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BOR - Papers in Press, published online ahead of print March 7, 2007.
Biol Reprod 2007, 10.1095/biolreprod.106.058578
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BIOLOGY OF REPRODUCTION 77, 9–17 (2007)
DOI: 10.1095/biolreprod.106.058578
© 2007 by the Society for the Study of Reproduction, Inc.

Impact of Dietary Fatty Acids on Oocyte Quality and Development in Lactating Dairy Cows1

Ali A. Fouladi-Nashta 3 4, Carlos G. Gutierrez 5, Jin G. Gong 6, Philip C. Garnsworthy 4, and Robert Webb 2 4

School of Biosciences,4 Sutton Bonington Campus, The University of Nottingham, Loughborough, Leicestershire LE12 5RD, United Kingdom Departamento de Reproducción,5 Facultad de Medicina Veterinaria, UNAM, Ciudad Universitaria, 04510 Mexico Roslin Institute,6 Roslin, Midlothian, Edinburgh EH25 9PS, Scotland, United Kingdom

ABSTRACT

The purpose of this study was to examine the effects of level of rumen inert fatty acids on developmental competence of oocytes in lactating dairy cows. Estrous cycles were synchronized in 22 cows on a silage-based diet supplemented with either low (200 g/day) or high (800 g/day) fat. A total of 1051 oocytes were collected by ultrasound-guided ovum pickup (OPU) in seven sessions/cow at 3–4 day intervals. Oocytes were matured, fertilized, and cultured to the blastocyst stage in vitro. Embryo quality was assessed by differential staining of Day 8 blastocysts. The high-fat diet reduced numbers of small and medium follicles. There was no effect on the quality of oocytes (grades 1–4) or cleavage rate. However, high fat significantly improved blastocyst production from matured (P < 0.005) and cleaved (P < 0.05) oocytes. Blastocysts from the high-fat group had significantly more total, inner cell mass and trophectoderm cells than the low-fat group (P < 0.05). Regression analysis showed negative effects of milk yield (P < 0.001), dry matter intake (P < 0.001), metabolizable energy intake (P < 0.005), and starch intake (P < 0.001) on blastocyst production in the low-fat group but not in the high-fat group. Within the low-fat group, blastocyst production was negatively related to growth hormone (P < 0.05) and positively related to leptin (P < 0.05). The low-fat group had higher nonesterified fatty acids than the high-fat group (P < 0.05). In conclusion, higher milk yields were associated with reduced developmental potential of oocytes in cows given a low-fat diet. Provision of a high-fat diet buffered oocytes against these effects, resulting in significantly improved developmental potential.

cattle, diet, embryo, embryo development, fatty acids, follicular development, lactation, oocyte, oocyte development, ovum pickup, ovum transport

INTRODUCTION

Intensive genetic selection for increased milk production, coupled with technological improvements in nutrition, has led to significant increases in milk yield in cows in recent decades. However, this increase in milk output per cow has been accompanied by a worldwide decline in cow fertility [1]. The decline in pregnancy rate to a single artificial insemination has been reported to be approximately 0.45%–1% per annum [2, 3]. High-yielding dairy cows are typically in a state of negative energy balance postpartum because the amount of energy required for maintenance both of metabolic function and milk production exceeds the amount of energy cows consume. Insufficient energy supply results in poor reproductive performance, which includes a delay in the onset of estrous cycles postpartum [4–7] and a reduction in oocyte quality [8, 9], resulting in low conception rates and a high rate of early embryonic death [10].

Embryo mortality during the preimplantation period is a key contributor to the reduced fertility in cattle, with up to 40% of total embryo loss occurring between Days 8–17 of pregnancy [11]. This stage of embryonic loss coincides with the inhibitory effects of interferon tau (IFN{tau}), produced by trophectoderm cells of embryos, on prostaglandin release from the uterus [12–14]. This suggests that a proportion of embryos are unable to inhibit prostaglandin F2{alpha} (PGF2{alpha}) release leading to regression of the CL, reduction in progesterone production [15], and termination of pregnancy [12, 14]. Hence the quality of the preimplantation embryo, and its ability to communicate with the cells of the uterus, are critical for the establishment and continuation of pregnancy.

Short-term changes in plane of nutrition have been shown to have a direct effect on ovarian follicular dynamics in cattle, without any changes in circulating concentrations of gonadotropins [16, 17]. It has been hypothesized that endocrine and metabolic signals that regulate follicular growth also influence oocyte development either through changes in hormone/growth factor concentrations in follicular fluid or via granulosa-oocyte interactions [18]. For example, as well as regulating follicular growth [19], short-term changes in dietary energy intake influence both oocyte morphology and developmental potential [20–23]. The majority of these studies have been carried out in sheep, beef cows, or heifers; thus far, the effects of dietary fatty acids on oocyte quality in lactating dairy cows, which are under greater metabolic pressure, have not been investigated fully.

Fatty acids are rich sources of energy and have important roles in the structure and function of biological membranes [24]. Dietary fats influence reproductive function, either by increasing energy balance [4] or by actions on reproductive processes that are not related to energy [25]. Supplementation of the diet with fat has been shown to increase the total number of follicles and to stimulate growth and size of the preovulatory follicle [26, 27]. In addition, increased availability of fatty acid precursors is coupled with increased steroid and eicosanoid secretion, which can alter ovarian and uterine function and affect embryo implantation [28]. Maternal dietary fat also influences amniotic fluid and fetal intestinal membrane structural lipid in the rat [29].

Oocytes in cows and sows contain a high level of fatty acids. For example, fatty acid content of immature cow oocytes has been estimated to be 63 µg per oocyte, with phospholipids accounting for 25% of all fatty acids [30]. Saturated fatty acids account for less than 30% of the total fatty acid composition. Tetraenoic acids are not found in small and large follicles [31]. The proportion of linoleic acid is significantly lower in follicular fluid from large follicles (31.1% ± 1.2% of total fatty acids) than from small follicles (34.8% ± 0.7% of total fatty acids), and there is a significant inverse correlation between follicle diameter and the percentage of linoleic acid in follicular fluid [31]. There are also seasonal changes in the developmental capacity of bovine oocytes that might be related to changes in fatty acid composition of oocytes [32]. These changes in the fatty acid profile of follicular fluid and oocytes were related to ambient temperature, but may also reflect seasonal changes in dietary ingredients.

Fatty acids are available for use as an energy source during oocyte maturation and the extended period of embryo development before implantation in the cow; hence alteration in the fatty acid composition of bovine oocytes might improve maturation [33] and subsequent embryo development. There is negligible information on the effects of dietary fat levels on oocyte quality, and subsequent development in vitro, for high-yielding lactating dairy cows. In the current study, the impact of two levels of supplementation of rumen inert fat in the diets of lactating dairy cows was investigated. Ovarian activity was assessed by recording numbers of follicles by ultrasonography, and oocyte quality was evaluated by morphological classification and by the rate of development after in vitro maturation and fertilization. If a clear effect of changing dietary fatty acids on bovine oocyte quality could be demonstrated, this would provide new insights into the underlying factors influencing oocyte quality and the mechanisms involved.

MATERIALS AND METHODS

Animals and Experimental Design

This experiment was conducted in accordance with the requirements of the Home Office Animals (Scientific Procedures) Act 1986.

Twenty-two dairy cows (parity 1–3) were selected and allocated equally to two groups receiving a total mixed ration (TMR) diet containing rumen inert fat (Megalac, Volac International, Royston, UK) at either 200 g/day (low fat) or 800 g/day (high fat) (Table 1) on Days 40–60 after calving. The fatty acid composition of the rumen inert fat was 47% palmitic, 5% stearic, 38% oleic, 9% linoleic, and 1% linolenic acids. Animals were allocated to groups on the basis of milk yield, days after calving, parity, and body condition score (BCS). The animals were fed on the experimental diets through a robotic feeding system for 14 days before start of ovum pickup (OPU) on Day 13 of the experiment. Dry matter intake (DMI), metabolizable energy intake (MEI), starch, and fat intakes were calculated from TMR intakes and analysis of the diets. Estrous cycles were synchronized by using a progesterone releasing implant (controlled internal drug release [CIDR] device, InterAg, Hamilton, New Zealand) on Day 2 followed by injection of PGF2{alpha} on Day 10 and removal of CIDR on Day 11. OPU sessions started on Day 13 and were repeated seven times with a 3- to 4-day interval between each OPU (Figure 1).


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  		TABLE 1. Composition and chemical analysis of the low- and high-fat diets.


Figure 01
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  		FIG. 1. An outline of the experimental design indicating initiation of dietary treatment, synchronization of estrus, and ovum pickup (OPU). Dairy cows were fed on experimental diets for 14 days before the start of OPU. Estrous cycles were synchronized with a CIDR on day 2 followed by injection of PGF2{alpha} on Day 10 and removal of the CIDR on Day 11. OPU sessions were started on Day 13 and repeated seven times with either 3- or 4-day intervals between OPU sessions. Immediately prior to performing each OPU, blood samples were collected from each cow for measurement of hormones and metabolites.

Immediately prior to each OPU, blood samples were obtained from each cow. These were analyzed for serum concentrations of amino acids, hormones (insulin, glucagon, insulin-like growth factor 1 [IGF1], growth hormone [GH], leptin, and progesterone) and metabolites (glucose, ß-hydroxybutyrate [BHB], nonesterified fatty acids [NEFA], albumin, urea, magnesium, inorganic phosphate, and globulin).

Chemicals

All chemicals were purchased from Sigma (Dorset, UK) unless otherwise stated.

Collection of Oocytes by OPU

OPU was performed with an ultrasound scanner (Medison Sonovet 600, Marl, Germany) fitted with a 6.5 MHz transvaginal probe. Before oocyte collection, cows were premedicated with 0.8 ml acepromacine (i.v.) and an epidural injection of 6–10 ml of 2% lignocaine. The rectum was then emptied and the perineal region was washed with surgical soap. Follicular aspiration was performed with a 60-cm, 18 G needle (Arnolds Veterinary Products, Arnolds, Harlescott, UK) at 60 mm Hg vacuum pressure.

Follicular fluid was collected into flushing medium (0.25 mg/ml heparin in phosphate buffered saline [PBS] supplemented with 5% heat-inactivated fetal calf serum [HIFCS]). The contents of the collection tubes were passed through an embryo filter (Em-Con, 75-µm pore size, Cook, Queensland, Australia) and washed with flushing medium. All collected oocytes were graded (Grades 1–4) morphologically based on the number and intensity of the cumulus cells and homogeneity of the ooplasm, as previously described [34]. Examples of oocytes from the four categories are presented in Figure 2.


Figure 02
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  		FIG. 2. Examples of the four grades of oocytes (Grades l–4) are shown, with Grade 1 being assessed as the highest quality oocyte. A) Grade 1. Compact cumulus (>4–5 layers) with a homogeneous ooplasm. B) Grade 2. Compact cumulus of one to two layers with homogeneous ooplasm having a coarse appearance. C) Grade 3. Less compact cumulus (slightly expanded cumulus) with irregular ooplasm containing dark clusters. D) Grade 4. Denuded oocyte or expanded cumulus, irregular ooplasm.

In Vitro Maturation, Fertilization, and Embryo Culture

Prior to culture, oocytes were washed twice in oocyte washing medium (Tissue Culture Medium 199 [TCM199] with Earles salts [Gibco, Grand Island, NY], 75.0 mg/L kanamycin monosulfate [Sigma-Aldrich, Poole, Dorset, UK], 7.08 g/L Hepes [pH 7.8, osmolarity 279 mOsmol/kg H2O]) supplemented with 10% HIFCS. Groups of oocytes recovered from each OPU for each cow were cultured separately. All oocytes were cultured in 35-mm culture dishes (Nunc, Denmark) containing 10 µl maturation medium per oocyte (TCM 199 supplemented with 10 µg FSH/ml [follitropin; Bioniche Animal Health, Belleville, ON], 10 µg LH/ml [leutropin; Bioniche Animal Health], 1 mg estradiol/ml, 50 µg gentamicin/ml, and 10% HIFCS) under mineral oil in a humidified atmosphere of 5% CO2 in air at 39°C. In vitro-matured oocytes were fertilized with frozen sperm from a single bull, as previously described [35]. Briefly, motile sperm were prepared after 45 min of swim up in calcium-free medium followed by centrifugation at 300 x g at room temperature and resuspension of the pellet in fertilization medium. The cumulus oocyte complexes (COCs) were gently pipetted to remove adhering granulosa cells and to break up aggregated COCs. Disaggregated COCs were then washed once in oocyte wash medium and transferred into 45-µl microdrops of fertilization medium containing sperm (106 sperm/ml) and cultured for 24 h at 39°C in a humidified incubator of 5% CO2 in air. After 24 h, all presumptive zygotes were denuded from cumulus cells and cultured in 5 µl/embryo of SOFaaci (synthetic oviductal fluid medium supplemented with amino acids, sodium citrate, and myoinositol; [36]) supplemented with 4 mg/ml of fatty acid-free BSA and cultured at 39°C in a humidified incubator with 5% O2, 5% CO2, and 90% N2. The culture was continued up to Day 8, and medium was renewed every 2 days. The number of cleaved embryos and development to the blastocyst stage were recorded.

Differential Staining of Blastocysts

Day 8 blastocysts were differentially stained for counting cells in the inner cell mass (ICM) and trophectoderm (TE) compartments. Briefly, TE cells were permeablized and stained by incubating embryos in a 0.2% solution of Triton X-100 in SOF–BSA containing 30 µg propodium idodide (PI)/ml for 20s. Immediately after, embryos were washed twice in SOF culture medium and fixed in ice-cold methanol containing 10 µg/ml bisbenzimide (Hoescht 33342) for 10 min. This allows fixation of embryos as well as staining for counting cells. Embryos were then transferred into a 50:50 solution of methanol:glycerol followed by mounting in small droplets of glycerol and examination under a Leica epiflourescent microscope (Leica, Germany). Differentially stained blastocysts are visualized with distinct TE (red) and ICM (blue) compartments (Figure 3).


Figure 03
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  		FIG. 3. Example of a differentially stained blastocyst. A simple and fast method (see Materials and Methods) was used to differentially stain blastocysts for counting numbers of total cells, inner cell mass (ICM), and trophectoderm (TE) cells. Lines indicate cell types.

Hormone and Growth Factors Assays

Plasma insulin concentrations were measured by using an I125-labeled insulin double-antibody radioimmunoassay based on the method of Starr et al. and [37] modified according to Sinclair et al. [38]. Assay sensitivity was 2.2 mIU/L and intra- and interassay coefficients of variation were 4.0% and 8.4%, respectively. Plasma glucagon was measured using an I125-labeled-glucagon double-antibody radioimmunoassay kit supplied by Linco Research Inc. (St. Charles, MO). The sensitivity of the assay was 40.5 pg/ml and the intra- and interassay coefficients of variation were 8.8% and 8.3%, respectively.

Insulin-like growth factor-1 (IGF1) was measured after removal of insulin-like growth factor binding protein (IGFBP) by size-exclusion high performance liquid chromatography, as described by Gutierrez et al. [39]. The sensitivity of the assay was 0.11 ng/ml and the intra-assay coefficient of variation was 3.8%.

GH was measured as described by Lovendahl et al. [40]. The sensitivity of the assay was 1.2 ng/ml. The interassay coefficient of variation was 11.3% and the mean intra-assay coefficient of variation was 3.9%.

Plasma leptin concentration was determined by the method of Blache et al. [41]. The primary antibody, normal emu serum, and ovine anti-emu second antibody were kindly provided by Dr. Blache (University of Western Australia, Perth, Australia); ovine leptin was kindly supplied by Dr. Keisler (University of Missouri, Columbia, MO) and was iodinated in-house [42]. The detection limit for leptin was 0.2 ng/ml, and the interassay coefficients of variation for low, medium, and high controls were 13.0%, 3.9%, and 15.3%, respectively. The corresponding intra-assay coefficients of variation were 9.1%, 12.2%, and 13.7%, respectively.

Plasma progesterone concentration was determined by direct radioimmunoassay [43]; assay sensitivity was 0.3 ng/ml, and intra- and interassay coefficients of variation were 5.7% and 9.4%, respectively.

Analysis of Metabolites and Amino Acids

Plasma concentration of urea was measured by an enzymatic glutamate dehydrogenase method (Randox Laboratories Ltd., Antrim, UK); interassay coefficient of variation was 10.5%. Plasma concentration of BHB was measured by using a kit (ß-hydroxybutyrate procedure 310-UV, Sigma Diagnostics Poole, Dorset, UK); interassay coefficient of variation was 4.8%. Plasma concentration of NEFA was measured by the method of Duncombe [44]; interassay coefficient of variation was 7.3%. Plasma concentrations of total protein, albumin, globulin, phosphorus, and magnesium were measured using kits (Bayer Healthcare Diagnostics, Newbury, UK); interassay coefficients of variation were 5%–10%. Plasma concentration of glucose was measured using a kit (HK; Sigma) in combination with a glucose standard (GLU GOD-Perid; Roche Diagnostics GmbH, Mannheim, Germany); interassay coefficient of variation was 8.9%.

Amino acid analysis was performed as previously described by Wiseman et al. [45]. Briefly, samples were oxidized with a hydrogen peroxide/formic acid/phenol mixture. Excess oxidation reagent was decomposed with sodium metabisulfite. The oxidized samples were hydrolyzed with 6 M hydrochloric acid for 24 h. The hydrolysate was adjusted to pH 2.2, centrifuged, and filtered. Amino acids were separated by ion exchange chromatography (Biochrom 20+ Amino Acid Analyser, Biochrom Ltd, Cambridge, UK) and determined after reacting with ninhydrin by using photometric detection at 570 nm (440 nm for proline).

Statistical Analyses

All data analysis was conducted using Genstat 8 (Lawes Agricultural Trust, Rothamstead, UK). Milk yield, live weight, intake, and plasma composition data were analyzed by analysis of variance with dietary treatment as the main effect. OPU session and cow were nested within treatment to allow for repeated measures. Follicle, oocyte, and embryo data were analyzed by GLM regression analysis allowing for effects of individual cows, utilizing a Poisson distribution for data involving counts and a binomial distribution with logit function for data involving proportions. The effects of milk yield, live weight, dry matter starch and fat intake, MEI, and blood metabolites on the proportion of blastocysts developing from oocytes undergoing in vitro fertilization (IVF) were analyzed by logistic regression. Mean data for individual cows were used to fit linear effects and diet interactions for each variable. Pearson correlation coefficients were generated for associations of milk yield, feed intake, and live weight with peripheral concentrations of hormones and metabolites. Data were analyzed using logistic regression analysis. Note that logistic regression calculates changes in the log odds of the dependent variable, not changes in the dependent variable itself. The predicted log odds ratio for any particular value of X can be translated back into a predicted blastocyst rate. Thus, for milk yield (X) = 35 in the low-fat group, the predicted log odds ratio (OR) would be log [OR] = 3.27 + (–0.152 x 35) = –2.0585. The corresponding predicted odds ratio would be: OR = exp(log [OR]) = exp (–2.0585) = 0.127645. The corresponding predicted blastocyst rate would be: OR / (1 + OR) = 0.127645 / (1 + 0.127645) = 0.113.

RESULTS

No difference was observed in milk yield, live weight, DMI, MEI, and starch intake between the two dietary groups (Table 2). The high-fat group had significantly higher intake of fat (P < 0.001) compared with the low-fat group (Table 2).


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  		TABLE 2. Effect of level of dietary fatty acids on milk production, live weight, and intake of dietary ingredients.

Follicle Development and Oocyte Quality and Development

Total and mean numbers of follicles observed and aspirated within the three size groups (small, medium, large), numbers of oocytes collected, matured, and fertilized in vitro are presented in Table 3. Animals in the low-fat group had a significantly (P < 0.01) higher total number of follicles than in the high-fat group (Table 3). This significant difference was maintained for both small (7.1 ± 0.3 vs. 6.1 ± 0.3; P < 0.05) and medium follicles (9.3 ± 0.4 vs. 7.9 ± 0.3; P = 0.05). A total number of 1051 oocytes were aspirated with more oocytes being recovered from the low-fat group (P < 0.05). However, there was no significant difference in oocyte recovery rate between the two groups (Table 3).


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  		TABLE 3. Comparison (mean ± SEM) between follicular development and oocyte recovery in the low- and high-fat dietary groups.

Number and percentage of oocytes in the different grades (Grade 1–4) are presented in Table 4. The percentage of Grade 2 oocytes collected from the high-fat group was higher than from the low-fat group (P = 0.08). Although a higher number of grade 3 oocytes were collected from the low-fat group, this difference did not reach statistical significance. The percentage of oocytes which cleaved to >4 cell stage by 48 h after fertilization and percentage of cleaved oocytes that developed to the blastocyst stage by Day 8 are presented in Table 4. Oocytes recovered from the high-fat group had a higher cleavage rate (72.0% ± 2.2% vs. 66.6% ± 2.1%; P = 0.076) and significantly higher rate of development (27.4% ± 2.2% vs. 19.4% ± 1.8%, P < 0.005) than oocytes from the low-fat group. The difference in rate of blastocyst production from the number of cleaved embryos was also significant (38.0% ± 2.8% in the high-fat group vs. 29.1% ± 2.5% in the low-fat group; P < 0.05).


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  		TABLE 4. Effect (mean per cow per OPU ± SEM) of dietary fats on oocyte quality and development.

Oocytes recovered from the high-fat group produced blastocysts with higher numbers of total, ICM, and TE cells than the low-fat group (P < 0.05). No difference was observed in the ratio of ICM to TE cells (Table 5).


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  		TABLE 5. Effect (mean ± SEM) of dietary fat on embryo quality.

Relationships with Hormone Concentrations and Production Characteristics

There was no difference between treatments in concentrations of GH, insulin, IGF1, leptin, progesterone, and metabolites in the peripheral circulation except for NEFA, which was significantly higher in the low-fat group than in the high-fat group (0.28 vs. 0.22, SED 0.026 mmol/L; P < 0.05) (Table 6).


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  		TABLE 6. Effect of dietary fat on peripheral concentrations (mean ± SED) of hormones and metabolites.

Logistic regression analyses for assessing the effects of production parameters and peripheral hormones and metabolites on blastocyst yield from IVF are presented in Table 7 and Figure 4. Effects were different in the two dietary groups. Milk yield, DMI, MEI, starch, and fat intakes significantly affected blastocyst yield in the low-fat group (P < 0.001; Table 7); the rate of blastocyst production from IVF oocytes reduced with increasing milk production (Figure 4, A–C). In the high-fat group, none of these parameters affected blastocyst yield. In the low-fat group, milk yield, DMI, MEI, starch and fat intakes, GH, blood urea nitrogen, and NEFA were also negatively related to blastocyst yield whereas insulin and leptin were positively related to blastocyst yield.


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  		TABLE 7. Effect of diet and blood metabolites on blastocyst production from IVF oocytes.a


Figure 04
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  		FIG. 4. Effects of dietary fat level on responses in blastocyst yield to milk yield, dry matter intake (DMI) and starch intake and metabolic hormones (insulin, growth hormone [GH], leptin). Lines show relationships predicted by logistic regression. Blastocyst yields are back-transformed from predicted log odds ratios. Equations and significance levels are shown in Table 7.

Differences in Production Variables

Correlations of hormones and metabolites with milk yield, DMI, starch and fat intake, MEI and live weight, and metabolic factors are presented in Table 8. There were significant negative correlations between leptin concentration and DMI, starch intake, and MEI (P < 0.05). Negative correlation was also observed between fat intake and NEFA concentration in the blood (P < 0.05). Concentration of BHB was positively correlated with milk yield and starch intake. In addition, positive correlations were observed between milk yield and DMI (r = 0.429; P < 0.001), starch intake (r = 0.923; P < 0.001), fat intake (r = 0.429; P = 0.05) and MEI (r = 0.879; P < 0.001). Furthermore, significant positive correlations were observed between DMI and starch intake (r = 0.983; P < 0.001), fat intake (r = 0.519; P < 0.05), and MEI (r = 0.987; P < 0.001). Fat intake was positively correlated with MEI (r = 0.647; P < 0.005), but a negatively correlated with NEFA (r = –0.446; P < 0.05).


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  		TABLE 8. Correlations of milk yield, feed intake, and live weight with peripheral concentrations of hormones and metabolites.*

Serum Amino Acid Concentrations

Mean (± SED) serum concentrations of essential amino acids are shown in Table 9. There was no significant difference between cows fed high-fat versus low-fat diets for any of the amino acids measured. However, there were tendencies for higher concentrations of glutamic acid and alanine in the high-fat group than the low-fat group.


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  		TABLE 9. Effect of dietary fat level on peripheral concentrations (mean ± SED) of amino acids.

DISCUSSION

The main objective of this study was to determine the effect of level of dietary fatty acids during short-term feeding on the developmental potential of oocytes in high-yielding dairy cows. Large numbers of oocytes were recovered by OPU from cows allocated to both dietary groups. The overall cleavage rate of oocytes from the high-fat group was significantly higher than the low-fat group (Table 4). Importantly, the higher level of dietary fat significantly increased rate of blastocyst production from both IVF oocytes and cleaved embryos (Table 4). In addition, the higher level of dietary fat improved embryo quality through increases in total and TE cell numbers in blastocysts (Table 5).

The beneficial effects of increasing dietary fat are supported by previous observations that supplementation of fatty acids in diets influences concentrations of prostaglandins, steroid hormones, and growth factors [26, 46–49]. The resulting enhancement of follicular growth and luteal activity can improve fertility of dairy cows. There are also recent reports showing beneficial effects of feeding rumen bypass fatty acids during the dry period prior to parturition, and continuing into the postpartum period, on postpartum dynamics of PGF secretion, uterine health, and reproductive performance [50, 51], embryo development [52], and subsequent pregnancy rate [53] in dairy cows. These effects indicate complex interactions between nutrient availability and metabolic adaptation.

In the study reported here, cows fed a low-fat diet had a higher number of ovarian follicles. Accordingly, more oocytes were recovered from this group. Fat supplementation has been shown to stimulate follicular growth and increase total number of follicles [26, 54, 55]. Diets containing calcium salts of long-chain fatty acids have also been shown to increase the number and size of corpora lutea [56], possibly through increased concentrations of cholesterol in both plasma and follicular fluid [57]. Increase in size of preovulatory follicles influences concentration of progesterone from the resultant CL, and this is important for maintenance of pregnancy [58]. In the current experiment, there was a tendency for higher progesterone in the high-fat group, although this did not reach significance (Table 6).

The type and level of fatty acid supplementation in the diet may influence their effect on the reproductive system. For example, feeding diets high in linoleic acid during the prepartum period delays parturition in sheep [59] and increases the incidence of retained placenta in cattle [60]. In contrast, increased availability of {alpha}-linolenic acid during the postpartum period improved the rate of pregnancy in cattle [61]. Increasing levels of dietary fat increased the number of large follicles on Day 14 postpartum in dairy cows compared with a low-fat diet [55]. However, cows fed on a moderate fat diet had higher peak plasma estradiol during the first follicular wave and a shorter interval to first ovulation than cows fed on a high-fat diet [55]. In addition, an increase in oocyte fatty acid composition of cows fed different fat levels has been reported to be important for oocyte competence [22] and to be responsible for differences in fertilization and developmental potential [33]. As demonstrated in the current experiment, inclusion of higher levels of inert rumen fat in the diet may impair ovarian follicular development (Table 3) whilst improving oocyte quality (Tables 4 and 5).

In the present study, a higher percentage of Grade 2 oocytes were collected from the high-fat group. This grade of oocyte has been correlated with morphological atresia in the marginal cumulus cells, corresponding to preovulatory follicles having higher developmental competence to the blastocyst stage, possibly due to higher cytoplasmic maturation [62]. We have recently reported higher blastocyst yield from this grade of oocytes collected from follicles with slightly raised caspase-3 activity [63]. These follicles also had low levels of IGFBP-5 expression, indicating that they may not be atretic in any meaningful sense but, rather, may have a lower level of cell proliferation. The absence of IGFBPs other than IGFBP-3 in bovine preovulatory follicles may allow for increased bioavailability of IGFs, which is important for oocyte maturation [17].

In the present study, concentration of NEFA in the peripheral serum was significantly higher in the low-fat group, although the difference was small. Mean NEFA concentrations were lower than values seen during severe negative energy balance. Using mean milk yields and energy intakes, energy balance was calculated to be –8 MJ/day for the low-fat group and +1 MJ/day for the high-fat group. During the NEB period in postpartum dairy cows NEFA concentrations in serum increase [64], which may impair oocyte quality through increased NEFA concentrations in follicular fluid [65]. In vitro, the developmental potential of oocytes after maturation is significantly reduced in the presence of NEFA [66]. Furthermore, elevated levels of NEFA have a detrimental effect on both bovine [67] and human [68] granulosa cell proliferation and steroid production in vitro. One possible explanation for the positive effects of rumen inert fat found in the present study is that reduced NEFA concentrations had beneficial effects for follicular health and oocyte quality, leading to improved oocyte developmental potential.

Although there was no difference in mean concentrations of hormones and metabolites between the two dietary groups, significant relationships were found between hormones and metabolites and the number of blastocysts yielded from IVF (Figure 4). The results demonstrate a negative effect of milk yield, DMI, MEI, and starch intake, but a positive effect of insulin and glucagon on oocyte quality as assessed by blastocyst yield. Higher fat in the diet mitigated the adverse effects of high milk yield and high starch on blastocyst yield. Insulin plays a central role in metabolism by stimulating the utilization of glucose in peripheral tissues, such as muscle and adipose tissue, and promoting accumulation of reserves of glycogen and lipid [69]. High insulin is associated with pseudo-maturation and reduced rate of blastocyst production [35]. Insulin is increased by diets with high starch content [70] and is often decreased by diets with high fat content [56, 71, 72]. Therefore, increased fatty acid intake may neutralize the adverse effects of high insulin on oocyte development.

Plasma concentrations of total amino acids and individual amino acids did not vary between diets. Therefore, although amino acids have been shown to enhance embryo development, cell numbers in blastocysts, and oocyte maternal mRNA [73], the positive effects of rumen inert fat observed in the present study were not linked to changes in amino acids.

The total number of cells, number of TE cells, and ICM cells were higher in the high-fat group (Table 5), suggesting better quality blastocysts that may improve their subsequent development [74]. This effect is highlighted in a recent review by Thatcher et al. [51] that suggested a significant improvement in first service conception rate by including a fatty acid source (Megalac) in the diet. Embryo quality significantly influences production of IFN{tau} by bovine trophectoderm cells [13], which is the primary signal necessary for maternal recognition of pregnancy [75] and establishment of pregnancy.

Higher levels of dietary fat may influence either fatty acid content of cell membranes or cytoplasm of oocytes, thus affecting their developmental competence. Reports indicate seasonal variation in concentrations of fatty acids in follicular fluid [32]; this has been correlated with changes in supply of dietary sources of fatty acids and also low fertility of dairy cows during the summer [76]. Increased levels of polyunsaturated fatty acids during the winter may also influence the chilling resistance of ovine oocytes [77]. Oocytes gradually gain developmental potential during follicular growth and development if an optimal microenvironment is maintained. This is due to an improvement in cytoplasmic maturation through synthesis of vital ingredients, including proteins and mRNAs that are crucial for future development after fertilization [78]. Therefore inclusion of high fat in the diet may benefit oocytes not only during growth and development in vivo, but also during the short period of oocyte maturation, and may also benefit embryos during the preimplantation period.

In conclusion, the specific role of dietary fat on the structure and function of oocytes, and the mechanisms through which the developmental potential of oocytes is improved, require further investigation. However, this study clearly shows the beneficial effects of increased level of dietary fat on the developmental potential of oocytes in the high-yielding dairy cow.

ACKNOWLEDGMENTS

The authors would like to thank Mr. M. Mitchell, Mr. N. Saunders, and Mrs. H. Russell for their assistance during OPU and sample preparation, and also during insulin, glucagon, leptin, and GH assays. We acknowledge Mrs. D. Li for amino acid analysis.

FOOTNOTES

3Current address: Reproduction Research Group, The Royal Veterinary College, Hawkshead Lane, Hatfield AL9 7TA, United Kingdom. Back

1Supported by a strategic grant (LS3306: Increasing dairy cow fertility through the precise control of nutrition) from The Department for Environment Food and Rural Affairs (Defra), United Kingdom. Back

Correspondence: 2FAX: 44 11 595 16069; e-mail: bob.webb{at}nottingham.ac.uk

Received: 1 November 2006.

First decision: 7 December 2006.

Accepted: 21 February 2007.

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