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,3
* Department of Animal Sciences, and
Department of Veterinary and Biochemical Sciences, Penn State University, University Park, PA 16801
Arm and Hammer Animal Nutrition Group, Church & Dwight Co. Inc., Princeton, NJ 08543
2 Corresponding author: bionaz{at}uiuc.edu
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ABSTRACT |
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 TOP ABSTRACT ACKNOWLEDGEMENTSREFERENCES |
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agonist) and rosiglitazone (ROSI, specific PPAR
agonist). The gene expression of specific PPAR-responsive genes such as carnitine palmitoyl transferase-1 (CPT1A) and acetyl coenzyme A oxidase (ACOX1) and of PPAR-responsive gene lipoprotein lipase (LPL) were analyzed using real-time reverse transcription PCR. It was found that CPT1A exhibited a significant increase in cells treated with WY, whereas the ACOX1 gene expression was not altered. The LPL gene expression showed an increase in response to ROSI. Interestingly, LPL was almost undetectable in MDBK cells not treated with ROSI. The potency of different fatty acids in activating PPAR as assessed by CPT1A mRNA abundance in MDBK cells was also tested. The mRNA of CPT1A (2.5- to 1.4-fold) was significantly increased by fatty acids in the order of palmitate > linolenate > linoleate > conjugated linoleate, and oleate. The results demonstrated MDBK cells to be responsive to PPAR agonists and thus a promising model to evaluate the role of PPAR in bovine cells. In addition, fatty acids were proven to have a different potency in modulating expression of CPT1A through PPAR.
Key Words: Madin-Darby bovine kidney cell • peroxisome proliferator-activated receptor • fatty acid • gene expression
In nonruminant species, peroxisome proliferator-activated receptors (PPAR) have been widely studied for their key role in many biological functions, particularly for their pivotal role in lipid metabolism (Desvergne et al., 2006). The sensitivity of PPAR to fatty acids (FA) suggests them to be important players in nutrition (Fekete and Brown, 2007). The PPAR family presents 3 subtypes called PPAR, PPAR, and PPAR
and β. The PPAR and PPAR are the most studied for their central role in lipid and glucose metabolism. Unsaturated FA, such as oleic (C18:1), linoleic (C18:2), and linolenic (C18:3), are potent activators of PPAR in non-ruminant species (Desvergne and Wahli, 1999; Wahle et al., 2003). Reports describing dose-dependent responses of PPAR to FA administration have been published for humans and rodents; however, no data are available in dairy cows. Furthermore, PPAR are known to have a diverse sensitivity to different FA (Vanden Heuvel, 1999).
The effect of PPAR agonists in vivo have been studied in the goat (Cappon et al., 2002), but not in bovine, in which in vivo studies are extremely costly and time-consuming. Therefore, preliminary studies to identify potency and doses of FA in activating PPAR are deemed necessary. Use of an in vitro system has the capacity of determining which FA are the most effective as PPAR agonist(s) and which concentrations more potently affect those nuclear receptors with a downstream effect on target genes and metabolism. Ruminants can only be treated with small amounts of FA using oral administration due to intolerance by rumen microorganisms (Jenkins, 1993). Because one of the interesting physiological effects of dietary FA is the regulation of gene expression (Pégorier et al., 2004), the amount of a specific FA should conceivably be based on the concentration and composition that induce a positive function. To our knowledge, no information exists on the type and concentration of FA that activate PPAR in ruminants. Because in vivo experiments with animals are time-consuming, expensive, and difficult to interpret, the use of an in vitro bovine cell culture model was explored.
The objectives of this study were as follows: (1) to evaluate the responsiveness of Madin-Darby bovine kidney cells (MDBK) to specific PPAR agonists and (2) to test the activation of PPAR in a dose-response manner by specific FA. Results could provide data to improve formulation of rumen-protected FA for the bovine to modulate the stimulatory effect of the PPAR.
Kidney cells in mammals express both the PPAR gene (PPARA) and the PPAR gene (PPARG; Desvergne and Wahli, 1999). The only bovine kidney cell line available is the MDBK (American Type Culture Collection #CCL-22) that was derived from a kidney of an apparently normal adult steer by Madin and Darby (1958). The MDBK cell line used was at passage 230 and was donated by Kristen Stanton (National Animal Disease Center, Ames, IA). Cells were cultivated in a high-glucose complete Dulbecco modified Eagle’s medium without pyruvate (#SC45000-304, VWR, West Chester, PA); the same medium was used for the treatment.
To our knowledge, the responsiveness to PPAR agonists by MDBK cell line has not been explored. To investigate the responsiveness of the MDBK cells to PPAR agonists, Wy-14643 (WY) and rosiglitazone (ROSI) were utilized. The MDBK cells were treated with 50 µM of Wy-14643 and 10 µM of rosiglitazone in a 6-well plate (#353046, BD Biosciences, San Jose, CA) with dimethyl sulfoxide (DMSO; 0.1% vol/vol; #MX1458-6, VWR) as control. The doses of the compounds were determined from several studies in other cell lines and systems. For example, the concentration of these compounds to activate PPAR has been examined in rat, mouse, and human cell lines (Belury et al., 1998; Bility et al., 2004) as well as monkey kidney cells (COS-1; Vanden Heuvel et al., 2006). The fact that we observed altered gene expression in the bovine cells further supports that the concentrations chosen were appropriate. Cells were washed once with 3 mL of phosphate buffer (#16777-247, VWR) and harvested at 0 (control), 6, 12, 18, and 24 h of culture with 1 mL of TRI Reagent (#93289, Sigma, St. Louis, MO). Three replicates were used for each time point. The experimental design was as follows: 3 treatments (WY, ROSI, CTR) x 6 time points x 3 biological replicates (for each time point).
A second experiment was performed in which the MDBK cells were treated with palmitic acid (C16:0), oleic acid (C18:1 cis-9), linoleic acid (C18:2 cis-9,cis-12), linolenic acid (C18:3n-3), and conjugated linoleic acid (CLA; mixture of cis-9, trans-11- and trans-10, cis-12-octadecadienoic acids; all from Sigma, catalog #P0500, O1383, L1268, L2376, and O5507, respectively) at 5 concentrations (10, 25, 50, 100, and 200 µM). All FA except palmitate were diluted in DMSO. Palmitate was insoluble in DMSO and was solubilized in ethanol after saponification with an equimolar amount of NaOH. A baseline control (98 µM or 1 µL/mL of DMSO for the unsaturated FA; 152 µM or 1 µL/mL of ethanol for palmitate) was performed in triplicate. The experimental design was as follows: 5 (FA) x 5 (concentrations) x 3 (biological replicates). Cells were transferred to 6-well plates and were allowed to recover overnight before the treatment that was performed for 24 h. Cells were harvested as described above.
Total RNA was immediately extracted following the protocol suggested by Sigma and was prepared for real-time reverse transcription PCR (quantitative PCR). Briefly, 200 µL of chloroform was added to 1 mL of TRI Reagent + cells, vortexed (15 s), and centrifuged for 15 min at 12,000 x g. The supernatant was removed in a new RNase-DNase-free tube, and 0.5 mL of isopropanol was added and mixed by inverting the tube a few times. The mixture was set to room temperature for 10 min and then centrifuged for >30 min at 12,000 x g and 4°C. The resulting pellet was washed with 75% ethanol, centrifuged, and the dry pellet was resuspended in nuclease-free water. The RNA quantification was performed by spectrophotometer and diluted to obtain 1 µg/µL of final concentration. The reverse transcription was performed using High Capacity cDNA Archive Kit (#4322171, ABI, Foster City, CA) with 1 µg of RNA in 50 µL of total RT solution following the instructions of the manufacturer. The program was 25°C for 10 min and 37°C for 2 h. The cDNA was stored at –20°C.
The quantitative PCR was performed using SYBR Green Master Mix (#4309155, ABI) in a 25-µL reaction (5 µL of cDNA + 1.5 µL of 300 nM forward primer + 1.5 µL of 300 nM reverse primer + 12.5 µL of SYBR Green Master Mix + 4.5 µL of DNase-RNase-free water) with a 5-point 10-fold dilution standard curve. Standard curve was obtained by pooling part of total RNA from all samples. The final data were obtained transforming the cycle threshold (Ct) values using the standard curve.
All primers used were previously published (Table 1
) except for lipoprotein lipase (LPL) that was designed using the free available software Primer3. The PCR product of the primer pair was tested running a 2% agarose gel containing ethidium bromide to check the presence of a single product and primer-dimer. The LPL PCR product was sequenced to verify correct gene amplification.
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The MDBK cells expressed >25-fold greater mRNA abundance of PPARG when compared with PPARA (calculated by 
Ct; median Ct was 23.2 and 28.0, respectively). Both CPT1A and ACOX1 have been shown to be PPAR-specific downstream genes in nonruminant animals (Varanasi et al., 1996; Desvergne et al., 2006). For this reason, the temporal expression of the 2 genes were tested in cells treated with WY, a potent and specific PPAR agonist (Desvergne and Wahli, 1999). The LPL gene has been demonstrated to be activated by PPAR (Desvergne and Wahli, 1999), and the temporal expression of the gene was measured in cells treated with ROSI. All treatments were compared with the DMSO control.
The expression of PPARA and PPARG was not affected by PPAR agonists (data not shown). The MDBK cells exhibited a response to PPAR agonists shown by the increase in expression of CPT1A and LPL (2.5-fold and >200-fold compared with time 0 at 24 and 18 h, respectively; Figure 1). Both genes demonstrated a significant activation at 24 h of incubation compared with time 0 and DMSO treatment. Furthermore, MDKB cells treated with 50 µM of WY for 24 h showed a similar magnitude of CPT1A gene expression induction when compared with 100 µM of the same drug used in mouse primary hepatocytes (Guo et al., 2006). Although the MDBK cells not treated with rosiglitazone presented a very low expression of LPL (almost undetectable; Ct >35), they had a large response in LPL mRNA expression with ROSI treatment, confirming LPL to be a PPAR target gene also in the bovine, as reported for other species (Desvergne and Wahli, 1999). The same concentration of rosiglitazone for 24 h in fetal rat primary brown adipocytes showed a >2-fold increase of LPL (Teruel et al., 2005). The low abundance of LPL in MDBK cells was unexpected, because LPL activity is reported to be elevated in the kidney in nonruminant species (Ruge et al., 2004); however, the low mRNA abundance of LPL in nontreated cells may be due to the very low concentration of FA in the culture medium. The DMSO treatment showed a large effect on LPL induction (
50-fold at 12 h incubation; P < 0.01). The very low mRNA abundance of LPL and the high response to DMSO precluded the use of the gene to test the activation of LPL by fatty FA unless another solvent is utilized. Nevertheless, the increase in LPL expression by ROSI was remarkable and is shown in Figure 1 (lower panel).
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agonists in nonruminant species (Tugwood et al., 1992; Duplus and Forest, 2002). In MDBK cells, expression of ACOX1 was not significantly activated by the PPAR agonist WY (data not shown). The lack of activation of ACOX1 in MDBK cells could be a consequence of cell line immortalization that requires cell adaptation to the in vitro environment. However, the data are puzzling, because MDBK cells present a relative high ACOX1 mRNA abundance. To verify if bovine ACOX1 is a PPAR-responsive gene, an in vivo investigation may be needed or another bovine cell line should be tested.
Results from the previous experiments demonstrated CPT1A to be an appropriate gene to investigate the activation of PPAR. Based on this observation, we measured CPT1A perturbation, in MDBK cells after treatment with 5 different FA, in 5 increasing concentrations for 24 h.
The 16:0, 18:1 cis-9, and 18:2 cis-9, cis-12 FA were chosen based on their high concentrations in bovine plasma (Rukkwamsuk et al., 1999), whereas 18:3n-3 was chosen because its concentration tends to increase in plasma of cows on pasture (Chilliard and Ferlay, 2004). All these FA are well established as PPAR agonists in mice (Desvergne and Wahli, 1999). Bovine plasma presents a noteworthy concentration of CLA (Loor et al., 2005) that originates from ruminal biohydrogenation of polyunsaturated FA (Beam et al., 2000) and by the endogenous activity of steroyl-CoA desaturase on absorbed 18:1 trans-11 (Griinari et al., 2000). Beside the demonstrated negative effect of 18:2 trans-10, cis-12 on expression of genes involved in milk fat synthesis in mammary gland (Peterson et al., 2004), the CLA proved to be a potent PPAR agonist in mice (Takahashi et al., 2003).
Results of the dose-response experiments of the 5 FA in activating PPAR are shown in Figure 2. The CPT1A gene expression was increased by all the FA at different doses (overall effect P < 0.01), except oleic acid (P = 0.25). None of the FA were significantly effective at a doses <100 µM. At 100 µM concentration, only 3 FA showed a significant effect: palmitate was the most potent [2.5-fold vs. control (CTR)] followed by 18:3n-3 and CLA (>1.5- and 1.4-fold vs. DMSO control, respectively). At 200 µM, 18:3n-3 showed the strongest effect (2.5-fold vs. DMSO control) followed by palmitate (2.2-fold vs. CTR); C18:2 cis-9, cis-12; CLA; and C18:1 cis-9 (2.1-, 1.8-, and 1.4-fold vs. DMSO control, respectively). Palmitate at 200 µM tended to have a negative effect on cell survival (almost 1/3 of the cells were detached from the plate after 24 h of incubation). Thus, this effect may have confounded data for CPT1A. The CLA showed a negative effect on CPT1A expression at 10-µM concentration (–2.0-fold vs. CTR; P < 0.01); this could be a consequence of the high concentration of DMSO in the control (98 µM or 1 µL/mL), which corresponded to the dose of DMSO + FA used to obtain the maximum FA concentration (200 µM). In preliminary results, DMSO has been shown to increase CPT1A mRNA abundance in a dose-dependent manner (data not shown). As a consequence, the induction of CPT1A by unsaturated FA could be underestimated.
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(Desvergne and Wahli, 1999). In mice and human, oleic, linoleic, and CLA have been shown to be potent activators of PPAR (Moya-Camarena and Belury, 1999; Khan and Vanden Heuvel, 2003; Kota et al., 2005). In our study, the response to the unsaturated FA was less effective; however, all of the tested FA showed a positive trend in increasing CPT1A expression, and all of them, except for oleic acid, significantly activated PPAR at a dose of 200 µM. At the same dose, oleic acid exhibited a tendency to increase CPT1A mRNA abundance. This suggests that greater concentrations need to be evaluated.
Overall, the results for the time course experiment suggest that MDBK cells are a promising model to investigate the effect of FA on PPAR. Data from the time point trial using specific PPAR agonists suggest that the minimum incubation time of the cells with FA should be 18 h. The CPT1A appears to be a good target gene to test the PPAR activation in MDBK cells. Nevertheless, we are aware that to obtain reliable data, more than one gene should be used to evaluate the activation of PPAR and that primary cells (i.e., bovine hepatocytes) should be a more appropriate model to infer in vivo effects. The LPL showed a tremendous response to ROSI, but the low expression of the gene in MDBK cells remains an issue.
The dose-response experiment allows us to conclude that palmitate is a potent PPAR activator in MDBK cells, as are linolenic and linoleic acids. In our system, 100 µM was the minimum concentration that affected PPAR activation for 16:0, 18:3, and CLA. The oleic and linoleic acids required higher doses (200 µM). The maximum concentration used in the experiment was 200 µM based on previous publications, most of which were performed in mice, a species with an extremely high sensitivity to PPAR activation (Lai, 2004). Our data showed that the sensitivity of MDBK cells for specific PPAR agonists, such as Wy-14643 and ROSI, appears to be comparable to rodents. Moreover, differently than nonruminant species, the PPAR in MDBK cells tended to be less responsive to unsaturated FA compared with palmitate, the only saturated FA tested. It is tempting to propose that bovine PPAR are more sensitive to saturated FA than unsaturated; maybe this is an evolutionary adaptation, because ruminants present high concentrations of saturated FA in plasma. To verify this hypothesis, the bovine MDBK cells should be treated with stearate. However, as mentioned above, the different potency in activating CPT1A expression, between palmitate and unsaturated FA, could be due to the different control group used. More data are required to confirm those preliminary results.
The data generated in these experiments support a role for FA as a candidate for nutrient modulation of metabolism in the bovine. Potential applications might lie in improved nutrition and methods to deal with metabolic problems of dairy cows, especially during the transition from pregnancy to lactation. In fact, PPAR have been suggested to be important for this critical period in dairy cows (Drackley, 1999), and a very recent report uncovered a positive effect of treatment of transition cows with thiazolidinedione, a specific PPAR activator (Smith et al., 2007).
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ACKNOWLEDGEMENTS |
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FOOTNOTES |
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3 Current address: Department of Molecular and Comparative Pathobiology, Johns Hopkins School of Medicine, Baltimore, MD 21205.
Received for publication October 17, 2007. Accepted for publication March 12, 2008.
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REFERENCES |
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9 desaturase. J. Nutr. 130:2285–2291., -β, and -, liver x receptor-β, and retinoid x receptor-. Toxicol Sci. 92:476–489.This article has been cited by other articles:
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  B. J. Thering, M. Bionaz, and J. J. Loor Long-chain fatty acid effects on peroxisome proliferator-activated receptor-{alpha}-regulated genes in Madin-Darby bovine kidney cells: Optimization of culture conditions using palmitate J Dairy Sci, May 1, 2009; 92(5): 2027 - 2037. [Abstract] [Full Text] [PDF]
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