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J. Anim Sci. 2007. 85:1675-1686. doi:10.2527/jas.2007-0022
© 2007 American Society of Animal Science
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ANIMAL GROWTH, PHYSIOLOGY, AND REPRODUCTION

Effects of dietary supplementation with flax during prepuberty on fatty acid profile, mammogenesis, and bone resorption in gilts1,2

C. Farmer*,3, H. V. Petit*, H. Weiler{dagger} and A. V. Capuco§

* Agriculture and Agri-Food Canada, Dairy and Swine R & D Centre, PO Box 90, Lennoxville Stn., Sherbrooke, QC J1M 1Z3, Canada; and {dagger} School of Dietetics and Human Nutrition, McGill University, Ste-Anne-de-Bellevue, QC H9X 3V9, Canada; and § USDA-ARS, Bovine Functional Genomics Lab, Beltsville, MD 20705


   Abstract
Top
 Abstract
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 
The possible role of dietary flax on pre-pubertal development of mammary glands and bone resorption was investigated in gilts. Fifty-seven gilts were fed 1 of 4 diets from 88 d of age until slaughter (d 212 ± 1). Diets were control without flax (n = 14); 10% flaxseed supplementation (n = 13); 6.5% flaxseed meal supplementation (n = 15); and 3.5% flaxseed oil supplementation (n = 15). All diets were isonitrogenous and isocaloric. Jugular blood samples were obtained on d 78 and 210 to establish the fatty acid profile and to determine the concentrations of prolactin, estradiol, and cross-linked N-telopeptides of type I collagen. At slaughter, the mammary glands were excised, parenchymal and extraparenchymal tissues were dissected, and the composition of the parenchymal tissue (protein, fat, DM, and DNA) was determined. Histochemical analyses of the mammary parenchyma were performed, and fatty acid profiles in the extraparenchymal tissue were evaluated. Dietary flax increased (P ≤ 0.001) the concentrations of PUFA and decreased those of SFA (P < 0.01) and MUFA (P ≤ 0.001) in plasma and extraparenchymal tissues, which was largely due to the inclusion of 10% flaxseed or 3.5% flaxseed oil (P ≤ 0.01) but not 6.5% flaxseed meal. Circulating concentrations of prolactin and estradiol were unaltered by treatments (P > 0.1), but concentrations of cross-linked N-telopeptides of type I collagen tended to be greater (P < 0.1) in flax-supplemented gilts. The DM content of parenchymal tissue was the only mammary compositional value affected, showing an increase with flax addition (P < 0.05). No change (P ≥ 0.1) in the bromodeoxyuridine labeling index or estrogen receptor localization was observed with treatments. Dietary supplementation with flax as seed, meal, or oil, therefore, brought about the expected changes in the fatty acid profile but had no beneficial effects on mammary development or bone resorption.

Key Words: bone resorption • flaxseed • mammary development • mammary gland • pig • prepubertal female


   INTRODUCTION
Top
 Abstract
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 
Milk yield of sows is a major determinant of piglet growth and is largely affected by the number of milk secretory cells present at the onset of lactation (Head et al., 1991Go). Feeding management of gilts during the early phase of mammary cell accretion (from 90 d of age until puberty; Sorensen et al., 2002Go) has an impact on mammary development (Farmer et al., 2004Go; Sorensen et al., 2006Go). Flaxseed is known to be the richest source of secoisolariciresinol diglycoside, which is a precursor for lignan formation. Lignans, in turn, influence endogenous hormone concentrations (Hutchins et al., 2001Go) and exhibit estrogenic activities (Adlercreutz et al., 1987Go). Estrogens are mediators of mammary gland development and differentiation in rodents (Russo et al., 1999Go) and are necessary for mammogenesis in gilts (Kensinger et al., 1986Go). Flaxseed was shown to alter mammary development in rodents, largely due to its estrogenic properties (Tou and Thompson, 1999Go). The consumption of a diet that is rich in PUFA may also alter mammary gland morphology and increases the number of estradiol receptor binding sites in the mammary gland of virgin mice (Hilakivi-Clarke et al., 1998Go). Feeding long chain PUFA also had beneficial effects on bone metabolism in rats (review by Watkins et al., 2001Go), yet a diet rich in flaxseed oil did not alter bone mass in piglets (Weiler and Fitzpatrick-Wong, 2002bGo).

The current study was undertaken to determine whether dietary supplementation with flaxseed during the prepubertal period affects mammogenesis, bone resorption, or both in gilts and, if so, whether this effect is related to the increased supply of lignans or to the oil component of flaxseed. Fatty acid profiles in blood and in mammary extraparenchyma were established to substantiate any possible effects of these diets on other measured variables and to clearly demonstrate the differential effects of the addition of flax as seed, meal, or oil on these profiles.


   MATERIALS AND METHODS
Top
 Abstract
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 LITERATURE CITED
 
Animals and Treatments

Animals were cared for according to a recommended code of practice (Agriculture and Agri-Food Canada, 1993Go). Fifty-seven Yorkshire x Landrace gilts were fed a commercial diet until 82 d of age. They were then divided into 4 nutritional regimens: 1) control without flax (n = 14), 2) 10% flaxseed supplementation (FS, n = 13), 3) flaxseed meal supplementation equivalent to regimen 2 on an oil-basis (FSM, n = 15), and 4) flaxseed oil supplementation equivalent to regimen 2 on an oil-basis (FSO, n = 15). Experimental diets were incorporated in the commercial diet in increasing daily amounts (25, 50, 50, 75, and 75% over 5 d) to reach 100% of the targeted dietary level on d 88. All diets were isonitrogenous and isocaloric based on calculated values (Table 1Go) and were fed ad libitum until slaughter at 212 ± 1 d (mean ± SE) of age. Representative feed samples were taken twice weekly throughout the experiment for compositional analyses (shown in Table 1Go). Fresh feed was given once daily, at 1300, and individual feed consumption was recorded daily from 83 d of age.


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  		Table 1. Actual ingredients, and calculated and analyzed composition of the 4 diets on an as-fed basis1
 
Gilts were housed in individual weaner pens (1.5 x 1.5 m) from d 90 to 123 and were then transferred to individual gestation pens (1.5 x 2.4 m) until d 151, at which time they were moved to individual gestation stalls (0.6 x 2.1 m) with a boar present in the room. Estrous behavior was evaluated daily beginning on d 151 by allowing physical contact between the gilts and a mature boar. All gilts had begun cycling before slaughter. Gilts were weighed and their backfat thickness was measured ultrasonically at the last rib (Scanmatic SM-1, Medimatic, Hellerup, Denmark) on d 86, 150, and 210. The experiment took place between November 2003 and April 2004.

Blood Sampling and Assays

Jugular blood samples were obtained at 0800 on d 78 and 210 to determine concentrations of estradiol, prolactin, and cross-linked N-telopeptides of type I collagen (NTx, d 210 only), and to establish a profile of plasma fatty acids (samples from 8 randomly selected animals per treatment were used). Samples collected for the prolactin and NTx assays were left at room temperature for 4 h, stored overnight at 4°C, centrifuged the following day, and serum was then harvested. Samples for estradiol and fatty acids measurements were collected in EDTA-containing tubes (Becton Dickinson and Cie, Rutherford, NJ), which were put on ice and centrifuged within 20 min, and plasma was immediately recovered. Serum and plasma samples were frozen at –20°C until they were assayed.

Concentrations of prolactin were determined using a previously described RIA (Robert et al., 1989Go). The radio-inert prolactin was donated by A.F. Parlow (US National Hormone and Pituitary Program, National Institute of Diabetes and Digestive and Kidney Diseases, Torrence, CA), and the first antibody to prolactin was purchased from Research Products International (Mt. Prospect, IL). Parallelism of a pool of serum from lactating sows with the standard curve was demonstrated. Average recovery, calculated by the addition of various doses of the radio-inert prolactin to 50 µL of a pooled sample, was 103.4%. Sensitivity of the assay was 1.5 ng/mL. Six samples of a representative pool of serum were carried in duplicates in all assays to calculate the CV. The intraassay CV, calculated from the mean values of the pools within each assay, was 3.9%. The interassay CV, calculated from the mean values of the pools obtained for different assays, was 7.5%.

Estradiol was measured using a commercial RIA kit (Comp. Diagnostic Systems Lab. Inc., Webster, TX). All samples were assayed in a single assay, with an intraassay CV of 1.21%. Concentrations of NTx were determined using a commercial ELISA kit (Osteomark NTx serum ELISA assay, Wampole Laboratories, Princeton, NJ) validated for swine serum. The assay was performed as recommended by the supplier, except that serum samples were diluted 1:75 (vol/vol) in assay buffer. The accuracy of the kit controls was 97% of the expected values, and agreement between duplicate samples ranged from 91 to 97%. Sensitivity for this assay was 3.2 mM bone collagen equivalents.

Fatty acids were extracted according to the method described by Cruz-Hernandez et al. (2004)Go, and preparation of plasma fatty acid methyl esters was carried out as described by Folch et al. (1957)Go. Fatty acid methyl ester profiles were measured at a split ratio of 1:60 on a Hewlett-Packard 6890N chromatograph (Hewlett-Packard Ltée, Montréal, Québec, Canada) equipped with an autosampler (Model 7683B), a flame ionization detector, and a Supelco SP-2380, fused silica, capillary column (60 m x 0.25 mm, 0.2-µm film thickness). The column parameters were as follows: an initial column temperature of 148°C was maintained for 5 min, the temperature was then programmed at 7.25°C/min to 240°C and remained at this temperature for 6.31 min. Injector and detector temperatures were 260 and 300°C, respectively. The carrier gas was helium at 0.8 mL/min. Hydrogen flow to the detector was 40 mL/min, airflow was 450 mL/min, and the flow of He2 make-up gas was 30 mL/min. Fatty acid peaks were identified using pure methyl ester standards (#GLC-617, Nu-Chek Prep Inc., Elysian, MN) and Agilent ChemStation software [version B.01.01(164), Hewlett-Packard Ltée]. All chemicals and solvents were analytical grade.

Mammary Gland Measurements and Organ Sampling

At slaughter, the mammary glands were excised from the abdominal wall of all gilts, and extraparenchymal tissue was obtained from the fourth cranial mammary glands for determination of the fatty acid profile, as described previously for plasma. These samples were stored at –80°C until analysis. The mammary glands were then stored at –20°C until dissection and analyses for tissue composition. Frozen mammary glands were trimmed of skin and teats and subsequently stored at –20°C. They were then cut into 2-cm-thick slices, and the mammary parenchymal tissue from each slice was dissected from the surrounding adipose (i.e., the extraparenchymal tissue) at 4°C. Parenchymal and extraparenchymal tissue weights were recorded. Mammary parenchyma was defined as containing epithelial tissue, but it also contained a large amount of adipose and connective tissues because the mammary ducts and alveolar tissues are embedded within loose connective tissue of the mammary fat pad in gilts of such young age.

Parenchymal tissue from all sliced and dissected glands was homogenized, and a representative sample was used for determination of composition by chemical analysis. The DNA content of parenchymal tissue was evaluated using a method based on fluorescence of a DNA stain (Labarca and Paigen, 1980Go). The DM, protein, and lipid contents were also measured (AOAC, 1998Go). Six gilts per treatment received an i.v. injection of 5 mg/kg (diluted in saline at a concentration of 20 mg/mL and a pH of 8 to 8.5) of 5-bromo-2'-deoxyuridine (BrdU; Sigma Chemical, St. Louis, MO) in the marginal ear vein, 2 to 3 h before slaughter. Three cubes of parenchyma (~3 x 3 mm) from these gilts were obtained from 2 regions within the gland and used to determine the rate of cell proliferation using BrdU incorporation. Tissues were collected from the second cranial mammary gland and were fixed in 10% formalin in phosphate-buffered saline at 4°C for 24 h before being transferred to 70% ethanol until further processing. They were then dehydrated through a graded series of ethanol to 100% ethanol and embedded in paraffin according to standard techniques. Embedded tissue was later sectioned at 5-µm for immunohistochemical staining and evaluation.

Ovaries were also collected from all gilts, and the number of corpora lutea was determined. A 5-g sample from the left lateral lobe of the liver was obtained from animals that were not previously injected with BrdU. These were immediately frozen in liquid nitrogen and stored at –80°C until measures of fatty acids were performed.

Total lipids were extracted from the liver samples as described by Folch et al. (1957)Go and were then homogenized before fatty acids were measured according to the methodology of Mollard et al. (2005)Go. Only values for fatty acids that are involved in bone metabolism are reported.

Immunohistochemistry

Slides were dewaxed in xylene and hydrated in a graded series of ethanol to PBS (pH 7.4). Tissue sections were quenched with 3% H2O2 in PBS for 10 min and then washed in PBS (3 x 2 min). Microwave antigen retrieval was then used. Briefly, the tissue sections were heated in a microwave at high power (650 W) in 400 mL of 10 mM citrate buffer (pH 6.0) for 5 min, remained undisturbed for 5 min, and then were microwaved for an additional 5 min. Slides remained in the buffer for a 30-min cooling period. They were then washed in PBS (3 x 2 min) and blocked with 5% nonimmune goat serum in PBS (30 min) before overnight incubation at 4°C with primary antibody. Brightfield microscopic detection of BrdU-labeled cells was performed as described previously (Capuco et al., 2002Go). Briefly, after overnight incubation with BrdU antibody (1:50 vol/vol dilution; clone BMC 9318, MAB3424, Chemicon International Inc., Temecula, CA), tissue sections were washed in PBS. Sections were incubated at room temperature with biotinylated secondary antibody, washed in PBS, and then incubated with a steptavidin-peroxidase-conjugate (Histostain SP kit, Zymed Laboratories, San Francisco, CA). After washing in PBS, sections were incubated with diaminobenzidine, counterstained with hematoxylin, and mounted with Permaslip (Alban Scientific Inc., St. Louis, MO).

To quantify the number of BrdU-labeled epithelial cells, photographs of the stained tissue sections were captured as digital images. A slide was prepared and processed from each of 2 parenchymal regions per gilt. For each slide, 10 tissue areas were photographed with a Spot digital camera (Diagnostic Instruments Inc., Sterling Heights, MI) on a Zeiss Axioskop microscope (Carl Zeiss Inc., Thornwood, NY) with the 40x objective. The number of BrdU-labeled epithelial cells and the total number of epithelial cells were determined manually. At least 1,500 epithelial cells were scored per gilt (average ± SE = 4,049 ± 289).

Immunohistochemical localization of estrogen receptor-alpha was performed on 3 randomly-selected gilts per dietary treatment using a rabbit polyclonal antibody [sc-787 at 1:100 dilution (vol/vol), Santa Cruz Biotechnology Inc., Santa Cruz, CA]. Tissue sections were prepared for immunohistochemistry as described above. After overnight incubation with the primary antibody, sections were washed in PBS (3 x 2 min at room temperature) and then incubated with second antibody-horse radish peroxidase conjugate (30 min at room temperature). Sections were washed in PBS and then were incubated with diaminobenzidine, counterstained with hematoxylin, and mounted with Permaslip.

Statistical Analyses

The MIXED procedure (SAS Inst. Inc., Cary, NC) was used for statistical analyses. The univariate model used for mammary gland and ovarian variables included the effect of nutritional treatment, with the residual error being the error term used to test the main effects of treatment. Analyses on weights of mammary extraparenchymal and parenchymal tissues were done using the BW of the gilts at slaughter as a covariate. Repeated measures ANOVA with the factors treatment (the error term being gilt within treatment) and day of age (the residual error being the error term) and the treatment x day of age interaction were carried out on backfat thickness, BW, feed intake, and hormonal and fatty acids data. Orthogonal contrasts set a priori were used to compare nutritional regimens in the following manner: control vs. all flax-based diets (FS, FSM, and FSO), FSM (no oil) vs. FS and FSO (oil), and FS (oil and lignans) vs. FSO (oil). Data were corrected with a logarithmic transformation (using natural logarithms) when the variances were not homogeneous. The estrogen receptor data were categorical (visual scoring index from 1 to 4) and were therefore analyzed with the Cochran-Mantel-Haenszel test (Stokes et al., 1995Go). Data in the tables and figures are presented as least squares means ± the maximal SEM.


   RESULTS
Top
 Abstract
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 LITERATURE CITED
 
Performance, Hormonal, and NTx Data

Feed intake of gilts had a tendency (P = 0.091) to be greater in flax-supplemented gilts, averaging 2.81, 2.99, 2.90, and 2.94 ± 0.07 kg/d for control, FS, FSM, and FSO, respectively, over the whole experimental period (data not shown). There was an effect of week on feed intake with values increasing as gilts got older (P < 0.001), and there was no treatment x week interaction (P = 0.99). Gilts were heavier and had more backfat as they got older (P < 0.001), but there were no effects (data not shown) of treatment or treatment x day on BW (P = 0.93 and 0.80 for treatment and treatment x day, respectively) and backfat thickness (P = 0.53 and 0.51 for treatment and treatment x day, respectively). Average weights of gilts across all treatment groups were 45.6 ± 0.65, 107.1 ± 1.23, and 163.4 ± 2.41 kg at 86, 150, and 210 d of age, respectively. Backfat thicknesses for the same days were 6.8 ± 0.1, 13.9 ± 0.4, and 20.3 ± 0.7 mm, respectively. The number of corpora lutea on the ovaries were similar across treatments (data not shown) averaging 8.1 ± 1.3 and 9.0 ± 1.5 on the right (P = 0.31) and left (P = 0.55) ovaries, respectively. Hormonal and NTx data are presented in Table 2Go. There were no significant effects of dietary treatments on concentrations of estradiol, prolactin, or NTx.


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  		Table 2. Circulating concentrations of estradiol, prolactin, and cross-linked N-telopeptides of type I collagen (NTx) in crossbred gilts fed 1 of 4 diets from 88 d of age until slaughter (d 212 ± 1)1
 
Fatty Acid Profiles

Tables 3Go and 4Go show the profiles of various fatty acids in the plasma and in mammary extraparenchymal tissue, respectively. Plasma concentrations of the various fatty acids measured did not vary between groups before the onset of treatments (d 78, data not shown), however, following treatments (d 210), concentrations of numerous fatty acids were altered by dietary treatments. The inclusion of flax in the diet of gilts increased (P ≤ 0.001) concentrations of PUFA in plasma while decreasing those of SFA (P < 0.01) and MUFA (P ≤ 0.001) fatty acids. These effects were seen with the FS or FSO, but not FSM diets, as shown by this significant contrast (P ≤ 0.01). Concentrations of PUFA were greater in plasma of gilts fed FSO than in plasma of gilts fed FS (P < 0.01), yet numerical differences were very small. When looking at fatty acids on an individual basis, there were some notable changes. The only SFA affected by dietary treatment (P ≤ 0.001) was C16:0, with the decrease in its concentration being due mainly to the inclusion of FS or FSO in the diet. Feeding flax-seed products decreased (P < 0.05) C18 MUFA concentrations in plasma. Moreover, all C18 MUFA decreased (P ≤ 0.001) with the inclusion of FS or FSO in the diet, and the comparison of FSO with FS induced only minor changes. In contrast, dietary supplementation with FSO increased (P ≤ 0.001) plasma C20:1 compared with FS. The response of individual PUFA to dietary treatments varied, and C18:2cis-9,cis-12, which was present in greatest amount, increased (P < 0.05) with the addition of flax. The PUFA present in second greatest amount, C20:4cis-14 (arachidonic acid), was lowered (P ≤ 0.001) by dietary supplementation with flax. Feeding FSO resulted in the greatest concentrations of C20:5cis-17 (eicosapentaenoic acid, EPA), C22:5cis-19 (docosapentaenoic acid, DPA), and C22:6cis-19 (docosahexaenoic acid, DHA). On the other hand, feeding FS led to the greatest concentrations of C18:3cis-15 (alpha linolenic acid). Some individual n-6 (omega-6) fatty acids concentrations were altered by flax. The one present in the largest amount (C18:2cis-9,cis-12) increased (P < 0.05), whereas the other ones that were affected decreased (P < 0.05, Table 3Go). Feeding flax increased (P ≤ 0.001) plasma concentrations of total n-3 fatty acids and had no effect on total n-6 fatty acids concentrations (P = 0.14), thus resulting in a decrease in the ratio of n-6 to n-3 fatty acids. This decrease in the n-6 to n-3 fatty acids ratio was more important with the FS and FSO diets than with FSM.


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  		Table 3. Plasma fatty acid composition at 210 d of age in gilts fed 1 of 4 diets from 88 d of age until slaughter (d 212 ± 1)1
 

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  		Table 4. Fatty acid composition of mammary extraparenchymal tissue in gilts fed 1 of 4 diets from 88 d of age until slaughter (d 212 ± 1)1
 
Changes in mammary extraparenchymal tissue were similar to those seen in plasma, with SFA (P < 0.01) and MUFA (P ≤ 0.001) decreasing and PUFA increasing (P ≤ 0.001) in response to dietary addition of flax. Once more, the effect was mainly due to the addition of FS or FSO compared with FSM. Not all SFA were affected by flax supplementation, but those that were always decreased with the addition of flax (P ≤ 0.001). The same was true for MUFA. In the case of PUFA, there were more specific fatty acid responses. Feeding flax decreased (P < 0.001) concentrations of arachidonic acid and increased those of EPA (P = 0.001) and DPA (P = 0.001), and tended to increase those of DHA (P = 0.072). Feeding FSO resulted in the greatest concentrations of EPA, DPA, and DHA in mammary extraparenchymal tissue of gilts. As was the case for plasma values, the n-6/n-3 fatty acids ratio in mammary extraparenchymal tissue decreased with flax (P ≤ 0.001) and this was most drastic with FS and FSO addition. The n-6 fatty acids were once again unaltered (P = 0.23) by treatments, whereas the n-3 fatty acids increased (P ≤ 0.001), largely due to FS and FSO.

Flax addition had no effect on hepatic concentrations of C18:2 (P = 0.70) and DHA (P = 0.25, Table 5Go). However, feeding flax increased hepatic concentrations of linolenic acid and EPA (P < 0.01), and feeding FS or FSO decreased those of arachidonic acid (P < 0.01). Gilts fed FSO had lower concentrations of arachidonic acid, linolenic acid, EPA, and DHA (P < 0.05) in hepatic tissue than those fed FS.


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  		Table 5. Fatty acid composition of hepatic tissue in gilts fed 1 of 4 diets from 88 d of age until slaughter (d 212 ± 1)1
 
Mammary Gland Data. Composition of mammary tissue at slaughter and BrdU incorporation are shown in Table 6Go. The DM content was the only compositional variable of mammary glands that was affected, and it increased with dietary flax (P < 0.05). The BrdU labeling showed that nearly all paraffin sections had ducts with secretions, and many also had additional branching and alveolar development in regions of the gland. However, there was considerable variability among gilts, and there did not seem to be any relationship to dietary treatments (P = 0.1). Visual evaluation of estrogen receptor staining (Figure 1Go) indicated that estrogen receptor-alpha was present in epithelial cells and some fibroblasts but not in those adipocytes that were present within the parenchymal region of the gland. There were few or no estrogen receptors in the stromal cells immediately around the epithelium, yet stromal cells that were positive for estrogen receptor could be found in the dense connective tissue. Epithelial cells positive for estrogen receptor were evident in mammary tissue from all dietary treatments, but there was no treatment difference (P = 0.30). Gilts in estrus (± 2 d) were omitted from these analyses to reduce sample variation, largely due to changes in circulating estrogens at the time of estrus.


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  		Table 6. Composition and BrdU incorporation of mammary glands from crossbred gilts fed 1 of 4 diets from 88 d of age until slaughter (d 212 ± 1)1
 

Figure 1
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  		Figure 1. Immunohistochemical detection of bromodeoxyuridine-labeled cells and cells expressing estrogen receptor-alpha (ER{alpha}). Panel A: Cells that incorporated bromodeoxyuridine in vivo exhibit strong nuclear labeling. Examples of labeled epithelial cells are indicated by arrows. Panels B and D: Cells expressing ER{alpha} exhibited strong nuclear staining. The ER{alpha}-positive cells were primarily epithelial cells (open arrows), although a portion of cells within the dense connective tissue (DCT) were ER{alpha}-positive (filled arrows). Cells in the loose connective tissue (LCT) and adipocytes were ER{alpha}-negative. Panel C: Nonspecific staining in the absence of primary antibody (negative control) is depicted. Magnification bar = 50 µm.

 

   DISCUSSION
Top
 Abstract
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
LITERATURE CITED
 
The use of supplementary flaxseed in swine diets to improve carcass quality via alterations in fatty acid profiles of various tissues and organs is well documented (Cunnane et al., 1990Go; Enser et al., 2000Go; Kouba et al., 2003Go) and is corroborated by the present findings, whereby feeding flaxseed or flaxseed oil diets increased the concentrations of n-3 PUFA in plasma, fat, and liver of growing gilts. The reported beneficial effects of flax on fatty acid profiles were further defined in the current study by also showing decreases in SFA, MUFA, and n-6 fatty acids in plasma, using a more complete fatty acid profile than previous studies (Kouba et al., 2003Go). The greater and lesser concentrations of PUFA and MUFA, respectively, in plasma of gilts fed FS and FSO than in plasma of those fed FSM demonstrate that these effects were largely due to the inclusion of FS or FSO, but not FSM. The fact that these effects were mainly due to the oil fraction of flax (i.e., FS or FSO, but not FSM) corroborates reported findings that fatty acid profiles in various swine tissues reflect the fatty acid profile of the diet (Kouba et al., 2003Go). Indeed, in the present case, the FSM diet contained animal fat in order to be isoenergetic to the other diets. Changes in fatty acid profiles of extraparenchymal mammary tissue were similar to those seen in plasma and corroborate previous findings in adipose tissue (Specht-Overholt et al., 1997Go; Kouba et al., 2003Go). One difference, however, between results in plasma and extraparenchymal mammary tissue was the finding that concentrations of many SFA were lowered by flax-based diets in extraparenchymal tissue, whereas only one was affected in plasma. Reasons for this difference are not known, but these fatty acids represent a very small portion of the total SFA, and similar changes were seen in plasma and extraparenchymal tissue for the predominant SFA, C16:0. On the other hand, the lowered MUFA in mammary extraparenchyma with dietary flax may be linked to alterations in lipogenic enzyme activities. Indeed, Kouba et al. (2003)Go showed that the reduced MUFA in backfat of flax-fed pigs was due to a reduction in stearoyl-CoA-desaturase, an enzyme that generates MUFA from SFA. In an earlier study, Kouba and Mourot (1998)Go also noted that when pigs were fed control and experimental diets that were isocaloric (as was the case in the current study), the activities of malic enzyme and glucose-6-phosphate-dehydrogenase, the main enzymes involved in supplying NADPH for the reductive biosynthesis of fatty acids, were increased in adipose tissue by dietary linoleic acid. This further supports the possible role of enzymes in determining the fatty acid response to supplemental dietary fat. It is interesting to note that, irrespective of the diet, the concentration of MUFA was much greater and that of PUFA much lower in extraparenchymal mammary tissue compared with plasma, but that the total of the 2 was similar. The greater MUFA is likely because fat depots are the main sites for stearoyl-CoA-desaturation (Kouba et al., 2003Go). In agreement with our findings, these last authors also noticed that plasma of pigs fed control or experimental diets contained a greater percentage of PUFA and lower percentages of SFA and MUFA compared with adipose and muscle tissues.

The fact that growth rate and backfat thickness of gilts were not affected by dietary treatments corroborates previous findings in finishing barrows using 5 or 10% ground flaxseed (Sasaki et al., 2005Go) and in growing gilts fed 6% crushed flaxseed (Kouba et al., 2003Go). Flaxseed oil was recently shown to reduce palatability of diets for piglets (Solá-Oriol et al., 2006Go), yet no data on daily feed intake were reported. Present results clearly indicate that feed intake is not lowered by the inclusion of 10% flax as seed, meal, or oil in the diet of growing gilts.

Over the last decade, a beneficial role of n-3 fatty acids for bone metabolism was suggested (Watkins et al., 2003Go), yet most of these studies used fish oil as a source of fatty acids. Dietary supplementation with arachidonic acid also showed beneficial (Weiler, 2000Go) or no effect (Weiler and Fitzpatrick-Wong, 2002aGo) on bone mineral status of piglets. More recently, the effect of flaxseed oil on bone development of growing mice was investigated, and even though concentrations of linolenic acid, EPA, and DHA were increased in serum, there was no effect on bone mass or strength (Cohen and Ward, 2005Go). Present results concur with these findings because increases in plasma concentrations of these fatty acids were also observed with dietary flax supplementation, whereas concentrations of NTx, which is a highly bone-specific parameter of bone collagen degradation and serves as an indicator of bone resorption (Weiler et al., 2001Go), were not decreased. Weiler and Fitzpatrick-Wong (2002b)Go also noted that dietary flaxseed oil caused a reduction in the n-6/n-3 ratio and increased plasma DHA in piglets, which was associated with less bone resorption (decreased NTx) and normal bone growth. Reasons for the discrepancy between NTx results from the latter study and the current study are likely related to the rapid bone modeling present during early development compared with that in growing gilts. Nonetheless, the fact that there was no significant change in NTx concentrations in gilts from the current study leads us to think that the supplementary flax did not alter bone resorptive activity, although enhancing n-3 PUFA in the tissues.

The present finding that dietary flax, whether fed as seed, meal, or oil, had no beneficial impact on mammary development of gilts is novel and indicates that neither the alteration in fatty acid profile nor the likely presence of secoisolariciresinol diglycoside had an effect. Effects of dietary fatty acids should have been most noted with FS or FSO diets and effects of phytoestrogens with FS or FSM diets. The small increase in DM content of parenchymal tissue seen with flax-based diets is most unlikely to have any effect on lactogenic potential of the mammary glands. Because gilts were at different stages of the estrous cycle when slaughtered, our ability to detect mammogenic effects was somewhat blunted. Nonetheless, it is apparent that the estrogenic activity from the lignans was insufficient to have an impact on mammogenesis. Indeed, concentrations of estradiol were unchanged, and there seemed to be no effect on expression of estrogen receptor in mammary tissue. This is in agreement with findings of Tou and Thompson (1999)Go, where significant changes in mammary structures of virgin mice were only seen when serum estradiol concentrations were increased by dietary flax, namely, after a longer period of treatment (i.e., gestation, lactation and postweaning compared with postweaning only) and using a higher dose (10 vs. 5% FS). The dose and the timing of exposure had an impact on mammary development in their study, with a 5% flax addition having no effect. These authors stated that the different endocrine effects produced by the low vs. high FS doses may have been responsible for the different effects on the mammary gland. It could therefore be that the 10% FS supplementation in the current study was not sufficient to bring about anticipated changes. Developmental stage of the animals when subjected to flax supplementation may also be an important factor in determining its effect, given that Tan et al. (2004)Go noted an enhancement of mammary gland morphogenesis when rats suckled dams that were fed a 10% flaxseed diet throughout gestation and lactation. It could therefore be that exposure to flaxseed at an earlier stage of development in swine may have some effects. Ip et al. (2001)Go also reported that age (i.e., mammary gland proliferative status) altered the response of mammary epithelial cells to CLA, with a loss in sensitivity being apparent as the rats got older. It is important to keep in mind that gilts in the present trial varied in the stages of their estrous cycle and that this could have masked subtle changes due to the experimental diets.

To the best of our knowledge, the localization of estrogen receptor in cells in mammary tissue of gilts was never documented, and present findings are the first to report that estrogen receptors are mostly found in epithelial cells, but not adipocytes, of pubertal gilts. This corroborates findings in humans (Anderson et al., 1998Go), rats (Russo et al., 1999Go), and cattle (Capuco et al., 2002Go), in which estrogen receptors are localized within a subpopulation of mammary epithelial cells, but differs from those in mice, in which estrogen receptors are expressed in epithelial and stromal cells of mammary tissue (Shyamala, 1997Go).

In conclusion, there is no indication that dietary supplementation with 10% flaxseed or its equivalent in flaxseed meal or flaxseed oil stimulates mammary development or bone metabolism in growing gilts. However, the expected beneficial changes in fatty acid profiles in plasma and mammary extraparenchymal tissue were seen with the flaxseed and flaxseed oil diets because of the difference in oil composition of the diets. In conclusion, dietary supplementation with flax increased the concentrations of PUFA and decreased the n-6/n-3 ratio in blood and mammary extraparenchyma, but had no detrimental effects on mammogenesis or bone resorption in young gilts.


   Footnotes
 
1 Lennoxville Dairy and Swine R & D Centre contribution No. 918. Back

2 The authors thank L. Thibault, S. Horth, L. Veilleux, L. Marier, S. Dallaire, and Frédéric Morel for their invaluable technical assistance; A. Hummel and D. L. Wood for assistance with immunohistochemistry, the staff of the Swine Complex, especially D. Morissette, C. Boudreau, E. Bérubé, and T. Marsh for care and slaughter of the animals, S. Méthot for statistical analyses, and R. Martineau for scientific advice while writing the manuscript. Sincere thanks to Shur-Gain (Brossard, Canada) for supplying the feed for this project and to M. Vignola for formulating the diets. Back

3 Corresponding author: farmerc{at}agr.gc.ca

Received for publication January 11, 2007. Accepted for publication March 19, 2007.


   LITERATURE CITED
Top
 Abstract
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 LITERATURE CITED
 


Adlercreutz, H., K. Höckerstedt, C. Bannwart, S. Bloigu, E. Hämäläinen, T. Fotsis, and A. Ollus. 1987. Effect of dietary components, including lignans and phytoestrogens, on enterohepatic circulation and liver metabolism of estrogens and on sex hormone binding globulin (SHBG). J. Steroid Biochem. 27:1135–1144.[CrossRef][Medline]

Agriculture and Agri-Food Canada. 1993. Recommended code of practice for the care and handling of farm animals—Pigs. Publ. No. 1898E. Agric. Agri-Food Canada, Ottawa, Ontario, Canada.

Anderson, E., R. B. Clarke, and A. Howell. 1998. Estrogen responsiveness and control of normal human breast proliferation. J. Mammary Gland Biol. Neoplasia 3:23–35.[CrossRef][Medline]

AOAC. 1998. Official Methods of Analysis International. 16th ed. Vol. 2. Assoc. Off. Anal. Chem., Arlington, VA.

Capuco, A. V., S. Ellis, D. L. Wood, R. M. Akers, and W. Garrett. 2002. Postnatal mammary ductal growth: Three-dimensional imaging of cell proliferation, effects of estrogen treatment, and expression of steroid receptors in prepubertal calves. Tissue Cell 34:143–154.[CrossRef][Medline]

Cohen, S. L., and W. E. Ward. 2005. Flaxseed oil and bone development in growing male and female mice. J. Toxicol. Environ. Health. Part A 68:1861–1870.[CrossRef][Medline]

Cruz-Hernandez, C., Z. Deng, J. Zhou, A. R. Hill, M. P. Yurawecz, P. Delmonte, M. M. Mossoba, M. E. R. Dugan, and J. K. G. Kramer. 2004. Methods for analysis of conjugated linoleic acids and trans-18:1 isomers in dairy fats by using a combination of gas chromatography, silver-ion thin layer chromatography/gas chromatography, and silver-ion liquid chromatography. J. AOAC Int. 87:545–562.[Medline]

Cunnane, S. C., P. A. Stitt, S. Ganguli, and J. K. Armstrong. 1990. Raised omega-3 fatty acid levels in pigs fed flax. Can. J. Anim. Sci. 70:251–254.

Enser, M., R. I. Richardson, J. D. Wood, B. P. Gill, and P. R. Sheard. 2000. Feeding linseed to increase the n-3 PUFA of pork: Fatty acid composition of muscle, adipose tissue, liver and sausages. Meat Sci. 55:201–212.[CrossRef]

Farmer, C., D. Petitclerc, M. T. Sorensen, M. Vignola, and J. Y. Dourmad. 2004. Impacts of dietary protein level and feed restriction during prepuberty on mammogenesis in gilts. J. Anim. Sci. 82:2343–2351.[Abstract/Free Full Text]

Folch, J., M. Lees, and G. H. Sloane-Stanley. 1957. A simple method for the isolation and purification of total lipids from animal tissues. J. Biol. Chem. 226:497–509.[Free Full Text]

Head, R. H., N. W. Bruce, and I. H. Williams. 1991. More cells might lead to more milk. Page 76 in Manipulating Pig Production III. E. S. Batterham, ed. Australasian Pig Sci. Assoc., Attwood, Australia.

Hilakivi-Clarke, L., A. Stoica, M. Raygada, and M. Martin. 1998. Consumption of a high-fat diet alters estrogen receptor content, protein kinase C activity, and mammary gland morphology in virgin and pregnant mice and female offspring. Cancer Res. 58:654–660.[Abstract/Free Full Text]

Hutchins, A. M., M. C. Martini, A. Olson, W. Thomas, and J. L. Slavin. 2001. Flaxseed consumption influences endogenous hormone concentrations in postmenopausal women. Nutr. Cancer 39:58–65.[CrossRef][Medline]

Ip, C., Y. Dong, H. J. Thompson, D. E. Bauman, and M. Ip. 2001. Control of rat mammary epithelium proliferation by conjugated linoleic acid. Nutr. Cancer 39:233–238.[CrossRef][Medline]

Kensinger, R. S., R. J. Collier, and F. W. Bazer. 1986. Effect of number of conceptuses on maternal mammary development during pregnancy in the pig. Domest. Anim. Endocrinol. 3:237–245.[CrossRef]

Kouba, M., M. Enser, F. M. Whittington, G. R. Nute, and J. D. Wood. 2003. Effect of a high-linolenic acid diet on lipogenic enzyme activities, fatty acid composition, and meat quality in the growing pig. J. Anim. Sci. 81:1967–1979.[Abstract/Free Full Text]

Kouba, M., and J. Mourot. 1998. Effect of a high linoleic acid diet on {Delta}9-desaturase activity, lipogenesis and lipid composition of pig subcutaneous adipose tissue. Reprod. Nutr. Dev. 38:31–37.[Medline]

Labarca, C., and K. Paigen. 1980. A simple, rapid, and sensitive DNA assay procedure. Anal. Biochem. 102:344–352.[CrossRef][Medline]

Mollard, R. C., H. R. Kovacs, S. C. Fitzpatrick-Wong, and H. A. Weiler. 2005. Low levels of dietary arachidonic and docosahexaenoic acids improve bone mass in neonatal piglets, but higher levels provide no benefit. J. Nutr. 135:505–512.[Abstract/Free Full Text]

Robert, S., A. M. B. de Passillé, N. St-Pierre, P. Dubreuil, G. Pelletier, D. Petitclerc, and P. Brazeau. 1989. Effect of the stress of injection on the serum concentrations of cortisol, prolactin, and growth hormone in gilts and lactating sows. Can. J. Anim. Sci. 69:663–672.

Russo, J., X. Ao, C. Grill, and I. H. Russo. 1999. Pattern of distribution of cells positive for estrogen receptor-{alpha} and progesterone receptor in relation to proliferating cells in the mammary gland. Breast Cancer Res. Treat. 53:217–227.[CrossRef][Medline]

Sasaki, K., S. K. Baidoo, and Q. Yang. 2005. Effect of ground flaxseed on the performance and carcass traits of finishing pigs. J. Anim. Sci. 83(Suppl. 1):165.

Shyamala, G. 1997. Roles of estrogen and progesterone in normal mammary gland development—Insights from progesterone receptor null mutant mice and in situ localization of receptor. Trends Endocrinol. Metab. 8:34–39.[Medline]

Solá-Oriol, D., E. Roura, and D. Torrallardona. 2006. Palatability of diets with different oil and fat sources in piglets. J. Anim. Sci. 84(Suppl. 1):242.

Sorensen, M. T., C. Farmer, M. Vestergaard, S. Purup, and K. Sejrsen. 2006. Mammary development in prepubertal gilts fed restrictively or ad libitum in two sub-periods between weaning and puberty. Livest. Sci. 99:249–255.[CrossRef]

Sorensen, M. T., K. Sejrsen, and S. Purup. 2002. Mammary gland development in gilts. Livest. Prod. Sci. 75:143–148.[CrossRef]

Specht-Overholt, S., J. R. Romans, M. J. Marchello, R. S. Izard, M. G. Crews, D. M. Simon, W. J. Costello, and P. D. Evenson. 1997. Fatty acid composition of commercially manufactured omega-3 enriched pork products, haddock, and mackerel. J. Anim. Sci. 75:2335–2343.[Abstract/Free Full Text]

Stokes, M. E., C. S. Davis, and G. G. Koch. 1995. Categorical data analysis using the SAS system, 2nd ed. SAS Institute Inc., Cary, NC.

Tan, K. P., J. Chen, W. E. Ward, and L. U. Thompson. 2004. Mammary gland morphogenesis is enhanced by exposure to flaxseed or its major lignan during suckling in rats. Exp. Biol. Med. 229:147–157.[Abstract/Free Full Text]

Tou, J. C. L., and L. U. Thompson. 1999. Exposure to flaxseed or its lignan component during different developmental stages influences rat mammary gland structures. Carcinogenesis 20:1831–1835.[Abstract/Free Full Text]

Watkins, B. A., Y. Li, H. E. Lippman, and S. Feng. 2003. Modulatory effect of omega-3 PUFA on osteoblast function and bone metabolism. Prostaglandins Leukot. Essent. Fatty Acids 68:387–398.[CrossRef][Medline]

Watkins, B. A., H. E. Lippman, L. Le Bouteiller, Y. Li, and M. F. Seifert. 2001. Bioactive fatty acids: Role in bone biology and bone cell function. Prog. Lipid Res. 40:125–148.[CrossRef][Medline]

Weiler, H. A. 2000. Dietary supplementation of arachidonic acid is associated with higher whole body weight and bone mineral density in growing pigs. Pediatr. Res. 47:692–697.[Medline]

Weiler, U., S. Finsler, U. Wehr, and R. Claus. 2001. Comparison of blood markers for the longitudinal monitoring of osteoclastic activity in the pig. J. Vet. Med. A 48:609–618.[CrossRef]

Weiler, H. A., and S. Fitzpatrick-Wong. 2002a. Dietary long-chain PUFA minimize dexamethasone-induced reductions in arachidonic acid status but not bone mineral content in piglets. Pediatr. Res. 51:282–289.[Medline]

Weiler, H. A., and S. C. Fitzpatrick-Wong. 2002b. Modulation of essential (n-6):(n-3) fatty acid ratios alters fatty acid status but not bone mass in piglets. J. Nutr. 132:2667–2672.[Abstract/Free Full Text]


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C. Farmer and H. V. Petit
Effects of dietary supplementation with different forms of flax in late-gestation and lactation on fatty acid profiles in sows and their piglets
J Anim Sci, August 1, 2009; 87(8): 2600 - 2613.
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