J. Anim Sci. 2008. 86:3014-3022. doi:10.2527/jas.2008-1086
© 2008 American Society of Animal Science
Impacts of linseed meal and estradiol-17β on mass, cellularity, angiogenic factors, and vascularity of the jejunum1
M. R. O’Neil,
G. P. Lardy,
L. P. Reynolds,
J. S. Caton and
K. A. Vonnahme2
Department of Animal Sciences, North Dakota State University, Fargo 58105
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Abstract
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To evaluate the estrogenic potential of the phytoestrogen secoisolariciresinol diglycoside (SDG) found in linseed meal (LSM) on jejunal mass, cellular proliferation, vascularity, and expression of angiogenic factors and their receptors, 48 ovariectomized ewes (54.6 ± 1.1 kg) were fed a diet containing 12.5% LSM for 0, 1, 7, or 14 d and implanted with estradiol-17β (E2) for 0, 6, or 24 h before tissue collection. Angiogenic factors and receptors measured included vascular endothelial growth factor (VEGF), VEGF receptor-1 (FLT), VEGF receptor-2 (KDR), fibroblast growth factor (FGF), FGF receptor 2 IIIc (FGFR), angiopoietin (ANG)-1, ANG-2, ANG receptor (Tie-2), endothelial nitric oxide synthase (eNOS), and soluble guanylate cyclase (sGC). There was a LSM x E2 interaction (P = 0.003) on the jejunal cellular proliferation index. Jejunal cellular proliferation increased (P < 0.002) in ewes not fed LSM and implanted with E2 for 6 or 24 h compared with ewes implanted for 0 h but did not increase when LSM was fed for 1, 7, or 14 d. Neither feeding LSM nor implanting ewes with E2 altered vascular area density (P > 0.75) or vascular surface area (P > 0.29) of the jejunal villi. Expression of mRNA for the angiogenic factors VEGF, FGF, FGFR, ANG-1, ANG-2, and Tie-2 were not altered (P > 0.33) by feeding LSM or implanting ewes with E2. Implanting ewes with E2 for 6 h increased (P = 0.04) eNOS expression compared with ewes implanted for 0 h. Feeding LSM and implanting ewes with E2 interacted to alter mRNA expression of FLT (P = 0.04), KDR (P < 0.001), and sGC (P = 0.04). When LSM was fed for 1, but not 0, 7, or 14 d, expression of FLT mRNA decreased (P < 0.03) when ewes were implanted with E2 for 24 h compared with ewes implanted for 0 or 6 h. Expression of KDR mRNA was suppressed in ewes fed LSM for 1 (P = 0.03) or 7 d (P = 0.0007) and implanted with E2 for 24 h compared with ewes implanted for 0 h. When LSM was fed for 14 d, implanting ewes for 6 h increased (P = 0.04) KDR expression compared with ewes implanted for 0 h. Ewes fed LSM for 0 and 1 d experienced an increase in sGC mRNA expression when implanted for 6 h (P = 0.001) compared with ewes implanted for 0 h. When implanted for 24 h, levels were similar (P = 0.80) to those observed when ewes were implanted for 0 h. Expression of sGC was not altered by E2 when LSM was fed for 1, 7, or 14 d (P > 0.11). The impacts of E2 and LSM on nutrient uptake and growth during physiologically important time points are unknown.
Key Words: angiogenesis • cellular proliferation • estrogen • linseed meal • phytoestrogen
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INTRODUCTION
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Lignans are diphenolic compounds with estrogen-like activity (Axelson et al., 1982
). Flaxseed (Linum usitatissimum) contains high quantities of the lignan secoisolariciresinol diglycoside (SDG) within the hull of the seed. The intestinal microflora (Axelson and Setchell, 1981) and gastrointestinal enzymes, such as phlorizin hydrolase (Day et al., 2000), metabolize SDG to the mammalian lignans enterodiol and enterolactone, which are then able to interact systemically with estrogen receptors (Axelson et al., 1982; Rickard et al., 1996; Begum et al., 2004).
Functional E2 receptors are present in intestinal epithelial cells (Thomas et al., 1993; Pfaffl et al., 2003) and feeding genistein, a phytoestrogen in soybeans, altered the intestinal crypt cell proliferation rate of piglets (Chen et al., 2005). Recent observations that estrogen receptors 
and β are present in the bovine jejunum (Pfaffl et al., 2003) suggest that estrogen or estrogenic compounds may influence nutrient uptake. As a variety of feedstuffs and forages contain varying amounts of phytoestrogens, there is a possibility that they may alter normal function of E2 in cycling or pregnant livestock. There are limited data on impacts of E2, or estrogenic compounds, on cellularity and vascularity of the gastrointestinal tract, particularly in the ruminant. We hypothesized that linseed meal (LSM), which contains SDG, would influence jejunal mucosal cellularity and vascularity in ruminants. Our objectives were to determine how length of LSM feeding and E2 exposure affect cellularity and vascularity of the jejunum in addition to the expression of the angiogenic factors vascular endothelial growth factor (VEGF), VEGF receptor-1 (FLT), VEGF receptor-2 (KDR), fibroblast growth factor (FGF), FGF receptor 2 IIIc (FGFR), angiopoietin (ANG)-1, ANG-2, ANG receptor (Tie-2), endothelial nitric oxide synthase (eNOS), and soluble guanylate cyclase (sGC) in ovariectomized ewes.
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MATERIALS AND METHODS
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All experimental protocols were approved by the North Dakota State University Institutional Animal Care and Use Committee.
Animals and Treatments
Ewes were housed indoors at the North Dakota State University Animal Nutrition and Physiology Center for the duration of the experiment (early December through mid January). Light-dark cycles were timer-controlled and were set to coincide with sunrise and sunset at this location. Forty-eight multiparous ewes (54.6 ± 1.1 kg initial BW) of mixed breeding were fed in groups of 9 to 11 in pens measuring 3.04 x 3.73 m, ovariectomized via midventral laparotomy (Reynolds et al., 1998a), and allowed to recover for at least 28 d before treatments were initiated. During the recovery period, ewes were fed a SDG-free diet (control diet; Table 1) until initiation of treatment. The feeding of a SDG-free diet following ovariectomy was carried out to ensure that any circulating endogenous estrogens as well as any dietary SDG was cleared from the body before treatments were initiated. Ewes were fed to meet their requirement for maintenance based on pen average BW (NRC, 1985). Ewes were weighed every 2 wk, had ad libitum access to water, and were fed once daily at 0800 h throughout the experiment.
Immediately before initiation of treatment, ewes were moved to individual pens measuring 1.52 m x 1.87 m. Ewes were weighed and stratified by BW into blocks (i.e., the 12 lightest ewes were assigned to block 1, the next 12 lightest ewes were assigned to block 2, etc.). Once stratified, ewes within a block were assigned randomly to each treatment group. Treatments were arranged as a 3 x 4 factorial. Main effects consisted of length of exposure to E2 via subcutaneous implant (0, 6, or 24 h) and length of feeding a diet containing 12.5% LSM (0, 1, 7, or 14 d; as-fed basis; Table 1
). The LSM diet was designed to only contain SDG from the inclusion of 12.5% LSM. Laboratory analysis of the LSM used to formulate the LSM diet indicated the SDG content of the LSM diet to be 14.53 mg SDG/g of feed (DM basis). Control and LSM diets were formulated to be similar in CP and NE. Both diets were pelleted and ewes were limit-fed to meet NEm requirements based on ewe metabolic BW (56 kcal/kg0.75; NRC, 1985). Ewes were either fed the control diet throughout the experiment (i.e., 0 d LSM feeding) or were fed the LSM diet for 1, 7, or 14 d before tissue collection. Ewes within each duration of LSM feeding were implanted with E2 for 0, 6, or 24 h before tissue collection. Thus, each treatment group consisted of 4 ewes. Ewes receiving E2 implants were locally anesthetized and implanted subcutaneously in the auxiliary region as validated by Johnson et al. (1997a). Implanted ewes each received 2 Silastic implants (3.35 mm i.d. x 4.65 mm o.d. x 15 mm length; Dow Corning, Midland, MI) each containing 50 mg of E2 (Sigma-Aldrich, St. Louis, MO). After implantation, the incision was closed with surgical staples and a topical antibiotic was applied to the incision site. The implants remained until tissue collection, at which time their presence was confirmed.
Tissue Collection
Exactly 1 h before tissue collection, ewes were injected via jugular venipuncture with bromodeoxyuridine (BrdU; Aldrich, Milwaukee, WI; 5 mg/kg of BW; Zheng et al., 1996; Johnson et al., 1997a,b). At time of tissue collection, ewes were stunned via captive bolt and exsanguinated. The viscera were removed from the body cavity and the jejunum was located as described previously (Scheaffer et al., 2004; Reed et al., 2007). Briefly, the distal 300 cm of the jejunum was divided into two 150-cm sections. The cranial 150-cm section was carefully removed with mesentery and with vasculature intact, placed in PBS, and immediately transported to the laboratory. To obtain the mass of the entire jejunum, the remaining jejunum was stripped of its contents, separated from the mesentery, and weighed. The mass of the caudal 150-cm section of the jejunum was doubled to account for the cranial 150-cm section that was previously removed for perfusion.
Approximately 10 cm of the proximal end of the cranial 150-cm section of jejunum was subsampled and snap-frozen in liquid N2 and stored at –70°C for later mRNA quantification. The remaining section of jejunum was fixed via vascular perfusion by methods adapted from Soto-Navarro et al. (2004) and Reed et al. (2007). Briefly, hemostats were used to isolate approximately 2 vascular arcades within the mesentery. A primary branch of the mesenteric artery was then dissected from the mesentery and catheterized using polyethylene tubing (PE-160; o.d. = 1.56 mm, i.d. = 1.14 mm; Becton Dickinson, Sparks, MD). Once catheterized, vessels were flushed with approximately 10 mL of PBS followed by perfusion with approximately 3 mL of Evans blue dye (0.05% in PBS, wt/vol; Sigma Chemical Co.) to allow visualization of the vascular area being perfused. Once the field was visualized, the Evans blue was flushed from the vessels with 10 to 20 mL of PBS. After flushing with PBS, the jejunum was perfuse-fixed with 10 to 15 mL of Carnoy’s solution (60% ethanol, 30% chloroform, and 10% glacial acetic acid). Following perfuse-fixing, the mesentery was carefully trimmed from the jejunum and the jejunum was immersed in Carnoy’s solution. Following fixation, the tissues were embedded in paraffin, sectioned to 4 µm, and affixed to glass slides for staining (Luna, 1968).
Diet and Tissue Analyses
Diet samples were analyzed for DM, ash, N, Ca, P (methods 930.15, 942.05, 990.02, 968.08, and 965.17, respectively; AOAC, 1990), ADF, and NDF (Ankom, Fairport, NY). In vitro OM digestibility was determined on diet samples by a modified procedure of Tilley and Terry (1963), in which samples were centrifuged (1,000 x g; time = 15 min), and the supernatant fluid was discarded before the addition of pepsin.
Jejunal mucosa was analyzed for DNA, RNA, and protein content as previously reported (Bradford, 1976; Johnson et al., 1997b). A single sample was used as a control in the DNA assay, an assay standard solution (12.5 mg of RNA/mL) was used as a control in the RNA assay, and the Accutrol normal control (Sigma-Aldrich) was used as a control in the protein assay. Intra- and interassay CV were 4.87 and 5.97%; 7.08 and 0.69%; 5.11 and 1.13% for DNA, RNA, and protein assays, respectively. The RNA:DNA ratio was used as an index of cellular activity, whereas the protein:DNA ratio was used as an index of hypertrophy.
Jejunal mRNA expression of VEGF, FLT, KDR, eNOS, sGC, FGF, FGFR2 IIIc, ANG-1, ANG-2, and Tie-2 was determined using quantitative real time-PCR as adapted from Redmer et al. (2005) and Vonnahme et al. (2006) following capillary electrophoresis of total cellular RNA using an Agilent 2100 Bioanalyzer (Agilent Technologies, Wilmington, DE) to determine quantity and quality of extracted total cellular RNA.
Perfuse-fixed jejunal tissues were sectioned to 4 µm and affixed to glass slides. Two staining protocols were used for jejunal tissues from each ewe to determine vascularity and cellular proliferation. Slides evaluated for cellular proliferation were deparaffinized in gradiated ethanol, rinsed in distilled water, bathed in 3% H2O2 for 5 min, rinsed in distilled water, bathed in 2 N HCl for 30 min, rinsed with distilled water, and then bathed in PBS with Triton X-100 (Sigma-Aldrich) for 10 min followed by a 10-min bath in PBS without Triton X-100. At this time, the slides were moved to a Dako Autostainer (Dako, Carpinteria, CA). The following staining protocol was programmed into the autostainer (unless it is specified, it can be assumed that the reagent was blown from the slide before the next step rather than completely rinsed from the slide): Tris buffered saline (TBS) rinse; 10-min protein block (serum free protein block; Dako); 60-min incubation with primary antibody (Envision mouse anti-BrdU diluted 1:150 with antibody diluent; Dako); TBS rinse; 30-min incubation with rabbit anti-mouse labeled polymer (Dako); TBS rinse and TBS was blown from the slide; 5-min rinse in PBS and PBS was blown from the slide; 10-min incubation with Vector SG substrate (Vector Labs, Burlingame, CA). The slides were then removed from the autostainer and rinsed in distilled water before being bathed in nuclear fast red for 15 min. This was followed by a final rinse, passage through graded ethanol, and mounted with coverslips. Slides evaluated for vascularity were prepared as described by Reed et al. (2007). Briefly, slides were deparaffinized via gradiated ethanol, rinsed in distilled water, and stained with periodic acid Schiff’s reagent for approximately 10 min.
Digital images of tissues were collected for both vascularity and proliferation data using a Nikon DXM 1200 digital camera (Fryer, Chicago, IL) and a image analysis software package (Image-Pro Plus, version 5.0; MediaCybernetics Inc., Silver Spring, MD). Proliferation was determined by image analysis of BrdU-stained cells in the crypt region of 6 fields for each ewe. The percentage of BrdU-stained nuclei of all nuclei present in the crypt region was determined to obtain a proliferation index. The middle third of individual jejunal villi was used to determine measurements of jejunal vascularity. Measures of vascular area density, number density, surface density, and area per capillary were collected from 10 jejunal villi per ewe and determined via image analysis.
Statistics
Data for ewe jejunal cellularity, cellular proliferation, vascularity, and angiogenic factor expression were analyzed as a 3 x 4 factorial in a randomized complete block design using PROC MIXED (SAS Inst. Inc., Cary, NC). To calculate jejunal mass as a percentage change from 0-h E2, the difference in jejunal mass of each ewe from the mean jejunal mass of the respective 0-h E2 groups within each duration of LSM was divided by the mean organ mass of the respective 0-h E2 group. Model statement included effects of block (based on initial BW), length of LSM feeding, length of E2 exposure, and the interaction of LSM feeding and E2 exposure. Fixed variables were length of LSM feeding and length of E2 exposure. The diagonal covariate structure was used. Data are presented as least squares means ± SEM. Mean separations were performed via LSD, which were protected by an overall treatment F-test at P = 0.05. Differences were considered significant if P 
0.05 unless otherwise stated.
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RESULTS
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Ewe BW were not different (P > 0.37) at the beginning or the end of the experiment, which averaged 52.1 ± 0.99 and 51.5 ± 0.97 kg, respectively (data not shown). Neither feeding LSM nor implanting ewes with E2 altered jejunal mass (g or g/kg; P 
0.11; Table 2), or the percentage of the jejunum that was mucosa (P 0.15). Although there was no interaction between LSM and E2 (P = 0.18) and no effect of E2 (P = 0.47), there was an effect of LSM (P < 0.001) on the percentage change of jejunal mass from 0-h E2 exposure (Figure 1). When ewes were fed LSM for 1 d, jejunal mass as a percentage from the 0-h E2 increased (P = 0.01) compared with when LSM was not fed. By 7 d of LSM feeding, jejunal mass decreased (P = 0.005) compared with feeding LSM for 1 d, but did not differ (P = 0.14) from 0-d LSM feeding. By 14 d of LSM feeding, jejunal mass as a percentage of the 0-h E2 group was reduced as compared with 0 d (P < 0.001) and 1 d (P < 0.001), and was similar (P = 0.11) to the 7-d group (Figure 1).
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Table 2. Effects of feeding linseed meal (LSM) and implanting ewes with estradiol-17β (E2) on jejunal weight, percentage mucosa, jejunal mucosal DNA, RNA, protein, RNA:DNA, and protein:DNA ratios, as well as vascular area density, surface density, number density, and area per capillary in jejunal villi1
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Figure 1. Effect (P < 0.0001) of linseed meal on jejunal mass as a percentage change from 0-h estradiol-17β (E2; difference in jejunal mass from the mean jejunal mass of the respective 0-h E2 groups within each duration of linseed meal divided by the mean jejunal mass of the respective 0-h E2 group). Error bars are SEM. a–cMeans not bearing a common letter differ (P 0.01).
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Neither feeding LSM, implanting ewes with E2, nor their interaction altered (P 0.11) mucosal DNA or RNA concentrations, RNA:DNA, or protein:DNA ratios (Table 2). However, there was a LSM x E2 interaction (P = 0.02) in mucosal protein concentration. For ewes fed LSM for 0 d, implanting with E2 for 6 h led to greater (P = 0.01) protein concentrations than implanting for 24 h. For ewes fed LSM for 1 d, implanting with E2 for 24 h led to greater (P = 0.02) protein concentrations than implanting for 6 h. There was no effect (P 0.47) of duration of E2 exposure in protein concentration when LSM was fed for 7 d. For ewes fed LSM for 14 d, implanting ewes with E2 for 6 h decreased (P = 0.02) protein concentration compared with no implant.
There was a LSM x E2 interaction (P = 0.003) on the cellular proliferation index within the crypt region of the jejunum (Figure 2). When ewes were fed LSM for 0 d, implanting E2 for 24 h increased the labeling index (BrdU-labeled cells/total cells within the crypt region of the jejunum) when compared with ewes implanted with E2 for 0 or 6 h (P = 0.002 and P < 0.001, respectively). However, when LSM was fed for 1, 7, or 14 d the increase in the labeling index seen in response to implanting ewes with E2 for 24 h was not observed. Furthermore, ovariectomized ewes fed LSM for any duration of time had reduced (P < 0.01) percentage of cellular proliferation when exposed to E2 for 24 h compared with ewes that were not fed LSM.
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Figure 2. Interaction between estradiol-17β (E2) and linseed meal (P = 0.003) on jejunal crypt cellular proliferation index (%). Black bars indicate 0-h E2 exposure; gray bars indicate 6-h E2 exposure; white bars indicate 24-h E2 exposure. Error bars are SEM. a–cMeans not bearing a common letter differ (P 0.05).
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There were no interactions (P 0.26) of LSM feeding or E2 exposure observed for any measurement of vascularity within the jejunum (Table 2). Neither feeding LSM nor implanting ewes with E2 altered measurements of vascular area density, vascular surface density, or area per vessel (P 0.20; Table 2). There was a tendency (P = 0.07) for 6-h E2 exposure (522 ± 44) to decrease vascular number density compared with 0 h (666 ± 47) and 24 h (631 ± 44) E2 exposure, which did not differ.
There was no effect (P 0.26) of LSM or E2 on FGF, FGFR, ANG-1, ANG-2, or Tie-2 expression (Table 3). Although mRNA expression of VEGF (Table 3) remained unaltered due to feeding LSM (P = 0.51) or implanting with E2 (P = 0.36), mRNA expression of its receptors, KDR and FLT, were altered as a result of an interaction of feeding LSM and implanting with E2 (P < 0.001 and P = 0.04, respectively). In the case of KDR, when ewes were implanted with E2 for 0 h, feeding LSM for 1 and 14 d decreased (P = 0.005 and P < 0.001, respectively) mRNA expression, whereas feeding LSM for 7 d increased (P = 0.02) expression compared with ewes fed LSM for 0 d. Additionally, implanting ewes with E2 did not alter (P > 0.10) mRNA expression when LSM was fed for 0 d but decreased expression with 24-h E2 exposure when ewes were fed LSM for 1 (P = 0.03) and 7 d (P < 0.001). In contrast, mRNA expression of KDR increased (P = 0.04) when ewes were implanted with E2 for 6 h when LSM was fed for 14 d. Expression of FLT mRNA was not altered in response to E2 when LSM was fed for 0 (P > 0.66), 7 (P > 0.08), or 14 d (P > 0.30) but decreased when ewes were implanted for 24 h compared with ewes implanted for 0 or 6 h (P = 0.03 and P = 0.004, respectively) when LSM was fed for 1 d. Feeding LSM and implanting with E2 also interacted (P = 0.04) to alter the expression of sGC.
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Table 3. Effects of feeding linseed meal (LSM) and implanting ewes with estradiol-17β (E2) on jejunal expression of angiogenic factors vascular endothelial growth factor (VEGF), fms-like tyrosine kinase (FLT; VEGF receptor-1), kinase insert domain-containing receptor (KDR; VEGF receptor-2), fibroblast growth factor (FGF), fibroblast growth factor receptor 2 IIIc (FGFR2 IIIc), angiopoietin 1 (ANG-1), angiopoietin 2 (ANG-2), angiopoietin receptor (Tie-2), endothelial nitric oxide synthase (eNOS), and soluble guanylate cyclase (sGC)1
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Implanting ewes with E2 for 6 h resulted in an increased (P = 0.001) sGC mRNA expression when LSM was fed for 0 d compared with ewes implanted for 0 h. When ewes were implanted with E2 for 24 h, expression of sGC mRNA returned to levels similar (P = 0.80) to those observed when ewes were implanted for 0 h. Feeding LSM for 1, 7, or 14 d resulted in sGC levels that were unaltered (P > 0.11, P > 0.54, and P > 0.54, respectively) due to duration of E2 implant. Implanting ewes with E2 tended (P = 0.10) to alter expression of eNOS mRNA with eNOS mRNA expression being greater (P = 0.04) in ewes implanted with E2 for 6 h (0.22 ± 0.031) compared with ewes implanted for 0 h (0.13 ± 0.029). Expression of eNOS mRNA after 24 h of E2 (0.18 ± 0.25) was similar to levels observed when ewes implanted for 0 (P = 0.18) and 6 h (P = 0.36). There was no effect (P 0.33) of LSM or E2 on FGF, FGFR, ANG-1, ANG-2, or Tie-2 expression levels.
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DISCUSSION
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This study confirmed that LSM and E2 can interact with, or affect, jejunal mass, cellularity, cellular proliferation within the crypt region of the jejunum, and angiogenic factor expression in the jejunal mucosa. Although phytoestrogens have been shown to alter cellular proliferation in the digestive tract (Chen et al., 2005), this is the first study in which LSM has been shown to interact with E2 to affect cellular proliferation in the jejunum.
Data from this study demonstrated that feeding LSM for an extended period of time in the presence of E2 decreased jejunal mass. In addition, jejunal cellular proliferation after 24 h of E2 exposure decreased when LSM was fed, indicating that longer LSM feeding may be anti-estrogenic in the jejunum. To our knowledge this is the first study showing that E2 alone is capable of increasing cellular proliferation within the gut. Estrogen concentrations increase dramatically in females during estrus and pregnancy, indicating that gastrointestinal tract function may be altered during these physiological states. Shurson et al. (2003) reported that pregnant sows gained BW more efficiently than nonpregnant sows. Scheaffer et al. (2004) reported that both vascularity percentage and total microvascular volume of the jejunum were increased in pregnant ewes compared with nonpregnant ewes, indicating that the jejunum increases absorptive capacity in response to the increasing nutritional demands of pregnancy, yet the physiological signals for this phenomenon are not known. Perhaps the elevated concentrations of E2 during pregnancy signal increased nutrient absorption in the small intestine. The present study did not investigate what effect prolonged exposure to E2 (longer than 24 h) would have on cellular proliferation or vascularity of the gut. Prolonged E2 exposure, such as during pregnancy, on gut function requires further investigation in addition to its interaction with LSM because it is evident from the present study that LSM is capable of interacting with E2.
Magness et al. (1998) observed an increase in blood flow to the small bowel of ovariectomized ewes following a long exposure to E2 (i.e., 10 d) but not during short-term duration (i.e., 2 h). It is therefore not surprising that changes in vascularity measurements were not observed after only 6 or 24 h of exposure to E2 in the current study. Perhaps vascularity measurements in the jejunum would be detectable if a longer exposure to E2 occurred. Moreover, increases in uterine vascularity in ovariectomized ewes are not increased until after 24 h (Johnson et al., 1997b; Reynolds et al., 1998a). Although vascularity measurements were not altered following LSM feeding periods of up to 14 d, further research may be necessary to determine the effects of feeding LSM to subjects exposed to E2 for prolonged periods. The tendency for a reduction in vascular number density without a change in other vascularity measurements at 6 h of E2 exposure is unclear. In the current study, the increase in proliferation index following exposure to E2 was ablated when LSM was fed. Perhaps feeding LSM would similarly ablate an increase in blood flow to the small bowel as observed by Magness et al. (1998) in response to prolonged E2 exposure.
The inability of E2 exposure to alter VEGF expression in the jejunum is in contrast to data which uterine VEGF expression is elevated in cyclic ewes after 4, 8, and 24 h of being implanted with E2 (Reynolds et al., 1998b). Suzuma et al. (1999) also noted an increase in VEGF mRNA expression in cultured bovine retinal endothelial cells 24 h after E2 exposure. This difference in VEGF expression patterns in response to E2 is likely due to the difference in tissue type. Angiogenesis is common in wound healing and developing tissues as well as in organs of the reproductive tract, but not common in adult tissue (Fotsis et al., 1993; Reynolds and Redmer, 1998). It is therefore feasible that nonreproductive tissues may respond differently in angiogenic factor expression to E2 exposure than reproductive tissues. Uterine tissue obtained from ewes in the current study responded similarly to previous studies (Reynolds et al., 1998b) by exhibiting an increase in caruncular VEGF mRNA expression after 6 h of being implanted with E2 irrespective of the length of time they were fed LSM (O’Neil et al., 2006).
Suzuma et al. (1999) noted an increase in KDR mRNA expression in bovine retinal endothelial cells after exposure to E2 for 6 to 24 h. This was not observed in the present study in jejunal tissue when LSM was fed for 0 d. Furthermore, a decrease in KDR mRNA expression in response to exposure to E2 for 24 h was observed when LSM was fed for 1 or 7 d, whereas implanting ewes fed LSM for 14 d increased expression. Expression patterns of VEGF mRNA due to feeding LSM were also peculiar with expression increased when LSM was fed for 7 d but decreased when LSM was fed for 1 or 14 d in ewes not implanted with E2. Perhaps this can be explained due to rumen transit time or ruminal metabolism of the SDG in that 24 h is not enough time for the bacteria to adapt to metabolizing SDG or the LSM still residing in the rumen. The response observed after 14 d of feeding may be related to downregulation of E2 receptors or some type of negative feedback. Although FLT mRNA expression was altered less by LSM and E2, it did respond similarly when LSM was fed for 1 d with a decrease in expression in ewes implanted for 24 h. These data indicate that in the jejunum angiogenesis may be altered by altered expression of FLT and KDR rather than by altering expression of VEGF. Moreover, when VEGF binds to FLT, increases in vascular permeability occur (Shibuya, 2001). These data conflict somewhat with the hypothesis that phytoestrogens are mainly antiangiogenic (Fotsis et al., 1993; Kruse et al., 1997; Dubey et al., 2000). Furthermore, these data also indicate that phytoestrogens are capable of ablating the angiogenic properties of E2 (Cullinan-Bove and Koos, 1993; Suzuma et al., 1999).
Weiner et al. (1994) observed an increase in eNOS expression in response to elevated concentrations of E2. Rosselli et al. (1995) also observed an increase in circulating levels of nitric oxide in postmenopausal women supplemented with E2. These reports agree with the present study where 6 h of E2 exposure tended to cause an increase in eNOS expression. Although feeding LSM for any duration did not appear to alter eNOS expression, it did suppress an increase in sGC mRNA expression observed after 6 h of E2 exposure when LSM was fed for 0 d. These differences may be tissue-, species-, or age-specific; the present study conflicts with data from Krumenacker et al. (2001) who reported that sGC mRNA expression decreased in immature rat uteri 3 h after an injection of E2, whereas expression in the lung, liver, and vascular tissue remained unchanged. Although we did not detect any alterations in vascularity of the intestinal villi, mRNA expression of eNOS was increased after 6 h of E2 implant and FLT, KDR, and sGC were altered by an interaction of E2 and LSM.
Although the present study examined the expression on angiogenic factor mRNA, it should be noted that this may not correlate with phenotypic expression of the gene product (Rehfeld, 1998). Further research may be useful to determine if the changes in mRNA expression due to interactions of LSM and E2 result in altered protein expression. Because angiogenesis is, for the most part, restricted to the female reproductive organs in adult organisms (Fotsis et al., 1993; Reynolds and Redmer, 1998), it is necessary to determine how LSM affects angiogenesis in the ovary, follicle, corpus luteum, and uterus of the adult female in addition to the placenta of the pregnant female. Additionally, the effect of LSM on angiogenesis of the developing conceptus and adolescent also need to be examined.
In summary, E2 increases jejunal cellular proliferation. However, when adult, ovariectomized ewes are fed LSM and exposed to E2, this increase in jejunal cellular proliferation decreases. Furthermore, although there was little influence of short-term E2 or LSM exposure on jejunal vascularity, E2 and LSM influence jejunal KDR, FLT, and sGC gene expression.
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Footnotes
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1 The authors thank Dakota Gold Marketing (Sioux Falls, SD) for graciously donating the dried distillers grains used to formulate the diets, the laboratory of Clifford Hall for analyzing samples for se-coisolariciresinol diglycoside, and M. L. Johnson, J. D. Kirsch, K. C. Kraft, Z. C. Hall, J. M. Evoniuk, K. R. Freeberg, S. L. Julius, M. A. Ward, P. P. Borowicz, R. M. Weigl, M. Marchello, C. E. Oliver, D. M. Larson, and J. J. Reed (all from North Dakota State University, Fargo) for assistance in collection of tissue and data. 
2 Corresponding author: kim.vonnahme{at}ndsu.edu
Received for publication April 4, 2008.
Accepted for publication June 12, 2008.
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LITERATURE CITED
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AOAC. 1990. Official Methods of Analysis. Vol. I. 15th ed. Assoc. Off. Anal. Chem., Arlington, VA.
Axelson, M., and K. D. R. Setchell. 1981. The excretion of lignans in rats—Evidence for an intestinal bacterial source for this new group of compounds. FEBS Lett. 123:337–342.[CrossRef][Medline]
Axelson, M., J. Sjövall, B. E. Gustafsson, and K. D. R. Setchell. 1982. Origin of lignans in mammals and identification of a precursor from plants. Nature 298:659–660.[CrossRef][Medline]
Begum, A. N., C. Nicolle, I. Mila, C. Lepierre, K. Nagano, K. Fukushima, S. Heinonen, H. Adlercreutz, C. Rémésy, and A. Scalbert. 2004. Dietary lignans are precursors of mammalian lignans in rats. J. Nutr. 134:120–127.[Abstract/Free Full Text]
Bradford, M. M. 1976. A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248–254.[CrossRef][Medline]
Chen, A.-C., M. A. Berhow, K. A. Tappenden, and S. M. Donovan. 2005. Genistein inhibits intestinal cell proliferation in piglets. Pediatr. Res. 57:192–200.[CrossRef][Medline]
Cullinan-Bove, K., and R. D. Koos. 1993. Vascular endothelial growth factor/vascular permeability factor expression in the rat uterus: Rapid stimulation by estrogen correlates with estrogen-induced increases in uterine capillary permeability and growth. Endocrinology 133:829–837.[Abstract/Free Full Text]
Day, A. J., F. J. Cañada, J. C. Díaz, P. A. Kroon, R. Mclauchlan, C. B. Faulds, G. W. Plumb, M. R. A. Morgan, and G. Williamson. 2000. Dietary flavonoid and isoflavone glycosides are hydrolysed by the lactase site of lactase phlorizin hydrolase. FEBS Lett. 468:166–170.[CrossRef][Medline]
Dubey, R. K., M. Rosselli, B. Imthurn, P. J. Keller, and E. K. Jackson. 2000. Vascular effects of environmental oestrogens: Implications for reproductive and vascular health. Hum. Reprod. Update 6:351–363.[Abstract/Free Full Text]
Fotsis, T., M. Pepper, H. Adlercreutz, G. Fleischmann, T. Hase, R. Montesano, and L. Schweigerer. 1993. Genistein, a dietary-derived inhibitor of in vitro angiogenesis. Proc. Natl. Acad. Sci. USA 90:2690–2694.[Abstract/Free Full Text]
Johnson, M. L., D. A. Redmer, and L. P. Reynolds. 1997a. Effects of ovarian steroids on uterine growth, morphology, and cell proliferation in ovariectomized, steroid-treated ewes. Biol. Reprod. 57:588–596.[Abstract]
Johnson, M. L., D. A. Redmer, and L. P. Reynolds. 1997b. Uterine growth, cell proliferation and c-fos proto-oncogene expression throughout the estrous cycle in ewes. Biol. Reprod. 56:393–401.[Abstract]
Krumenacker, J. S., S. M. Hyder, and F. Murad. 2001. Estradiol rapidly inhibits soluble guanylyl cyclase expression in rat uterus. Proc. Natl. Acad. Sci. USA 98:717–722.[Abstract/Free Full Text]
Kruse, F. E., A. M. Joussen, T. Fotsis, L. Schweigerer, K. Rohrschneider, and H. E. Völcker. 1997. Inhibition of neovascularization of the eye by dietary factors exemplified by isoflavonoids. Ophthalmologe 94:152 (Abstr.).[CrossRef][Medline]
Luna, L. G. 1968. Manual of Histological Staining Methods of the Armed Forces Institute of Pathology. McGraw-Hill, New York, NY.
Magness, R. R., T. M. Phernetton, and J. Zheng. 1998. Systemic and uterine blood flow distribution during prolonged infusion of 17β-estradiol. Am. J. Physiol. 275:H731–H743.[Medline]
NRC. 1985. Nutrient Requirements of Sheep. Sixth ed. National Academy Press, Washington, DC.
O’Neil, M. R., G. P. Lardy, L. P. Reynolds, J. S. Caton, M. L. Johnson, and K. A. Vonnahme. 2006. Effects of estradiol (E2) and linseed meal (LSM) on caruncular angiogenic factors in ovariectomized (OVX) ewes. Biol. Reprod. 75(Suppl. 1):132 (Abstr.)
Pfaffl, M. W., I. G. Lange, and H. H. Meyer. 2003. The gastrointestinal tract as target of steroid hormone action: Quantification of steroid receptor mRNA expression (AR, ERalpha, ERbeta and PR) in 10 bovine gastrointestinal tract compartments by kinetic RT-PCR. J. Steroid Biochem. Mol. Biol. 84:159–166.[CrossRef][Medline]
Redmer, D. A., R. P. Aitken, J. S. Milne, L. P. Reynolds, and J. M. Wallace. 2005. Influence of maternal nutrition on messenger RNA expression of placental angiogenic factors and their receptors at midgestation in adolescent sheep. Biol. Reprod. 72:1004–1009.[Abstract/Free Full Text]
Reed, J. J., M. A. Ward, K. A. Vonnahme, T. L. Neville, S. L. Julius, P. P. Borowicz, J. B. Taylor, D. A. Redmer, A. T. Grazul-Bilska, L. P. Reynolds, and J. S. Caton. 2007. Effects of selenium supply and dietary restriction on maternal and fetal body weight, visceral organ mass and cellularity estimates, and jejunal vascularity in pregnant ewe lambs. J. Anim. Sci. 85:2721–2733.[Abstract/Free Full Text]
Rehfeld, J. F. 1998. The new biology of gastrointestinal hormones. Physiol. Rev. 78:1087–1108.[Abstract/Free Full Text]
Reynolds, L. P., J. D. Kirsch, K. C. Kraft, D. L. Knutson, W. J. Mc-Claflin, and D. A. Redmer. 1998a. Time-course of the uterine response to estradiol-17β in ovariectomized ewes: Uterine growth and microvascular development. Biol. Reprod. 59:606–612.[Abstract/Free Full Text]
Reynolds, L. P., J. D. Kirsch, K. C. Kraft, and D. A. Redmer. 1998b. Time-course of the uterine response to estradiol-17β in ovariectomized ewes: Expression of angiogenic factors. Biol. Reprod. 59:613–620.[Abstract/Free Full Text]
Reynolds, L. P., and D. A. Redmer. 1998. Expression of the angiogenic factors, basic fibroblast growth factor and vascular endothelial growth factor, in the ovary. J. Anim. Sci. 76:1671–1681.[Abstract/Free Full Text]
Rickard, S. E., L. J. Orcheson, M. M. Seidl, L. Luyengi, H. H. S. Fong, and L. U. Thompson. 1996. Dose-dependent production of mammalian lignans in rats and in vitro from the purified precursor secoisolariciresinol diglycoside in flaxseed. J. Nutr. 126:2012–2019.[Abstract/Free Full Text]
Rosselli, M., B. Imthurn, P. J. Keller, E. K. Jackson, and R. K. Dubey. 1995. Circulating nitric oxide (nitrite/nitrate) levels in postmenopausal women substituted with 17β-estradiol and norethisterone acetate. Hypertension 25:848–853.[Abstract/Free Full Text]
Scheaffer, A. N., J. S. Caton, D. A. Redmer, and L. P. Reynolds. 2004. The effect of dietary restriction, pregnancy, and fetal type in different ewe types on the fetal weight, maternal body weight, and visceral organ mass in mature ewes. J. Anim. Sci. 82:1826–1838.[Abstract/Free Full Text]
Shibuya, M. 2001. Structure and function of VEGF/VEGF-receptor system involved in angiogenesis. Cell Struct. Funct. 26:25–35.[CrossRef][Medline]
Shurson, G. C., G. W. Libal, J. Crenshaw, C. R. Hamilton, R. L. Fisher, D. D. Koehler, and M. H. Whitney. 2003. Impact of energy intake and pregnancy status on rate and efficiency of gain and backfat changes of sows postweaning. J. Anim. Sci. 81:209–216.[Abstract/Free Full Text]
Soto-Navarro, S. A., T. L. Lawler, J. B. Taylor, L. P. Reynolds, J. J. Reed, J. W. Finley, and J. S. Caton. 2004. Effect of high-selenium wheat on visceral organ mass, and intestinal cellularity and vascularity in finishing beef steers. J. Anim. Sci. 82:1788–1793.[Abstract/Free Full Text]
Suzuma, I., M. Mandai, H. Takagi, K. Suzuma, A. Otani, H. Oh, K. Kobayashi, and Y. Honda. 1999. 17β-Estradiol increases VEGF receptor-2 and promotes endothelial cells. Invest. Ophthalmol. Vis. Sci. 40:2122–2129.[Abstract/Free Full Text]
Thomas, M. L., X. Xu, A. M. Norfleet, and C. S. Watson. 1993. The presence of functional estrogen receptors in intestinal epithelial cells. Endocrinology 132:426–430.[Abstract/Free Full Text]
Tilley, J. M. A., and R. A. Terry. 1963. A two-stage technique for the in vitro digestibility of forage crops. J. Br. Grassl. Soc. 18:104–111.
Vonnahme, K. A., D. A. Redmer, E. Borowczyk, J. J. Bilski, J. S. Luther, M. L. Johnson, L. P. Reynolds, and A. T. Grazul-Bilska. 2006. Vascular composition, apoptosis, and expression of angiogenic factors in the corpus luteum during prostaglandin F2- induced regression in sheep. Reproduction 131:1115–1126.[Abstract/Free Full Text]
Weiner, C. P., I. Lizasoain, S. A. Baylis, R. G. Knowles, I. G. Charles, and S. Moncada. 1994. Induction of calcium-dependent nitric oxide synthases by sex hormones. Proc. Natl. Acad. Sci. USA 91:5212–5216.[Abstract/Free Full Text]
Zheng, J., M. L. Johnson, D. A. Redmer, and L. P. Reynolds. 1996. Estrogen and progesterone receptors, cell proliferation and c-fos expression in the ovine uterus during early pregnancy. Endocrinology 137:340–348.[Abstract]
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Abstract
INTRODUCTION
