hormones in mares during pregnancy. It is postulated that secondary
CL remained in the ovaries of pregnant mares longer than necessary as a fail-safe mechanism to help ensure a successful outcome of pregnancy (Discafani et al., 1995).
Certain progestins seem to be necessary for maintaining pregnancy in the mare, altering ionic permeability across myometrial muscle cell membranes, increasing the resting membrane potential, and lowering cellular conduction and excitability (Peariman,
1948). Most progestins also serve as precursors for other steroids. Plasma progestins reflect physiological activity of the placenta and fetus since P4 is not present in significant amounts in mid- to late gestation in the feto-placental circulation. As P4 decreases during the second half of pregnancy, relaxin reduces uterine contractility.
There may be other progestagens present that are biologically active in suppressing uterine activity to prevent abortion (Short, 1959).
Predominant steroids in mare plasma in late gestation are 20a-Scx and a-diol, and are found in high concentrations of 2000 to 2400 ng/ml and 100 to 350 nglml respectively (Holtan et al.,
1991). They are first detected between 30 to 60 days of gestation, increasing gradually, and reaching highest levels 30 days prior to parturition
(Holtan et al., 1975; 1991). Burns and Fleeger (1975) found that total plasma progestagens increased to 25 ng/ml at 90 days before parturition and then to 60 ng/ml 10

days prior to birth, dropping to 58 ng/ml less than 5 days pre-partum. Holtan et al.
(1991) stated that 5a-DHP and 3-5a are of maternal origin, and are related to CL function since they both follow a pattern similar to P4 in early pregnancy. It is not until late in gestation when they are both produced by the feto-placental unit. Both 5a-DHP and 33-5a have a rather slow, constant increase the last third of gestation, with a rapid increase the last 20 days of gestation. Progestin 5a-DHP concentration in maternal plasma in late pregnancy is similar to P4 concentration in early pregnancy (Pashen and
Allen, 1979a). However, when 5a-DHP increases around 50 to 70 days of gestation and
P4 decreases around that time, this suggests initiation of placental steroid production
(Burns and Fleeger, 1975). Progestin 5u-DHP is known as a potent androgen and acts via testosterone receptors in the fetal prostate (Short, 1959), and is of placental origin
(Pashen and Allen, 1979a); it can also act via P4 receptors derived from the feto-placental unit (Short, 1959). Its values were found to be much higher in Thoroughbred mares than in ponies, implying genetic differences between different breeds of horses in the quantity of P4 metabolites (Hamon et al., 1991). Plasma concentration of progestins was found to be higher in mares carrying twins than in those carrying single foals (Morgenthal and van
Niekerk, 1991). Holtan et al. (1991) determined that no 5f3-pregnanes, 20u- and 203hydroxy-4-pregnen-3-one could be detected in plasma throughout gestation. In sheep,
20a- and 20-hydroxy-4-pregnen-3-one were detected, but not so in mare plasma (Miller and Holtan, 1997).
According to Holtan et al. (1991), the differential in progestin quality and quantity throughout pregnancy in equids most likely represented different sources and/or metabolic pathways. Miller and Holtan (1997) stated that due to high steroid concentrations (10 to 100 times higher in feces than in plasma), and a greater diversity of steroids (15 to 19 progestins in feces versus 10 to 12 progestins in plasma), only a selected few could be useful in determining placental function.

Estrogens are important in priming uterine tissues prior to labor and lactogenesis in the mare. Estrogens cause a tense uterine tone in early pregnancy between 10 to 50 days of gestation (Berg and Ginther, 1978). Estrogens have been found to stimulate oxytocin (OT) receptors in the sheep uterus, and help with distribution and rate of blood flow through uterine and placental blood vessels (Pashen and Allen, 1979b). Estrogens of conceptal origin are key factors in regulating carbohydrate and amino acid flux from the maternal system into the uterine lumen and conceptus. This mechanism assures a continuous source of nutrition in the pre-attachment period (Zavy et al., 1984).
There have been eight estrogens isolated from urine, the predominant being estrone (E1), equilin, and equilenin (Cox, 1975). When they were identified by Nett et al.
(1973), radioimmunoassays (RIAs) showed cross-reactions between estradiol
(Ei713) and other estrogens. The most abundant steroid hormones in plasma of pregnant mares are
E1 and equilin, which increase dramatically at the same time as hypertrophy of steroidogenic tissues in fetal gonads occurs (Merchant-Larios, 1979). Equilin plateaus after 180 days of gestation and is <100 ng/ml in plasma. Equilin has less biological activity than E1, and is more active than equilenin. The 173-hydroxy derivative of equilin is never found in urine (Cox, 1975). Daels et al. (1991a) stated that E173 is the most potent of all estrogens, but is found in low concentrations in circulation as compared to
E1.
Measuring
E1 in serum and plasma is a good indicator of fetal viability after 44 days of gestation. Normally,
E1 concentration increases between 34 to 40 days of pregnancy in serum and urine, and is predominantly secreted by the fetus and fetal membranes (Kasman et al., 1988). Lasley et al. (1990) also determined that measuring estrogen conjugates (EC) after 70 days of gestation was a direct test to determine pregnancy, since estrogens increase between 70 to 150 days parallel to fetal development.
On the other hand, Monfort et al. (1991) did a study with Przewalski horses and determined that an increase in urinary excretion of EC around 6 weeks of gestation indicated established pregnancy, and did not actually reflect fetal presence and viability until endometrial cup production ceased around 90 days of gestation. If fetal death occurred, endometrial cups would continue producing eCG.
7

According to Jeffcott et al. (1987), there was a rise in total estrogens of ovarian origin between 35 to 40 days of gestation, and high E1S concentrations were present due to a sulfo-transferase system in the endometrium of the pregnant mare. The ovary itself contributes significantly to EC concentrations between 20 to 70 days of gestation, while increasing levels at 33 to 39 days are due to the change in rate of ovarian estrogen synthesis (Daels et al., 1990).
Endometrial cups are a striking feature of equine pregnancy. These small, ulcerlike, endometrial outgrowths are formed between 36 to 38 days of gestation and are arranged in a circumferential pattern at the base of the pregnant horn (Steven, 1982). The annulate portion of the trophoblast and chorionic girdle invaded the endometrium during formation of the endometrial cups. The cups themselves are made of large decidual-like cells of fetal origin (Allen, 1975) rather than of maternal origin as previously stated by
Allen and Moor (1972). There is no physical integration between fully formed cups and fetal membranes (Allen and Stewart, 1978). They contain lipids and other materials to synthesize and secrete eCG and other substances (Yamauchi, 1975). Cups reach maximum size of 2 to 10 cm in diameter, and 1 to 3 cm in width by 70 days of gestation
(Allen and Stewart, 1978). According to a study by Douglas and Ginther (1975), endometrial cups reached maximum weight around 60 days of gestation, and did not decrease significantly in weight until 180 days of pregnancy. In general, the cups were desquamated completely by 100 days of pregnancy, and 4 g of necrotic tissue remained until 140 to 160 days of gestation.
Width and development of the progenitor chorionic girdle appears to be governed by paternal genotype. For example, a narrower chorionic girdle present in donkey and mule conceptuses results in formation of smaller endometrial cups. If mule embryos are transferred to a recipient, this transfer results in the development of a small chorionic girdle, giving rise to small endometrial cups at 36 to 38 days of gestation (Allen et al.,
1993). It was shown by Meadows et al. (1995) that failure of the donkey-in-horse

chorionic girdle to invade in vivo was not attributable to any defect in girdle cells themselves, but rather to the lack of some intrauterine stimulus.
Similar studies done by Allen and Short (1997) involving transferring donkey embryos to horse mares resulted in development of a small chorionic girdle that failed to invade the endometrium to form endometrial cups. Approximately 70% of pregnancies were aborted between 80 to 85 days in conjunction with delayed and abnormal placental attachment, along with vigorous maternal cell-mediated reaction against the xenogeneic donkey trophoblast. Normally, invasive trophoblast cells initiated the attachment and interdigitation of the non-invasive placenta for fetal sustenance, as well as the modulation of the materno-fetal immunological interaction to enable survival of the antigenically foreign fetus in the uterus.
According to Allen (1982), after about day 80 of pregnancy, endometrial cups become increasingly pale and begin to release a sticky, honey-colored secretion that adheres to the surface of the overlying allantochorion. This exocrine phase indicates the beginning of steady degeneration that finally results in necrotic cup tissue due to poor vascularization (Enders et al., 1995; Lea et al., 1995), and exocrine coagulum being sloughed from the surface of the endometrium between days 120 to 150 (Allen, 1982).
As endometrial cups mature, lymphocytes increase and accumulate in the endometrial stroma at the cup periphery. Allen (1980) noted the leukocyte reaction to endometrial cups was greater in mares bred to jack donkeys as compared to those bred to studs, and the reaction was also greater in jenny donkeys carrying a stallion conceptus.
On a molecular level of endometrial cups, Lea et al. (1995) discovered that endometrial tissue from the gravid horn of the uterus had a population of chromotrope 2R positive cells that contained transforming growth factor-132 (TGF-2) mRNA transcripts in the periphery of the cups. This factor suppressed potential cytotoxic activities of activated killer T cells CD8 lymphocytes. After invasion, mature cups down-regulated expression of the major histocompatibility complex (MHC) class I antigen, so its expression was turned off by 45 days of gestation. As a result, greater number of cells positive for mRNA encoding for TGF-2 occurred around 40 days of gestation rather than at a later stage. This is important for hybridization of maternal leukocytes in the

region of the developing cups, and maintenance of the cups until their degeneration around 80 to 100 days of gestation.
10
Equine chorionic gonadotropin (eCG), a highly glycosylated heterodimeric protein, expresses both FSH-like and LH-like biological activities in non-equine species.
The hormone eCG is the primary luteotrophic agent in the first half of equine pregnancy along with pituitary gonadotrophins LH and FSH, which stimulate secondary follicular growth necessary for formation of secondary CL (Stewart and Allen, 1979). In mares treated with eCG that have functional CL, production of androstenedione, estrogens, and progestins increased, proving eCG is luteotropic (Daels et al., 1998).
The hormones above, like all glycoproteins, possess a similar a-subunit polypeptide chain, while eCG, LH, and FSH have a different -subutht polypeptide chain
(Combarnous et al., 1991). Equine a-subunit shows about 80% homology with that of most mammals, differing only in transposition of tyrosine and histidine at positions 87 and 93 respectively (Hoppen, 1994). The a- and 13-subunits of eCG can be detected as early as 30 days of gestation in equine placental membranes (McDowell et al., 1993).
The hormone eCG first appears in peripheral blood between 37 to 41 days after ovulation, and its concentration reaches highest levels between 55 to 70 days of gestation.
It decreases and disappears from serum between 100 to 140 days of gestation (Allen et al., 1993). There are two types of eCG isolated from pregnant mare serum that differ in amino acid and carbohydrate composition. Low titer eCG (LT) has less carbohydrates than high titer eCG (HT), which has more phenylalanine and serine. The HT is also more biologically active than LT and has a longer half-life. The LT resembles eCG isolated from trophoblastic tissue. There is also a difference between pony and horse mares regarding eCG synthesis and chemical composition (Stewart and Allen, 1979).
A study by Albrecht et al. (1997) showed that eCG stimulated luteal androgen and estrogen production, increasing plasma concentration 2 to 3 fold. Secretion of eCG

increased luteal estrogen synthesis by transcriptional up-regulation of cytochrome
P450 17a-hydroxylase/17,20-lyase expression from the primary CL, suggesting that availability of aromatizable androgens could be rate-limiting in luteal estrogen synthesis prior to eCG production.
It was believed eCG bound the same luteal receptor as equine LH, but with a
11 lower affinity. For example, eCG only has 4% of the binding activity of LH to LH receptors. Also, eCG does not exhibit a stimulating effect to testosterone production in rat Leydig cells like LH (Combarnous et al., 1991).
In another study by Albrecht et al. (1997), there was a 24 hour delay in luteal androgen and estrogen production after eCG administration, indicating that steroidogenic enzymes must be synthesized first. This finding suggests that eCG influences luteal steroidogenesis at the transcriptional level.
In a study by Allen et al. (1993), eCG concentration in serum of pregnant mares and jenny donkeys carrying normal interspecies and hybrid interspecies pregnancies suggested production of eCG was influenced by parental gene imprinting. Interspecies pregnancies resulted in different size, secretion activity, and lifespan of endometrial cups.
Main function of endometrial cups is to secrete eCG in high concentrations (Allen et aL,
1973). Production of eCG ceases before the endometrial cups become completely degenerate; but hormone levels present in the maternal blood persist for some time
(Steven, 1982). Fetal genotype was found to be influential on eCG levels in maternal blood (Allen et al., 1973). According to Allen (1982), eCG tended to be higher in smaller pony breeds than in larger and draft breeds. He noted this inverse effect of mare size could be due to a dilution effect of greater blood volume in larger mares. Allen and
Stewart (1978) noted that if the mare had twins, eCG concentrations would double, while eCG levels decreased as mare age and parity increased.
Some other significant hormones in mare pregnancy include prolactin (PRL) and relaxin. Prolactin was found to be luteotropic in several species. In rat luteal cells, PRL promoted P4 secretion during short incubations in the presence of lipoproteins (Mennon et al., 1985). Low PRL levels (4.58 to 0.06 nglml) were used to maintain P4 secretion in humans. In mares that aborted between 28 to 44 days of gestation, there was no correlation found between plasma PRL and P4. However, in another study, there was a

positive conelation in pregnant mares between plasma P4 and PRL levels between 14 to 37 days of pregnancy (Schwab et al., 1990).
Relaxin is produced by the placenta and OT stimulates its secretion; there is no direct evidence that relaxin inhibits uterine activity in the mare. Its main function is to
12 soften the cervix of the mare pre-partum (Liggins and Thorburn, 1993). Relaxin can be utilized to diagnose pregnancy since it is high in concentration between 75 to 80 days of gestation, remaining elevated until its concentration sharply increases at the expulsive stage of labor (Stabenfeldt and Hughes, 1987).
According to Beckers et al. (1998), the mare has no placental lactogens, which are significant in ruminants to stimulate mammogenesis, fetal growth, and maternal metabolism. Instead, the mare has pregnancy-associated glycoproteins (PAG) to aid in these functions.
The feto-placental relationship is one of interdependence (Cottrill et al., 1991).
The resident fetus manipulates maternal biochemistry to suit its own personal living standards (Allen, 1984). The placenta is essential in hormone production. From earliest days of pregnancy, trophoblastic cells are veritable producers of a diversity of hormones.
This endocrine function is particularly evident in the human placenta, as well as in the mare, which produces protein and steroid hormones in large quantities such as estrogens,
P4 and other progestins, eCG, and relaxin (Beaconsfield and Villee, 1979).
Based on studies by Holtan et al. (1979) using OVX mares, the feto-placental unit is capable of producing enough progestins to support pregnancy due to various metabolic pathways as early as 50 days of pregnancy. The placenta produces P4 around 85 days of pregnancy (Knowles et al., 1993); however, it is not detected in plasma after four months of gestation (Ganjam et al., 1975). In the mare, the major source of P4 changes gradually from the ovaries to the placenta between days 50 to 70; concentration of plasma P4 reaches maximal levels around day 100 of gestation (Vivrette, 1994) and decreases from day 150 to 180 (Holtan et aL, 1975).
As reviewed by Thorburn (1993), the placenta, endometrium, and fetal

13 adrenals contain side-chain cleavage enzyme (P450 scc) in late pregnancy. It cleaves the side chain from cholesterol and is the rate-limiting enzyme in synthesizing P5.
Progesterone (P4) and its metabolites are synthesized in the placenta and endometrium as well, reaching the myometrium by diffusion.
Pashen and Allen (1979b) stated that converting P5 to P4 via 3-HSD was a crucial step in synthesizing most steroids including progestins. The placenta has 3-HSD activated from an early stage of pregnancy, and is able to use maternal cholesterol to synthesize PS and P4, therefore the fetus is not involved in P4 synthesis since the placenta produces adequate amounts of it starting around 50 days of gestation. However, blocking 3-HSD activity did not block metabolism of PS to other progestins, thus an alternate route of metabolism existed. However, Shideler et al. (1982) found no data to show the placenta produced P4 prior to 50 days of gestation.
The placenta also contains 3-oxidase, 3-hydroxylase, and 3-hydroxysteroid dehydrogenase (3-HSD), while 5a-reductase and 20a-hydroxylase are present in the endometrium. The placenta and endometrium have enzymes that are able to convert P5, which is of fetal origin, to 5a-DHP, and then to 5a-pregnanes, such as 3-5a and -dioI
(Schutzer and Holtan, 1996; Chavatte et al., 1997).
As for estrogen synthesis, Raeside et al. (1973) discovered the placenta could not produce estrogens from C21 steroids, but was able to aromatize androgens to estrogens via other methods. It could not convert P5, P4, and 17a-hydroxylated derivatives to estrogens either. The mare is capable of producing ring f unsaturated estrogens, such as equilin. Ring t unsaturated estrogens are biosynthesized by the pathway not involving triterpene (C30) squalene, the sterol cholesterol (Tait et al., 1983). Pashen and Allen
(1979a) found the
E17 precursor was dehydroepiandrosterone (DHEA) sulfate.
Bhavnani et al. (1969) found DHEA was the precursor for E1, but not for equilin.
Bhavnani et al. (1969; 1975) isolated 313-5a, f3f-dio1, and 3a-diol along with '4C-labeled
E1, E17, equilin, and equilenin, which indicated they shared a similar biosynthetic pathway up to isopentenylpyrophosphate using the radiolabeled precursor. Bhavnani and
Short (1973) delineated the classical pathway of neutral and phenolic steroid biosynthesis: acetate to squalene to cholesterol to a pregnene derivative to androgens, and

finally to estrogens. They noted
E1 was formed from DHEA sulfate, since 7H-DHEA,
14C-androstenedione, 3H-cholesterol, and '4C-acetate were converted to classical estrogens, and only '4C-acetate was transformed to ring unsaturated estrogens. Ring
14 unsaturated estrogens formed after bifurcation in the classical pathway of steroid biosynthesis. This occurred after formation of mevalonic acid, and before formation of squalene. The precursor was 1 -14C-isopentenylpyrophosphate. Ring unsaturated estrogens appear in urine around the fourth month of gestation, reach maximal levels around 8 to 9 months of pregnancy, and are not present following birth; thus, placental or fetal enzymes are less likely to be involved.
Ainsworth and Ryan (1970) discovered goat placental tissue was able to synthesize estrogens from androstenedione in vitro, while Ainsworth (1972) found the presence of steroid sulfate synthesis in the placenta of cattle, guinea pigs, and sheep.
Channing (1969) reported androstenedione and testosterone could be converted to
E1 and
E17 in equine granulosa cell cultures with 15% horse serum, 30% medium '199,' and
55% Hank's solution. The same author also found acetate could be converted to P4 and
E17, and this pathway was similar to the one in luteal tissue.
In another study, it was discovered equine granulosa cell cultures secreted mainly
P4 and 20a-hydroxy-5-pregnen-3-one, even though ovarian cell types differed quantitatively rather than qualitatively in their pattern of steroid secretion. Granulosa cells were found to have all enzymes necessary for biosynthesis of P4 and estrogen
(Channing and Grieves, 1969).
The horse conceptus produces a variety of estrogens and androgens that are involved in embryonic signaling. Estrogens produced increase around 12 to 20 days post-ovulation. Marsan et al. (1987) discovered two cell types, high and low density cells, present in the conceptus that have different steroidogenic potentials. For example, low density cells (LDC) primarily produced P4, and high density cells (HDC) produced
E17J3.
Holtan et al. (1991) have shown major steroids entering the placenta from fetal
diol. The fetus delivers P5 in large amounts via the umbilical artery to the placenta for

metabolism. Progestin 5cc-DHP was one of the major steroids in umbilical and uterine
15 veins, but not in their respective arteries (Hamon et al., 1991). 3- and 2O-
Hydroxylation predominate in the feto-placental unit. The placenta converts P5 to P4, which is found in high amounts in the umbilical vein, but no P4 is found in the uterine vein, which contains 5a-reduced metabolites of P4 and P5; therefore, what little amount of P4 secreted by the uterus is metabolized before returning to the uterus, and 5areductase activity is present in the placenta (Hamon et al., 1991; Holtan et al. 1991). In cows and ewes, radiolabeled substrate revealed the major site of 5a-reductase was in the endometrium along with 20a-hydroxylase, thus 2Oc-5a and a-dio1 were found in the circulation. As a result, a-diol could be a direct metabolite of P5, or it could be indirectly synthesized from P5 via P4 and 5a-DHP (Hamon et aL, 1991).
Holtan et al. (1991) measured the concentration of 8 progestins in fetal circulation, and determined the total amount of progestins in fetal arteries or maternal veins were similar, but higher than those found in fetal veins or maternal arteries. This difference in the total amount and concentration of progestins indicated metabolism by the placenta from the fetus to the mare and vice-versa.
Regarding location of progestins in fetal and maternal circulation, P4 was seldom located in the uterine vein and never in the maternal artery. Progestins 3-5u and 3-dio1 were found in fetal vessels, while a-dio1 was located in maternal vessels, therefore, 3and 2O-hydroxylation was occurring in the feto-placental unit, and 20a-hydroxylation was present on the maternal side. Progestin 20a-5a was found in highest concentration in the uterine vein at 738 ng/ml, thus it was produced by the endometrium. It was unclear why reduced pregnanes predominated (Holtan et al., 1991).
The equine fetal liver is involved in converting Cl 9 steroids to substrates that can be further metabolized into estrogens in the placenta (Pashen and Allen, 1979b). By day
100 of gestation, the fetal liver increases its enzyme activity in the transformation of
DHEA to 7a-hydroxy-DHEA, which is transported to the placenta where it is further metabolized and aromatized to equilin and equilenin. Phenolic and ring unsaturated estrogens in pregnant mare plasma and urine increase around 180 to 210 days and 210 to
240 days respectively. Fetal liver also converts PS to P5and -diol; P4 to 3-5a

16 and 3I3-diol; and 3-5a to 5a-DHP, 20x-5a, and 2O[-5a (Schutzer and Holtan, 1996;
Chavatte et al., 1997).
Comparing a few differences between mare and jenny steroidogenesis, Heap et al.
(1991) noted that in the donkey conceptus, aromatase activity was the same in all fetoplacental tissues, except it was lower in the horse conceptus. There was a higher proportion of labeled precursor incorporated into E173 by extra-embryonic tissues in the donkey than in the horse.
Estrogens and progestins are excreted significantly in bile and reabsorbed in the gastrointestinal tract (G1T), especially in humans. Intestinal microflora and mucosa contribute significantly to levels of active hormones in the organism due to biliary excretion of these metabolites (Adlercreutz et al., 1979). This may be the case in the mare (Miller and Holtan, 1997).
Adlercreutz et al. (1979) identified and quantified 12 estrogens in human feces along with P4 metabolites that were found in unconjugated, monosulfate, disulfate, and glucuronide form in feces. There were more 3-5a-pregnane isomers and 17a-pregnane derivatives in pregnant feces than in bile due to 3-hydroxysteroid oxidoreduction and 16dehydroxylation of biliary pregnanes by intestinal bacteria, such as Bacteroides fragilis, which converted
E1 to E17. Such studies would be significant to determine if this affects mare steroidogenesis and steroid excretion.
With regard to other steroid biosynthesis in the mare, active cortisol is converted to inactive cortisone by 11 -hydroxysteroid dehydrogenase (11 f3-HSD) in the placenta, which protects the fetus from excess maternal cortisol (Chavatte et al., 1995).
Embryological Development of the Equine Fetus
Survival of the foal at birth begins at conception and depends on normal intrauterine development, pre-partum maturation of organ systems vital for extra-uterine life, and an uncomplicated delivery at full term (Rossdale, 1993). Following a few days after conception, Barer and First (1983) noticed the conceptus spend 144 hours at the ampullary-isthmic junction of the oviduct before entering the uterus. An important

function of the conceptus at this point, as noted by Hershman and Douglas (1979), is the blastocyst must be present in the uterus by day 12 post-ovulation in order to prevent
17 luteal regression in the mare. Allen (1980) and Enders et al. (1988) found only fertilized eggs enter the uterus around 5 to 6 days post-ovulation, and unfertilized eggs remain in the fallopian tubes. Previous reports have suggested that the blastocyst reaches 2 mm in diameter around 9 days post-ovulation and grows rapidly (3.4 mm/day) to reach 24 mm by day 16 of gestation (van Niekerk and Allen, 1975; Ginther, 1992). The blastocyst remains spherical until 50 days of gestation (Bazer and First, 1983). More recent research by Allen (2000) states that the blastocyst becomes more elongated after 21 days of pregnancy. The conceptus is mobile in the uterus from the time of entry into the uterus to 15 days of gestation, traveling to all portions of the uterus 10 to 20 times per day.
Mobility decreases by day 15 of gestation and ceases by day 17 (Gastal et al., 1996).
This process, called the "maternal recognition of pregnancy," prevents luteolysis and subsequent early embryonic loss.
Fixation generally occurs in the caudal part of the uterine horn near the bifurcation of the uterine body and horn around day 16 post-ovulation (Enders and Liu,
1991). Gastal et al. (1996) noted that the larger the embryonic vesicle, the sooner fixation will occur. It was shown that more than 80% of conceptuses implanted in the horn opposite to the one occupied by a previous pregnancy. Allen (1984), however, found placentation was random in maiden mares.
In general, an increase in uterine tone during fixation is associated with a decrease in uterine diameter; therefore, the expanding embryonic vesicle is fixed in the bend in the caudal portion of the uterine horn. Side of cord attachment is the same as the side of fixation (Ginther, 1994).
The conceptus is surrounded by an acellular glycoprotein capsule until 22 days of gestation, relying on endometrial gland secretions for its sustenance (Stewart et al.,
1995). Its heartbeat can also be detected via ultrasound on day 22 of gestation (Squires et al., 1988). Griffin and Ginther (1991) noted that the early fetus (69 to 81 days) was quite mobile and changed positions and location in allantoic fluid several times per hour.
Regarding organogenesis pertinent to this project, fetal adenohypophysis (anterior pituitary, AP) was identified on an ultrastructural level by 100 days of gestation.

Neurohypophysis (posterior pituitary, PP) normally develops from the floor of the diencephalon, and adenohypophysis forms from Rathke's Pouch. The AP is linked functionally to the thyroid gland, gonads, and adrenal cortex via production of trophic hormones. In fetal life, these relationships are complicated by feedback of hormones secreted by the placenta and maternal endocrine organs. For example, in rabbits, little placental hormones are produced, thus fetal pituitary glands are responsible for development of the male genital tract, glycogen storage, and adrenal function (Samuel et al., 1975a).
From the point of phylogeny, the placenta is a device that evolved when animals ceased to be oviparous and became viviparous. In placental animals, protection of the fetus is supplied by the walls of the mother's abdomen and brood pouch, and by amniotic fluid, which provides flotation in a water jacket and prevents desiccation. The placenta takes over the nutritive function by establishing a connection with parental circulation, supplying food substances, and providing developmental building blocks to the fetus
(Ramsey, 1982).
The placenta of the mare is classically described as being non-invasive in its activity, epitheliochorial in its structure, and diffuse in its architecture (Allen, 1982). In general, the placenta consists of six tissue layers from the outermost to the innermost respectively: maternal endothelium, connective tissue, maternal epithelium, trophoblast, fetal connective tissue, and fetal endothelium (Leiser and Kaufmann, 1994). Collins
(1993) stated that all mammalian placentas develop convolutions at the materno-fetal interface based on various layers present. For example, pigs have the simplest type of interdigitation. In the horse, there is no trophoblastic invasiveness, decidual cell reaction, or formation of the syncytio-trophoblast, which is present in ruminants (Leiser and
Kaufmann, 1994). The large yolk sac is an integral part of the survival of the embryo.
However, early on, the equine embryo develops an unusual glycocalyx capsule that completely surrounds the embryo prior to the formation of the zona pellucida. This capsule functions to accumulate proteins and other components of the internal

endometrial glands or "uterine milk." It becomes dysfunctional around 50 days of gestation (Jones, 1993).
The three main compartments of the placenta in the mare are the amnion, allantois, and chorion. The amnion begins to develop around 16 days post-fertilization,
19 and the amnionic cavity is completely formed by 21 days of gestation (Jones, 1993). The amnion is an epithelial layer derived from embryonic ectoderm by folding. It achieves support from the mesenchyme surrounding the fetus (Leiser and Kaufmann, 1994). The allantois emerges around 25 days of pregnancy, while it begins fusion with the chorion to develop the allantochorion (Jones, 1993). The pig and domestic ruminant allantois takes form of an elongated T-shaped sac, which reaches the extremities of the extra embryonic coelom by passing between the chorion and one side of the amnion. In the mare, relationships of the allantois are different. Equine allantois grows from the hindgut into the extra embryonic coelom, and expands dorsally over the amnion. In this manner, the future umbilicus is subdivided into two distinct yet continuous parts (Steven, 1982). The allantoic sac stores fetal urine via the urachus (Jones, 1993). The chorion is the epithelial layer derived from the outer blastocystic wall (Leiser and Kaufmann, 1994). The band that completely encircles the blastocyst is known as the chorionic girdle. Cells of the chorionic girdle atrophy and leave a scar. The actual girdle separates from the conceptus at 40 days of gestation and invades the endometrium. Cells then migrate into the stroma between the uterine glands to establish endometrial cups (Jones, 1993).
Placentation in equids occurs at a later time compared to that in other large mammals, in part because of the unique blastocyst capsule that envelops the conceptus until day 23 of gestation (Stewart et al., 1995). The equine blastocyst retains a roughly spherical shape until about day 21 of gestation (Allen, 2000). Only after about day 40 of pregnancy does any intimate and stable contact occur between the allantochorion and the endometrium (Bjorkman, 1973; Allen, 2000). In early pregnancy, except in regions of developing endometrial cups, the trophoblast lays in simple apposition to the uterine epitheium. As allantoic vessels begin to vascularize the chorion, short finger-like evaginations of the chorion develop along their course, until day 40 of pregnancy when most of the membrane surface is covered by rudimentary primary viii (Steven, 1982).
During the next 20 days, blunt, finger-like viii of the ailantochorion form a tight-fitting

interdigitation with the villi of the endometrium (Allen, 2000). Attachment between the uterine epithelium and a wide band of trophoblast in the equatorial region of the chorionic sac is established around 46 days of gestation. At this time, the trophoblast consists of tall columnar epithelium supported by a collagenous basal lamina (Samuel et
20 al., 1976). By day 56, the allantochorion has expanded from the base of the pregnant uterine horn into the uterine body, and by day 77, it has extended into the contra lateral non-pregnant horn. As it continues to do so, a band of modified trophoblast develops in a relatively avascular region of the chorion just before the leading edge of the allantois
(Steven, 1982).
Between day 60 and 150 of pregnancy, the maternal epithelium is greatly reduced in height. This modification indicates a change in the requirements of the fetus for oxygen and metabolites as gestation increases. By day 100 of gestation, primary villi develop secondary branches (Steven, 1982) marking a complex feto-maternal interdigitation both at a macro- and microvillous level over the entire endometrial surface
(Allen, 1982). Formation of tertiary villi, a distinctive feature of a mature equine placenta called microcotyledons, is completed by day 150 of gestation (Samuel et al.,
1974; 1975b), but recent research suggests this process is completed by 120 days of pregnancy (Allen, 2000). Microcotyledons are believed to represent principal sites of gas and small molecule exchange (Steven, 1982). Their arrangement is consistent with the counter-current blood flow system in the mare and other species (Silver, 1984).
There is intense pinocytotic activity by trophoblast cells as well as development of rough endoplasmic reticulum (RER) and cellular organelles during the first 200 days of gestation. By 300 days of gestation, smooth ER (SER) is developed. The HSD enzymes, which play a role in steroid interconversions in trophoblast tissues, are correlated with the development of SER. These changes suggest trophoblast cells are more highly involved in synthetic processes as the time of pregnancy progresses (Samuel etal., 1976; 1977).
On a molecular level of placentation, the mare is an excellent model to study cellular events associated with early pregnancy because uterine and embryonic tissues can be collected easily for experimental purposes (Brady et al., 1993). In-situ hybridization results of Stewart et al. (1995) supported the proposed model of girdle

21 development, and provided strong evidence that a component of mesoderm-derived mitogenic stimulus was hepatocyte growth factor-scatter factor (HCG-SF) and its receptor, proto-oncogene c-met. The HCG-SF was strongly expressed in allantoic mesoderm, and c-met was expressed strongly in trophoblast tissues. For example, HCG-
SF induced epithelial differentiation of mesenchyme in chorionic girdle cells during embryonic development.
According to Lennard et al. (1995), in-situ hybridization techniques and Northern blotting revealed transforming growth factor
Ii
(TG931), a multifunctional growth factor, was responsible for regulating endometrial and/or trophoblast growth and differentiation during placentation. It could also influence fetal development, via placental transfer, at later stages of gestation.
Northern blot and in-situ hybridization techniques performed by Lennard et al.
(1998) have demonstrated an increase in mRNA encoding epidermal growth factor (EGF) in endometrial tissues of mares, coincident with interdigitation between the allantochorion and endometrium during placentation. The EGF was found to play an important role in placental and fetal growth throughout gestation. High EGF expression was shown to be in the epithelium of endometrial glands until about day 250 of gestation.
Binding of EGF was high in fetal membranes before implantation (day 30 to 34 of pregnancy), and in the fully developed placenta (day 150 to 250 of gestation). Since EGF could be secreted in the uterine lumen, it could stimulate maternal or fetal epithelial layers of the placenta, and be transferred to the developing fetus. An unusual feature about EGF was that it was expressed as a large, membrane bound precursor molecule that underwent proteolytic cleavage at the membrane level to release growth factors.
Upregulation of EGF could be due to increasing levels of eCG, estrogens, and progestagens.
Schorpp-Kistner (1999) discovered JunB, an immediate early gene product and member of the AP- 1 transcription factor family, seemed to be essential to the embryo in establishing proper vascular interactions with maternal circulation. The JunB was involved in multiple signaling pathways regulating genes that established a proper fetomaternal circulatory system, since placentation requires extensive angiogenesis to begin transplacental exchange between the mother and fetus. The trophoblast is rich in

angiogenic growth factors (VEGF) and proliferin secreted by trophoblastic giant cells that stimulate uterine neovascularization.
Unlike in the mare, human mechanism of placentation uses VEGF and placental
22 growth factor (P1GF). Both are used to induce endothelial cell proliferation (Iruela-
Arispe, 1997). McDowell et al. (1995) described another important protein found in the mare endometrium that plays an important role in placentation. Retinol binding protein
(RBP) is a specific carrier protein for retinol, which is involved in hematopoiesis and differentiation of the retina during embryogenesis. The RBP is produced by the conceptus and endometrium, has acid phosphatase activity toward P-nitrophenyl phosphate; E17f3 and P4 control changes in it. Total protein concentrations increase with advanced luteal function and decrease with a decrease in P4. RBP was also found in the endometrium of cows, ewes, and sows, although there were some species differences in the effect of ovarian steroids on its production.
Badinga et al. (1994) studied uterine expression of mRNA encoding antileukoproteinase (ALP) and its role in elastase and cathepsin G protease inhibitor in feto-maternal interactions in pregnant horses, cattle, rats, and mice. They found abundance of ALP mRNA in the equine endometrium, which increased between 125 to
170 days of gestation, and decreased by 215 days of pregnancy. The ALP expression established distinct uterine-placental interactions. Increase in ALP levels was due to an increase in
E17 concentrations. Both ALP and uterine cathepsin G protease inhibitor were important in preventing enzymatic degradation of the uterine-placental interface.
With brief regard to placental physiology, the placenta in general behaves like a lipid membrane towards exogenous compounds such as environmental pollutants, drugs, and other ingested chemicals. In contrast, all substances essential for fetal development
(apart from respiratory gases) cross the placenta against an electrochemical gradient.
This is a process of considerable specificity. For example, the placenta differentiates between those proteins being transported to the fetus and those that undergo proteolysis in placental tissue (Beaconsfield and Villee, 1979).

23
Sexual differentiation of the fetus occurs around 39 to 45 days. Gonadogenesis occurs at the genital ridge with migration of gonocytes from the primitive hindgut. Sex chromosome complement of gonocytes has no influence on sex determination and gonadal blastema cells, which manifest their sex chromosome complement during the development of gonads, although eCG may influence stimulation of gonadal hypertrophy
(Walt et al., 1979).
Early cytodifferentiation of somatic cells into steroidogenic cells takes place before sex differentiation of gonads. Seminiferous cords of testis are completely segregated from steroidogenic tissue by a basal lamina, and in the medullary of the ovary, steroidogenic cells are differentiated inside epithelial cords containing germ cells.
Females remain undifferentiated for a longer time, and no steroidogenic tissue is found before folliculogenesis takes place. Mesothelium, mesonephros, and mesenchyme participate in establishing undifferentiated gonads in the horse and rat, but with certain differences, such as no formation of undifferentiated gonadal blastema in the horse. Fetal ovaries are enclosed in a cortical layer surrounded by germinal epithelium around 40 to
45 days of gestation. Oocytes are present between 90 to 120 days of gestation deeper within the cortex of the ovary (Walt et al., 1979).
Allen (1980) noted that fetal gonads enlarge around 100 days of gestation and are often larger than maternal ovaries between 230 to 250 days; therefore, their growth is paralleled by a sharp increase in conjugated
E17 in mare blood and urine. The reason for gonadal increase around 250 days of gestation is due to hypertrophy and hypoplasia of
Leydig cells, which are predominant cell types in both sexes; these play a role in steroid biosynthesis or detoxification of xenobiotic compounds. Hay and Allen (1975) stated that despite gonadal enlargement, there seems to be no change in the ultrastructure of gonads between 100 to 250 days of pregnancy. However, Merchant-Larios (1979) found the absence of ultrastructure differentiation of steroidogenic cells did not indicate steroidogenesis was not occurring. After 300 days of gestation, gonads regress and degenerative changes occur in interstitial cells, but no marked ultrastructural differences

from younger gonads are evident, except that dense bodies are more numerous, larger,
24 and less regular in shape (Merchant-Larios, 1979).
Gonads contain lots of SER in the third month of gestation, indicating they are highly developed and function in steroid synthesis, lipid storage, and detoxification
(Gonzalez-Angulo et al., 1975). Hydroxylating activity of cytochrome P450 system is observed in fetal testes, therefore biosynthesis of steroids is present (Rogerson et al.,
1993). Despite this, Flood and Marrable (1975) found fetal gonads lack HSD enzymes, but a variety of these are found in trophoblast tissue as early as 13 days post-ovulation.
Fetal adrenal glands and kidneys have limited HSD activity, unlike in sheep which use
3f3-HSD substrates in mid-gestation. The fetal liver in the horse has limited 313-HSD.
However, HSDs are located in the trophoblast, uterine epithelium, and endometrial cups, therefore lots of steroid conversions are occurring. Although the cups contain HSDs, this does not mean they have important steroidogenic function; instead, this indicates their line of descent.
Gonads have a high content of z5-steroids and DHEA (Hay and Allen, 1975) along with 5,7-dienes, 3-hydroxy-5,7-pregnadien-20-one, and 3J3-hydroxy-5,7androstadien-l7-one, indicating the presence of a 5,7-diene pathway to synthesize equilin in the placenta (Tait et al., 1983). The DHEA is converted to estriol (E3) and other estrogens by the placenta as well (Hay and Allen, 1975). When gonads regress around
280 days of gestation, there is a decrease in DHEA output, causing a fall in estrogen levels in mare plasma (Pashen and Allen, 1979b).
In other studies, guinea pig, cattle, and rabbit ovaries were able to convert testosterone to E17f3 at the same age that testes began to secrete testosterone. There was no difference found with respect to steroids secreted by the fetal ovaries or fetal testes
(Merchant-Larios, 1979). Complete fetal gonadectomy (GX) results in a rapid decrease in maternal estrogens, but only to one-half the total amount if only one fetal gonad is removed (Raeside et al., 1973). In subsequent work, Pashen and Allen (1979a) reported similar reductions in maternal estrogens following fetal GX, but found that progestins remained elevated. From this same work, they reported lower 13,14-dihydro-15-oxo-
PGF-2a (PGFM) at term after GX, and thus reduced uterine contractions and difficult

births in some mares (Pashen and Allen, 1979b). Barnes et al. (1975) indicated fetal estrogen and progestin levels were much higher than those found in the mare in late gestation.
25
Early pregnancy loss (EDL), defined by Ginther (1985b) as "...degeneration and resorption of debris and placental fluids and expulsion of the embryonic vesicle through the cervix," is of major concern to horse breeders. Approximately 12.5% of mares diagnosed pregnant on equine brood farms are victims of EDL. Ginther (1985a) stated pregnancy loss occurs between 7 to 17% in individual mares, and abortion is less likely to occur after 60 to 75 days of pregnancy. However, fetal death is more likely in maiden mares, geriatric, and/or malnourished mares between 25 to 31 days of gestation (Irwin,
1975; Ginther, 1985a; Woods et al., 1987; Woods, 1989). van Niekerk and van Niekerk
(1998a) stated embryonic loss depended on a variety of factors such as etiology of disease, endometrial cysts, twin pregnancies, environmental conditions, history, chromosomal abnormalities, and others. In general, most pregnancy failures resulted from abnormal fetal development or abnormal uterine environmental conditions, rather than primarily due to P4 deficiency or CL insufficiency (Forde et al., 1987; Stabenfeldt and Hughes, 1987). Irvine et al. (1990) supported this finding because pregnancy loss between 17 to 42 days of gestation was rarely coincident with a fall in plasma P4, since
P4 generally decreased after abortion. Darenius et al. (1991) noted the most critical time for pregnancy loss was up to 4 to 5 weeks of pregnancy. Good management and minimal stress were suggested as preventive methods.
During early pregnancy, low or decreasing P4 concentrations were used to diagnose "at risk" mares. However, Jeffcott et al. (1987) stated measuring maternal P4 concentration is often unreliable to diagnose embryonic death because pseudopregnancy could occur, thus high P4 levels could be present even during fetal loss. They suggested monitoring fetal well being via
E1 concentrations after 40 days of gestation instead.
Consistent concentrations of >4 ng/ml of P4 were associated with embryo survival. II P4 concentrations were low on day 4 of gestation, this aided in differentiating

luteal insufficiency from premature, uterine-induced luteolysis until day 5 of pregnancy when the primary CL was most susceptible to luteolysis (Ginther, 1985a;
Ginther et al., 1985). This could be caused by inadequate stimulation of granulosa cells at ovulation by LH, resulting in improper CL formation due to being fed low quality protein lacking essential amino acids necessary for proper LH production (van Niekerk and van Niekerk, 1998b). However, colic, navicular disease, acute babesiosis, and treatment with prednisolone and anabolic steroids could cause a dramatic drop in overall progestins in mares (Ginther, 1 985a; Villahoz et al., 1985).
According to Ginther (1985a), there was a positive relationship between luteal phase P4 concentrations and pregnancy rate in cattle and sheep. Short interovulatory intervals due to luteal regression also occurred repeatedly in mares.
Bell and Bristol (1987) found no difference between pregnancy loss occurrence between mares treated with exogenous progestins and
E17 as compared to control mares.
With respect to pregnant mares, mean P4 levels were lower on day 14 post-ovulation in mares that were not pregnant, and in mares that resorbed their pregnancies between day
14 and 21 of gestation.
Bergfelt et al. (1992) found luteal regression resulted in pregnancy loss occurring before day 20 of gestation. Low P4 concentrations and decreased CL diameters were present on day 12, 15, and 18 of fetal loss. A decrease in P4 concentration before or on the day of fetal death has been associated with acute endometritis. Daremus et al. (1987) noted when early fetal death occurred, the embryonic sac collapsed and its content spread into the uterus. According to Irvine et al. (1990), a decrease in P4 concentration was rarely a cause of pregnancy loss.
However, Bergfelt et al. (1992) associated early embryonic loss with a decrease in
P4 due to endometritis, failure of the conceptus to block luteolysis, and primary luteal insufficiency or regression. Ginther et al. (1985) noted that primary CL insufficiency was inadequately documented as a cause of fetal loss, and the primary CL was maintained if embryonic loss was >20 days of gestation rather than earlier. Bergfelt et al.
(1992) found the resurgence of primary CL, along with an increase in size and P4 production, occurred in many mares between 35 to 40 days of gestation due to eCG release into the circulation.

27
Exogenous hormone treatment is indicated for cases with documented luteal insufficiency and P4 deficiency, and a history of abortion. Pregnancy loss is considerably greater during the first trimester, especially between days 0 to 39, and usually if P4 levels are less than 2.0 ng/ml in the blood (McKinnon et al., 1988a). As a result, P4 therapy is initiated after breeding and often continued to mid- or late-gestation.
Although treatment is expensive at $4 per mare per day, and since current diagnostic tests
(radioimmunoassays (RIAs)) are inconclusive, owners often choose treatment. However, using RIAs could indicate problem mares that have low or decreasing total progestins preceding infectious or non-infectious abortions. If treatment is initiated and no clear rationale of treatment termination is available, current recommendations state stopping treatment at 80, 100, or 150 days, or simply continuing treatment until term (Rossdale et al., 1991). Current treatment of choice is the synthetic progestin Regu-Mate® (17x-
Allyl- 17-hythoxyestra-4,9, 11 -trien-3-one; altrenogest; Hoeschst-Roussel, Agri-Vet Co.;
Sommerville, NJ 08876). Contraindication to P4 therapy includes bacterial uterine infection, since the uterus has limited ability to clear infections under P4 influence
(Wilker et al., 1992).
In order to determine the effectiveness of ReguMate® on early pregnancy maintenance, Voller et al. (1991) worked with nine pregnant pony mares. Regu-Mate® was administered orally at 0.044 mgfkg to each mare daily starting day 4 post-ovulation.
An injection of PGF2 (3 mg, IM) was given on day 10 post-ovulation to each mare to regress the primary CL. This resulted in decreased P4 levels (<1 ng/ml), which remained at this level until secondary CL formed. Only 4 out of 9 mares maintained pregnancy regardless of treatment. Their secondary CL formed between days 32 and 85 of pregnancy, along with increasing P4 levels; the other 5 mares had resorbed fetuses despite Regu-Mate® treatment. When
PGF2a was administered to each mare at days 65 and 75 of gestation, abortion did not result and accessory CL did not regress completely
(Voller et al., 1991).
ReguMate® also did not affect accessory ovulation or accessory

luteal function when compared to non-treated pregnant controls (Parry-Weeks and
Holtan, 1987). However, repeated injections of
PGF2a
(5 mg, IM, BID) resulted in abortion despite Regu-Mate® treatment. Schideler et al. (1982) also discovered 1 ml
Regu-Mate®/50 kg was not sufficient to maintain pregnancy in OVX mares beyond about day 40 of gestation.
Contrary to previous studies, no new secondary CL were formed with Regu-
Mate® treatment, indicating this particular treatment was inhibitory until day 70 of gestation (Daels et al., 1991a).
In a study by Naden et al. (1990), Regu-Mate® administration to pregnant mares resulted in enlarged citori that had no effect on the functional reproductive performance in fillies born to treated mares. Thus, ReguMate® treatment was deemed safe to use in pregnant mares. Similarly, Ballet al. (1992) found administration of 450 mg of Regu-
Mate® to pony mares daily between 0 to 6 days after ovulation did not significantly affect early embryonic development, as compared to control mares.
In another study, Regu-Mate® given the last 20 to 30 days of gestation did not increase the length of gestation. In the same study, ReguMate® increased mammary development due to an increase in exogenous progestin levels (Chavatte et al., 1997).
Daels et al. (1991b) induced fetal loss via endotoxemia during 21 to 35 days of gestation. Mares were treated with ReguMate® daily to compensate for the loss of endogenous P4 secretion. Plasma P4 decreased 24 hours post-injection with an endotoxin, but daily treatment with ReguMate® prevented embryo loss in mares <2 months pregnant. If Regu-Mate® was discontinued at 40 days of gestation and P4 concentration was <1 ng/mll, fetal death would result 4 days later. Continuation of treatment was recommended until 70 days of pregnancy. If mares were OVX at day 30 of gestation, pregnancy was maintained with 22 to 44 mg/day of Regu-Mate® given up to
100 days of pregnancy.
A study by Jackson et al. (1986) stated exogenous progestins would alter serum concentrations of P4 in the mare. The study indicated Regu-Mate® did not alter endogenous P4 levels if 22 mg were administered daily (P0).

29
Knowles et al. (1993) performed a study with pregnant OVX mares, administering 100 mg Regu-.Mate® daily for the first 30 days of gestation. Endogenous
P4 levels were measured at 91 days of gestation in pregnant OVX mares. Progesterone
(P4) concentration was >1 ng/ml in blood, indicating placental P4 secretion at that time, as compared to intact mares whose increase in P4 concentration between 60 to 100 days was of ovarian origin.
Ganjam and Kenney (1975) performed a study on OVX mares and gave them 150 mg (IM, once daily) of exogenous P4. They measured P4 plasma levels and found it took
21 days to increase P4 levels in plasma to 6 to 7 ng/ml, and when the dose was doubled, it took 14 days to increase it to that level if they were treated daily. Since it is common practice to put a mare on Regu-Mate® at 200 to 250 mg every 10 to 30 days to prevent abortion, the study showed this was an inadequate measure, since it took 14 days at 300 mg administered daily to obtain desired plasma P4 levels. It was recommended administering 500 mg every other day starting at day 40 of gestation, and continuing this until a week prior to parturition.
Other uses for ReguMate® were to allow the extension of the donor-recipient ovulation synchrony required for successful non-surgical embryo transfer in mares
(Parry-Weeks and Holtan, 1987), to control estrous cycling in mares during the late transitional phase, and to do the same in normally cycling mares (Squires et al., 1983).
The latter is an effective method to reduce erratic sexual behavior typical of the transition period. Only mares with considerable follicular activity (several 20 to 30 mm follicles) should be selected for exogenous progestin treatment (Squires et al., 1984).
Other preparations administered to problem mares include melengestrol acetate, proligestone, chlormadinone acetate, hydroxy-progesterone caproate, and medroxyprogesterone acetate, which are all synthetic progestins. There are anecdotal reports of their use for pregnancy maintenance, but experimental evidence of effectiveness is poor or lacking (Wilker et al., 1992). For example, hydroxyprogesterone caproate is widely used in Australia to prevent early pregnancy loss, but is ineffective due to a lack of binding affinity for equine P4 receptors in the endometrium (Allen, 1993).
Instead, ReguMate® is the treatment of choice in most instances. Despite Regu-Mate' s®

popularity and effectiveness, Allen (1982) found it tended to hasten the demise of existing CL, and did not increase fetal survival rates.
30
Holtan et al. (1979) noted humans, horses, and sheep OVX at 50 days of gestation did not abort, since the placenta produced adequate P4 to maintain pregnancy. This could be due to the fact that the myometrium continues to be bathed locally by P4 and its metabolites, because the placenta synthesizes these at that time. They also found if some mares were OVX before 50 to 70 days of gestation, they would abort. However, if OVX between 140 to 210 days of gestation, fetuses were carried to term, implicating ovaries were not needed at this stage to maintain pregnancy.
Endogenous P4 in OVX pregnant mares was >1 ng/ml by 91 days of gestation, indicating placental secretion of P4. Serum concentrations of progestins in ovary-intact pregnant mares were greater than those in OVX mares until 130 days of gestation, and from then on they were the same. In conclusion, ovaries were a significant source of progestins until 130 days of gestation, and the feto-placental unit was the source of progestin production from 100 days of gestation (Knowles et al., 1994).
Regarding estrogen levels in OVX mares, Daels et al. (1990) found mean conjugated estrogen (CE) concentrations increased in a constant linear manner from 17 ng/mg creatinine (Cr) on day 20 of gestation to 291 ng/mg Cr on day 70 of pregnancy, as compared to intact mares, which had a mean concentration of 201 to 172 ng/mg Cr between 20 to 33 days of gestation respectively, and reached maximal levels at 1066 ng/mg Cr on day 39 of pregnancy. There was a decrease from that point to 637 ng/mg Cr at 44 days of gestation. There was a linear increase to 1191 ng/ml on day 70 of gestation.
These changes were due to the rate of ovarian synthesis of estrogens.
In a study by Squires et al. (1974), P4 concentration was measured in plasma in 17 pregnant, 6 non-pregnant (used as controls), and 12 HX mares from 7 to 19 days of gestation and following HX, and 32 to 140 days of pregnancy and following HX.
No difference was found between pregnant and HX mares on days 7 to 19 of pregnancy or following HX. In mares that were HX at >19 days, there was a linear decrease in P4

levels. Progesterone (P4) concentrations increased between 32 to 44 days in pregnant
31 mares and continued to increase from >44 days to 90 days of gestation, then decreased from 150 to 180 days of pregnancy. However, there were differences between mares and
P4 concentrations. Ovulation was detected in 2 out of 12 HX mares 140 days following
HX. It occurred after the primary CL regressed based on a decrease in P4 to 1.6 ng/ml before ovulation, and with an increase to 8 ng/ml post-ovulation. This study also noted
CL regression occurred earlier in HX mares than in pregnant mares. Secondary CL failed to form from anovulatory follicles in FIX mares. The conclusion in this study was that eCG contributed to the maintenance, increased steroidogenesis, and formation of secondary CL, but was not essential for the growth of follicles during early pregnancy in mares.
FIX mares had an interrupted pattern of plasma progestin production due to prolonged CL tissue lifespan. In non-pregnant mares, HX prolonged the lifespan of CL, since there was no uterus to produce PGF2a. The uterus is important for the initiation of regression of CL in normal, non-pregnant mares, but all HX mares had follicular activity despite high levels of circulating progestins (Stabenfeldt et al., 1974).

32
Objectives
Previous studies by Riad and Holtan (1993) have shown urinary hormones PdG and E1S in mares increased abruptly around
85 to 90 days of gestation. It could be concluded the feto-placental unit was functional at that time; therefore one objective of this study was to describe an accurate profile of total and individual progestins found in plasma and fecal samples taken every other day from 64 to
150 days of gestation.
Another goal was to correlate individual plasma to fecal progestins over time. The final objective was to determine an accurate time frame, using spline regression, of abrupt changes in plasma and fecal progestins, similar to findings for the urinary hormones.
This data may help define when the feto-placental unit is adequately functional to maintain pregnancy.
Animals (n = 4 pregnant mares) were housed at the Oregon State University (OSU)
Horse Center. Mares were two Shetland ponies and two Quarter Horses of various ages with no known reproductive pathology. Pony mares were bred artificially (A.I.) to the same Shetland stallion, and horse mares were bred A.I. to the same Quarter Horse stallion between June and July 1999, with the day of breeding being day 0 of pregnancy. Three mares were kept on grass pasture, while the fourth mare was kept in an outside run due to a foot injury. All subjects had water and trace minerals available at all times.
Sample Collection
Plasma and feces were collected from pregnant mares (n = 4) every other day from day 64 to 150 of gestation. Blood was collected by jugular venipuncture into

heparinized tubes (Vacutainers, ® Beckton & Dickinson; Rutherford, New Jersey
07070). Plasma was separated after centrifugation and stored at -18°C. Fecal samples
33 were stored at -18°C.
Ultrasound procedures were conducted as instructed by Ginther and Pierson
(1984a, 1984b), McKinnon et al. (1988b), and Squires et al. (1988). An ultrasound machine, model ALOCA 210 DX, was used along with a 5 MHz probe (Corometrics
Medical Systems Inc.; Wallingford, CT). Follicles, CL, and CH were counted and their diameters were measured with built-in calipers.
Deuterium (D4) labeled progestins (500 ng each) of D4-5a-DHP, D4-P5,
Deuterium labeled progestins were prepared as described by Dehennin et al. (1980).
Extraction methods followed established methods by Holtan et al. (1991) and Schutzer
(1995).
Standard curves were generated with ChemStation software (HP, version
G1034B) by adding 500 ng of each D4 standard to each of the five vials containing a range from 125 to 2000 ng of non-labeled standard in ovariectomized (OVX) mare plasma or gelding feces. Non-labeled standards were 5a-DHP, 513-DHP, P4, 3a-5a, 33-
5a, 2Oct-5x, P5-13f, 20f3-5a, c4E-diol, aa-diol, -diol, and 3a-diol. For each, a standard curve of the response ratios was developed using the D4 standard most similar in structure. The assumption is that similar hormones under the same extraction and derivatization procedures will be recovered in similar proportions. Quantitation consisted of calculating least squares linear regression of standards versus their D4-ISTD. Thus, least squares linear regression for each steroid within each assay would be calculated from the peak area ratios (unlabeled standards versus D4-ISTD).

Samples of 5.0 ml of plasma or 0.5 g of feces were diluted with deionized water (1:l,v:v). Samples were then passed through
C18
Sep-Pak® cartridges (PrepSep-
34
C18; Fisher Scientific; Fair Lawn, NJ 07410) using a vacuum system (Waters Company;
Milford, MA). Prior to sample washing,
C'8
Sep-Pak® carthdges were charged with methanol (MeOH) (HPLC grade; 2 ml) and rinsed with deionized water (2 ml). After the entire sample passed through the cartridges, it was rinsed again with deionized water (5 ml), followed by hexane (HPLC grade; 1 ml). Unconjugated steroids were extracted from the cartridge bed with anhydrous diethyl ether (4 ml) into 4-dram vials. The ether was evaporated under a stream of air at 60°C.
Ketones were derivatized to methoximes with 50 p.1 of 5% methoxyamine-HC1
(MOX; Sigma Chemical Company; St. Louis, MO) in silation grade pyridine (Regis
Technologies Inc.; Morton Grove, IL) at 60°C for 30 mm. in 4-dram vials with Teflon coated caps. Pyridine was subsequently evaporated under a stream of air at 80°C.
Samples were then incubated for 45 mm. at 80°C with 25 p.1 of N-methyl-N-(tbutyldimethylsilyl) trifluoroacetamide (MtBSTFA; Regis) and 50 p.1 of 5% diethylamine
(DEA; Sigma) in silation grade dimethylformamide (DMF; Regis) as the catalyst.
Derivatization of unhindered-hydroxyl groups to tert-butyldimethylsilyl (tBDMS) occurred at this step, and was stopped by adding 0.5 ml of deionized water to each sample.
Derivatives were extracted from the aqueous phase with anhydrous diethyl ether
(3 ml; 2X), and then concentrated under a stream of air. Samples were taken up with nundecane (Sigma; 30 p.1) and analyzed with GCIMS.
Sample Analysis
Laboratory facilities and equipment for assays, including a Hewlett Packard gas chromatograph (HP 5890) with autosampler (HP 7673) system interfaced with a mass spectrometer (HP 5971A) and 486/66 series microcomputer (Hewlett Packard;
Wilmington, DE; ChemStation Software, HP, version G1034B), were available in
Withycombe Hall 317 laboratory.

Each (1 tl) sample was injected (splitless mode) into the GC, which contained
35 a DB-5MS 30 mX 0.25 mm id fused silica capillary column (J & W Scientific; Folsom,
CA). Samples were then separated under the temperature program mode. Injection port temperature was isothermal at 250°C, while initial temperature of the GC oven was at
160°C. Upon injection, the GC oven temperature increased 25°C/mm. to 280°C, then
3°C/mm. to 3 10°C. The GC oven remained at 3 10°C for 10 mm.; total run time was 27.5
mm. The MS was operated in full scan mode for initial identification and selected ion mode (SIM) for routine quantitation.
Statistical Analysis
Eight different progestins were routinely identified and quantified in plasma and
fa-diol, and 4-diol in feces, while c43-diol and 20-5u were left out in plasma due to low sensitivity detection. Statistical analysis consisted of using analysis of variance
(ANOVA) with repeated measures split-plot design (SAS, version 8.10; SAS Institute
Inc.; Cary, NC). Since variances were not homogeneous, least squares means (LSM) were log-transformed. Main effects were mare, sample (plasma or fecal), and day (time in gestation), and the interactions were sample by day and mare by sample.
Pearson correlation method (SAS) was run on all progestins, except for 20-5a and c4-diol, to determine if there was a correlation between individual plasma and fecal progestins over time. A Pearson partial correlation method (SAS) was done on all progestins, except for 203-5x and a3-diol, to determine if there was a correlation between individual plasma and fecal progestins without the time effect.
To further study interactions between sample and day, a spline regression (Gauss-
Newton, SAS) method was used to determine a point of rapid increase of slopes in plasma and fecal progestins. Ratios of feces to plasma were also plotted to demonstrate interactions obtained from SAS results. This would indicate whether or not plasma or feces would increase at a different rate over time.

36
CHAPTER 4: RESULTS
The data examines progestins from 64 to 150 days of gestation in the mare. The data compared a variety of progestins and their concentrations present in plasma and feces in both pony and light horse mares. Sensitivity of the method used was about 0.5
ng/ml in plasma or 0.5 ng/g in feces. Mean coefficient of variation (CV) among assays was 27.1%, and within assays, CV was 12.4% as reported by Miller et al. (2000). Least squares means (LSM) ± standard error (SE) were used throughout (ANOVA general linear model for repeated measures, SAS) unless noted otherwise. Eight different progestins in plasma (5a-DHP, P4, 3a-5cz, 3f3-5a, 20a-5u, aa-diol, 33-diol, and a-
3-diol, and 13a-diol) were analyzed. Other progestins found, such as P5, P5-, and
51-
DHP, were not included in the analysis because they were only sporadically detected.
Plasma 20-5a and c43-diol were often below normal detection, thus they were disregarded in the analysis.
No statistical analysis or conclusive evidence correlating progestin levels and CL formation were performed, because it was difficult to identify new CL formation from the old CL formed. The ultrasound data collected was not reliable due to lack of repeatability of results and operator inexperience.
Table 1 shows the overall mean progestin concentrations from 64 to 150 days of gestation in mares (n = 4). Concentrations in feces were approximately 100 times higher than in plasma for total progestins, yet differed among individual progestins (Table 1 and
Figure 1). Progesterone (P4) concentration remained similar in both plasma and feces over time. Progestin 20a-5c and 5a-DHP concentrations were about 6 times higher in feces than in plasma. Progestin aa-diol and 33-5a concentrations were approximately
50 to 60 times higher in feces than in plasma. Progestin 3a-5a concentration was about
150 times higher in feces than in plasma. Progestin 33-diol concentration was

37 approximately 800 times higher in feces than in plasma, while kx-diol concentration was 100 times higher in feces than in plasma (Table 1).
Four progestins were chosen to depict different pauerns over time; these included
3(-5a, 2Oct-5a, 34-diol, and a-diol. Progestin 20x-5cx remained in highest concentration in plasma over time, while 13-diol was highest in feces (Table 1).
Because of heterogeneity of variance, and the great difference between plasma and feces, data were log-transformed to eliminate skewness. Figure 1 shows plasma and fecal LSM increase over time at a steady, linear rate. The slopes of the lines are the same for total progestins.
Table 1: Mean concentration of progestins in plasma (ng/ml) and feces (ng/g) of mares
(n 4) sampled from 64 to 150 days of gestation.
Progestina
5a-DHP
P4
3a-5a
20a-5ct
13a-diol
20-5a a-dio1
Total
Plasma LSMb
13.2
5.1
4.0
9.8
26.2
±
SEC
0.4
0.3
0.1
0.2
0.4
Fecal LSM
± SE
78.9
3.2
8.1
0.4
584.3
19.3
582.1
144.1
29.1
8.1
-diol
13.1
6.3
16.6
NAd
NA
95.0
0.5
0.2
0.3
NA
NA
1.6
602.3
5159.4
1635.9
217.0
490.4
9501.4
b a See List of Abbreviations for full systematic names
Least squares means
C d
Pooled standard error of means
Not analyzed
34.4
153.2
68.9
12.8
18.5
242.1
Table 2 depicts the analysis of variance for repeated measures for total progestins.
Mare effect was significant indicating a difference between individual animals. Sample
(plasma versus feces) effect was significant showing there was a difference between

plasma and fecal concentrations. Day was significant demonstrating a general increase over time, as observed in Figure 1.
Table 3 is a summary of individual progestin analyses performed in a similar manner as in Table 2. Note that day effect was not significant for P4 because there was a constant rate of change over time, unlike the other progestins. Sample effect was also not significant for P4 since there was no difference between plasma and fecal concentrations.
3.5
3
-*- Feces
-4-Plasma
Linear (Feces)
2.5
0
1.5
1
03
0
60 70
9816anO.O1O9x.731
80 90 100
Days
110 120 130 140 150
Figure 1: Mean concentration and pooled standard error (SE) of total progestins
(log) in plasma (bottom line, SE = 0.057) and feces (top line, SE = 0.069) from 64 to 150 days of gestation in mares (n = 4).

Table 2: ANOVA table for the log concentration of total progestins in plasma and feces from 64 to 150 days of gestation in mares (n = 4).
Source
Mare
Sampleb
DF
3
1
3
Type ifi SS MS F Value Pr>F
7.631
302.462
0.069
2.544
302.462
0.023
158.150
13213.800
Sa
5
1.420
NS Mare x
Sample
Day
Sample x
Day
Error
Total
37
37
28.448
0.3 19
0.769
0.009
222
303
3.571
342.499
0.016
a b
Significant (P0.05) or not (NS)
Mare x Sample used as error term
47.800
S
0.540
NS
39
Table 3: Summary table of individual plasma and fecal progestins as compared to total progestins using ANOVA analyses.
Source Total
P4 3cx-5a
Progestins DHP
Mare
Sample
Sample
Day
Sample x Day sa
S
NS
S
NS s
S
S s
NS
S
S NS
NS S a Significant (P0.05) or not (NS) s
S
S
S
S
3-5a
2Oct-5a aa-
4adiol diol diol
S
S
S
S
S
S
S
S
S
S
S
S
S
S
S
S
S
S
S
S
NS
S S
NS NS
There were positive correlations in the full Pearson analysis (r = >0.8290, P<0.01) for all progestins analyzed, except for P4, which was negatively correlated (r = -0.6993,
P<0.01) as shown by Figure 2. There were positive correlations in the partial Pearson analysis (r = <0.3745) for all progestins analyzed, except for P4, which was negatively correlated (r = -0.1041), but these low correlations were not significant (P>0.05).

I
' Os
1.2
1.4
10.6
0.4
0.2
0
60 70 80 90 100
Days
110 120 130 140 150
Figure 2: Mean log concentration and pooled standard error (SE) in plasma
(decreasing, SE 0.103) and fecal (increasing, SE = 0.113) progesterone (P4) from 64 to
150 days of gestation in mares (n = 4).
IiJ
Spline regression was used to determine the point of change between slopes in plasma and fecal progestins, and to portray an interaction between plasma or feces and
indicated a change at 112.7 days in plasma (Figure 3) and at 109 days in feces (Figure 4).
There was considerable variation between individual progestins and between plasma versus feces regarding when the increase in concentration occurred. Spline regression indicated both mean plasma and fecal 3-5a increased at 110 days of gestation (Figures 3 and 4 respectively). Visually, mean plasma 3-5a appeared to increase dramatically around 110 days of pregnancy, while its fecal counterpart increased around 90 days of gestation. Mean log-transformed plasma 3f-5cx appeared to increase visually around 104 days of gestation, and fecal values seemed to rise around 94 days of pregnancy.

41
80
70
60 so
20
10
0
60 70 80 90 100
Days
110 120 130 140 150
Figure 3: Mean concentration and pooled standard errors (SE) of plasma
3-hythoxy-5a-pregnan-20-one (3 j3-5cz, SE = 179.753), 20a-hydroxy-5a-pregnan-3-one
(20a-5a, SE = 49.9 19), 5a-pregnane-33,203-diol (343-dio1, SE 944.158), and
5x-pregnane-3j3,20a-diol (13a-diol, SE = 424.952) from 64 to 150 days of gestation in mares (n = 4). Spline regression (SR) indicates mean day (112.7) of increase for all progestins listed.
Spline regression indicated plasma 20a-5a increased at 114 days of gestation, and fecal increased at 108 days of pregnancy (Figures 3 and 4 respectively). Visually, mean plasma 20a-5a seemed to increase rapidly around 85 days of gestation, while its fecal counterpart increased around 94 days of pregnancy. Mean log-transformed plasma 20a-
5a appeared to increase visually around 85 days of gestation, and fecal values seemed to rise around 87 days of pregnancy.

42
12O
8000
4000
2000
0
60 70 80 90 100
Days
110 120 130 140 150
Figure 4: Mean concentration and pooled standard errors (SE) of fecal
3-hydroxy-5a-pregnan-20-one (313-5a, SE = 1.405), 20a-hydroxy-5a-pregnan-3-one
(2Ocx-5a, SE = 2.655), 5a-pregnane-33,20J3-diol (3f3-diol, SE = 1.463), and
5a-pregnane-33,20cx-diol (13u-diol, SE = 2.124) from 64 to 150 days of gestation in mares (n = 4). Spline regression (SR) indicates mean day (109) of increase for 3-5u and
20x-5a only; -diol and a-diol did not show a defined change.
Spline regression indicated the point of change in plasma 43-dio1 increased at
114.7 days of pregnancy, while a decided change in fecal I-diol could not be detected
(Figures 3 and 4 respectively). Visually, mean plasma 43-dio1 appeared to increase dramatically around 110 days of gestation, while its fecal counterpart increased around
104 days of pregnancy. Mean log-transformed plasma f3-diol increased visually around
75 and 115 days of gestation, and fecal values seemed to rise around 87 days of pregnancy.

Spline regression showed an increase in plasma a-diol around 112.2 days of pregnancy, but could not detect the change point in fecal ikt-diol (Figures 3 and 4 respectively). Visually, mean plasma 13a-diol appeared to increase rapidly around 75 days of gestation, while its fecal counterpart increased around 85 days of pregnancy.
Mean log-transformed plasma fa-diol increased visually around 75 days of gestation, and fecal values rose at 87 days of pregnancy.
Visually, mean plasma and fecal aa-diol appeared to increase around 90 days of pregnancy. Mean plasma and fecal 3a-5a increased around 85 days of gestation. Mean fecal c4-diol increased around 85 days of pregnancy. Mean fecal 203-5a increased around 82 days of gestation. No noticeable change was seen in plasma 5a-DHP, while its log-transformed value seemed to increase around 100 days of gestation. Mean fecal
5a-DHP increased around 96 days of gestation, while its log counterpart increased around 115 days of pregnancy.
Ratios of feces to plasma in 3-5a, 20a-5a, and 3a-dio1 were used to visually demonstrate the interactions between sample and day (Figure 5). There was a positive slope indicating an interaction between sample and day for 3-5a (Figure 5A); P4 and
43
3a-5a were similar. This showed a greater increase in feces over time than in plasma. A negative slope was observed for 20u-5a (Figure SB); aa-diol was similar. A negative slope indicated plasma was increasing at a faster rate than in feces. Progestin f3a-diol demonstrated a straight line when there was no interaction between sample and day
(Figure SC), meaning plasma and feces increased at the same rate over time; 5a-DHP and
-diol were similar.
Most differences among mares were seen in 3f-Sa, 20a-5a,
regarding a notable increase over time. As depicted by Figure 6, there was significant variation among different progestins, mares, and plasma versus feces regarding the point

I
I
60
B
70 SO
2O-5
Linr (2O.-5.)
90 100 slope = -
110 120 130 140 150
C)
0
60
2
2
2 c
70
2
1
1
SO
1u-dioI
90 100 i
110 120 130 140 150
1
60 70 80 90 100
Days
110 120 130 140 150
Figure 5: Mean ratio of feces to plasma (log) in 3-hydroxy-5a-pregnan-2O-one
(A, 313-5a), 20a-hydroxy-5cx-pregnan-3-one (B, 20u-5a), and 5a-pregnane-33,2Oa-dio1
(C, f3a-diol) from 64 to 150 days of gestation in mares (n = 4).

1500
1000
15
10
5
0
60
2500
30
25
20
70 80 90 100 110 120 130 140 150
45
2000
0
60 70 80 90 100
Days
110 120 130 140 150
Figure 6: Plasma (A) and fecal (B) 33-hydroxy-5a-pregnan-2O-one (33-5a) in individual mares from 64 to 150 days of gestation.

of increase, which was difficult to determine at times. Table 4 depicts the ranges of change points in time for spline regression for the four progestins, indicating the tremendous variability among individual mares. Similarly, log ratios showed this variability among individual mares, and did not increase the reliability in determining the change points for spline regression.
Table 4: Spline regression ranges for plasma and fecal 3-hydroxy-5a-pregnan-2O-one
(3f3-5a), 20a-hydroxy-5a-pregnan-3-one (20a-5a), 5a-pregnane-3f3,2O3-diol
(-diol), and 5u-pregnane-3f3,2Occ-diol (f3a-diol) in individual mares (n =4) from 64 to 150 days of gestation.
Progestin
3-5a
20a-5cx
3-dioI u-diol
Plasma (days)
80-115
109.1-119
78.45-129.4
95.49-118.4
Fecal (days)
110-117
108-109.2
None
None

47
CHAPTER 5: DISCUSSION
Overview
Aims of this research were to analyze samples taken during 64 to 150 days of gestation to establish data and comparisons between and among plasma and fecal progestins; to determine a time frame, using spline regression, when an increase would likely reflect a functional feto-placental unit; and to compare these findings to the trend seen with urinary hormones. The method (GC/MS) accurately separated progestins with similar structures and molecular make-up.
Progestin production in the mare is critical for the maintenance of pregnancy as well as in other species (Brown et aL, 1994; Miller and Holtan, 1997). Progesterone (P4) plays an essential role in placentation, embryo development, and placental functions. Profiling progestins, such as P4, in mid-gestation could be indicative of fetal well being and a successful pregnancy to term. Determining the time certain progestins increase rapidly may reflect placental progestin production. Finally, this study may provide some insight on the necessity of treating problem mares with Regu-Mate® until term, particularly regarding stopping treatment at a determined time.
Holtan et al. (1991) found sensitivity, range, accuracy, and precision of GC/MS to be at least equal to most immunoassays, including the added benefit of specificity. The
GC/MS allowed progestins to be accurately separated without cross-reactivity, such as P4 in both plasma and feces, than with RIAs. The GC/MS identified 13 total progestins, 8 of these in plasma and 10 in feces. All plasma progestins identified in this study had been previously identified by Holtan et al. (1991) using similar GCIMS techniques. Even with rigorous preparation, purification, and sensitivity of 0.5 ng/ml for plasma or 0.5 ng/g for feces before analysis, concentrations of P5, P5-1, and 5-DHP in both plasma and feces, and 203-5a and a-diol in plasma were often low or undetectable at times.

In the mare, the major source of P4 changes gradually from the ovaries to the placenta between days 50 to 70, reaches maximal levels around day 100 of gestation
(Vivrette, 1994), and decreases from day 150 to 180 (Holtan et al., 1975). However, no data has shown the placenta to produce P4 prior to 50 days of gestation (Shideler et al.,
1982). We detected P4 in both plasma and feces as early as 64 days of gestation, which could be of placental and/or ovarian origin, but at this point in research, it is difficult to distinguish the source, even though Holtan et al. (1991) found the presence of reduced pregnanes in plasma after 50 days of gestation to perhaps indicate placental production.
Furthermore, Burns and Fleeger (1975) stated the increase in progestin concentration between 30 to 60 days of pregnancy was of ovarian origin. On the other hand, Vivrette (1994) noted an increase in reduced pregnanes in plasma, such as 20u-5a and a-diol, between days 30 to 60 of gestation was due to production by the fetoplacental unit. These two progestins were found in high concentrations in plasma and feces respectively in our study as well. Perhaps these two could be used as indicators to detect successful development of the placenta in early gestation.
According to a study by Holtan et al. (1991), the predominant progestins in mare plasma were 20a-5u and u-diol; concentrations were around 25 ng/ml at 100 days of gestation in both, and at approximately 75 ng/ml and 50 ng/ml at 150 days of pregnancy respectively. Our findings showed mean 20a-5a and cx-diol were also found in highest concentration in plasma, but at half the concentration at 100 days of gestation and at similar levels at 150 days of pregnancy. Our results were also consistent with Burns and
Fleeger (1975) who found 5a-DHP increased around 50 to 70 days of gestation, while P4 decreased at that time. Progesterone (P4) decreased in plasma around 64 days of gestation and increased in feces (Figure 2), indicating luteal regression and feto-placental production respectively as reported by Short (1959) and Holtan et al. (1975).
Holtan et al. (1991) found plasma 3ct-5ct increased at the same rate as 3-5a in late pregnancy, as was the case from 64 to 150 days of gestation in our study. Plasma 3-
5a was almost twice as high in concentration as 3a-5a between 64 to 150 days of gestation, yet they remained parallel to each other. The same can be stated about aa-diol and f-dioI, which were almost equal in concentration in late pregnancy (Holtan et al.,

49
1991), but aa-diol was almost twice the concentration as -diol from 64 to 150 days of gestation. The data is supported by previous work done in this lab (Baumholtz, 1998).
ANOVA results in Table 2 for the log of total progestins showed all main effects were significant. Variation among individual mares, plasma versus feces, and time was expected, since this has been reported by Hamon et al. (1991). Despite mare effect being significant, this was not a major factor in our study, because it is already expected that there will be variation among individual mares (Hamon et al., 1991; Baumholtz, 1998).
Averaging tended to smooth out trends, thus changes in individual mares were overlooked.
Fecal progestins were 100 fold greater in concentration than plasma progestins
(Table 1), similarly to previous studies by Miller et al. (2000). Knowles et al. (1994) supported the idea of total progestins increasing from 100 to 160 days of gestation.
Interestingly, the range of concentrations between progestins in plasma versus feces, as depicted by Table 1, was anywhere from about 6 times the concentration of 20a-5ct in feces than in plasma, to approximately 800 times the concentration in fecal -diol than in plasma. According to Figure 1, log of total plasma and fecal progestins increased at a relatively similar rate, indicating either plasma or feces could be analyzed to obtain similar results, particularly in research with wild equids. Fecal samples would be ideal in these studies, since sample collection is safe, easy, and less stressful on the animal than blood collection. According to Miller and Holtan (1997), much higher concentrations were detected and a greater number of different progestins were present in feces of ewes than in the peripheral circulation, thus it would be ideal to utilize feces instead of plasma.
This trend would be applicable to certain progestins like 20a-5a, so further assays, such as developed by Squires et al. (1986), can be manufactured to measure these in the field to determine if placentation was successful.
According to Holtan et al. (1991), the differential in progestin quality and quantity throughout pregnancy in equids is most likely represented by the difference between

sources, increased 20a- and 2013-hydroxylation in the liver and gastrointestinal system, different metabolic pathways in the feto-placental unit, and variability among progestins and individual mares, as described by Hamon et al. (1991) and Chavatte et al. (1997).
50
Miller and Holtan (1997) found similar results stating the ewe placenta had a complex metabolism as well. The most likely explanation for this phenomenon, as discussed by
Adlercreutz et al. (1979) in humans, is that intestinal microflora and mucosa alter the activity, quantity, and excretion of progestins throughout gestation. However, more research needs to be conducted in the mare to determine the exact mechanism.
Interactions of mare by sample and sample by day were not significant for total progestins because over time and in individual mares, the trend for both plasma and feces was to increase at a constant rate over time. This was not the case when individual progestins were analyzed, as reported in Table 3. For example, compared to total progestins, all progestins were significant regarding the mare by sample interaction. This was so because plasma and fecal progestin concentrations differed among individual mares.
Like the total progestins, 5a-DHP did not have a significant sample by day interaction, since over time, plasma and fecal concentrations changed at a constant rate.
Progesterone (P4) was not significant regarding sample or day, because there was not much difference in concentration between plasma and feces, and there was a constant rate of change over time (Figure 2). Progestins P4, 33-5a, 3cx-5a, and 20a-5a were significant in sample by day interaction, unlike the total progestins, because over time there was a difference between plasma and fecal concentrations. These progestins showed this relationship well in plots of log ratios (Figure 5).
Even though progestins were highly positively correlated in relation to the time factor (full Pearson correlation) between plasma and feces, as described by Miller et al.
(2000) in ewes, they still changed at different rates over time. The only negative correlation found was between plasma and fecal P4. This meant plasma P4 was decreasing in concentration, while fecal P4 was increasing over time (Figure 2). The reason plasma P4 decreased over time was due to decreased ovarian and CL function, as described by Squires et al. (1974) and Ganjam and Kenney (1975). Other possibilities

include differences in solubility in plasma or feces, sample stability, physiological effects on metabolic routes, and bacterial action (Adlercreutz et al., 1979).
A partial Pearson correlation indicated none of the individual plasma and fecal progestins were related to each other when the time factor was taken out, since daily variation was such that no clear correlation was possible.
When log ratios were examined, compared to the increase observed in total progestins over time, individual progestins showed three different rate patterns-increase, decrease, or no change over time-caused by differences in steroidogenesis and metabolism, as described by Holtan et al. (1991). This is important when choosing specific progestin assays for diagnosing functional placentation, and whether to collect plasma or feces depending on the rate of change of that particular progestin located in that source. For example, 20a-5a would be ideal to analyze in plasma, since its
51 concentration in plasma increased much faster than in feces over time resulting in a negative ratio (Figure SB).
Preliminary data by Miller and Holtan (1997) indicated fecal progestins increased abruptly around day 80 of gestation, similarly to the study of urinary hormones increasing visually around 90 days of pregnancy in mares. Marked increase was noted in E1S at 80 days of gestation, reaching maximal levels of 143.3 pg/mg Cr on day 142 of pregnancy.
Its maximum concentration was reached between 140 to 170 days of gestation, and then decreased until parturition (Riad and Holtan, 1993). This study showed most progestins were sharply increasing beyond 120 days of pregnancy due to the time factor and increased fetal size, except for plasma P4. On the other hand, PdG remained stable until
80 days of gestation. Its profile paralleled blood values, as noted by Riad and Holtan
(1993). They determined the different patterns produced by the urinary hormones were caused by the differences between source and metabolism in the feto-placental unit. The sharp increase in PdG and El S thus reflected a mature, functional feto-placental unit.
This would probably be the case in our study. Observing a similar trend of increase in progestins, such as 33-5a, 2Oct-5a, and ac-diol in both plasma and feces, would indicate a mature placenta capable of maintaining pregnancy. Day of changeover from ovarian to
a-diol,

52 varied significantly from the 90-day change point in the urinary hormones, PdG and
E1S (Riad and Holtan, 1993). The point of changeover for both plasma and fecal progestins on average seemed to be around 111 days of gestation.
Spline regression indicated the mean day of increase in plasma for the four
-diol, and a-diol, was 112.7, and day 109 in feces.
Plasma fEkx-diol and fecal 20a-5a matched the mean values the closest, although the other two progestins were within this limit. Some spline regression values were unobtainable, because this procedure only works on linear equations when there are different slopes along the same lines. It is not effective on constant or curvilinear data, such as for fecal
3-diol and Ia-diol, which did not have a distinct change in the rate of increase, as depicted by the plotted ratio (Figure 5C). Overall range for the mean day of increase for the four progestins was from 110 to 115 days of pregnancy in plasma, while in feces the range was from 108 to 110 days of gestation. This wide range was due to individual progestin and mare variability, as well as differences in the metabolism of progestins, as described by Ganjam et al. (1975), Adlercreutz et al. (1979), and Hamon et al. (1991).
The possible reason spline regression indicated a change point around 111 days of gestation is because Allen (1982) and Steven (1982) stated that by day 100 of gestation, primary viffi develop secondary branches marking a complex feto-maternal interdigitation both at the macro- and microvillous level over the entire endometrial surface. Anatomically, this would mean the placenta is fully functional, coinciding with the time of increase and differentiation of progestins in this study due to increased vascularization and interdigitation between fetal and maternal endocrine tissues.
When considering this project, an afterthought was to review studies dealing with the question when is the mare fully capable of carrying a foal to term without ovarian or
Regu-Mate® support. Ginther (1985a) stated abortion was less likely to occur after 60 to
75 days of pregnancy. If treatment is initiated without cause and no clear rationale for treatment termination is available, this practice of supplemental progestin therapy is expensive at best and most likely unnecessary at worst. Recent work suggests initiating and continuing treatment until term (Rossdale et al., 1991). Knowles et al. (1994) later suggested 100 days of gestation was the most appropriate time to stop treatment, as

demonstrated by their previous studies (1993). Holtan et al. (1979) noted that if mares
53 were OVX between 140 to 210 days of gestation, fetuses were carried to term, implicating ovaries were not needed at this stage to maintain pregnancy. Despite Regu-
Mate's® popularity and effectiveness, Allen (1982) found it tended to hasten the demise of existing CL, and did not increase fetal survival rates. Although this study did not perform similar studies, our data suggest stopping treatment around 110 to 120 days of gestation, as reported by Discafani et al. (1995).
There was quite a lot of variation, not only between mean progestin concentrations and source, but also between individual mares, as reported by Hamon et al. (1991). It was often difficult to determine points of change in individual mares and progestins, as depicted by Figure 6. Individual mare variability among values could not be compared to urinary data because only mean values were known.
Future research, as summarized by Baumholtz (1998), needs to be conducted to understand the unique hormonal profile of the pregnant mare, as well as to understand source and biological activity of these different progestins. Increased knowledge will help in practical applications, such as developing pregnancy or pregnancy problem diagnostic tests, and developing hormonal treatment therapies for at-risk pregnancies as in the study by Brown et al. (1994). Increased knowledge in this field could also have effects on research in related fields or other species. Next step in this research would be to add miniature horses and draft mares, since Hamon et al. (1991) noted there was significant variation among different types and breeds of mares. Additionally, other equids such as donkeys, mules, and zebras could be studied. This is a relatively new area of study in equine reproduction and opens up many options for research. Comparative

54 studies between different species can be conducted to determine which progestins function best in order to develop specific assays that detect successful placentation.
For further progestin profile studies, it would be necessary to do a 24-hour progestin profile collecting plasma and fecal samples every half hour in several mares. In this way, we would be able to determine the extent of progestin level fluctuation during the day. This may explain why some progestins varied tremendously regarding concentration on a daily basis.
GCIMS is not practical for fieldwork or routine sampling, but is more suited toward research. Feces need to be extensively analyzed using GC/MS to determine all the progestins present throughout the entire gestation in the mare. The study of fecal progestins alone is far from clarified. Basic fecal progestin research with reference to diet, anestrus, diestrus, estrus, transitional mares, levels during the entire gestation, and other hormonal influences needs to be addressed before "mare side" tests of fecal progestins will be of value. The other question that begs to be addressed is which of the progestins identified in this study are mimicked or supplemented by the administration of
ReguMate.® Only then can specific assays selective for specific progestins be developed for routine sampling. Knowledge of which progestins change and specifically when these changes occur during gestation would tremendously aid in the development and interpretation of such immunoassays.

55
Adlercreutz, H., F. Martin, P. Jarvenpaa, and T. Fotsis. 1979. Steroid absorption and enterohepatic recycling. Contraception. 20(3):20 1-223.
Ainsworth, L. 1972. The cleavage of steroid sulfates by sheep and pig fetal liver, fetal kidney and placental preparations in vitro. Steroids. 19(6):741-750.
Ainsworth, L., and K.J Ryan. 1970. Steroid hormone transformations by endocrine organs from pregnant mammals VI. The conversion of4-androstene-3,17-dione to estrogens by goat placental preparations in vitro. Steroids. 16(5):553-559.
Albrecht, B.A., J.N. MacLeod, and P.F Daels. 1997. Differential transcription of steroidogenic enzymes in the equine primary corpus luteum during diestrus and early pregnancy. Biol. Reprod. 56(4):821-829.
Allen, W.R. 1975. The influence of fetal genotype upon endometrial cup development and PMSG and progestagen production in equids. J. Reprod. Fertil., Suppi.
23:405-413.
Allen, W.R. 1980. Hormonal control of early pregnancy in the mare. Vet. Clinics N.
Am.: L. Anim. Pract. 2(2):291-302.
Allen, W.R. 1982. Immunological aspects of the endometrial cup reaction and the effect of xenogeneic pregnancy in horses and donkeys. J. Reprod. Fertil., Suppl. 31:57-
94.
Allen, W.R. 1984. Aspects of early embryonic development in farm animals. 10th
Intern. Congr. Anim. Reprod. Artif. Insemin., Univ. illinois Urbana-Champaign,
USDA. 4:Xffl- 1-Xffl-9.
Allen, W.R. 1993. Progesterone and the pregnant mare: unanswered chestnuts. Equine
Vet. J. 25(2):90-91.
Allen, W.R. 2000. The physiology of early pregnancy in the mare. AAEP Proc.
46:33 8-354.
Allen, W.R., and R.M. Moor. 1972. The origin of the equine endometrial cups I.
Production of PMSG by fetal trophoblast cells. J. Reprod. Fertil. 29(2):313-316.
Allen, W.R., and R.V Short. 1997. Interspecific and extraspecific pregnancies in equids: anything goes. J. Hered. 88(5):384-392.

56
Allen, W.R., and F. Stewart. 1978. The biology of pregnant mare serum gonadotrophin (PMSG). Current Topics Vet. Med. 1:50-72.
Allen, W.R., D.W. Hamilton, and R.M. Moor. 1973. The origin of equine endometrial cups II. Invasion of the endometrium by trophoblast. Mat. Rec. 177(4):485-502.
Allen, W.R., J.A. Skidmore, F. Stewart, and D.F. Antczak. 1993. Effects of fetal genotype and uterine environment on placental development in equids. J.
Reprod. Fertil. 97(1):55-60.s
Badinga, L., F.J. Michel, M.J. Fields, D.C. Sharp, and R.C. Siminen. 1994. Pregnancyassociated endometrial expression of antileukoproteinase gene is correlated with epitheliochorial placentation. Mol. Reprod. Dev. 38(4):357-363.
Ball, B.A., P.G. Miller, and P.F. Daels. 1992. Influence of exogenous progesterone on early embryonic development in the mare. Theriogenology. 38(6): 1055-1063.
Barnes, RJ., P.W. Nathanielsz, P.D. Rossdale, R.S. Comline, and M. Silver. 1975.
Plasma progestagens and oestrogens in fetus and mother in late pregnancy. J.
Reprod. Fertil., Suppl. 23:617-623.
Baumholtz, H.M. 1998. Progestin profiles near partuntion in light horse, pony and miniature horse mares. M.S. Thesis. Oregon State Univ., Corvallis. 1-7 1.
Bazer, F.W., and N.L. First. 1983. Pregnancy and parturition. J. Anim. Sci. 57(2):425-
460.
Beaconsfield, P., and C.A. Villee. 1979. Placenta-a neglected experimental animal (1st
Ed.). Pergamon Press Ltd., NY.
Beckers, J.F., A. Zarrouk, E.S. Batalha, J.M. Garbayo, L. Mester, and 0. Szenci. 1998.
Endocrinology of pregnancy: chorionic somatomanimotropins and pregnancyassociated glycoproteins: review. Acta Vet. Hung. 46(2): 175-189.
Bell, R.J., and F. Bristol. 1987. Fertility and pregnancy loss after delay of foal oestrus with progesterone and estradiol-17. J. Reprod. Fertil., Suppl. 35:667-668.
Berg, S.L., and O.J. Ginther. 1978. Effect of estrogens on uterine tone and life span of the corpus luteum in mares. J. Anim. Sci. 47(l):203-208.
Bergfelt, D.R., J.A. Woods, and O.J. Ginther. 1992. Role of the embryonic vesicle and progesterone in embryonic loss in mares. J. Reprod. Fertil. 95(2):339-
347.

Bhavnani, B.R., and R.V. Short. 1973. Formation of steroids by the pregnant mare ifi.
57
Metabolism of '4C-squalene and 3H-dehydroisoandrosterone injected into the fetal circulation. Endocrinology. 92(3):657-666.
Bhavnani, B.R., L.J. Martin, and R.D. Baker. 1975. Formation of steroids by the pregnant mare V. Metabolism of 14C-isopentenylpyrophosphate and 3Hdehydroisoandrosterone injected into the fetus. Endocrinology. 96(4):! 009-1017.
Bhavnani, B.R., R.V. Short, and S. Solomon. 1969. Formation of estrogens by the pregnant mare I. Metabolism of 7-3H-dehydroisoandrosterone and 4-14Candrostenedione injected into the umbilical vein. Endocrinology. 85(6):1 172-
1179.
Bjorkman, N. 1973. Fine structure of the fetal-maternal area of exchange in the epitheliochorial and endotheliochorial types of placentation. Acta Anat., Suppl.
86: 1-22.
Brady, HA., R.C. Burghardt, J.W. Evans, T.L. Blanchard, D.D. Varner, and J.E.
Bruemmer. 1993. Model system for the study of uterine/trophoblast interactions in the mare. J. Equine Vet. Sci.
I 3(9):506-5 11.
Brown, J.L., S.K. Wasser, D.E. Wildt, and L.H. Graham. 1994. Comparative aspects of steroid hormone metabolism and ovarian activity in felids, measured noninvasively in feces. Biol. Reprod. 51 (4):776-786.
Burns, SJ., and J.L. Fleeger. 1975. Plasma progestagens in the pregnant mare in the first and last 90 days of gestation. J. Reprod. Fertil., Suppi. 23:435-439.
Channing, C.P. 1969. Studies on tissue culture of equine ovarian cell types: pathways of steroidogenesis. J. Endocrinol. 43(3):403-414.
Chanmng, C.P., and S.A. Grieves. 1969. Studies on tissue culture of equine ovarian cell types: pathways of steroidogenesis. J. Endocrinol. 43(3):391-402.
Chavatte, P., D.W. Holtan, J.C. Ousey, and P.D. Rossdale. 1997. Biosynthesis and possible biological roles of progestagens during equine pregnancy and in the newborn foal. Equine Vet. J., Suppi. 24:89-95.
Chavatte, P., P.D. Rossdale, and A.D. Tait. 1995. 11-hydroxysteroid dehydrogenase
(11-HSD) in equine placenta. Proc. Annu. Cony. Am. Assoc. Equine
Practitioners. 41:264-265.
Collins, M.H. 1993. Placentas and foetal health. Equine Vet. J., Suppl. 14:8-11.

Combarnous, Y., M.C. Maurel, T. Magallon, M. Chopineau, F. Lecompte, F.
Apparailly, C. Kuntz, S. Hofferer, N. Martinat, and F. Guillou. 1991. Structure-
58 function study of equine pituitary and placental gonadotrophins, eLH, eFSH and eCGIPMSG. J. Reprod. Fertil., Suppi. 44:637.
Cottrill, C.M., J. Jeffers-Lo, J.C. Ousey, A.J. McGladdery, S.W. Ricketts, M. Silver, and
P.D. Rossdale. 1991. The placenta as a determinant of fetal well being in normal and abnormal equine pregnancies. J. Reprod. Fertil., Suppi. 44:591-601.
Cox, J.E. 1975. Oestrone and equilin in the plasma of the pregnant mare. J. Reprod.
Fertil., Suppi. 23:463-468.
Daels, P.F., B.A. Albrecht, and H.O. Mohammed. 1998. Equine chorionic gonadotropin regulates luteal steroidogenesis in pregnant mares. Biol. Reprod. 59(5): 1062-
1068.
Daels, P.F., D.C. Ammon, G.H. Stabenfeldt, I.K.M. Liu, J.P. Hughes, and B.L. Lasley.
1991a. Urinary and plasma estrogen conjugates, estradiol and estrone concentrations in nonpregnant and early pregnant mares. Theriogenology.
35(5): 1001-1017.
Daels, P.F., G.H. Stabenfeldt, J.P. Hughes, K. Odensvik, and H. Kindahi. 1991b.
Evaluation of progesterone deficiency as a cause of fetal death in mares with experimentally induced endotoxemia. Am. J. Vet. Res. 52(2):282-288.
Daels, P.F., S. Shideler, B.L. Lasley, J.P. Hughes, and G.H. Stabenfeldt. 1990. Source of oestrogen in early pregnancy in the mare. J. Reprod. Fertil. 90(1):55-61.
Darenius, K., S. Einarsson, and H. Kindahi. 1991. Endocrine studies of early pregnancy loss in the mare; comparison within mares between pregnancy loss and consecutive pregnancy. J. Reprod. Fertil., Suppi. 44:726-727.
Darenius, K., H. Kindahi, and A. Madej. 1987. Clinical and endocrine aspects of early fetal death in the mare. J. Reprod. Fertil., Suppl. 35:497-498.
Dehennin, L., A. Reiffstech, and R. Scholer. 1980. Simple methods for the synthesis of twenty different, highly enriched deuterium labeled steroids, suitable as internal standards for isotope dilution mass spectrometry. Biomed. Mass. Spectro. 7:493-
499.
Discafani, C.M., E.L. Squires, and G.D. Niswender. 1995. Functional state of primary and secondary corpora lutea in pregnant mares. Biol. Reprod., Suppi. 1:293-297.
Douglas, R.H., and O.J. Ginther. 1975. Development of the equine fetus and placenta.
J. Reprod. Fertil., Suppl. 23:503-505.

Enders, A.C., and I.K.M. Liu. 1991. Lodgement of the equine blastocyst in the uterus from fixation through endometrial cup formation. J. Reprod. Fertil., Suppi.
44:427-438.
59
Enders, A.C., K.C. Lantz, I.K.M. Liu, and S. Schlafke. 1988. Loss of poiar trophoblast during differentiation of the blastocyst of the horse. J. Reprod. Fertil., Suppi.
83(1):447-460.
Enders, A.C., K.C. Lantz, S. Schlafle, and I.K.M. Liu. 1995. New cells and old vessels: the remodeling of the endometrial vasculature during establishment of endometrial cups. Biol. Reprod., Suppl. 1:181-190.
Evans, J.W., B. Anthony, H. Hintz, and L.D. Van Vieck. 1990. The Horse (2x Ed.).
W.H. Freeman & Co., NY.
Flood, P.F., and A.W. Marrable. 1975. A histochemical study of steroid metabolism in the equine fetus and placenta. J. Reprod. Fertil., Suppi. 23:569-573.
Forde, D., L. Keenan, J. Wade, M. O'Connor, and J.F. Roche. 1987. Reproductive wastage in the mare and its relationship to progesterone in early pregnancy. J.
Reprod. Fertil., Suppi. 35:493-495.
Ganjam, V.K., and R.M. Kenney. 1975. Peripheral blood plasma levels and some unique metabolic aspects of progesterone in pregnant and non-pregnant mares.
Proc. Annu. Cony. Am. Assoc. Equine Practitioners. 21:263-276.
Ganjam, V.K., R.M. Kenney, and G. Flickinger. 1975. Plasma progestagens in cyclic, pregnant and post-partum mares. J. Reprod. Fertil., Suppi. 23:441-447.
Gastal, E.L., M.O. Gastal, D.R. Bergfelt, and O.J. Ginther. 1997. Role of diameter differences among follicles in selection of a future dominant follicle in mares.
Biol. Reprod. 57(6):1320-1327.
Gastal, M.O., E.L. Gastal, K. Kot, and O.J. Ginther. 1996. Factors related to the time of fixation of the conceptus in mares. Theriogenology. 46(7):1 171-1180.
Ginther, O.J. 1985a. Embryonic loss in mares: incidence, time of occurrence, and hormonal involvement. Theriogenology. 23(1 ):77-89.
Ginther, O.J. 1985b. Embryonic loss in mares: nature of loss after experimental induction by ovariectomy or prostaglandin F2a. Theriogenology. 24(1):87-98.
Ginther, O.J. 1992. Reproductive Biology of the Mare-Basic and Applied Aspects
(21x1 Ed.). Equiservices.
Cross Plains, WI.

Ginther, OJ. 1994. Equine physical utero-fetal interactions: a challenge and a wonder for the practitioner. J. Equine Vet. Sci. 14(6):313-318.
Ginther, OJ., and R.A. Pierson. 1984a. Ultrasonic anatomy and pathology of the equine uterus. Theriogenology. 21 (3):505-5 16.
Ginther, OJ., and R.A. Pierson. 1984b. Ultrasonic anatomy of equine ovaries.
Theriogenology. 21 (3):47 1-483.
Ginther, OJ., M.C. Garcia, D.R. Bergfelt, G.S. Leith, and S.T. Scraba. 1985.
Embryonic loss in mares: pregnancy rate, length of interovulatory intervals, and progesterone concentrations associated with loss during days 11 to 15.
Theriogenology. 24(4):409-4 17.
Gonzalez-Angulo, A., P. Hernandez-Jauregui, and G. Martinez-Zedilo. 1975. Fine structure of the gonads of the horse and its functional implications. J. Reprod.
Fertil., Suppi. 23:563-567.
Griffin, P.G., and O.J. Ginther. 1991. Uterine and fetal dynamics during early pregnancy in mares. Am. J. Vet. Res. 52(2):298-306.
Hafez, E.S.E. 1993. Reproduction in Farm Animals
(6th
Ed.). Lea & Febinger, Philad.
Hamon, M., S.W. Clarke, E. Houghton, A.L. Fowden, M. Silver, P.D. Rossdale, J.C.
Ousey, and R.B. Heap. 1991. Production of 5a-dihydroprogesterone during late pregnancy in the mare. J. Reprod. Fertil., Suppl. 44:529-535.
Hay, M.F., and W.R. Allen. 1975. An ultrastructural and histochemical study of the interstitial cells in the gonads of the fetal horse. J. Reprod. Fertil., Suppi. 23:557-
561.
Heap, R.B., M.H. Hamon, and W.R. Allen. 1991. Oestrogen production by the preimplantation donkey conceptus compared with that of the horse and the effect of between-species embryo transfer. J. Reprod. Fertil. 93(1): 141-147.
Heistermann, M., U. Mohie, H. Vervaecke, L. van Elsacker, and J.K. Hodges. 1996.
Application of urinary and fecal steroid measurements for monitoring ovarian function and pregnancy in the bonobo
(Pan paniscus) and evaluation of perineal swelling patterns in relation to endocrine events. Biol. Reprod. 55(4):844-853.
Hershman, L., and R.H. Douglas. 1979. The critical period for the maternal recognition of pregnancy in pony mares. J. Reprod. Fertil., Suppl. 27:395-401.
Holtan, D.W., E. Houghton, M. Silver, A.L. Fowden, J. Ousey, and P.D. Rossdale.
1991. Plasma progestagens in the mare, fetus and newborn foal. J. Reprod.
Fertil., Suppi. 44:517-528.

Holtan, D.W., T.M. Nett, and V.L. Estergreen. 1975. Plasma progestins in pregnant, postpartum and cycling mares. J. Anim. Sci. 40(2):251-260.
Holtan, D.W., E.L. Squires, D.R. Lapin, and O.J. Ginther. 1979. Effect of ovariectomy on pregnancy in mares. J. Reprod. Fertil., Suppi. 27:457-463.
Hoppen, H.O. 1994. The equine placenta and equine choriomc gonadotrophin-an overview. Exp. Clin. Endocrinol. 102(3):235-243.
Iruela-Arispe, M.L. 1997. Normal placentation: a tale that requires an epithelial-toendothelial conversion. J. Clin. Investig. 99(9):2057-2058.
Irvine, C.H.G., P. Sutton, J.E. Turner, and P.E. Mennick. 1990. Changes in plasma progesterone concentrations from days 17 to 42 of gestation in mares maintaining or losing pregnancy. Equine Vet. J. 22(2):104-106.
Irwin, C.F.P. 1975. Early pregnancy testing and its relationship to abortion. J.
Reprod. Fertil., Suppi. 23:485-488.
Jackson, S.A., E.L. Squires, and T.M. Nett. 1986. The effect of exogenous progestins on endogenous progesterone secretion in pregnant mares. Theriogenology.
25(2):275-279.
Jeffcott, L.B., J.H. Hyland, A.A. MacLean, T. Dyke, and G. Robertson-Smith. 1987.
Changes in maternal hormone concentrations associated with induction of fetal death at day 45 of gestation in mares. J. Reprod. Fertil., Suppl. 35:46 1-467.
Jones, D. 1993. Formation, function, and clinical evaluation of the equine placenta.
Comp. Cont. Educ. Pract. Vet. 15(1):97-105.
Kasman, L.H., J.P. Hughes, G.H. Stabenfeldt, M.D. Starr, and B.L. Lasley. 1988.
Estrone sulfate concentrations as an indicator of fetal demise in horses. Amer. J.
Vet. Res. 49(2):184-187.
Knowles, J.E., E.L. Squires, R.K. Shideler, S.F. Tarr, and T.M. Nett. 1993.
Relationship of progesterone to early pregnancy loss in mares. J. Equine
Vet. Sci. 13(9):528-533.
Knowles, J.E., E.L. Squires, R.K. Shideler, S.F. Tarr, and T.M. Nett. 1994. Progestins in mid- to late-pregnant mares. J. Equine Vet. Sci. 14(12):659-663.
Lasley, B.L., D.C. Ammon, P.F. Daels, J.P. Hughes, C. Munro, and G.H. Stabenfeldt.
1990. Estrogen conjugate concentrations in plasma and urine reflect estrogen secretion in the nonpregnant and pregnant mare: a review. J. Equine Vet. Sci.
lO(6):444-448.

62
Lea, R.G., F. Stewart, W.R. Allen, I. Ohno, and D.A. Clark. 1995. Accumulation of chromotrope 2R positive cells in equine endometrium during early pregnancy and expression of transforming growth factor-2 (TGF-2). J. Reprod. Fertil.
103(2):339-347.
Leiser, R., and P. Kaufmann. 1994. Placental structure: in a comparative aspect. Exper.
Clin. Endocrinol. 102(3):122-134.
Lennard, S.N., C. Gerstenberg, W.R. Allen, and F. Stewart. 1998. Expression of epidermal growth factor and its receptor in equine placental tissues. J. Reprod.
Fertil.
1 12(1):49-57.
Lennard, S.N., F. Stewart, and W.R. Allen. 1995. Transforming growth factor i expression in the endometrium of the mare during placentation. Mol. Reprod.
Dev. 42(2):131-140.
Liggins, G.C., and G.P. Thorburn. 1993. Marshall's Physiology of Reproduction
Ed.). Chapman & Hall, London.
(4th
Marsan, C., A.K. Goff, J. Sirois, and K.J. Beneridge. 1987. Steroid secretion by different cell types of the horse conceptus. J. Reprod. Fertil., Suppi. 35:363-369.
Martin, J.L., A. Saltiel, and J.W. Evans. 1989. Progesterone synthesis by different types of corpora lutea during the first 198 days of pregnancy. J. Equine Vet. Sci.
9(2):84-87.
McDowell, K.J., M.H. Adams, and C.B. Baker. 1993. Chorionic gonadotropin a and subunit RNAs are present in equine placental membranes by day 30 of pregnancy.
J. Equine Vet. Sci. 13(9):515-518.
McDowell, K.J., M.H. Adams, K.M. Franklin, and C.B. Baker. 1995. Changes in equine endometrial retinol-binding protein RNA during the estrous cycle and early pregnancy and with exogenous steroids. Biol. Reprod. 52(2):438-443.
McKinnon, A.O., E.L. Squires, E.M. Carnevale, and M.J. Hermenet. 1988a.
Ovariectomized steroid-treated mares as embryo transfer recipients and as a model to study the role of progestins in pregnancy maintenance. Theriogenology.
29(5): 1055-1063.
McKinnon, A.O., E.L. Squires, and R.K. Shideler. 1988b. Diagnostic ultrasonography of the mare's reproductive tract. J. Equine Vet. Sci.
8(4):329-333.

Meadows, S., W.R. Allen, and D.F. Antczak. 1995. Factors affecting the maturation and invasiveness in vitro of chorionic girdle cells of different genotypes. Biol.
Reprod., Suppi. 1:171-180.
63
Mennon, M., H. Peegel, and K.M. Menon. 1985. Lipoprotein augmentation of human choriomc gonadrotropin and prolactin stimulated progesterone synthesis by rat luteal cells. J. Steroid Biochem. 22:79-84.
Merchant-Larios, H. 1979. Ultrastructural events in horse gonadal morphogenesis. J.
Reprod. Fertil., Suppl. 27:479-485.
Miller, C.W., and D.W. Holtan. 1997. Progestin analysis by gas chromatography/mass spectrometry in plasma and feces of horses, sheep, and cattle. Biol. Reprod.
56(1): 191.
Miller, C.W., H.H. Meyer, and D.W. Holtan. 2000. Fecal progestins in the early gestation ewe monitored by gas chromatography/mass spectrometry. Proc.
Amer. Soc. Anim. Sci. 5 1:79-82.
Monfort, S.L., N.P. Arthur, and D.E. Wildt. 1991. Monitoring ovarian function and pregnancy by evaluating excretion of urinary oestrogen conjugates in semi-freeranging Przewalski's horses (Equusprzewalskii). J. Reprod. Fertil. 91(1):155-
164.
Morgenthal, J.C., and C.H. van Niekerk. 1991. Plasma progestagen levels in normal mares with luteal deficiency during early pregnancy, and in twinning habitual aborters. J. Reprod. Fertil., Suppl. 44:728-729.
Naden, J., E.L. Squires, and T.M. Nett. 1990. Effect of maternal treatment with altrenogest on age at puberty, hormone concentrations, pituitary response to exogenous GnRH, oestrous cycle characteristics and fertility of fillies. J. Reprod.
Fertil. 88(1):185-195.
Nett, T.M., D.W. Holtan, and V.L. Estergreen. 1973. Plasma estrogens in pregnant and postpartum mares. J. Anim. Sci. 37(4):962-970.
Parry-Weeks, L.C., and D.W. Holtan. 1987. Effect of altrenogest on pregnancy maintenance in unsynchronized equine embryo recipients. J. Reprod. Fertil.,
Suppi. 35:433-438.
Pashen, R.L., and W.R. Allen. 1979a. Endocrine changes after fetal gonadectomy and during normal and induced parturition in the mare. Anim. Reprod. Sci. 2(3):271-
288.

Pashen, R.L., and W.R. Allen. 1979b. The role of the fetal gonads and placenta in steroid production, maintenance of pregnancy and parturition in the mare. J.
Reprod. Fertil., Suppl. 27:499-509.
Peariman, W.H. 1948. In the Hormones (6t1I Ed.). Academic Press, NY.
Raeside, J.I., R.M. Liptrap, and F.J. Milne. 1973. Relationship of fetal gonads to urinary estrogen excretion by the pregnant mare. Am. J. Vet. Res. 34(6):843-845.
Ramsey, E.M. 1982. The placenta-human and animal (1st Ed.). Praeger Publ., NY.
Riad, M.T., and D.W. Holtan. 1993. Urinary estrogens and progestins in pregnant pony mares. Proc.
13th
Equine Nutr. Physiol. Symp. Univ. of FL, Gainsville.
311.
Rogerson, F.M., J.R. Head, K. Imaishi, P. Suart, and J.I. Mason. 1993. Expression of cytochrome 17a-hydroxylase in the equine fetal gonad and adult adrenal cortex.
Biol. Reprod. 48:362.
Rossdale, P.D. 1993. Clinical view of disturbance in equine foetal maturation. Equine
Vet. J., Suppi. 14:3-7.
Rossdale, P.D., J.C. Ousey, C.M. Cottrill, P. Chavatte, W.R. Allen, and A.J. McGladdery.
1991. Effects of placental pathology on maternal plasma progestagen and mammary secretion calcium concentration and on neonatal adrenocortical function in the horse. J. Reprod. Fertil. 44:579-590.
Samuel, C.A., W.R. Allen, and D.H. Steven. 1974. Studies on the equine placenta I.
Development of the microcotyledons. J. Reprod. Fertil. 41(2):441-445.
Samuel, C.A., W.R. Allen, and D.H. Steven. l975a. Development of pituitary and adrenal glands in the fetal horse. J. Reprod. Fertil., Suppl. 23:553-556.
Samuel, C.A., W.R. Allen, and D.H. Steven. 1975b. Ultrastructural development of the equine placenta. J. Reprod. Fertil., Suppl. 23:575-578.
Samuel, C.A., W.R. Allen, and D.H. Steven. 1976. Studies on the equine placenta II.
Ultrastructure of the placental barrier. J. Reprod. Fertil. 48(2):257-264.
Samuel, C.A., W.R. Allen, and D.H. Steven. 1977. Studies on the equine placenta ifi.
Ultrastructure of the uterine glands and the overlying trophoblast. J. Reprod.
Fertil. 51(2):433-437.
Schorpp-Kistner, M., Z.Q. Wang, P. Angel, and E.F. Wagner. 1999. JunB is essential for mammalian placentation. EMBO J. 18(4):934-948.

Schutzer, W.E. 1995. Steroid transformations in the pregnant mare: the effect of exogenous progestins and trilostane on the progestin milieu-in vivo and in vitro.
M.S. Thesis. Oregon State Univ., Corvallis. Appendix.
65
Schutzer, W.E., and D.W. Holtan. 1996. Steroid transformations in pregnant mares: metabolism of exogenous progestins and unusual metabolic activity in vivo and in vitro.
Steroids. 61:94-99.
Schwab, C.A., J.W. Evans, and G.D. Potter. 1990. Prolactin and progesterone concentrations during early pregnancy and relationship to pregnancy loss prior to day 45. J. Equine Vet. Sci. 10(4):280-283.
Shideler, R.K., E.L. Squires, J.L. Voss, and D.J. Eikenberry. 1982. Exogenous progestin therapy for maintenance of pregnancy in ovariectomized mares. Proc. Annu.
Cony. Am. Assoc. Equine Practitioners. 27:211-219.
Short, R.V. 1959. Progesterone in blood IV. Progesterone in the blood of mares. J.
Endocrinol. 19:207-210.
Silver, M. 1984. Some aspects of equine placental exchange and foetal physiology.
Equine Vet. J. 16(4):227-233.
Sist, M.D., M.A. Youngblood, and J.F. Wiffiams. 1987. Using fecal estrone sulfate concentrations to detect pregnancies. Vet. Med. 82(10): 1036-1043.
Squires, E.L., and O.J. Ginther. 1975. Collection technique and progesterone concentration of ovarian and uterine venous blood in mares. J. Anim. Sci.
40(2):275-28 1.
Squires, E.L., C.P. Heesemann, S.K. Webel, R.K. Shideler, and J.L. Voss. 1983.
Relationship of altrenogest to ovarian activity, hormone concentrations and fertility of mares. J. Amm. Sci. 56(4):901-910.
Squires, E.L., A.O. McKinnon, and R.K. Shideler. 1988. Use of ultrasonography in reproductive management of mares. Theriogenology. 29(1):55-70.
Squires, E.L., T.M. Nett, G.J. Wiepz, and E.J. Mock. 1986. Use of an enzyme assay for determination of progesterone in broodmares. Proc. Annu. Cony. Am. Assoc.
Equine Practitioners. 32:565-570.
Squires, E.L., J.L. Voss, and R.K. Shideler. 1984. Use of altrenogest for the broodmare.
Proc. Annu. Cony. Am. Assoc. Equine Practitioners. 29(1):431-434.
Squires, E.L., B.C. Wentworth, and O.J. Ginther. 1974. Progesterone concentration in blood of mares during the estrous cycle, pregnancy and after hysterectomy. J.
Anim. Sci. 39(4):759-767.

Stabenfeldt, G.H., and J.P. Hughes. 1987. Clinical aspects of reproductive endocrinology in the horse. Comp. Cont. Educ. Pract. Vet. 9(6):678-686.
Stabenfeldt, G.H., J.P. Hughes, J.D. Wheat, J.W. Evans, P.C. Kennedy, and P.T. Cupps.
1974. The role of the uterus in ovarian control in the mare. J. Reprod. Fertil.,
Suppi. 37(2):343-351.
Steven, D.H. 1982. Placentation in the mare. J. Reprod. Fertil., Suppi. 31:41-55.
Stewart, F., and W.R. Allen. 1979. The binding of FSH, LH and PMSG to equine gonadal tissues. J. Reprod. Fertil., Suppi. 27:431-440.
Stewart, F., S.N. Lennard, and W.R. Allen. 1995. Mechanisms controlling formation of the equine chorionic girdle. Biol. Reprod., Suppl. 1:15 1-159.
Tait, A.D., S. Santikarn, and W.R. Allen. 1983. Identification of 3-hythoxy-5,7pregnadien-20-one and 3-hydroxy-5,7-androstadien- 17-one as endogenous steroids in the fetal horse gonad. J. Endocrinol. 99(1):87-92.
Thorburn, G.D. 1993. A speculative review of parturition in the mare. Equine Vet. J.,
Suppl. 14:41-49.
van Niekerk, C.H., and W.R. Allen. 1975. Early embryonic development in the horse. J.
Reprod. Fertil., Suppl. 23:495-498.
van Niekerk, F.E., and van C.H. Niekerk. 1998a. The effect of dietary protein on reproduction in the mare VI. Serum progestagen concentrations during pregnancy.
J. S. Afr. Vet. Assoc. 69(4):143-149.
van Niekerk, F.E., and C.H. van Niekerk. 1998b. The effect of dietary protein on reproduction in the mare
\Tll.
Embryonic development, early embryonic death, foetal losses and their relationship with serum progestagen. J. S. Afr. Vet. Assoc.
69(4): 150-155.
Villahoz, M.D., E.L. Squires, J.L. Voss, and R.K. Shideler. 1985. Some observations on early embryonic death in mares. Theriogenology. 23(6):915-924.
Vivrette, S. 1994. The endocrinology of parturition in the mare. Vet. Clinics of North
America: Equine Prac. 10(1):1-17.
Voller, B.E., L.C. Parry-Weeks, and D.W. Holtan. 1991. The effect of Regu-Mate,® a synthetic progestin, on early pregnancy maintenance, conceptus growth, and corpora lutea development in pregnant pony mares. J. Equine Vet. Sci.
1 1(1):46-.
50.

Walt, M.L., G.H. Stabenfeldt, J.P. Hughes, D.P. Neely, and R. Bradbury. 1979.
Development of the equine ovary and ovulation fossa. J. Reprod. Fertil., Suppi.
27:471-477.
67
Wilker, C., P.F. Daels, P.J. Burns, and B.A. Ball. 1992. Progesterone therapy during early pregnancy in the mare. Proc. Annu. Cony. Am. Assoc. Equine Practitioners.
37(1): 161-172.
Woods, G.L. 1989. Pregnancy loss: a major cause of subfertility in the mare. Equine
Practice. 1 1(5):29-32.
Woods, G.L., C.B. Baker, J.L. Baldwin, B.A. Ball, J. Bilinsky, W.L. Cooper, W.B. Ley,
E.C. Mank, and H.N. Erb. 1987. Early pregnancy loss in brood mares. J.
Reprod. Fertil., Suppl. 35:455-459.
Yamauchi, S. 1975. Morphology and histochemistry of the endometrial cup. J. Reprod.
Fertil., Suppl. 23:397-400.
Zavy, M.T., M.W. Vernon, D.C. Sharp ifi, and F.W. Bazer. 1984. Endocrine aspects of early pregnancy in pony mares: a comparison of uterine luminal and peripheral plasma levels of steroids during the estrous cycle and early pregnancy.
Endocrinology. 115(1):214-219.