QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) An erratum has been published Purchase Article View Shopping Cart Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Bonnet, M. Articles by Chilliard, Y. Search for Related Content PubMed PubMed Citation Articles by Bonnet, M. Articles by Chilliard, Y.

---

Article

Lipoprotein Lipase Activity and mRNA Are Up-Regulated by Refeeding in Adipose Tissue and Cardiac Muscle of Sheep^{1 ,2}

Muriel Bonnet^*, Christine Leroux, Yannick Faulconnier^*, Jean-François Hocquette^*, François Bocquier^*,3, Patrice Martin and Yves Chilliard^*4

^* INRA, Unité de Recherches sur les Herbivores, Theix, 63122 Saint-Genès-Champanelle, France; and INRA, Laboratoire de Génétique biochimique et de Cytogénétique, 78352 Jouy-en-Josas cedex, France

⁴To whom correspondence should be addressed.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Previous studies in rodents have shown that the lipoprotein lipase (LPL) regulation is complex and often opposite in adipose tissue (AT) and muscle in response to the same nutritional treatment. However, neither LPL responses nor the molecular mechanisms involved in the nutritional regulation have been studied in both AT and muscle of ruminant species. To explore this, we measured the LPL activity and mRNA levels in perirenal AT and cardiac muscle (CM) of control, 7-d-underfed or 14-d-refed ewes. Underfeeding decreased (P < 0.01) LPL activity both in AT (-59%) and CM (-31%), and these activities were restored (P < 0.01) by refeeding (AT, +248%; CM, +34%). Variations of LPL mRNA level measured by real-time reverse transcription-polymerase chain reaction or by Northern blot followed variations of LPL activity: underfeeding decreased AT- and CM-LPL mRNA levels (-58 and -53%, respectively), and refeeding restored (P < 0.01) them in CM (+117%) and increased them over the baseline in AT (+640%). Quantification of either 3.4- or 3.8-kb LPL mRNA levels revealed a predominant (P < 0.001) expression of the 3.4-kb mRNA in AT (60%) and of the 3.8-kb mRNA in CM (56%), without any preferential regulation of one of these mRNA species by the nutritional status. This work reveals a tissue-specific expression pattern of the ovine LPL gene and a pretranslational nutritional regulation of its expression, which is achieved in the same direction in perirenal AT and CM. The different regulation of CM-LPL between ewes and rats probably arises from peculiarities of ruminant species for nutrient digestion and absorption and liver lipogenesis.

KEY WORDS: • LPL gene expression • nutritional status • adipose tissue • cardiac muscle • sheep

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Lipoprotein lipase (LPL)⁵ , the rate-limiting enzyme in triglyceride-rich lipoprotein catabolism, provides triglyceride-derived fatty acids to adipose tissue (AT) for storage and to muscle for energy production. Starvation/refeeding studies in rodents have disclosed a reciprocal regulation of LPL between AT and oxidative muscles, emphasizing the role of LPL in partitioning triglyceride fatty acids between these tissues, according to the needs of the organism. Indeed, during starvation, LPL activity decreases in AT (Bergo et al. 1996 , Cryer and Jones 1979 , Doolittle et al. 1990 , Ladu et al. 1991 , Lee et al. 1998 , Semb and Olivecrona 1986 ) while it increases or remains unchanged in oxidative muscles, depending on the anatomical site, the fiber type composition, the duration of the experiment and animal species (for review see Borensztajn 1987 , Cryer and Jones 1979 , Ong et al. 1994 , Sugden et al. 1993 ). A reverse effect occurs in each tissue after refeeding. Studies performed at the molecular level have disclosed changes in the levels of LPL mRNA, immunoreactive mass and specific activity in response to starvation and refeeding (Bergo et al. 1996 , Doolittle et al. 1990 , Ladu et al. 1991 , Lee et al. 1998 , Ong et al. 1994 ). However, this common reciprocal regulation of LPL may not be a general rule since cardiac muscle-LPL activity was shown to be regulated in the same direction as the AT-LPL activity in starved pigs (Enser 1973 ) and was unaffected in starved guinea pigs (Enerbäck et al. 1988 , Semb and Olivecrona 1986 ). Hence, because no information is available regarding the nutritional regulation of muscle-LPL in ruminant species, as well as the molecular mechanisms involved, the first aim of this study was to examine the effects of underfeeding and refeeding treatments on LPL activity and mRNA levels, in both perirenal AT and cardiac muscle (CM) of ewes.

On the other hand, since there is only one gene encoding LPL (Kirchgessner et al. 1987 ), its nutritional regulation is probably tissue-specific but the molecular mechanisms involved remain unknown. Preliminary studies in humans (Ranganathan et al. 1995 ) and guinea pigs (Enerbäck et al. 1988 ) showed indeed that the expression pattern of the differently sized LPL mRNAs differed between AT and CM. This suggests that polyadenylation sites are used differently, and not randomly, in each tissue during post-transcriptional modifications of the primary LPL transcript, which could be involved in a putative tissue-specific pretranslational regulation of LPL gene expression. Nevertheless, it is not known whether changes in nutritional status affect this tissue-specific expression pattern. To answer to these questions, the second and third aims of this study were: i) to quantify the levels of each mRNA species, after we characterized the 3'untranslated region (3'UTR) terminus of the ovine LPL cDNA and ii) to investigate the effect of underfeeding and refeeding on the tissue-specific expression of these mRNA species. These objectives led us to develop a real-time quantitative reverse transcription-polymerase chain reaction (real-time RT-PCR) assay. This recent methodology has been used in medical applications, but rarely for analyzing the physiological variation of mRNA levels (Sloop et al. 1998 ), and we feel that it is more sensitive than Northern blot and less labor-intensive than the semiquantitative or competitive RT-PCR, to quantify LPL mRNA levels.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Animals and diets.

This study was carried out in accordance with the guidelines of the Animal Care Committee of INRA. Multiparous dry nonpregnant and ovariectomized Lacaune ewes (n = 25) were allotted to one of the three nutritional groups: control, underfed, refed. During the pre-experimental period, the 25 ewes received a control diet for 6 wk, providing 123% of their maintenance energy requirement (MER) calculated on the basis of 0.4 MJ metabolizable energy (ME) · d^-1 · kg body wt^-0.75 (INRA 1989 ). Five ewes (control group) were slaughtered while the remaining 20 ewes were underfed to 22% MER for 7 d. The underfed group (10 ewes) was slaughtered at the end of the underfeeding period, while the remaining ewes were refed, until slaughtering, at 190% MER for 14 d. The diet of the control group consisted of 78% hay and 22% barley grain. The diet of the underfed group consisted of 66% hay and 34% straw. The diet of the refed group consisted of 45% hay and 55% concentrate. The concentrate (expressed as g/kg) was corn, 190; sugar beet pulp, 300; soybean meal, 416; molasses, 20; fish meal, 50; and vitamin-mineral premix, 24. Vitamin-minerals premix (20 g/day) was added to the feed of each group (minerals, g/100 g: Ca, 15; P, 10; Mg, 2; Na, 3; S, 1; trace elements, mg/kg: Zn, 8000; Mn, 6000; I, 50; Co, 10; Se, 10; vitamins, mg/kg: retinyl acetate, 86; cholecalciferol, 1.25; -tocopherol, 134; thiamine hydrochloride, 21). Diets of control and underfed groups were offered at 1000 h, and that of refed group was divided into two equal portions given at 1000 and 1500 h. Ewes had free access to drinking water. Offered feeds and refusals were recorded daily so as to calculate daily intakes of each animal (Table 1 ). Ewes were slaughtered by exsanguination between 0900 and 0930 h, and samples of perirenal AT and CM were immediately either placed at 37°C for adipocyte volume determination or frozen in liquid nitrogen, pending measurements of DNA content, LPL activity and LPL mRNA assays.

Blood samples were collected from the jugular vein the day before slaughter, at 0900 and 1400 h for the determination of plasma insulin (INSI-PR RIA kit; CIS Bio International, Gif-sur-Yvette, France) and metabolites. Plasma levels of glucose, acetate, triglycerides, nonesterified fatty acids (NEFA) and ß-hydroxybutyrate (3-OH-butyrate) were determined with an ELAN auto-analyzer (Merck-Clévenot SA, Nogent-sur-Marne, France) by spectrophotometric enzymatic assays using specific kits (Glucose S-system 100; Merck-Clévenot SA; acid acetic kit, Boehringer Mannheim, Meylan, France; triglycerides GPO-trinder, Sigma, Saint-Quentin Fallavier, France; NEFA C WAKO, Unipath SA, Dardilly, France), except for 3-OH-butyrate which was assayed as described by Barnouin et al. (1986) .

LPL activity.

LPL (EC 3.1.1.34) activity was measured using an artificial emulsion containing ³H-triolein (Amersham, Les Ulis, France) after a detergent (Deoxycholate-Nonidet P40) extraction procedure (Faulconnier et al. 1994 ). Enzyme activity was expressed either on a tissue weight basis, i.e., as nmol of fatty acids released per minute and per gram of AT or CM, per total tissue (perirenal AT and CM), or on a cellular basis, i.e., per 10⁶ adipocytes for AT or per µg DNA for CM, after measurement of adipocyte volume (Robelin 1981 ) or CM-DNA tissue content (Labarca and Paigen 1980 ).

RNA extraction and Northern blot analysis.

Total RNA was extracted as described previously for both AT (Bonnet et al. 1998 ) and CM (Hocquette et al. 1998 ). For each sample, equal amounts of total RNA (40 and 30 µg for AT and CM, respectively) were resolved on a 6.5% formaldehyde-1% agarose gel, transferred to a nylon membrane (GeneScreen; NEN Life Science Products, Le Blanc Mesnil, France) and blotted with a [-³²P]-labeled goat LPL cDNA probe (Bonnet et al. 1998 ). Quantification was performed using a phosphoimager (Molecular Dynamic, Bondoufle, France) and the accompanying software. Quantification of LPL mRNA levels was corrected for variations of the amount of RNA loaded on each lane by using values of hybridization to the [-³²P] oligonucleotide probe for rat 18S ribosomal RNA (Hocquette et al. 1998 ).

Oligonucleotides.⁶

Oligonucleotides were provided by Genosys (Cambs, England) and Oligo express (Paris, France). TaqMan probes were provided by PE Applied Biosystems (Courtaboeuf, France). Primer 1 was designed for the RT step of the 3' rapid amplification of cDNA ends (3'RACE) experiment (see below). Primer 2 was chosen in a sequence segment strictly conserved between cows (Senda et al. 1987 ) and humans (Wion et al. 1987 ), and it was used simultaneously with primer 1 for the PCR step of the 3'RACE experiment (see below). Sequences of primers 3 to 7 and TaqMan LPL probe were deduced from the 3'RACE experiment and used then either to amplify a part of the ovine LPL cDNA 3'UTR (primers 3 and 4) or for the quantification of LPL mRNA (primers 4 to 7) by real-time RT-PCR (see below). Sequences of primers 8 and 9 and TaqMan CYCLO probe were deduced from the characterization of an ovine cyclophilin cDNA fragment and used then for quantification of the cyclophilin mRNA by real-time RT-PCR (see below).

LPL 3' RACE.

Characterization of the LPL cDNA 3'UTR was performed using the 3'RACE technique, essentially as described by Frohman et al. (1988) . From 4 µg total RNA, cDNA was primed with 10 pmol of primer 1 in a final volume of 20 µL containing 50 mmol/L Tris-HCl (pH 8.3), 75 mmol/L KCl, 3 mmol/L MgCl₂, 500 µmol/L of each dNTPs, 10 mmol/L dithiothreitol, 20 U of RNAsin (Promega, Charbonniéres, France) and 100 U of SuperScript reverse transcriptase (Gibco BRL; Life Technologies, Cergy Pontoise, France). After 45 min at 37°C, the cDNA was diluted to 50 µL with sterile water, and PCR was performed. The reaction mix (50 µL) consisted of 5 µL of 10 X PCR buffer (500 mmol/L KCl, 100 mmol/L Tris-HCl, 1% Triton X-100, pH 9.0), 3 µL of 25 mmol/L MgCl₂, 2.5 µL of 5 mmol/L dNTP mix, 25 pmol of each primer, 2 µL of DNA template and 1 U of Taq polymerase (Promega). PCR was conducted for 40 cycles (1 min at 94°C, 2 min at 65°C, 2 min at 72°C) in a DNA 480 thermal cycler (Perkin-Elmer Cetus, Courtaboeuf, France), using primers 1 and 2.

RACE products were phosphorylated with T4 DNA kinase (Pharmacia Biotec, Orsay, France) and cloned into the SmaI-digested and dephosphorylated pGEM-4Z. Escherichia coli DH5 competent cells (Gibco BRL; Life Technologies) were transformed to ampicillin resistance with the ligation reaction mix according to the manufacturer’s protocol. LPL recombinant plasmids were screened by PCR directly on colonies. Sequencing reactions were carried out on recombinant plasmid DNA prepared by the alkaline lysis method. The resulting products were analyzed on polyacrylamide gels using an ABI 373A automated DNA sequencer and the accompanying software (PE Applied Biosystem).

A clone containing a part of the ovine 3'UTR LPL cDNA, called poLPL, was isolated and sequenced. From this sequence, primers 3 and 4 were chosen for PCR amplification of the LPL cDNA distal segment. PCR products were sequenced in both strands.

Real-time quantitative RT-PCR.

Quantification of the levels of 3.4- and 3.8-kb mRNA species was performed by real-time RT-PCR, which utilizes the fluorescent TaqMan methodology and a 7700 Sequence Detector System (PE Applied Biosystems) (Heid et al. 1996 ). The strategy uses the 5'-nuclease activity of the Taq polymerase to cleave a 3'-blocked fluorogenic oligonucleotide probe designed to hybridize specifically to a LPL cDNA molecule. The probe is doubly labeled with two fluorochromes: a reporter and a quencher that absorbs the reporter fluorochrome emission. During the PCR extension phase, the cleavage of the probe results in an increase in the reporter fluorochrome emission (due to the separation of the two fluorochromes) which is plotted vs. PCR cycle number in order to generate an amplification curve for each sample. The first cycle during which the value of a normalized fluorescence parameter (Rn) was significantly higher than a fluorescence threshold was defined as the threshold cycle (C_t) (Fig. 1A ).

View larger version (27K): [in this window] [in a new window]   Figure 1. Polymerase chain reaction (PCR) product detection in real-time. A: Amplification plot from five copy numbers (135625, 67812, 33906, 2118, 529) of lipoprotein lipase (LPL) plasmid. For each concentration, the software calculated a normalized fluorescence parameter values: Rn, using the following equation: Rn = (Rn+) - (Rn-), where Rn+ and Rn- = emission intensity of reporter/emission intensity of passive fluorochrome (included in PCR buffer) for a tube containing all components (Rn+), and for a tube containing all the same components except the cDNA to be amplified (Rn-). This Rn value is plotted against the PCR cycle number. The first cycle during which the fluorescence exceeded a threshold limit (10 times the SD above the mean of baseline emission calculated from cycles 1 to 15) was defined as the threshold cycle (Ct). B: Ct values were plotted as a function of log of known added LPL plasmid copy number and defined. From these data a least-squares regression was calculated, as a calibration curve. Copy numbers for test samples were then automatically calculated from the corresponding CT values. Each point is the mean of triplicate PCR amplifications. Assay acceptability on the R2 value for the calibration curve was >0.98.

For the assays, cDNA was synthesized from 4 µg of total RNA in a final volume of 20 µL containing 10 pmol of oligo(dT)₁₈ and 100 U of SuperScript reverse transcriptase (Gibco BRL; Life Technologies), as described above. Then, cDNA was diluted to 1000 µL with sterile water and aliquoted. Amplification reactions (50 µL) contained cDNA sample (10 µL), 10 X PCR buffer A (5 µL, 500 mmol/L KCl, 100 mmol/L Tris-HCl, 0.1 mol/L EDTA, 600 mmol/L passive fluorochrome rhodamine; pH 8.3), 5 mmol/L MgCl₂, 200 µmol/L dATP, dCTP, dGTP, dUTP, 40 pmol of each primer, 10 pmol of probe, 1.25 U AmpliTaq Gold DNA polymerase (PE Applied Biosystem), and 0.5 U AmpErase uracil N-glycosylase. Either sum of 3.4- and 3.8-kb LPL mRNAs or only 3.8-kb LPL mRNA were amplified using TaqMan LPL probe and primer pairs 3–5 or 6–7 (see results, and Fig. 4B ). The cycling conditions included 2 min at 50°C and 10 min at 95°C for thermal activation of the AmpliTaq Gold DNA polymerase. Thermal cycling proceeded with 45 cycles at 95°C for 10 s and at 60°C for 2 min. Each assay was performed in triplicate.

View larger version (40K): [in this window] [in a new window]   Figure 4. Partial nucleotide sequence of ovine lipoprotein lipase (LPL) cDNA, and positioning of the primers used for the real-time reverse transcription-polymerase chain reaction (PCR) assays. A: Alignment of nucleotide sequence of a partial ovine cDNA (o) spanning 779 bp of the LPL mRNA 3'UTR with its bovine (b) and human (h) counterparts. Sequence accession numbers are M16966 and M15856 for bovine mammary gland (Senda et al. 1987 ) and human adipose tissue (Wion et al. 1987 ) sequences, respectively. Gaps (.) have been placed to maximize the similarity. Dashes (-) correspond to nucleotides that are identical to those of the ovine LPL sequence. The putative polyadenylation signals are dimmed. B: schematic representation of the ovine LPL cDNA 3'UTR (solid bar). Arrows indicate the position of the primers for PCR: i) to yield by 3'RACE (primers 1 and 2) the end of the 3'UTR of ovine LPL cDNA, that was subsequently cloned (poLPL clone), ii) to amplify (primers 3 and 4) the 779-bp fragment, that was subsequently sequenced, iii) to quantify levels of either the 3.4- plus 3.8-kb LPL mRNAs (primer 3 and 5) or the 3.8-kb LPL mRNA alone (primers 6 and 7). The position of the TaqMan LPL probe (framed) is also given. The open bar represents the last 70 bp of the coding sequence. The dimmed areas depict the two polyadenylation signals characterized in Figure 4A .

To decrease the effects of the variability in amount of starting total RNA or RT efficiency, the LPL mRNA copy number was normalized by the mRNA copy number of the constitutively-expressed cyclophilin gene. The cyclophilin copy number of AT and CM samples was determined using a calibration curve prepared by amplifying serial dilutions of a recombinant plasmid containing a part of the goat cyclophilin cDNA (Le Provost et al. 1996 ). Quantification assays for cyclophilin cDNA were performed by real-time RT-PCR using conditions described above for LPL mRNA. The TaqMan CYCLO probe and primers 8 and 9 were chosen in a sequence segment strictly conserved between goat and ewe (data not shown), after partial sequencing of ovine cyclophilin cDNA.

Statistical analysis.

Differences between two nutritional treatment groups were tested using the nonparametric Mann and Whitney-U test with differences considered significant when P < 0.05. Specific comparisons were made between: i) the control and the underfed groups, ii) the control and refed groups and iii) the underfed and refed groups.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Effects of nutritional status on empty body weight, AT- and CM-weight and cellularity, and plasma insulin and metabolites.

Empty body weight (i.e., body weight minus weight of digestive content), perirenal AT- and CM-weight and cellularity, were not significantly modified by nutritional status (Table 2 ).

LPL activity, expressed on a per cell basis, was significantly (P < 0.01) modulated by nutritional status, in the same direction but with different amplitudes in AT and CM (Fig. 2 ). Underfeeding (vs. control ewes) decreased AT- (-59%, P < 0.01) and CM- (-31%, P < 0.01) LPL activities. Refeeding (vs. underfed ewes) restored these activities in both tissues (AT: +248%, P < 0.01; CM: +34%, P < 0.01), to levels not different from those in the control group. Similar effects were observed when AT- and CM-LPL activities were expressed per gram of AT or CM, or per whole tissues (results not shown).

View larger version (17K): [in this window] [in a new window]   Figure 2. Effect of nutritional status on (A) adipose tissue and (B) cardiac muscle lipoprotein lipase activity in control (n = 5), underfed (n = 10) and refed (n = 10) ewes. Means (±SEM) with different superscripts are different, P < 0.01.

Northern blot analysis revealed two LPL mRNA species at 3.4 and 3.8 kb, both for AT and CM (Fig. 3A and B ). Scanning densitometry of CM Northern blots revealed a 78% (P < 0.01) increase in the LPL mRNA/18S rRNA ratio for refed vs. underfed ewes (Fig. 3B) . There was a slight increase in the 18S signal, and a dramatic increase in LPL mRNA levels in AT of refed ewes. However, it was not possible with Northern blot analysis to quantify precisely the effect of refeeding on AT-LPL mRNA (due to the very low level of expression in underfed ewes), nor on the respective levels of 3.4- and 3.8-kb LPL mRNA in AT and CM (due to the small difference in size and, hence, to the proximity between the two transcripts). To address these issues, we developed a real-time RT-PCR protocol based on the use of a TaqMan probe and primers chosen after sequencing the 3'UTR of the ovine LPL cDNA.

View larger version (51K): [in this window] [in a new window]   Figure 3. Northern blot analysis of the effects of refeeding on lipoprotein lipase (LPL) mRNA levels in ovine adipose tissue and cardiac muscle. Total RNA was extracted from tissues of five underfed and five refed ewes and analyzed by Northern blotting with 32P-labeled probes for LPL and 18S. A: representative Northern blot of total RNA from adipose tissue; B: representative Northern blot of total RNA from cardiac muscle; LPL mRNA and 18S rRNA levels from cardiac muscle were quantified by densitometry in underfed and refed ewes in order to calculate LPL mRNA/18S rRNA ratio; * P < 0.01.

Two polyadenylation signals were identified in the 779-bp fragment of the 3'UTR of the ovine LPL cDNA that we sequenced (Fig. 4A and B ). Comparison with other known LPL sequences revealed that the ovine sequence has 91 and 70% similarity with the bovine and human sequences, respectively (Fig. 4A ). Primer pairs 3–5 or 6–7, specific either for both the 3.4- and 3.8-kb LPL mRNAs, or only for the 3.8-kb LPL mRNA, were chosen to yield amplicons of 335 and 336 bp, respectively (Fig. 4B) . As a reliable quantification requires that primer pairs 3–5 or 6–7 allow amplification of the LPL cDNA with an equal efficiency, this was checked by amplifying 2118, 33906 and 133565 copies of the poLPL recombinant plasmid. Differences between the fluorescence values (Ct) obtained with either primer pair 3–5 or primer pair 6–7 were <3%. Indeed, fluorescence values (Ct) were 30.0 ± 0.2, 25.3 ± 0.1 and 23.1 ± 0.1 Ct with primer pair 3–5, and 30.3 ± 0.1, 26.1 ± 0.2 and 23.3 ± 0.2 Ct with primer pair 6–7, for the three known plasmid copy numbers, respectively.

Effect of nutritional status on LPL mRNA levels.

The AT level of 3.4-kb plus 3.8-kb LPL mRNAs (Fig. 5 ) tended to be decreased (P = 0.1; -58%, vs. control ewes) by underfeeding and was dramatically increased by refeeding (+640%, vs. underfed ewes, P < 0.01). Likewise, the CM level of 3.4-kb plus 3.8-kb LPL mRNAs (Fig. 5) was decreased (-53%, P < 0.01, vs. control ewes) significantly by underfeeding, and refeeding reversed this effect (+117%, P < 0.01, vs. underfed ewes). Therefore, the levels of 3.4- plus 3.8-kb LPL mRNAs changed in a similar manner in AT and CM during underfeeding and refeeding, although these effects were more marked in AT than in CM. Indeed, LPL mRNAs in AT increased over the baseline with refeeding, whereas they did not in CM. Similar ranges of variation were observed for levels of either the 3.4- or the 3.8-kb LPL mRNA, both in AT and CM (Table 4 ). However, the levels of each CM-LPL mRNA species did not decrease significantly during underfeeding, probably due to the small number (n = 5) of ewes in the control group.

View larger version (15K): [in this window] [in a new window]   Figure 5. Effect of nutritional status on (A) adipose tissue and (B) cardiac muscle 3.4- plus 3.8-kb lipoprotein lipase (LPL) mRNA levels, determined using a real-time quantitative reverse transcription-polymerase chain reaction protocol. LPL and cyclophilin mRNA levels were measured on tissues of control (n = 5), underfed (n = 10) and refed (n = 10) ewes in order to calculate LPL/cyclophilin mRNA ratio. Means (±SEM) with different superscripts differ, P < 0.01.

The level of the 3.4 kb mRNA, as a proportion of the sum of 3.4- plus 3.8-kb mRNAs, was significantly (P < 0.001) higher in AT (60%) than in CM (44%; Fig. 6 ). However, expression patterns were not significantly affected by nutritional status (Fig. 6) .

View larger version (48K): [in this window] [in a new window]   Figure 6. Expression pattern of the 3.4 and 3.8 kb lipoprotein lipase (LPL) mRNAs in sheep adipose tissue (AT) and cardiac muscle (CM). The levels of either the sum of 3.4- plus 3.8-kb LPL mRNAs or only the 3.8-kb species were assayed in AT and CM of control (C, n = 5), underfed (U, n = 10) and refed (R, n = 10) ewes by real-time reverse transcription-polymerase chain reaction, as described in the Materials and Methods section. The level of the 3.4-kb LPL mRNA was obtained by difference. Means (± SEM) with different letters differ, P < 0.001.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

We carried out mid-length (1 to 2 wk) nutritional treatments that were too short to induce significant variations of empty body weight, and AT and CM weight and cellularity, but sufficient to induce the variations of plasma metabolites and insulin that are expected during underfeeding-refeeding experiments in ruminants (for review see Chilliard et al. 1995 ). Refeeding decreased plasma NEFA arising from fat mobilization and increased plasma insulin and metabolites arising from food digestion: acetate, glucose and triglycerides, as well as postprandial (1400 h) 3-OH-butyrate arising from ruminal ketogenesis (dietary origin). The decrease in preprandial (0900 h) 3-OH-butyrate level after refeeding probably reflects a decreased hepatic ketogenesis from mobilized NEFA (for review see Chilliard et al. 1995 ).

Effect of nutritional status on LPL gene expression.

Our results in ovine species are in agreement with previous studies (Bonnet et al. 1998 , Chilliard et al. 1979 , DiMarco et al. 1981 , Tume et al. 1983 ) reporting that ruminant AT-LPL activity is down-regulated by underfeeding and up-regulated by refeeding, but demonstrate for the first time in sheep that nutritional status regulates CM-LPL in the same direction as AT-LPL activity, although with a lower magnitude.

Changes in AT-LPL activity, together with changes in the levels of LPL mRNA measured by real-time RT-PCR, suggest a pretranslational regulation of AT-LPL gene, which confirms previous Northern blot results on AT of 8-d-underfed ewes refed for 10 d (Bonnet et al. 1998 ). Nutritional studies in monogastric species have also demonstrated a pretranslational regulation of LPL expression in AT (Cooper et al. 1989 , Enerbäck et al. 1988 , Ladu et al. 1991 , Lee et al. 1998 ) although some post-translational regulation also occurs (Doolittle et al. 1990 , Lee et al. 1998 ). Furthermore, the similar range of variation for plasma insulin level (63% decrease with underfeeding, 171% increase with refeeding, Table 3 ) and LPL activity (59% decrease with underfeeding, 248% increase with refeeding, Fig. 2 ), and mRNA levels (58% decrease with underfeeding, 640% increase with refeeding, Fig. 5 ), suggests that dietary actions could be mediated at least in part by insulin in sheep, as in monogastric species (for review see Enerbäck and Gimble 1993 ).

The downregulation of ovine CM-LPL activity by a strong underfeeding treatment (22% of MER during 7 d) contrasts with results reported for monogastric species, except for those from the pig. In agreement with our 31%- decrease in CM-LPL activity, a 35%- decrease was indeed reported from 48 h-starved pigs (Enser 1973 ). In contrast, depriving guinea pigs of feed for 48 h did not cause any significant change in CM-LPL activity (Enerbäck et al. 1988 ). In starved rats, an increase (two- to four-fold) in CM-LPL activity was generally reported (Cryer and Jones 1979 , Doolittle et al. 1990 , Ladu et al. 1991 , Sugden et al. 1993 ), but a lack of variation was also described depending on experimental conditions, and noticeably for starvation duration (Borensztajn et al. 1970 , Ladu et al. 1991 ). Increases in CM-LPL activity were also measured in 24-h starved rabbits (+34%, Cryer and Jones 1979) and chickens (+250%, Husbands 1972). The up-regulation (by a factor of 1.4) of sheep CM-LPL activity during refeeding contrasts with the decrease (-57%) or the lack of variation observed when refeeding starved rats (Doolittle et al. 1990 , Sugden et al. 1993 ). Furthermore, our data strongly suggest that underfeeding and refeeding regulate ovine CM-LPL at a pretranslational level since the level of CM-LPL mRNA is modulated in a range similar to that of enzyme activity. This pretranslational regulation of ovine CM-LPL by nutritional status is in agreement with previous Northern blot results obtained in starved rats (Ladu et al. 1991 , Ong et al. 1994 ) and chickens (Cooper et al. 1989 ). Although previous studies have analyzed the possible effects of plasma glucose, glucagon, glucagon/insulin ratio or catecholamines (for review see Borensztajn 1987 ), the factor which mediates dietary actions on CM-LPL gene expression has not been identified.

Physiological significance of the parallel regulation of LPL in ovine adipose tissue and cardiac muscle.

In fed ruminants, muscles utilize mainly acetate and glucose as energy sources (Pethick 1984 ). During underfeeding the availability of these substrates decreases, whereas the plasma concentrations of NEFA mobilized from AT and of preprandial 3-OH-butyrate from hepatic origin increase (Table 3) . This allows the ruminant muscles to increase considerably the use of NEFA and ketone bodies (Pethick 1984 ), as suggested by the 82%- higher 3-hydroxybutyrate dehydrogenase activity in sheep than rat CM (Malki et al. 1992 ). An alternative metabolic adaptation is the recycling by the liver of plasma NEFA as triglyceride-rich lipoproteins. This pathway seems to be important in the starved rat in which plasma triglycerides slightly decreased (Sugden et al. 1993 ) or remained unchanged (Björntorp et al. 1982 ), thus maintaining substrate availability for CM-LPL. This contrasts with the sharp decrease of triglyceridemia in underfed ewes (present study), that is probably related to the low capacity of ruminant liver to synthesize fatty acids and to secrete triglycerides (Pullen et al. 1990 ). There is indeed a striking correlation (Fig. 7 ) across animal species between the response of the CM-LPL activity to starvation or underfeeding (Borensztajn et al. 1970 , Cryer and Jones 1979 , Doolittle et al. 1990 , Enerbäck et al. 1988 , Enser 1973 , Husbands 1972 , Ladu et al. 1991 , Sugden et al. 1993 and present results), and the hepatic ability to secrete triglycerides (Pullen et al. 1990 ). Although this striking correlation does not prove a cause and effect relationship, it does suggest that there is a direct or indirect link between the nature of available energy substrates in each species, and the interspecies differences in the response of CM-LPL activity to underfeeding.

View larger version (22K): [in this window] [in a new window]   Figure 7. Interspecies relationship between the ability of liver to secrete triglycerides (TG) and the change in cardiac muscle lipoprotein lipase (CM-LPL) activity during starvation or underfeeding. Secretion of TG was determined in vitro from liver slices incubated with [14C]palmitate and [3H]oleate (Pullen et al. 1990 ). The response of CM-LPL activity was measured in pigs (Enser 1973 ), sheep (present study), guinea pigs (Enerbäck et al. 1988 ), rabbits (Cryer and Jones 1979 ), rats (means ± SEM, from Borensztajn et al. 1970, Cryer and Jones 1979, Doolittle et al. 1990, Ladu et al. 1991, Sugden et al. 1993), and chickens (Husbands 1972 ).

Two LPL mRNA species of 3.4- and 3.8-kb were identified in both AT and CM, by Northern blot analysis, in agreement with previous results in ovine AT (Bonnet et al. 1998 ) and bovine AT or muscle (Bonnet et al. 1998 , Hocquette et al. 1998 ). In these studies a third 1.7-kb mRNA was also identified, but in very low amounts, which probably explains why it was not detected in present Northern blots. In other respects, the presence of the 1.7- and 3.4-kb LPL mRNA in ruminant species was explained previously by the characterization of two corresponding polyadenylation signals on the partial bovine LPL cDNA (Senda et al. 1987 ). We characterized the end of the 3'UTR of the ovine LPL cDNA, and identified a third polyadenylation signal, that explains the presence of the 3.8-kb LPL mRNA in ruminant tissues. Furthermore, this characterization of the 3'UTR sequence of ovine LPL cDNA allowed us to develop a real-time RT-PCR procedure to quantify precisely, the levels of either the 3.4- or the 3.8-kb LPL mRNA. Although both of them were present in AT and CM, the 3.4-kb mRNA dominated in AT, and the 3.8-kb in CM, confirming the tissue-specificity in the expression pattern of the two major LPL mRNAs that was suggested by qualitative observations in guinea pig (Enerbäck et al. 1988 ) and human (Ranganathan et al. 1995 ) tissues. Furthermore, the quantification of each mRNA species in sheep during underfeeding-refeeding shows, for the first time, the lack of a preferential regulation by the nutritional status of one of these mRNA in AT and CM, although total LPL mRNA levels were sharply affected.

In conclusion, we have shown that in sheep, like in pigs, AT- and CM-LPL are regulated in the same direction by nutritional status, contrary to the situation in rabbits, rats and chickens, which might be related to species differences in the liver’s ability to synthesize and to secrete fatty acids as triglyceride-rich lipoproteins. These species differences in LPL regulation may help to identify the mediator responsible for CM-LPL nutritional regulation, assuming that this mediator is the same, but differently regulated, depending on the animal species. It may be of practical interest to control LPL gene expression in muscle and AT, and the nutrient partitioning between tissues, to favor protein accretion instead of fat accretion. Furthermore, our data show a tissue-specific LPL gene expression in sheep, that it is not modulated by the nutritional status. Future studies at the translational level are needed to elucidate if this tissue-specific expression pattern of LPL mRNA has physiological importance.

	ACKNOWLEDGMENTS

 
We thank A. Ollier, J. P. Pezant and their team for the management of the animals, and G. Cuylle and his team for slaughtering the animals; J. Fléchet and N. Guivier for total RNA extraction and DNA measurement; M. Tourret for adipocyte volume and plasma insulin and metabolites measurements, as well as F. Le Provost for kindly providing us with the primers, goat sequence for cyclophilin and goat cyclophilin recombinant plasmid, and C. Giraud-Delville for performing sequencing.

	FOOTNOTES

 
¹ Presented in part at <<Journées Francophones de Nutrition>>, November 1997, Paris, France [Bonnet, M., Hocquette, J. F., Faulconnier, Y., Fléchet, J., Bocquier, F. & Chilliard, Y. (1998) Nutritional regulation of lipoprotein lipase activity and its messenger RNAs in ewe adipose tissue and heart. Reprod. Nutr. Dev. 38: 197 (abs.)]; and at <<3^rd French-British Meeting on Nutrition>>, Septembre-Octobre 1998, Nancy, France, [Bonnet, M., Faulconnier, Y., Leroux, C., Bocquier, F., Martin, P. & Chilliard, Y. (1999) Effect of refeeding on two mRNAs species of lipoprotein lipase in adipose tissue and cardiac muscle of the sheep. Proc. Nutr. Soc. 58: 108A (abs.)].

² This work was financially supported by an INRA grant for studies on <<Lipogenesis in farm animals>>.

³ Present address: ENSAM-INRA, Productions animales, 2 Place Viala, 34060 Montpellier Cedex 2, France.

⁵ Abbreviations used: AT, adipose tissue; CM, cardiac muscle; LPL, lipoprotein lipase; ME, metabolizable energy; MER, maintenance energy requirement; NEFA, nonesterified fatty acids; PDI, protein digestible in the intestine; RACE, rapid amplification of cDNA end; RT-PCR, reverse transcription-polymerase chain reaction; 3-OH-butyrate, 3-hydroxybutyrate; 3'UTR, 3'untranslated region.

⁶ Primer 1: 5'-TCAAGCTTCTGCAGGATCCTTTTTTTTTTTTTTTTT-3' Primer 2: 5'- GTATAGTGGCCAAATAGCACA -3' Primer 3: 5'-AACTAGTCAAGAGTGAGTGAAC-3' Primer 4: 5'-TTTGTAATAAAATGTTGTCAGTT-3' Primer 5: 5'-AGTAGAATGAATGCTGTGATTGACAT-3' Primer 6: 5'-TTCAGAGGCTATTACTGGAAATCC-3' Primer 7: 5'-CATTAATTCTCGGGATTTGGG -3' Primer 8: 5'-TCACCACCCTGACACATAAATCC -3' Primer 9: 5'-CAAGATGCCAGGACCTGTATG-3' TaqMan LPL probe: 5'-TTCCAGTGGTGCCGGAACACTCCTTC-3' TaqMan CYCLO probe: 5'-TCTCCCCATAGATGGACTTGCCACCAGT -3'

Manuscript received August 30, 1999. Initial review completed October 22, 1999. Revision accepted December 15, 1999.

	REFERENCES

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1. Barnouin J., El Idilbi N., Chilliard Y., Chacornac J. P., Lefaivre R. Micro-dosage automatisé sans déprotéinisation du 3-hydroxybutyrate plasmatique chez les bovins. Ann. Rech. Vet. 1986;17:129-139[Medline]

2. Bergö M., Olivecrona G., Olivecrona T. Forms of lipoprotein lipase in rat tissues: in adipose tissue the proportion of inactive lipase increases on fasting. Biochem. J. 1996;313:893-898

3. Björntorp P., Edström S., Kral J. G., Lundholm K., Presta E., Walks D., Yang M. U. Refeeding after fasting in the rat energy substrate fluxes and replenishment of energy stores. Am. J. Clin. Nutr. 1982;36:450-456[Abstract/Free Full Text]

4. Bonnet M., Faulconnier Y., Fléchet J., Hocquette J. F., Leroux C., Langin D., Martin P., Chilliard Y. Messenger RNAs encoding lipoprotein lipase, fatty acid synthase and hormone-sensitive lipase in the adipose tissue of underfed-refed ewes and cows. Reprod. Nutr. Dev. 1998;38:297-307

5. Borensztajn J. Heart and skeletal muscle lipoprotein lipase. Borensztajn J. eds. Lipoprotein Lipase 1987:133-148 Evener Publishers Chicago, IL.

6. Borensztajn J., Otway S., Robinson D. S. Effect of fasting on the clearing factor lipase (lipoprotein lipase) activity of fresh and defatted preparations of rat heart muscle. J. Lipid Res. 1970;11:102-110[Abstract]

7. Chilliard Y., Doreau M., Bocquier F., Lobley G. E. Digestive and metabolic adaptations of ruminants to variations in food supply. Journet M. Grenet E. Farce M. H. Thériez M. Demarquilly C. eds. Recent developments in nutrition of herbivores. Proceedings of the IVth International Symposium on the Nutrition of Herbivores 1995:329-360 INRA Editions Paris

8. Chilliard Y., Sauvant D., Morand-Fehr P. Goat mammary, adipose and milk lipoprotein lipases. Ann. Rech. Vet. 1979;10:401-403[Medline]

9. Cooper D. A., Stein J. C., Strieleman P. J., Bensadoun A. Avian adipose lipoprotein lipase: cDNA sequence and reciprocal regulation of mRNA levels in adipose tissue and heart. Biochim. Biophys. Acta 1989;1008:92-101[Medline]

10. Cryer A., Jones H. M. The distribution of lipoprotein lipase (clearing factor lipase) activity in the adiposal, muscular and lung tissue of ten animal species. Comp. Biochem. Physiol. 1979;63B:501-505

11. DiMarco N. M., Bietz D. C., Whitehurst G. B Effect of fasting on free fatty acid, glycerol and cholesterol concentrations in blood plasma and lipoprotein lipase activity in adipose tissue of cattle. J. Anim. Sci. 1981;52:75-82

12. Doolittle M. H., Ben-Zeev O., Elovson J., Martin D., Kirchgessner T. G. The response of lipoprotein lipase to feeding and fasting. J. Biol. Chem. 1990;265:4570-4577[Abstract/Free Full Text]

13. Enerbäck S., Gimble J. M. Lipoprotein lipase gene expression: physiological regulators at the transcriptional and post-transcriptional level. Biochim. Biophys. Acta 1993;1169:107-125[Medline]

14. Enerbäck S., Semb H., Tavernier J., Bjursell G., Olivecrona T. Tissue specific regulation of guinea-pig lipoprotein lipase; effect of nutritional state and tumor necrosis factor on mRNA levels in adipose tissue, heart and liver. Gene 1988;64:97-106[Medline]

15. Enser M. Clearing-factor lipase in muscle and adipose tissue of pigs. Biochem. J. 1973;136:381-385[Medline]

16. Faulconnier Y., Thévenet M., Fléchet J., Chilliard Y. Lipoprotein lipase and metabolic activities in incubated bovine adipose tissue explants. Effects of insulin, dexamethasone, and fetal bovine serum. J. Anim. Sci. 1994;72:184-191[Abstract]

17. Frohman M. A., Dush M. K., Martin G. R. Rapid production of full-length cDNAs from rare transcripts: amplification using a single gene-specific oligonucleotide primer. Proc. Natl. Acad. Sci. USA 1988;85:8998-9002[Abstract/Free Full Text]

18. Heid C. A., Stevens J., Livak K. J., Williams P. M. Real time quantitative PCR. Genome Res 1996;6:995-1001[Abstract/Free Full Text]

19. Hocquette J. F., Graulet B., Olivecrona T. Lipoprotein lipase activity and mRNA levels in bovine tissues. Comp. Biochem. Physiol. 1998;121:201-212

20. Husbands D. R. The distribution of lipoprotein lipase in tissue of the domestic fowl and the effects of feeding and starving. Br. Poult. Sci. 1972;13:85-90[Medline]

21. Institut. National de la Recherche Agronomique (INRA) Recommended allowances and feed tables. Jarrige R. eds. Ruminant Nutrition 1989 John Libley Eurotext Paris.

22. Kirchgessner T. G., Svenson K. L., Lusis A. J., Schotz M. C. The sequence of cDNA encoding lipoprotein lipase. J. Biol. Chem. 1987;262:8463-8466[Abstract/Free Full Text]

23. Labarca C., Paigen K. A. Simple, rapid, sensitive DNA assay procedure. Anal. Biochem. 1980;102:344-352[Medline]

24. Ladu M. J., Kapsas H., Palmer W. K. Regulation of lipoprotein lipase in adipose and muscle tissues during fasting. Am. J. Physiol. 1991;260:R953-R959[Abstract/Free Full Text]

25. Le Provost F., Lépingle A., Martin P. A survey of the goat genome transcribed in the lactating mammary gland. Mamm. Genome 1996;7:657-666[Medline]

26. Lee J. J., Smith J. P., Fried S. K. Mechanisms of decreased lipoprotein lipase activity in adipocytes of starved rats depend on duration of starvation. J. Nutr. 1998;128:940-946[Abstract/Free Full Text]

27. Malki M. C., Kante A., Demigne C., Latruffe N. Expression of R-3-hydroxybutyrate dehydrogenase, a ketone body converting enzyme in heart and liver mitochondria of ruminant and non-ruminant mammals. Comp. Biochem. Physiol. 1992;101B:413-420

28. Ong J. M., Simsolo R. B., Saghizadeh M., Pauer A., Kern P. A. Expression of lipoprotein lipase in rat muscle: regulation by feeding and hypothyroidism. J. Lipid Res. 1994;35:1542-1551[Abstract]

29. Pethick D. W. Energy metabolism of skeletal muscle. Gawthorne J. M. Baker S. K. Mackintosh J. B. Purser D. B. eds. Ruminant Physiology, Concepts and Consequences 1984:431-441 University of Western Australia Nedlands.

30. Pullen D. L., Liesman J. S., Emery R. S. A species comparison of liver slice synthesis and secretion of triacylglycerol from nonesterified fatty acids in media. J. Anim. Sci. 1990;68:1395-1399[Abstract]

31. Ranganathan G., Ong J. M., Yukht A., Saghizadeh M., Simsolo R. B., Pauers A., Kern P. A. Tissue-specific expression of human lipoprotein lipase. Effect of the 3`-untranslated region on translation. J. Biol. Chem. 1995;270:7149-7155[Abstract/Free Full Text]

32. Robelin J. Cellularity of bovine adipose tissue: developmental changes from 15 to 65 percent mature weight. J. Lipid Res. 1981;22:452-457[Abstract]

33. Semb H., Olivecrona T. Nutritional regulation of lipase in guinea-pig tissues. Biochim. Biophys. Acta 1986;876:249-255[Medline]

34. Senda M., Oka K., Brown W. V., Qasba P. K. Molecular cloning and sequence of a cDNA coding for bovine lipoprotein lipase. Proc. Natl. Acad. Sci. USA 1987;84:4369-4373[Abstract/Free Full Text]

35. Sloop K. W., Surface P. L., Heiman M. L., Slieker L. J. Changes in leptin expression are not associated with corresponding changes in CCAAT/Enhancer Binding Protein-. Biochem. Biophys. Res. Comm. 1998;251:142-147[Medline]

36. Sugden M. C., Holness M. J., Howard R. M. Changes in lipoprotein lipase activities in adipose tissue, heart and skeletal muscle during continuous or interrupted feeding. Biochem. J. 1993;292:113-119

37. Tume R. K., Thornton R. F., Johnson G. W. Lipoprotein lipase of sheep and rat adipose tissues. Aust. J. Biol. Sci. 1983;36:41-48[Medline]

38. Wion K. L., Kirchgessner T. G., Lusis A. J., Schotz M. C., Lawn R. M. Human lipoprotein lipase complementary DNA sequence. Science 1987;235:1638-1641[Abstract/Free Full Text]

This article has been cited by other articles:

				M. Bonnet, Y. Faulconnier, C. Leroux, C. Jurie, I. Cassar-Malek, D. Bauchart, P. Boulesteix, D. Pethick, J. F. Hocquette, and Y. Chilliard Glucose-6-phosphate dehydrogenase and leptin are related to marbling differences among Limousin and Angus or Japanese Black x Angus steers J Anim Sci, November 1, 2007; 85(11): 2882 - 2894. [Abstract] [Full Text] [PDF]

				C. Jurie, I. Cassar-Malek, M. Bonnet, C. Leroux, D. Bauchart, P. Boulesteix, D. W. Pethick, and J. F. Hocquette Adipocyte fatty acid-binding protein and mitochondrial enzyme activities in muscles as relevant indicators of marbling in cattle J Anim Sci, October 1, 2007; 85(10): 2660 - 2669. [Abstract] [Full Text] [PDF]

				S. Ollier, C. Robert-Granie, L. Bernard, Y. Chilliard, and C. Leroux Mammary Transcriptome Analysis of Food-Deprived Lactating Goats Highlights Genes Involved in Milk Secretion and Programmed Cell Death J. Nutr., March 1, 2007; 137(3): 560 - 567. [Abstract] [Full Text] [PDF]

				L. Bernard, J. Rouel, C. Leroux, A. Ferlay, Y. Faulconnier, P. Legrand, and Y. Chilliard Mammary Lipid Metabolism and Milk Fatty Acid Secretion in Alpine Goats Fed Vegetable Lipids J Dairy Sci, April 1, 2005; 88(4): 1478 - 1489. [Abstract] [Full Text] [PDF]

				S. H. Lee and K. L. Hossner Coordinate regulation of ovine adipose tissue gene expression by propionate J Anim Sci, November 1, 2002; 80(11): 2840 - 2849. [Abstract] [Full Text] [PDF]

				R. B. Eckhardt Genetic Research and Nutritional Individuality J. Nutr., February 1, 2001; 131(2): 336S - 339. [Abstract] [Full Text]

This Article Abstract Full Text (PDF) An erratum has been published Purchase Article View Shopping Cart Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar