J. Anim. Sci. 2005. 83:1914-1923
© 2005 American Society of Animal Science
Effects of grazing program and subsequent finishing on gene expression in different adipose tissue depots in beef steers1,2
J. W. Ross*,
T. K. Smith*,
C. R. Krehbiel*,3,
J. R. Malayer
,
U. DeSilva*,
J. B. Morgan*,
F. J. White*,
M. J. Hersom*,4,
G. W. Horn* and
R. D. Geisert*
* Department of Animal Science, Oklahoma Agricultural Experiment Station, and
and
Department of Physiological Sciences, College of Veterinary Medicine, Oklahoma State University, Stillwater 74078
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Abstract
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This experiment was conducted to examine the effects of grazing program and subsequent finishing on gene expression in adipose tissue from steers. Twenty Angus x Angus-Hereford steer calves (initial BW = 231 ± 25 kg) were allotted randomly to one of two winter grazing treatments: 1) grazing winter wheat pasture to achieve a high rate of BW gain (HGW); or 2) grazing dormant tallgrass native range (NR). Steers in the NR treatment were provided 0.91 kg•steer–1•d–1 of a 41% CP (as-fed basis) cottonseed meal supplement. Following the grazing period, steers were assigned randomly to feedlot pens. Steers were fed to a common endpoint of 1.27 cm of backfat between the 12th and 13th rib. Four steers from each treatment were slaughtered at the end of the grazing period, and the remaining steers from each treatment (n = 6) were slaughtered at the predetermined compositional endpoint. Intramuscular and s.c. fat samples were collected from LM sections of each steer at the 12th-/13th-rib interface on the left side. Pools of RNA were prepared for HGW and NR s.c. adipose tissue from steers slaughtered immediately after grazing. Suppression subtractive hybridization was performed followed by dot-blot hybridization screening to confirm differential expression of subtracted transcripts. Transcripts confirmed to be differentially expressed were subjected to dideoxy chain-termination sequencing. Quantitative reverse transcription PCR was performed on three differentially expressed clones: osteonectin, ferritin heavy chain, and decorin. Osteonectin, ferritin heavy chain, and decorin gene expression was greater (P < 0.05) in s.c. than in i.m. adipose tissue of finished steers. A depot x background interaction for osteonectin (P < 0.01) and ferritin heavy chain (P = 0.03) gene expression was observed for steers slaughtered after grazing, indicating that nutritional management can affect gene expression in adipose tissue depots differently. No differences resulting from prefinishing nutritional background (HGW or NR) were noted in osteonectin, ferritin heavy chain, or decorin gene expression in i.m. adipose tissue collected from finished steers, which might have resulted from feeding steers to the same compositional endpoint. Our data suggest that nutritional background alters gene expression in adipose depots, and that depots are influenced differently.
Key Words: Adipose Tissue • Beef Steers • Gene Expression • Stocker Programs
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Introduction
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Intramuscular fat (i.e., marbling) influences beef quality (Oishi et al., 2000
) and is positively related to the tenderness, juiciness, and flavor intensity of beef (Wheeler et al., 1994). A significant proportion of beef available to consumers is regarded as unacceptable because of toughness (Jeremiah et al., 1993). Understanding factors that govern marbling may help the beef industry to produce a consistently tender and palatable beef product.
Numerous efforts have been made to identify specific subsets of genes involved with preadipocyte differentiation. Smas and Sul (1993) indicated that preadipocyte factor-1 is a gene that is downregulated during preadipocyte differentiation. The novel APOBEC-1 target-1, a translational repressor, is decreased in i.m. fat as marbling scores increase in feedlot steers, suggesting that suppression of certain factors is crucial during the initiation of preadipocyte differentiation (Childs et al., 2002). Peroxisome proliferator-activated receptor gamma and CCAAT/enhancer binding protein alpha represent two families of transcription factors whose expression increases during preadipocyte differentiation (Cornelius et al., 1994; Hu et al., 1995). Although these data contribute greatly to our understanding of preadipocyte differentiation, we suggest that evaluating differential gene expression between different in vivo-derived adipose depots in growing and finishing cattle might further enhance our understanding of the various factors involved in regulating preadipocyte differentiation and subsequent fat deposition during commonly used growing and feeding programs.
The objective of the present investigation was to use suppression-subtractive hybridization (Diatchenko et al., 1996) to isolate genes that are differentially expressed between different fat depots of beef cattle during preadipocyte differentiation in response to differences in nutritional background. Identification and characterization of gene expression patterns during preadipocyte differentiation in different fat depots might provide a better understanding of the events required for i.m. fat deposition and provide future targets and management practices for improving the quality and palatability of beef products.
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Materials and Methods
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Animals and Management
Animals used in the present experiment represented a subset of steers used in Exp. 2 of a previous study (Hersom et al., 2004). Twenty fall-weaned Angus x Angus-Hereford steers (average initial BW = 231 ± 25 kg) were assigned randomly to one of two winter grazing treatments consisting of 1) grazing winter wheat pasture to achieve a high rate of BW gain (HGW; stocking density = 0.43 to 0.55 ha/steer; ADG = 1.10 kg/d; n = 10); or 2) grazing dormant tallgrass native range (NR; 0.63 ha/steer; ADG = 0.15 kg/d; n = 10). Steers grazing NR were provided a 41% CP (DM basis) cottonseed-meal supplement (0.91 kg•steer–1•d–1 (as-fed basis)). Winter grazing began on December 18, 2000, and ended on May 10, 2001, resulting in a 144-d grazing period. Following the grazing period, steers were transported to the USDA, ARS Grazinglands Research Laboratory in El Reno, OK. Steers were sorted by BW within winter grazing treatment and assigned randomly to feedlot pens. Within pen, steers were fed individually using the Calan Broadbent Feeding System (American Calan, Northwood, NH). Steers received a Revalor-S implant (Intervet, Millsboro, DE) and were vaccinated against infectious bovine rhinotracheitis, bovine virus diarrhea, parainfluenza, and respiratory syncytial virus (Titanium 5; Diamond Animal Health, Des Moines, IA). Steers were adapted over 4 wk to the final feedlot diet (Exp. 2 in Hersom et al., 2004) by replacing ground alfalfa hay with dry-rolled corn. Steers were fed once daily at 0800 and offered ad libitum access to water as described (Hersom et al., 2004).
Slaughter of Animals
At the end of the grazing phase, four animals from each treatment were slaughtered for adipose tissue collection. Steers were gathered from their respective pastures at 0700 on the day of slaughter and transported to the Oklahoma Food and Agricultural Products Research and Technology Center abattoir. Steers were stunned by captive bolt gun and killed by exsanguination. Following feedlot finishing, remaining steers in each treatment (n = 6) were slaughtered (as described for the end of the grazing phase) within 9 d of reaching a common endpoint of 1.27 cm of 12th-rib fat as determined by ultrasound (model 210, model UST-5021 probe; Aloka Co., Ltd., Wallingford, CT).
Collection of Adipose Tissue
Immediately following exsanguination, adipose samples were obtained from s.c. and i.m. adipose depots with sterilized instruments and working surfaces as previously described (Childs et al., 2002). Approximately 2 to 10 g of adipose tissue were collected from each depot of each animal. Subcutaneous and i.m. adipose were obtained by cutting through the hide of the animal on the left side at the 12th-/13th-rib interface. Subcutaneous adipose samples obtained were immediately labeled and snap frozen in liquid N2. Longissimus muscle sections were immediately transferred to a 4°C processing room for dissection as previously described (Childs et al., 2002). Muscle sections were rinsed in sterile Hank’s balanced salt solution (Gibco Invitrogen Corp., Grand Island, NY), and i.m. adipose was dissected from the muscle section being careful to remove all muscle fibers from the adipose tissue. Although care was taken, difficulties encountered in collecting i.m. adipose tissue from NR steers might have resulted in sample contamination with connective tissue. The i.m. adipose samples were labeled and snap frozen in liquid N2, and all samples were transferred to –80°C for storage.
RNA Extraction
The RNA was extracted using TRIzol reagent (Gibco Invitrogen Corp.) according to the manufacturer’s protocol. Briefly, 800 mg of adipose tissue were placed in a sterile centrifuge tube containing 8 mL of TRIzol reagent. Tissue samples were homogenized with a VirTis-hear homogenizer (The Virtus Co., Inc., Gardiner, NY) until all aggregates of adipose tissue had been disrupted. The samples were allowed to incubate at 22 to 25°C for 5 min. Following incubation, 2 mL chloroform were added to each tube, vortexed for 15 s, and incubated at 22 to 25°C for 3 min. Following this incubation, tubes were centrifuged at 4,750 x g for 30 min at 4°C. The upper aqueous phase was transferred to a fresh tube containing 5 mL of isopropyl alcohol, which was briefly vortexed and incubated at 22 to 25°C for 10 min. The samples were centrifuged at 11,000 x g for 20 min at 4°C. The supernatant fraction was discarded, and the remaining pellet was washed with 3 mL of 75% (vol/vol) ethanol by vortexing until the pellet had broken, followed by centrifugation at 11,000 x g for 15 min at 4°C. Ethanol was carefully removed, and the RNA pellet was allowed to air dry for 5 min at 22 to 25°C. The pellet was then resuspended in 200 µL of sterile H2O and stored at –80°C. Concentration of RNA in samples was estimated based on absorbance at a wavelength of 260 nm. The purity and integrity of RNA were determined using the 260:280 ratio and visualization on a 1.5%, ethidium bromide-stained agarose gel, respectively.
RNA Preparation
Pools of total RNA for suppression-subtractive hybridization were prepared from RNA extracted from the s.c. adipose tissue from the four HGW and four NR steers slaughtered after the backgrounding phase and before entering the feedlot phase. Forty micrograms of s.c. total RNA were composited for HGW and NR by combining 10 µg of s.c. adipose total RNA from each steer within each treatment.
Suppression-Subtractive Hybridization
Suppression-subtractive hybridization was performed using the Clontech PCR-Select cDNA subtraction kit (K-1804-1, Clontech Laboratories Inc., Palo Alto, CA). The procedure was conducted according to instructions in the Clontech PCR-Select cDNA subtraction kit user manual, as previously described by our laboratory (Mohan et al., 2002; Ross et al., 2003). Forward and reverse subtraction were performed for the s.c. adipose tissue comparison between HGW and NR. Driver and tester cDNA was produced from 20 µg of total RNA of s.c. adipose tissue from both HGW- and NR-fed steers following the manufacturer’s guidelines. Briefly, synthesized cDNA was digested with the restriction enzyme Rsa1, and the tester cDNA populations were divided into two tubes and ligated to both Adaptor 1 or Adaptor 2R. Before ligation at 16°C overnight, 2 µL from each adaptor ligation for each tester population were combined to serve as an unsubtracted control and were diluted into 1 mL sterile water following ligation. The subtractive hybridization was performed by adding 1.5 µL of driver cDNA to each tube, one containing 1.5 µL of Adaptor 1 and the other containing 1.5 µL of Adapter 2R-ligated tester cDNA (tester cDNA was approximately 30 times less concentrated than driver cDNA) in 1 µL of 4x hybridization buffer. Samples were denatured at 98°C for 1.5 min and then allowed to anneal at 68°C for 8 h. Following the first hybridization, the two samples were combined simultaneously with the excess addition of 1 µL of freshly denatured driver cDNA, and hybridization was continued at 68°C overnight. Products from the second hybridization were diluted in 200 µL of dilution buffer (20 mM HEPES, pH 8.3; 50 mM NaCl; and 0.2 mM EDTA), heated at 68°C for an additional 7 min, and stored at –20°C.
PCR Amplification of Subtracted Products
Two rounds of PCR amplification were performed on both the subtracted and unsubtracted products. In the first amplification, double-stranded cDNA with both adaptor sequences were exponentially amplified. The first PCR was performed for each subtracted product from the second hybridization and from the diluted un-subtracted driver cDNA from both the forward and reverse subtractions. Diluted cDNA (1 µL) for each subtracted sample and unsubtracted tester control was added to 24 µL of PCR master mix prepared using the reagents in the kit. This reaction mix was then incubated at 75°C for 5 min to extend the adaptors, and thermal cycling was immediately commenced: 94°C for 25 s, followed by 27 repetitive cycles of 94°C for 10 s, 66°C for 30 s, and 72°C for 1.5 min. Products resulting from this PCR amplification were diluted 1:10 with sterile water. One microliter of the diluted products from the first PCR amplification was added to 24 µL of secondary PCR master mix containing nested primers to ensure specific amplification of only those products that contained both adaptors. The second PCR was then conducted using the following conditions: 94°C for 10 s; 68°C for 30 s; and 72°C for 1.5 min. The number of cycles for the secondary PCR was optimized through the visualization of the secondary PCR products on a 2.0% agarose gel; optimal was determined by the maximum number of cycles before distinct bands appeared as smears. The optimal number of cycles for the second PCR was 8 cycles for s.c. HGW unsubtracted control; 10 cycles for s.c. HGW subtracted tester; 14 cycles for s.c. NR unsubtracted control; and 7 cycles for s.c. NR subtracted tester. Products from both the primary and secondary PCR amplification were analyzed on a 2% agarose gel.
Cloning of Subtracted cDNA
Following the second round of PCR amplification, subtracted products for each tester cDNA population were cloned using the TOPO TA cloning kit for sequencing (Invitrogen, Carlsbad, CA). Subtracted products were cloned into the pCR4-TOPO vector supplied in the kit and transformed into One Shot TOP10 chemically competent (DH5
-T1R) Escherichia coli cells (Invitrogen). Transformed cells were cultured overnight at 37°C on Luria Broth (Fisher Scientific, Pittsburgh, PA) agar plates containing carbanocillin, X-gal (5-brom- 4-chloro-3-indoyl-ß-D-galactopyranoside) and isopropyl-ß-D-thio-galactopyranoside for blue/white colony screening. Ninety-six plasmid colonies from each tester were randomly selected and cultured in terrific broth (Becton Dickinson Microbiology Systems, Sparks, MD) containing carbanocillin for 16 to 20 h at 37°C. An aliquot of terrific broth following culture was frozen at –80°C (final concentration of 30% glycerol), while plasmid DNA was extracted from the remaining culture using the Wizard SV96 plasmid DNA purification system (Promega Corp., Madison, WI) and eluted into 100 µL of sterile nuclease-free water.
Differential Screening and Sequence Determination
To confirm that cDNA inserts present in the subtracted products were indeed representative of uniquely expressed transcripts, all clones were subjected to differential screening as previously described by Ross et al. (2003). Plasmids that contained confirmed differentially expressed genes were cultured, and plasmid DNA was extracted using the Wizard Plus mini-prep DNA purification system (Promega Corp.) and eluted in 30 µL of nuclease-free water. The DNA was subjected to dideoxy chain termination sequencing (model 373A automated sequencer, Applied Biosystems, Foster City, CA) at the Oklahoma State University Recombinant DNA/Protein Resource Facility. The basic local alignment search tool (Altschul et al., 1990) was used to determine sequence homology of each differentially expressed template.
Quantitative One-Step Reverse-Transcription PCR
Based on current literature suggesting their involvement in preadipocyte differentiation in other tissues, three differentially expressed clones of interest (osteonectin, ferritin heavy chain, and decorin) were evaluated using quantitative reverse-transcription (RT) PCR as previously described by our laboratory (Hettinger et al., 2001; Ross et al., 2003). Individual samples from both s.c. and i.m. adipose tissue were evaluated for the three clones of interest. Gene expression for the selected transcripts was assayed using a dual-labeled probe designed to have a 5' reporter dye (6-FAM) and a 3' quenching dye (TAMRA). Sequence specific primers and probes are presented for each selected gene in Table 1. Reverse transcription and PCR amplification of 100 ng of total RNA was performed in 25-µL reactions using the Quantitect RT-PCR probe kit (Qiagen, Valancia, CA). Each reaction contained 2 µL of total RNA (50 ng/µL) and 23 µL of master mix, using the reagents provided in the kit and the designed primers (400 nM final concentration of each) and fluorescence-labeled probe (200 nM final concentration). Amplification was conducted using the ABI PRISM 7700 sequence detection system (PE Applied Biosystems, Foster City, CA). Thermal cycling conditions were as follows: 50°C for 30 min and 95°C for 10 min; followed by 40 repetitive cycles of 95°C for 15 s and a combined annealing/extension phase of 60°C for 1 min. Continuous fluorescent data collection was obtained. To correct for any variation in the amount of RNA loaded into each well, 18S ribosomal RNA was also determined for each sample (18S ribosomal control kit; PE Applied Biosystems).
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Table 1. Primer and probe sequences for quantitative reverse-transcriptase polymerase chain reaction of decorin, osteonectin, and ferritin heavy chain in adipose tissue
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Following RT-PCR, quantification of gene amplification was done by determining the cycle threshold (CT) based on fluorescence detected in the geometric region of the semilog view of the amplification plot. Relative quantification of target gene expression was evaluated by using the comparative CT method (Hettinger et al., 2001; Ross et al., 2003). The 
CT value was determined by subtracting the target CT of each sample from its respective ribosomal 18S CT value. Calculation of CT involved using the highest sample CT value as an arbitrary constant to subtract from all other CT sample values. Fold-changes in gene expression (presented in figures) of the target gene are equivalent to 2–Ct. For each transcript evaluated, an RNA pool constructed from random samples was generated and serial dilutions of this sample were assayed to measure the accuracy of the differences detected of known dilutions (i.e., every twofold dilution was equivalent to an increase in one cycle before crossing the threshold in the geometric region of the semilog view of the amplification plot). Furthermore, the pooled sample was used to conduct a no-reverse-transcriptase control to confirm the specific amplification of cDNA created from the RNA provided in the reaction, and that genomic DNA was not influential in the results.
Statistical Analyses
The Mixed procedure of SAS (SAS Inst. Inc., Cary, NC) for a completely randomized design was used to analyze the quantitative RT-PCR CT values. An unequal number of replications resulted from unequal recovery of RNA from each sample and the complete utilization of some samples to optimize and conduct the suppression–subtractive hybridization experiment. Satterthwaite’s approximation was included in the statistical model to correct for heterogeneity of variances. Following suppression subtractive hybridization, differences between HGW and NR steers in genes that were differentially expressed in s.c. adipose tissue collected immediately after grazing were confirmed with a model that contained grazing program (Table 2). For the three target genes of interest (decorin, osteonectin, and ferritin heavy chain), effects of slaughter date (after grazing vs. finished), grazing program (HGW vs. NR), and adipose depot (s.c. vs. i.m.), as well as all two-way interactions, were evaluated. Least squares means ± SE representing the CT values are presented. Least squares means were compared using LSD protected by a significant (P < 0.05) F-value. Differences in gene expression were considered significant at P 
0.05.
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Table 2. Effect of grazing program on decorin, osteonectin, and ferritin heavy chain gene expression confirming suppressive subtractive hybridization of subcutaneous adipose tissue
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Results
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Performance response and carcass characteristics of steers from different backgrounds slaughtered after grazing or at finishing have been previously reported in Exp. 2 of Hersom et al. (2004). Briefly, during the winter grazing period, ADG was 1.10 and 0.15 kg/d, whereas at the end of winter grazing, BW was 395 and 257 kg, 12th-rib fat thickness was 0.69 and 0.01 cm, and marbling score was 275 and 0 (0 = devoid; 100 = practically devoid; 200 = traces; 300 = slight) for HGW and NR steers, respectively. After finishing, 12th-rib fat thickness (1.37 and 1.55 cm, respectively) and marbling score (405 and 406, respectively) did not differ between HGW and NR steers (Hersom et al., 2004).
Suppression-Subtractive Hybridization
Ninety-six randomly selected template clones from each subtracted product of the comparison were differentially screened to confirm unique expression. Of the total 192 colonies selected, 47 were confirmed to carry a differentially expressed transcript. Clones confirmed differentially expressed were subjected to dideoxy chain termination sequencing to determine the putative identity of each (Table 3). Among the genes identified and allegedly expressed higher in HGW steers as opposed to NR steers were histone H3.3A, elongation factor 1, cathepsin-like genes, ferritin heavy chain, decorin, and osteonectin. The 12 template clones expressed higher in NR steers possessed high homology to pancreatic anionic trypsinogen, satellite DNA, and an unannotated gene sequenced during bovine high throughput genome sequencing. As indicated in Table 3, several of the 47 clones expressed the same transcript.
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Table 3. Putative identity of subtracted clones following high-gain wheat vs. native range subtraction of subcutaneous adipose tissue
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Quantitative RT-PCR
Based on their homology and identity to known sequences, osteonectin, decorin, and ferritin heavy chain were selected to undergo quantitative RT-PCR to confirm differential gene expression. As indicated through suppression subtractive hybridization, osteonectin (P < 0.001) and ferritin heavy chain (P < 0.02) gene expression in s.c. adipose tissue was greater in HGW than in NR steers slaughtered following grazing, whereas the difference for decorin approached significance (P < 0.11; Table 2
). These differences represent 2.4-, 13-, and 3.9-fold increases in gene expression for decorin, osteonectin, and ferritin heavy chain, respectively, in HGW vs. NR grazing programs.
Decorin gene expression was not affected by depot x grazing program (P < 0.76), depot x slaughter (P < 0.75), or grazing program x slaughter (P < 0.23) interactions (data not shown). Decorin gene expression was affected by both adipose depot (P < 0.001) and time of slaughter (P < 0.005), and tended (P < 0.06) to differ between grazing programs (Table 4). These differences in gene expression represent 1.5-, 2.6-, and 2-fold greater decorin gene expression in HGW, s.c. and prefeed steers compared with NR, i.m., and finished steers, respectively.
Adipose depot (s.c. vs. i.m.) x grazing program (HGW vs. NR) interactions were observed for osteonectin (P < 0.001) and ferritin heavy chain (P < 0.01) gene expression (Table 5). Subcutaneous fat from steers grazing HGW had approximately 8.5-fold greater osteonectin gene expression than i.m. fat from HGW steers, while expressing 10- to 15-fold more osteonectin mRNA compared with either adipose depot of NR steers. Ferritin heavy chain gene expression was similar to that of osteonectin, with greatest gene expression occurring in s.c. adipose tissue of HGW steers. Ferritin heavy chain gene expression was four- to fivefold greater in s.c. adipose tissue from HGW steers compared with s.c. fat from NR steers and i.m. fat from steers grazing either HGW or NR.
The effect of slaughter (prefeed vs. finished) interacted with adipose depot (s.c. vs. i.m.) affecting both osteonectin (P < 0.001) and ferritin heavy chain (P < 0.002) gene expression (Table 6). Osteonectin gene expression was greater (approximately 1.8-fold; P < 0.01) in s.c. adipose tissue compared with i.m. adipose tissue from steers following grazing. This difference was amplified following high-grain feeding as s.c. fat had approximately 8.5-fold greater (P < 0.001) osteonectin gene expression than i.m. fat. Overall, high-grain feeding significantly increased expression for osteonectin in both adipose depots compared with expression levels directly after grazing; however, the change in expression levels was greater in s.c. depots (12-fold increase) compared with i.m. depots (2.5-fold increase). Similarly, ferritin heavy chain gene expression was greatest in s.c. adipose tissue of finished steers compared with s.c. adipose following grazing (P < 0.001) and i.m. fat from before (P < 0.001) or after finishing (P < 0.003). Unlike osteonectin, the increase in ferritin heavy chain gene expression following high-grain feeding occurred only in s.c. adipose tissue, increasing approximately sevenfold following the finishing period compared with expression levels directly after grazing.
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Discussion
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Using suppression-subtractive hybridization, we identified decorin as a gene expressed greater in s.c. adipose tissue from HGW steers compared with NR steers slaughtered before finishing. Quantitative RT-PCR confirmed numerical increases for decorin gene expression in s.c. adipose tissue from HGW compared with NR steers. Decorin is a leucine-rich proteoglycan of low molecular mass, whose core protein is substituted with a chondroitin sulfate chain (Hardingham and Fosang, 1992; Kresse et al., 1993). Decorin participates in cell proliferation and differentiation (Santra et al., 1995), and it is thought to be involved in tissue morphogenesis (Nishimura et al., 2002). Calvo et al. (1991) reported that differentiating 3T3-L1 preadipocytes produced a small chondroitin/dermatin sulfate proteoglycan with a structure similar to decorin. Proteoglycans, such as decorin, are often involved in the formation of fibrous connective tissue, and they may be a component of the extracellular matrix, giving preadipocytes the ability to change morphologically and functionally, a process hypothesized to occur during preadipocyte differentiation into adipose tissue (Calvo et al., 1991). The production and accumulation of small proteoglycans, such as decorin, is 50 to 70% less in 3T3-L1 fibroblasts stimulated to differentiate into adipocytes compared with control fibroblasts (Musil et al., 1991). Our data suggested lower decorin gene expression in i.m. adipose tissue compared with s.c. adipose, and decorin gene expression also was lower in finished steers than in steers slaughtered before finishing. When the results of 3T3-L1 cell models and the present data are viewed collectively, the decrease of decorin gene expression following finishing may be associated with the simultaneous accumulation of i.m. adipose, which is characterized by decreased decorin gene expression.
Similar to decorin, osteonectin (also known as secreted protein acidic and rich in cysteine; SPARC) also is a component of the extracellular matrix. Osteonectin is a nonstructural component, and it is involved in modulating cell-matrix interactions (Bornstein, 1995). Osteonectin gene expression increases in obese mice compared with their lean counterparts and in obesity-prone mice consuming a high-fat diet for 10 wk compared with similar mice consuming a control diet (Tartare-Deckert et al., 2001). These results indicate that osteonectin/SPARC is upregulated during obesity and may affect key functions of adipose tissue. In addition, osteonectin-null mice developed an increased number of s.c. fat deposits without a change in BW (Bradshaw et al., 2003), suggesting that although osteonectin is not required for adipose synthesis, it may be involved with extracellular matrix rearrangements associated with obesity. Our data suggest that osteonectin expression in cattle adipose tissue is increased in response to high energy levels in the diet. Osteonectin gene expression was greater in HGW vs. NR and finished vs. prefeed for both i.m. and s.c. adipose tissues. The slaughter x depot and grazing program x depot interactions indicate that high-energy diets had a much more dramatic affect on osteonectin gene expression in s.c. depots than in i.m. depots, albeit i.m. adipose also was characterized by greater osteonectin gene expression in response to high-energy diets. Therefore, it is logical that osteonectin was expressed greater in s.c. adipose from HGW steers than from NR steers slaughtered after grazing, which might help explain why osteonectin gene expression was greater in adipose from finished steers than in steers slaughtered before finishing. Cell-to-cell contact as adipocytes become confluent seems to be required for differentiation from preadipocytes into mature adipocytes, and the extracellular matrix of cells has been shown to play a critical part in cell differentiation and proliferation (Ruoslahti, 1988). Therefore, it seems logical that both osteonectin, as well as decorin, are involved with fat deposition during high-energy grazing (wheat pasture) and finishing of beef steers.
Ferritin is an intracellular protein that is largely involved with cellular iron homeostasis. Differentiation of 3T3-L1 cells into adipocytes is associated with a consistent increase of ferritin mRNA (Festa et al., 2000). It has been concluded that ferritin interrupts the reaction sequence that results in lipid peroxidation and cell damage by sequestering iron ions from the site of oxygen radical formation, and that increased synthesis of ferritin protects adipocytes from lipid peroxidation (Festa et al., 2000). Our data suggest that ferritin heavy chain gene expression in s.c. adipose is regulated by energy, as its expression was greater in HGW and finished steers compared with NR and prefed steers, respectively, whereas neither grazing or finishing diet affected its expression in i.m. adipose tissue. It might be suggested that ferritin heavy chain is involved with the differentiation and accumulation of specific adipose depots, such as s.c. This concept is supported by the carcass data from the present study, which indicated both s.c. and i.m. adipose tissue accretion was occurring in HGW vs. NR and finished vs. prefeed steers (Hersom et al., 2004). Interestingly, ferritin heavy chain gene expression changes are specific for s.c. adipose tissue, indicating that morphological differences exist between different fat depots that are necessary for their accumulation.
In the present experiment, it seems that gene expression of decorin, osteonectin, and ferritin heavy chain in adipose tissue from beef steers was influenced by nutrition and adipose tissue depot. As described by Smith et al. (2000), i.m. adipose tissue has been distinguished from other fat depots by its location within perimysial connective tissue alongside myofibers (Moody and Cassens, 1968), rate of fatty acid biosynthesis (5 to 10% of the rates observed in s.c. adipose tissue; Smith and Crouse, 1984; Lin et al., 1992), effects of starvation (Smith et al., 1998), and carbon source for fatty acid biosynthesis (Smith and Crouse, 1984). In addition, several studies (Smith and Crouse, 1984; Miller et al., 1991; May et al., 1994) have reported greater lipogenic enzyme activities in s.c. adipose tissue than in i.m. adipose tissue. Therefore, our data are consistent with previous data, which indicate that i.m. and s.c. adipose tissue are metabolically distinct. It is interesting to note that none of the enzymes reported to have greater activities in s.c. vs. i.m. adipose tissue were identified as different by suppression-subtractive hybridization; however, the technique of suppression-subtractive hybridization is aimed at determining differences in expression patterns among low- to moderate-abundance mRNA. Highly abundant gene products usually associated with specific tissues, such as adipose, would be eliminated during suppression-subtractive hybridization.
Differences in gene expression between i.m. and s.c. adipose tissue were observed in steers slaughtered both before and after finishing. It should be noted that steers in the present experiment were not implanted during grazing, but were implanted once at the beginning of finishing with 24 mg of estradiol and 120 mg of trenbolone acetate. It is possible that implanting influenced our results, although it is generally accepted that anabolic implants have no effect on fat deposition in cattle. Greater rates of protein accretion and no change in fat accretion seem to result in the higher lean:fat observed in implanted cattle (Johnson et al., 1996), which also is supported by several studies that showed implantation with a combination of estradiol and trenbolone acetate had little or no effect on s.c. adipose thickness (Perry et al., 1991; Bartle et al., 1992; Johnson et al., 1996). As a result of the increased protein deposition, implanting alters i.m. lipid amount and composition through a dilution effect with increased LM size (Duckett et al., 1999). Torii et al. (1998) showed that fibroblast-like cells prepared from skeletal muscle of beef cattle could differentiate into adipocytes in the presence of peroxisome proliferator-activated receptor gamma-specific ligands. If skeletal muscle contains cells that can be induced to undergo adipose differentiation, then the implant during finishing might have influenced our results; however, steers from both grazing programs were treated similarly during grazing and finishing in the present experiment.
Conclusions
It can be concluded from the present experiment that gene expression in adipose tissue can be regulated similarly (i.e., osteonectin) or differently (i.e., ferritin heavy chain) in different adipose depots of beef steers. As suggested by Smith et al. (2000), because gene expression in different adipose tissue depots might be regulated differently throughout their accretion during grazing or finishing, it may be possible to influence fat deposition of fat depots independently of each other. Although different grazing programs seem to affect gene expression of adipose tissue, feeding steers to a common compositional endpoint seemed to minimize differences in adipose tissue gene expression. Nonetheless, specific factors noted in the present study that are likely to be involved in the regulation of adipose tissue accretion seem to be affected by diet.
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Footnotes
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1 Approved for publication by the Director, Oklahoma Agric. Exp. Stn. This research was supported by the Oklahoma Agric. Exp. Stn. under Project H-2438, the Oklahoma Beef Industry Council check-off dollars, and the Cooperative State Research, Education, and Extension Service, USDA, under Agreements No. 99-34198-7481 and 2001-34198-10403. 
2 The authors thank M. Ashworth for help with sample analyses and K. Poling for help with animal care and sample collection. The authors also thank the Oklahoma State Univ. Recombinant DNA/Protein Resource Facility for DNA sequencing.
4 Current address: Dept. of Anim. Sci., Rm 231 E. Anim. Sci. Bldg. 459, Univ. of Florida, Gainesville 32611-0910.
3 Correspondence: 208 Anim. Sci. Bldg. (phone: 405-744-8857; fax: 405-744-7390; e-mail: kclinto{at}okstate.edu).
Received for publication October 21, 2004.
Accepted for publication April 18, 2005.
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Literature Cited
|
|---|
Altschul, S., W. Gish, W. Miller, E. Myers, and D. Lipman. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403–410.[Medline]
Bartle, S. J., R. L. Preston, R. E. Brown, and R. J. Grant. 1992. Trenbolone acetate/estradiol combinations in feedlot steers: Dose-response and implant carrier effects. J. Anim. Sci. 70:1326–1332.[Abstract]
Bornstein, P. 1995. Diversity of function is inherent in matricellular proteins: An appraisal of thrombospondin 1. J. Cell. Biol. 130:503–506.[Free Full Text]
Bradshaw, A. D., D. C. Graves, K. Motamed, and E. H. Sage. 2003. SPARC-null mice exhibit increased adiposity without significant differences in overall body weight. Proc. Natl. Acad. Sci. USA 100:6045–6050.[Abstract/Free Full Text]
Calvo, J. C., D. Rodbard, A. Katki, S. Chernick, and M. Yanagishita. 1991. Differentiation of 3T3-L1 preadipocytes with 3-isobutyl-1-methylxanthine and dexamethasone stimulates cell-associated and soluble chondroitin 4-sulfate proteoglycans. J. Biol. Chem. 266:11237–11244.[Abstract/Free Full Text]
Childs, K. D., D. W. Goad, M. F. Allen, D. Pomp, C. Krehbiel, R. D. Geisert, J. B. Morgan, and J. R. Malayer. 2002. Differential expression of NAT1 translational repressor during development of bovine intramuscular adipocytes. Physiol. Genomics 10:49–56.[Abstract/Free Full Text]
Cornelius, P., O. A. MacDougald, and M. D. Lane. 1994. Regulation of adipocyte development. Annu. Rev. Nutr. 14:99–129.[Medline]
Diatchenko, L., Y. F. Lau, A. P. Campbell, A. Chenchik, F. Moqadam, B. Huang, S. Lukyanov, K. Lukyanov, N. Gurskaya, E. D. Sverdlov, and P. D. Siebert. 1996. Suppression subtractive hybridization: A method for generating differentially regulated or tissue-specific cDNA probes and libraries. Proc. Natl. Acad. Sci. USA 93:6025–6030.[Abstract/Free Full Text]
Duckett, S. K., D. G. Wagner, F. N. Owens, H. G. Dolezal, and D. R. Gill. 1999. Effect of anabolic implants on beef intramuscular lipid content. J. Anim. Sci. 77:1100–1104.[Abstract/Free Full Text]
Festa, M., G. Ricciardelli, G. Mele, C. Pietropaolo, A. Ruffo, and A. Colonna. 2000. Overexpression of H ferritin and up-regulation of iron regulatory protein genes during differentiation of 3T3-L1 preadipocytes. J. Biol. Chem. 275:36708–36712.[Abstract/Free Full Text]
Hardingham, T. E., and A. J. Fosang. 1992. Proteoglycans: Many forms and many functions. FASEB J. 6:861–870.[Abstract]
Hettinger, A. M., M. R. Allen, B. R. Zhang, D. W. Goad, J. R. Malayer, and R. D. Geisert. 2001. Presence of the acute phase protein, bikunin, in the endometrium of gilts during estrous cycle and early pregnancy. Biol. Reprod. 65:507–513.[Abstract/Free Full Text]
Hersom, M. J., G. W. Horn, C. R. Krehbiel, and W. A. Phillips. 2004. Effect of live weight gain of steers during winter grazing: I. Feedlot performance, carcass characteristics, and body composition of beef steers. J. Anim. Sci. 82:262–272.[Abstract/Free Full Text]
Hu, E., P. Tontonoz, and B. M. Spiegelman. 1995. Transdifferentiation of myoblasts by the adipogenic transcription factors PPAR
and C/EBP. Proc. Natl. Acad. Sci. USA 92:9856–9860.[Abstract/Free Full Text]
Jeremiah, L. E., A. K. W. Tong, S. D. M. Jones, and C. McDonell. 1993. A survey of Canadian consumer perceptions of beef in relation to general perceptions regarding food. J. Cons. Stud. Home Econ. 17:13–37.
Johnson, B. J., P. T. Anderson, J. C. Meiske, and W. R. Dayton. 1996. Effect of a combined trenbolone acetate and estradiol implant on feedlot performance, carcass characteristics, and carcass composition of feedlot steers. J. Anim. Sci. 74:363–371.[Abstract/Free Full Text]
Kresse, H., H. Hausser, and E. Schonherr. 1993. Small proteoglycans. Experentia 49:403–414.[Medline]
Lin, K. C., H. R. Cross, and S. B. Smith. 1992. Esterification of fatty acids by bovine intramuscular and subcutaneous adipose tissues. Lipids 27:111–116.[Medline]
Margareto, J., J. Gomez-Ambrosi, A. Marti, and J. A. Martinez. 2001. Time-dependent effects of a high-energy-yielding diet on the regulation of specific white adipose tissue genes. Biochem. Biophys. Res. Commun. 283:6–11.[Medline]
May, S. G., J. W. Savell, D. K. Lunt, J. J. Wilson, J. C. Laurenz, and S. B. Smith. 1994. Evidence for preadipocyte proliferation during culture of subcutaneous and intramuscular adipose tissues from Angus and Wagyu crossbred steer. J. Anim. Sci. 72:3110–3117.[Abstract]
Miller, M. F., H. R. Cross, D. K. Lunt, and S. B. Smith. 1991. Lipogenesis in acute and 48-hour cultures of bovine intramuscular and subcutaneous adipose tissue explants. J. Anim Sci. 69:162–170.[Abstract]
Mohan, M., S. Ryder, P. L. Claypool, R. D. Geisert, and J. R. Malayer. 2002. Analysis of gene expression in bovine blastocyst produced in vitro using suppression-subtractive hybridization. Biol. Reprod. 67:507–513.
Moody, W. B., and R. G. Cassens. 1968. A quantitative and morphological study of bovine longissimus fat cells. J. Food Sci. 33:47–55.
Musil K. J., A. Malmstrom, and J. Donner. 1991. Alteration of proteoglycan metabolism during the differentiation of 3T3-L1 fibroblasts into adipocytes. J. Cell Biol. 114:821–826.[Abstract/Free Full Text]
Nishimura, T., E. Futami, A. Taneichi, T. Mori, and A. Hattori. 2002. Decorin expression during development of bovine skeletal muscle and its role in morphogenesis of intramuscular connective tissue. Cells Tissues Organs 171:199–214.[Medline]
Oishi, M., Y. Taniguchi, K. Nishimura, T. Yamada, and Y. Sasaki. 2000. Characterization of gene expression in bovine adipose tissue before and after fattening. Anim. Genet. 31:166–170.[Medline]
Perry, T. C., D. G. Fox, and D. H. Beermann. 1991. Effect of an implant of trenbolone acetate and estradiol on growth, feed efficiency, and carcass composition of Holstein and beef steers. J. Anim. Sci. 69:4696–4702.[Abstract]
Ross, J. W., M. D. Ashworth, A. G. Hurst, J. R. Malayer, and R. D. Geisert. 2003. Analysis and characterization of differential gene expression during rapid trophoblastic elongation in the pig using suppression subtractive hybridization. Reprod. Biol. and Endocrinol. 1:23. Available: http://www.RBEj.com/content/1/1/23. Accessed May 7, 2005.
Ruoslahti, E. 1988. Structure and biology of proteoglycans. Annu. Rev. Cell Biol. 4:229–255.[Medline]
Santra, M. T., T. Skorski, B. Calabretta, E. C. Lattime, and R. V. Iozzo. 1995. De novo decorin gene expression suppresses the malignant phenotype in human colon cancer cells. Proc. Natl. Acad. Sci. USA 92:7016–7020.[Abstract/Free Full Text]
Smas, C. M., and H. S. Sul. 1993. Pref-1, a protein containing EGF-like repeats, inhibits adipocyte differentiation. Cell 73:725–734.[Medline]
Smith, S. B., and J. D. Crouse. 1984. Relative contributions of actate, lactate, and glucose to lipogenesis in bovine intramuscular and subcutaneous adipose tissue. J. Nutr. 114:792–800.
Smith, S. B., K. C. Lin, J. J. Wilson, D. K. Lunt, and H. R. Cross. 1998. Starvation depresses acylglycerol biosynthesis in bovine subcutaneous but not intramuscular adipose tissue homogenates. Comp. Biochem. Biophys. 120B:165–174.
Smith, S. B., D. K. Lunt, and M. Zembayashi. 2000. Intramuscular fat deposition: The physiological process and the potential for its manipulation. Pages 1–12 in Plains Nutr. Council Spring Conf. Publ. No. AREC 00-22. Texas A&M Res. and Ext. Center, Amarillo.
Tang, Q. Q., and M. D. Lane. 1999. Activation and centromeric localization of CCAAT/enhancer-binding proteins during the mitotic clonal expansion of adipocyte differentiation. Genes Dev. 13:2231–2241.[Abstract/Free Full Text]
Tartare-Deckert, S., C. Chavey, M.-N. Monthouel, N. Gautier, and E. V. Obberghen. 2001. The matricellular protein SPARC/osteonectin as a newly identified factor up-regulated in obesity. J. Biol. Chem. 276:22231–22237.[Abstract/Free Full Text]
Torii, S. I., T. Kawada, K. Matsuda, T. Matsui, T. Ishihara, and H. Yano. 1998. Thiazolidinedione induces the adipose differentiation of fibroblast-like cells resident within bovine skeletal muscle. Cell Biol. Int. 22:421–427.[Medline]
Wheeler, T. L., L. V. Cundiff, and R. M. Koch. 1994. Effect of marbling degree on beef palatability in Bos taurus and Bos indicus cattle. J. Anim. Sci. 72:3145–3151.[Abstract]
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Abstract
Introduction
