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Lipoprotein lipase activity and mRNA levels in bovine tissues
Comparative Biochemistry and Physiology Part B 121 (1998) 201 – 212
Lipoprotein lipase activity and mRNA levels in bovine tissues Jean-Franc¸ois Hocquette a,*, Benoıˆt Graulet a, Thomas Olivecrona b a
INRA, Laboratoire Croissance et Me´tabolismes des Herbi6ores, Theix, 63122 Saint-Gene`s Champanelle, France b Department of Physiological Chemistry, Uni6ersity of Umea˚, S-901 87 Umea˚, Sweden Received 20 March 1998; received in revised form 3 July 1998; accepted 13 August 1998
Abstract Lipoprotein lipase (LPL) in cattle has been extensively studied in adipose tissue, milk and mammary gland, but only to a limited extent in muscles. Therefore, we have adapted our in vitro LPL assay method for the measurement of LPL activity and describe, for the first time, sensitive procedures to quantify LPL activity and mRNA levels in bovine muscles. In vitro activation of bovine LPL activity is approximately 5-fold greater with rat than with bovine sera for heart and muscles, but not for adipose tissues. Values of LPL activity are in the upper range of those previously reported for rat or bovine tissues. With rat serum as activator, LPL activity in the heart of seven calves (662–832 mU g − 1) is at least 3-fold lower than in the rat heart (2150 – 2950 mU g − 1, PB 0.05). LPL activity is higher in bovine heart and oxidative muscles (412 – 972 mU g − 1), except the diaphragm, than in mixed or glycolytic muscles (33–154 mU g − 1, P B0.05). The levels of LPL transcripts are positively related to LPL activity in bovine tissues, including muscles and adipose tissues. © 1998 Elsevier Science Inc. All rights reserved. Keywords: Lipoprotein lipase activity; Gene expression; LPL; Muscles; Adipose tissues; Bovine
1. Introduction Triacylglycerols (TG) are unloaded from chylomicrons and very low density lipoproteins through hydrolysis by lipoprotein lipase (LPL) on capillary endothelial surfaces, where circulating lipoproteins bind briefly. Fatty acids liberated by LPL are available for tissues as energy sources especially in muscles, or for storage in the form of TG in adipose tissues. It is generally assumed that the rate-limiting factor for energy delivery from lipoproteins is the amount of active LPL available at the endothelium which is correlated to LPL activity in tissue cells (for review, see [7,14,29]). This may be true in most physiological conditions [30] except in case of fat overload [32]. Therefore, tissue LPL and its regulation became the subject of active investigation in Abbre6iations: LPL, lipoprotein lipase; TG, triacylglycerol; ICDH, isocitrate dehydrogenase. * Corresponding author. Tel.: + 33 473 624253; fax: + 33 473 624639; e-mail: [email protected]
rodents and humans (for review, see [29]) but also in ruminants (for review, see [9]). The study of LPL is indeed of particular interest in tissues of meat-producing ruminants, since LPL controls TG partitioning between adipose tissues and muscles, thereby increasing fattening or providing energy in the form of fatty acids for muscle growth. Some quantitative studies at the whole animal level have suggested an important function of LPL activity in muscle tissues with regard to the total TG removal capacity of the body in both single-stomached (for review, see [7]) and ruminant species (for review, see [33]). Thus, in the adult sheep at maintenance, 55–60% of the total amount of free fatty acids originate from hydrolysis of circulating TG by LPL, and the skeletal muscle mass and the heart together could utilize approximately 40% of the non-esterified fatty acid entry rate (for review, see [33]). For this reason, it seems to us very important to study LPL activity, not only in adipose tissues, but also in muscles and heart, to get a better knowledge of the control of TG partitioning between these tissues in meat-producing cattle.
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To our knowledge, LPL has been little studied in muscles of ruminant species, despite numerous studies in adipose tissues ([6]; for review, see [9]) or milk (for review, see [31]). However, the presence of LPL activity 8 – 17-fold lower in beef heart compared to that in rat heart has been previously reported [12,24]. As in the rat, bovine LPL activity is expected to be lower in skeletal muscles than in heart [23]. Therefore, the first objective of our work was to develop a more sensitive assay of LPL activity in bovine skeletal muscles using an improved protocol [4]. In addition, the expression of the LPL gene in bovines has been only studied in mammary gland [22,37] and adipose tissue [6] and, LPL transcripts were barely detectable in beef heart [24]. Therefore, the second objective of this work was to characterize and quantify the mRNAs coding for LPL in various bovine tissues, especially in muscles.
2. Material and methods
seter (MA), diaphragm (D), rectus abdominis (RA), longissimus thoracis (LT) at the level of the 6th rib, semitendinosus (ST) from the medial portion, tensor fasciae latae (TFL), cutaneus trunci (CT) from the thick part, perirenal adipose tissue (PAT), omental adipose tissue (OAT) and subcutaneous adipose tissue (SCAT). Tissue samples were frozen in liquid nitrogen within 10 min of exsanguination and stored at − 80°C for subsequent analysis. Frozen samples were pulverized in liquid nitrogen. Others tissues (heart, lungs, kidney, spleen, small intestine and liver) as well as PAT were taken from five-week-old preruminant crossbred Friesian-Holstein male calves. Brain, adrenal glands, ovaries and mammary gland were taken from non-lactating pregnant cows. Samples of PAT and masseter muscle were taken from 230–260-day-old foetus. Muscle tissues (heart, soleus, extensor digitorum longus, epithroclearis) were taken after an overnight fast from 2–4-month-old male Sprague-Dawley rats (Iffa-Credo, L’Arbresle, France) and were treated as bovine tissues.
2.1. Reagents 2.3. Analytical techniques [3H]triolein (1.85– 2.96 GBq mmol − 1) and Hyperfilms MP were supplied by Amersham (Bucks, UK). [a-32P]dCTP ( \ 111 TBq mmol − 1) and [g-32P]dATP ( \ 111 TBq mmol − 1) were purchased from ICN Biochemicals (Irvine, CA). Guanidium thiocyanate was obtained from Fluka (Ronkonkoma, NY). Genescreen membranes were from Du Pont-NEN (Boston, MA). T4 polynucleotide kinase was supplied by New England Biolabs (Beverly, MA). Saturated phenol, chloroform/ isoamyl alcohol (49:1 v/v), agarose and nonaprimer labeling kit were purchased from Applige`ne (Illkirch, France). RNA molecular weight markers were from BRL (Bethesda, MD). Other reagents were from Sigma (St. Louis, MO). Antibodies to LPL purified from bovine milk were raised in rabbits [8].
2.2. Animals and experimental design Seven Montbe´liard male calves, tied and housed in individual stalls were used. Animal characteristics were described previously [19]. Briefly, from weaning (107– 128 days of age) to slaughter (170 days of age), calves were fed individually concentrate (80%) and hay (20%) according to a feeding pattern designed to allow an average daily gain of 1300 g. Composition of the concentrate was dehydrated alfalfa (30%), sugar beet pulp (40%), barley (14%) soybean meal (11%) and mineral compound (5%). The concentrate contained 16.2% of protein and 1.8% of fat. Slaughtering was done at 09:00 h after an overnight fast by stunning (captive-bolt pistol) and exsanguination. Tissue samples (50 – 100 g) from the following tissues were taken as described [18,19]: heart (H), mas-
Tissue protein or DNA contents and isocitrate dehydrogenase (ICDH) activity were measured as previously described [18,19].
2.4. Assay of LPL acti6ity LPL activity was assayed after detergent extraction as previously described [4]. Fresh tissue (six to seven tissue samples corresponding to approximately 4 g) or frozen tissue powder were homogenised at 4°C in 9 ml per g of the following buffer: ammonia–HCl (25 mM), pH 8.2 containing EDTA (5 mM), Triton X-100 (0.8% w/v), SDS (0.01% w/v), heparin (5000 IU l − 1) and peptidase inhibitors [pepstatin A (1 mg ml − 1), leupeptin (10 mg ml − 1), and aprotinin (0.017 TIU ml − 1)]. The inclusion of protease inhibitors and/or elimination of Ca2 + inhibit loss of LPL activity induced by frozen storage [36]. Insoluble material was removed by centrifugation (20000× g for 20 min at 4°C). LPL assay was performed as below. The substrate emulsion was Intralipid 10% (120 mM of TG) into which a trace amount of [3H]triolein (approximately 14 MBq) had been incorporated by sonication (75 W, 10 min). The emulsion was stored at 4°C and used for approximately two weeks since blank values increase beyond 10–15 days. Incubation medium was prepared from 10 ml emulsion, 10 ml heat-inactivated serum as a source of apolipoprotein CII, 60 ml of deionized water and 100 ml of the incubation buffer which contained 12% fatty acid free bovine serum albumin, 0.02% standard heparin, 0.2 M NaCl and 0.3 M Tris–HCl, pH 8.5. The total volume was adjusted to 200 ml with the
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sample volume diluted 5-fold in water (20 ml, i.e. 4 ml of tissue homogenate). The final substrate concentration was 6 mM. Incubation was carried out for 60 min at 25°C in a shaking water bath. The reaction was stopped by addition of distilled water (0.5 ml) and 2 ml of isopropanol/heptane/H2SO4 (48:48.3:1 v/v/v). Total lipids were extracted and fatty acids separated from TG as follows: after centrifugation (2000× g for 3 min, 4°C), a sample of the upper phase (800 ml) containing total lipids was transferred to new tubes into which 1 ml alkaline ethanol (ethanol 95%/water/2 M NaOH, 500:475:25 v/v/v) and 3 ml heptane were added. After a second centrifugation, the upper heptane phase containing unhydrolysed TG was discarded. A new extraction was performed with 3 ml of heptane. Finally, an aliquot (800 ml) of the remaining alkaline ethanol phase containing fatty acids was counted. Extraction efficiency of fatty acids was determined to be 41.89 0.53% (mean 9S.E., n = 20) and was similar to that observed for rat tissues [4]. All incubations were performed in triplicate. LPL activity in mU per g tissue wet weight (1 mU =1 nmol of fatty acids released per min) was calculated taking into account dilution of sample, extraction efficiency of fatty acids, incubation time, specific activity of the substrate and tissue/volume ratios in homogenates.
2.5. Quantification of mRNA coding for LPL Total RNA was isolated and analysed as previously described [18,19]. For Northern analysis, RNA aliquots (40 mg) were denatured in a solution containing 2.2 M formaldehyde and 50% formamide (v/v) by heating at 65°C for 10 min, size-fractionated by 1.5% agarose gel electrophoresis, and electrophoretically transferred to Genescreen membranes. The bovine LPL probe was labeled by random priming with 32P using the nonaprimer labeling kit. Prehybridization and hybridization to the LPL probe were performed at 42°C for 2–4 h and 16–20 h, respectively, in solutions containing 45% deionized formamide [19]. Then, the membranes were washed twice for 5 min in 2X SSPE at room temperature (1X SSPE=0.15 M NaCl, 10 mM NaH2PO4, 1 mM EDTA, pH 7.4), once for 30 min in 2X SSPE/2% SDS at 55°C, once for 15 min in 1X SSPE/2% SDS at 55°C and once for 15 min in 0.1X SSPE/2% SDS at 55°C. Hyperfilm MP were exposed to the membranes for 2 – 15 days at − 80°C with two intensifying screens. Quantification was performed with scanning densitometry (Hoeffer, San Francisco, CA). Preliminary Northern blot experiments were performed with various levels of RNA loaded on the gel from 10 to 50 mg. Membranes were hybridized with the LPL probe and also with a probe for the rat 18S ribosomal RNA [19]. Levels of LPL mRNA were positively correlated with levels of 18S rRNA (r = 0.895,
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n = 7, PB 0.01, data not shown). Therefore, under standard conditions (40 mg of total RNA), quantification of LPL mRNA was corrected for variations of the amount of RNA loaded on each lane by using values of hybridization to the 18S rRNA probe [19]. Results were expressed in arbitrary densitometric units per mg of total RNA loaded on the gel.
2.6. Statistical analysis Results were expressed as means 9 S.E. of four to seven animals. Significant differences between enzyme activities or mRNA levels in two different tissues from the same animals were evaluated by paired Student’s t-test. 3. Results
3.1. De6elopment of the LPL assay procedure LPL activity in bovine heart was at maximum at pH 8.5 as in rat samples [4]. A minimum of 10% of albumin and of 0.02% of heparin in the incubation buffer was necessary to observe maximum LPL activity. The amount of substrate (6 mM in the incubation medium) was saturating since LPL activity was at a maximum between 6 and 9.3 mM for both heart and adipose tissue. LPL activity was linear with time and similar between assays at 25°C but not at 37°C (Fig. 1A) as noted previously [4]. Similar results were obtained with adipose tissues (results not shown). The assay was linear between 0.2 and 4 ml of tissue homogenate, equivalent to 20–400 mg of fresh tissue (Fig. 1B) independent of tissue (heart, muscles or adipose tissue). Beyond this point, a partial inhibition of LPL activity occurred. Therefore, LPL activity was routinely assayed at 25°C for 60 min with 4 ml of homogenate in order to measure the highest level of activity in the linear range [27]. Under these standard conditions, hydrolysis did not exceed 10% of total amount of substrate for any sample. Finally, the intra-assay variation was low (9 5–10%), whereas inter-assay variations were higher (13–30% for the samples with the lowest activity and less than 10% for tissues which exhibit the highest activity).
3.2. Specificity of the measured LPL acti6ity As in human adipose tissue [20], LPL activity was completely inhibited in presence of a high concentration (100 mM) of sodium deoxycholate because this detergent desorbs the enzyme from lipid droplets [28]. In contrast, only a partial inhibition of LPL was observed in the presence of 0.5–1 M NaCl: − 18 to − 21% for adipose tissue and − 31 to −39% for heart and skeletal muscles.
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Fig. 1. Biochemical characteristics of the LPL assay in bovine heart. (A) Effect of incubation time on the velocity of the enzyme reaction. Results are expressed in mmol of fatty acids released for the indicated time. Four assays were done at two different temperatures demonstrating the absence of replication at 37°C but showing the linearity of the assay at 25°C with respect to time. (B) Effect of the amount of tissue homogenate on the measurement of LPL activity. Results are expressed in nmol of fatty acids released per min for the indicated amounts of homogenate. In all experiments, 1 g of fresh tissue was homogenized in 9 ml of buffer. The incubation medium (200 ml) was prepared from 10 ml emulsion, 10 ml serum, 60 ml deionized water, 100 ml of the incubation buffer and 20 ml of diluted sample.
Since LPL binds to heparin, tissue homogenates of bovine muscles were passed through a column of heparin-sepharose. After elution with a NaCl gradient, only one peak of LPL activity was detected at the expected concentration of NaCl suggesting the existence of only one lipase assayed by our protocol. Finally, lipase activity was completely abolished in presence of polyclonal antibodies against bovine LPL, which may be considered as the best test of specificity [8].
than with bovine serum (Fig. 2). In contrast, LPL activity in adipose tissues was similar with the two sources of sera (ratio of 1.09 0.02; mean9 S.E. of 16 samples; Fig. 2). Nevertheless, levels of LPL activity assayed with rat serum were positively correlated with levels of LPL activity measured with bovine serum for both adipose tissues (r= 0.99, n = 24, PB 0.001) and muscles (r= 0.76, n= 28, P B 0.01) as shown in Fig. 2. From all these experiments, we decided to use the best batch of rat serum for LPL assay in standard conditions.
3.3. Effect of serum acti6ator on LPL acti6ity 3.4. Effect of tissue processing on LPL acti6ity For LPL assay, one must also add serum to the incubation medium to provide an activator (apo CII) to obtain maximum activity. Basal LPL activity (i.e. without serum) was between 19 and 52% of maximum activity for heart, skeletal muscles or adipose tissues. This relatively high and variable basal LPL activity may be explained by the presence of small amounts of blood serum in the tissue which might differ among samples. To activate LPL, the optimum amount of serum was found to be 5% by volume independent of the source of serum (rats or bovines). But great variations in LPL activity were observed depending on the source of serum in interaction with the source of tissue extract (Table 1). Therefore, we pooled the best rat sera and the best bovine sera to obtain two homogeneous sources of activators [4,27]. When these two batches of sera were tested in the same assay, rat serum was more effective than bovine serum to activate LPL in heart and skeletal muscles. In one representative experiment, LPL activity was 5.59 0.27-fold higher (mean9S.E. of 28 samples) with rat
At the time of slaughter of the animals, it is often impossible to perform LPL assays immediately. Therefore, we tested different conditions of processing and storage of samples. In a first experiment, two procedures were studied: the first batch of tissues was frozen in liquid nitrogen, stored at − 80°C, pulverized in liquid nitrogen and homogenized in the appropriate buffer on the day of assay; the second batch of tissues was frozen into flasks with 9 ml of homogenisation buffer for LPL assay (containing detergents and heparin to protect LPL against any degradation) per g tissue wet weight and stored at − 80°C until subsequent homogenisation after complete thawing on the day of assay. LPL activity in adipose tissues was 1.49 0.13-fold higher for samples frozen within homogenisation buffer than for tissue powder (mean9 S.E., n= 6, PB 0.07). In contrast, LPL activity in heart and masseter muscle were 1.49 0.08- and 2.7 90.38-fold higher, respectively, for tissue powder (mean9S.E., n=8, PB 0.05).
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Table 1 LPL activity in the presence of bovine or rat sera from different donors as source of activator Source of rat sera
Donor Donor Donor Donor Donor
1 2 3 4 5
LPL activity in assay 1 (%) Heart
Omental adipose tissue
100 68.4 84.8 77.2 60.8
100 90.4 110.0 90.4 81.9
Source of bovine sera
Donor Donor Donor Donor
A B C D
LPL activity in assay 2 (%) Heart
Omental adipose tissue
100 54.8 51.6 80.0
100 54.6 62.2 63.2
LPL activity in two samples was measured in triplicate with rat sera from five different donors (assay 1) or with bovine sera from four different donors (assay 2) to compare the effects of the source of serum. Results of LPL activity were first calculated in mU per g tissue wet weight as usual and then expressed within the same column relative to the values obtained for donors 1 and A for assays 1 and 2, respectively.
In a second experiment, we showed that, in heart, one or two of freeze/thaw cycles induced a 10–20% decrease in LPL activity. After six cycles, a 30% decrease was observed. In adipose tissues, a 20 –40% decrease in LPL activity was detected after only one cycle and a 40–60% decrease was observed after two to six cycles of freezing/thawing suggesting that LPL is less stable in adipose tissue than in heart. Thus, we recommend that LPL should be assayed on fresh samples, and, if this is not possible, that samples or tissue homogenates should be frozen only once.
3.5. LPL acti6ity and gene expression in 6arious bo6ine tissues LPL activity was barely detectable in small intestine, kidney, spleen, adrenal glands and lung from young calves. The LPL activity in liver was very low and even undetectable in some assays (Table 2). For these tissues, blank values might represent more than 50% of total
values. Thus, absolute values for LPL activity were considered unreliable. In contrast, LPL activity was higher in heart and adipose tissues (Table 2). Northern blot analysis revealed the presence of three different-sized species of LPL mRNAs in bovine tissues (Fig. 3). The sizes of the two major mRNAs were approximately 3.5–3.8 and 3.2–3.4 kb. The 3.5–3.8 kb species seemed to be the most abundant transcript in many, but not all, experiments. The minor mRNA species (1.7 kb), was detected at a low level in tissues which expressed the highest level of LPL transcripts. Only the two major mRNA species were quantified by scanning densitometry since the third mRNA species was minor and not always detected. In agreement with the results for activity, levels of LPL mRNAs were high in heart and adipose tissues from calves even during foetal life (Fig. 3). In contrast, LPL mRNAs occurred in very low levels in spleen, adrenal glands and ovaries, barely detectable in lung, small intestine and kidney and undetectable in liver and brain.
3.6. LPL acti6ity and gene expression in eight bo6ine muscles
Fig. 2. Effect of serum activator on LPL activity in bovine tissues. Tissue LPL activity was measured with either bovine serum or rat serum in the same assay in heart, masseter (MA), omental adipose tissue (OAT) and perirenal adipose tissue (PAT). LPL activity measured with rat serum was plotted against LPL activity measured with bovine serum.
Eight different muscles from an homogeneous group of calves were characterized using ICDH activity, characteristic of oxidative metabolism [18,19] (Fig. 4). LPL activity assayed with rat serum was higher in heart and masseter (the most oxidative muscles) than in other skeletal muscles (Fig. 4). Similar results were observed with bovine serum as activator although absolute values were lower than those with rat serum (449922, 2569 39 and 29–77 mU g − 1 tissue wet weight for heart, masseter and other skeletal muscles, respectively). Surprisingly, LPL activity in the diaphragm was as low as in glycolytic muscles although ICDH activity was higher than in other skeletal muscles of the carcass. LPL activity was also expressed in mU per mg of DNA and per mg of protein. Similar results and similar differences among muscle tissues were observed whatever the method of expression of the results and the source of serum activator used in the assay.
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Table 2 Range of LPL activity in various bovine tissues Liver
Small intestine
Kidney
Spleen
Adrenal glands
Lung
Heart
Perirenal adipose tissue
0–58
137–340
50–250
58–184
140–202
80–208
473–1465
580–3560
Hearts and adipose tissues were obtained from 12 1–6-month-old calves. No significant effect of age was found. Other tissues were taken from five 5-week-old preruminant calves. Adrenal glands were taken from cows. LPL activity was measured in triplicate with different sera (5% v/v) as source of activator in many different assays for heart and adipose tissue and in two different assays for other tissues. The highest and the lowest values in mU per g tissue wet weight obtained in the different assays for the different animals are indicated.
LPL activity levels in rat heart and soleus (2150– 2950 and 1160–1350 mU g − 1 wet tissue, respectively) were higher (PB0.05) than in bovine heart and masseter (662–832 and 412 – 972 mU g − 1 wet tissue, respectively). LPL activity was also higher in rat extensor digitorum longus and epithroclearis, the most glycolytic muscles in the rat (130 – 190 mU g − 1 wet tissue) than in RA, TFL, ST, LT and CT (33 – 154 mU g − 1 wet tissue), the most glycolytic muscles in the calf [18,19]. LPL mRNA levels expressed per mg of total RNA were highest in oxidative muscles (Fig. 5). However, the level of LPL mRNA in masseter was approximately 2-fold lower than that in heart, despite similar LPL activity levels (Figs. 4 and 5). When results of LPL mRNA levels were expressed per g wet tissue weight, differences between muscles were amplified (60.69 23.4, 21.794.1, 0.5– 8.4 arbitrary units for heart, masseter and other skeletal muscles, respectively) since yields of extracted RNA were higher in oxidative muscles, especially in heart as in rats [23] and in bovines [18,19].
3.7. LPL acti6ity and gene expression in three bo6ine adipose tissues LPL activity was higher in internal adipose tissues (OAT and PAT) than in SCAT (Fig. 4). Similar results were observed with bovine serum as activator (971 9 218, 2589 121 and 12469180 mU g − 1 tissue wet weight for OAT, SCAT and PAT, respectively). However, since protein and DNA contents were approximately 2- and 1.5-fold higher in SCAT, respectively, than in internal adipose tissues, differences between SCAT and internal adipose tissues were amplified when results of LPL activity were expressed per mg of protein or DNA [27]. LPL mRNA levels were higher in PAT and OAT than in SCAT (Fig. 6). LPL mRNAs in SCAT were barely detectable in some experiments (Fig. 6A), but quantifiable in most. However, the level of LPL mRNA in PAT was approximately 1.5-fold higher (P B 0.05) than in OAT, despite similar LPL activity (Figs. 4 and 6). Similar results were observed when data of LPL mRNA levels were expressed per g wet tissue weight, since yields of extracted RNA were similar in the three adipose tissues.
4. Discussion
4.1. Extraction and assay of LPL in bo6ine tissues It might seem a trivial task to develop assays of LPL in bovine tissues since many methods have been reported in the literature for LPL from milk or adipose tissues of ruminants [15,36,40]. However, by studying LPL in bovine muscles, we demonstrated that conditions for assay of LPL have to be set up and critically evaluated for each new application. Depending on the methods used to extract and to assay the enzyme, the activity recorded for muscles could vary by an order of magnitude. Factors needing special attention for the bovine muscles were sample storage, extraction procedures and the source of serum to provide activator. Tissue extracts of acetone-ether powders were widely used in the earlier literature [27], but have been found to give low recovery. Most investigators now favour homogenates in buffers with heparin and detergents which help to stabilize the enzyme, to extract it from intracellular vesicles and to dissociate it from heparin sulfate proteoglycans [4]. The earlier recommendation was to store tissue pieces frozen in this buffer until they were homogenized for assay [4]. In bovine, this worked well for adipose tissue, but not for muscles since the muscle pieces need to be pulverized in liquid nitrogen. As substrate, we used radioactive triolein that was incorporated into a commercial phospholipid-stabilized TG emulsion. This provides high sensitivity, but substantial between-assay variation, especially when different emulsion preparations are used [27]. However, the intra- and inter-assay variations of our assay were in agreement with other data in rodents with the same assay procedure [4] or in cow adipose tissues with another procedure [15]. Tissue samples may contain other lipase activities. In the present system, the substrate with long-chain TG in a lipoprotein-like emulsion virtually eliminates the contribution of non-specific esterases with water-soluble esters as substrate, and the high pH minimizes the contribution from lysosomal enzymes [27]. The specificity of our assay was demonstrated by the fact that, (i) heparin-agarose columns bound the lipase activity which then eluted as a single peak in the position
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Fig. 3. Northern blot analysis of the tissue distribution of LPL mRNAs in bovine tissues. Total RNA was extracted from perirenal adipose tissue (PAT), foetal perirenal adipose tissue (fPAT), liver (Li), small intestine (SI), kidney (Ki), spleen (Sp), adrenal glands (AG), brain (Br), lung (Lu), ovaries (Ov), heart, (H), foetal heart (fH) and masseter muscle (MA). RNAs (40 mg) was then analyzed by Northern blot using the bovine LPL cDNA. Films were exposed one (for adipose tissues) or seven days (for the other tissues) to the membranes.
characteristic for LPL [17], and (ii) polyclonal antibodies against bovine milk LPL suppressed the lipase activity almost completely. The commonly used criterion to differentiate LPL from other lipases, i.e. its inhibition by NaCl, is not a very reliable tool since it is very dependent on the composition of buffers (for review, see [11]) or on the conditions of incubation [3]. At 37°C, the inactivation by NaCl is rapid and almost complete. At 4 or 10°C, and with albumin, the enzyme works well at 1.5 M NaCl (for review, see [28]). In our assay at 25°C, 1 M NaCl caused only partial loss of activity and this differed with the source of tissue extract. The pH-dependency of the reaction depends on an interplay of factors [4] such as pH effects on the binding of LPL to lipid droplets and on the active site reaction (for review, see [28]). However, LPL extracted from human muscle showed about the same activity from pH 7.9–8.7 [26], whereas the highest level of activity for LPL from bovine muscles was obtained between pH 9 and 9.5 [24]. For the present assay system, small variations in pH of the incubation medium might affect the activity and hence be a source of interassay variability. This is the reason why in our laboratory, a large batch of incubation medium is prepared, and used in all assays to minimize inter-assay variability.
4.2. Differential effect of bo6ine or rat sera on LPL acti6ation Apo CII provided by the serum binds to the lipid– water interface and to the enzyme. With soluble model substrates, apo CII has little or no effect (for review, see [17]). With emulsified substrates and in monolayer systems, the effect of apo CII varies widely, and this appears to relate to the ‘quality of the interface’ [3]. There are examples of systems where LPL exerts full activity without apo CII and others where the enzyme has little or no activity without apo CII. Hence, it appears that the role of apo CII is to restore the activity in situations where other factors suppress it [3]. Particu-
larly important is the presence of other lipid-binding proteins, for instance, apolipoproteins such as apo CIII or fragments of these apolipoproteins, and also unrelated proteins [4]. A striking finding here was the 5-fold higher activation of the lipase activity with rat compared to bovine serum for heart and muscles, but not for adipose tissue. We tentatively ascribe this to the presence of much more protein in the muscle extracts, and note that haemoglobin strongly suppresses LPL activity [3]. Another possibility is that there are differences between the lipase in muscle and in adipose tissue as suggested by kinetic [26] or immunological studies [38]. There is, however, only one gene for LPL [21] but LPL undergoes a wide range of posttranscriptional [21] or posttranslational [13] modifications that may occur in a tissue related-fashion. It may seem paradoxical that rat serum was more efficient than bovine serum for LPL activation. Although species-specific immunological or enzymatic properties of the LPL molecule alone have been reported [8,40], there is no evidence for species specificity in the LPL–apo CII interaction. In model systems, a similar activation of bovine LPL was obtained using apo CII or analogues from as widely different animals as cows, humans, rats, guinea pigs, chicken and salmon. Conversely, human apo CII has been shown to activate LPL from many different animals including cows and rats [2,25]. In agreement with this, the region in apo CII that interacts with the enzyme is quite strictly maintained across bovine and other species in the C-terminal third of the molecule required for LPL activation [5]. Hence it appears unlikely that our observations relate to a particularly good fit between the bovine enzyme and rat apo CII. These differential effects of bovine and rat sera on LPL activation may be linked to the absolute and relative concentrations of apo CII and other apolipoproteins (for instance, apo CIII) which might differ between rats and cows. In addition, the shift of apolipoproteins from plasma lipoproteins of the added activator to the lipid–water interface depends on sev-
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Fig. 4. Metabolic activity of various muscles and adipose tissues from seven 170-day-old ruminant calves. Values are means 9 S.E. of four to seven observations. Isocitrate dehydrogenase (ICDH) activity was measured spectrophotometrically in muscle tissue homogenates and expressed in U g − 1 tissue wet tissue (1 U =1 mmol of substrate used per min). LPL activity was measured in tissue homogenates with rat serum as source of activator and expressed in mU g − 1 tissue wet weight (1 U = 1 mmol of fatty acid released per min). A value followed by a superscript differs significantly (PB 0.05) from all other values not followed by the same superscript. H, heart; MA, masseter; D, diaphragm; RA, rectus abdominis; TFL, tensor fasciae latae; LT, longissimus thoracis; ST, semitendinosus; CT, cutaneus trunci; OAT, omental adipose tissue; SCAT, subcutaneous adipose tissue; PAT, perirenal adipose tissue.
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Fig. 5. Northern blot analysis of the LPL mRNAs distribution in muscles from seven 170-day-old ruminant calves. Total RNA was extracted from bovine muscles and then analyzed by Northern blot using the bovine LPL cDNA (A) and the 18S rRNA probe (B) to verify that equivalent amounts of total RNA were loaded on each lane. Quantification of LPL mRNA and of 18S rRNA levels was performed by scanning densitometry. Different films exposed 1–8 days to the same membrane were used for quantification to be sure that the signals were within the linear range of the films depending on the samples and the time of exposure. Results of LPL mRNA levels were corrected for variations in 18S RNA levels and were then expressed in densitometric arbitrary units per mg of total RNA loaded on the gel relative to the level in longissimus thoracis (C). Values are means 9S.E. of five to seven observations (C). A value followed by a superscript differs significantly (P B0.05) from all other values not followed by the same superscript. H, heart; MA, masseter; D, diaphragma; RA, rectus abdominis; TFL, tensor fasciae latae; LT, longissimus thoracis; ST, semitendinosus; CT, cutaneus trunci.
eral factors including the relative concentrations of HDL and VLDL and the level of TG in the serum activator [39]. Therefore, the fact that the proportions of circulating VLDL and HDL differ markedly between rats and ruminants [41] might also induce differences in activation of LPL by the two types of sera. There are also differences in activation of LPL by apo CII from different species that relate to the interaction with the lipid interface rather than with the enzyme [2]. In support of this, there are rather large differences among apo CII from different species in the N-terminal twothirds of the molecule which is responsible for interaction with the lipid– water interface [5]. It may be this type of difference that made rat serum a much better source of activator for the LPL in bovine muscle samples. Whatever the reasons for the observed differences, we used rat serum in our assay to obtain high values for LPL activity in samples from muscle, and to reduce interassay variability. To enhance reproducibility, it is advisable to prepare a large batch and freeze this in aliquots.
4.3. Comparison of LPL acti6ity in similar tissues by different procedures When we applied the present method to rat heart, we obtained values (2150 – 2950 mU g − 1) similar to those reported using a similar procedure for LPL assay [8]. These values are higher than those reported by most investigators (150 [16], 500 [23], 950 – 1250 [12] mU g − 1, but similar to those reported by others (1300 – 6500 mU
g − 1 [34]). The values we obtained for LPL activity in bovine heart with rat (749 mU g − 1) or bovine (449 mU g − 1) serum as activator were much higher than those previously reported (90 mU g − 1 [24]). Our values for LPL in calf adipose tissues (372–923 mU g − 1) are similar to those in adipose tissues of steers (500–833 mU g − 1 [35]), but higher than those of cows (133 mU g − 1 [15]) or sheep (85–109 mU g − 1 [40]). The values for LPL activity in skeletal muscles from our calves (33–154 mU g − 1) are higher than those for sheep (6–14 mU g − 1 [40]; 15–55 mU g − 1 [36]) or human muscles (5–32 mU g − 1 [26]). Thus, LPL activity levels registered in this study are in the upper range of values reported in the literature for LPL activity in rat or ruminant tissues. Some of these differences may be explained by physiological parameters of the animals such as age or feeding [8,34,36], but the sensitivity and the quality of the assay procedure is of prime importance especially with tissues which exhibit a low activity such as bovine muscles. The LPL activity in bovine heart was 3.4–4.8-fold lower than that in rat heart depending on the serum used. This difference is not unexpected due to differences in sizes and, hence, in metabolic rate between the two studied species. However, a much higher difference (8–17-fold) was previously reported [12,24].
4.4. Detection of LPL transcripts in bo6ine tissues The detection of three different-sized species of LPL mRNAs in bovine tissues (the two larger species being present in significant amounts for quantification) was in
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Fig. 6. Northern blot analysis of the LPL mRNAs distribution in adipose tissues from seven 170-day-old ruminant calves. Total RNA was extracted from bovine adipose tissues and then analyzed by Northern blot using the bovine LPL cDNA (A) and the 18S rRNA probe (B) as described in Fig. 5. Results of LPL mRNA levels were corrected for variations in 18S RNA levels and were then expressed in densitometric arbitrary units per mg of total RNA loaded on the gel relative to the level in omental adipose tissue (C). Values are means9S.E. of four to seven observations (C). A value followed by a superscript differs significantly (P B0.05) from all other values not followed by the same superscript. OAT, omental adipose tissue; SCAT, subcutaneous adipose tissue; PAT, perirenal adipose tissue.
agreement with the previous observations in bovine mammary gland [22,37] and adipose tissue [6]. In contrast, it was reported that the primary LPL mRNA detected in beef heart was in amounts too low for accurate quantification [24]. This may be explained by methodological differences between the two studies: (i) we used an homologous probe instead of a mouse LPL cDNA, (ii) labelling was performed by random priming rather than by the nick translation method, (iii) total RNA were denaturated with 6.6% formaldehyde instead of 3.2%. Finally, the small differences reported for the size of the LPL transcripts in bovine tissues among laboratories [6,22,24,37]; and this study) could be explained by the different standards used for calculation (DNA or RNA kb ladders, ribosomal RNAs). However, polymorphisms affecting mRNA sizes among tissues and/or species or strains [21] cannot be excluded. Except in liver and brain, LPL mRNA transcripts were observed in all tissues examined albeit at very different levels. In addition, LPL mRNA levels were higher in growing calves than in foetuses as described in the rat [21]. As in mouse, human [22] and rat tissues [21,23], levels of LPL transcripts were higher in bovine heart and oxidative skeletal muscles which use fatty acids as energy source than in muscles composed of fast-twitch white fibers which use glucose as fuel [23]. Great differences in metabolism among adipose sites have been previously reported in pigs [1] and calves [18]. In the cow, LPL activity may also differ among adipose tissues from various anatomical sites [10]. These differences are mainly related to differences in LPL mRNA levels among adipose sites (Fig. 6). Our results also suggest that the higher the LPL
activity, the higher the adipocyte size as previously described in rats and humans [14]. The LPL system is subject to regulation at many levels involving gene expression, synthesis/degradation, intra- and intercellular translocation, activation/inactivation by serum factors. However, LPL activity depends primarily on a balance between rates of synthesis and degradation (for review, see [11]). Our results support this last idea since differences in LPL activity among a variety of bovine tissues are mainly reflected by differences in LPL mRNA levels, as in the rat [23]. However, other mechanisms of regulation may also be involved in the fasted rat [23] and in the bovine since LPL activity did not always parallel mRNA levels among tissues in rats [23] and in cattle (this study). In conclusion, although LPL specific activity and gene expression in bovine muscles are significantly lower than that found in rat muscles, we have developed sensitive procedures to assay LPL and to quantify LPL mRNA levels in bovine tissues. Acknowledgements We thank Dr D. Bauchart (INRA, Laboratoire Croissance et Me´tabolismes des Herbivores, France) for helpful discussions and Dr Y. Furuichi (Nippon Research Center, Kamakura, Japan) for kindly providing us the LPL bovine cDNA. We also thank Nicole Guivier and Samia Brazi for their expertise and contributions, Robert Jailler and his group for the management of the animals and Gilbert Cuylle and his group for the management of the slaughterhouse. This study was supported in part by grant 13x-727 from the Swedish Medical Research Council.
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References [1] Anderson DB, Kauffman RG, Kastenschmidt LL. Lipogenic enzyme activities and cellularity of porcine adipose tissue from various anatomical locations. J Lipid Res 1972;13:593–9. [2] Andersson Y, Lookene A, Shen Y, Nilsson S, Thelander L, Olivecrona G. Guinea pig apolipoprotein CII: expression in E. coli, functional studies of recombinant wild-type and mutated variants and distribution on plasma lipoproteins. J Lipid Res 1997;38:2111 – 24. [3] Bengtsson G, Olivecrona T. Lipoprotein lipase: some effects of activator proteins. Eur J Biochem 1980;106:549–55. [4] Bengtsson-Olivecrona G, Olivecrona T. Assay of lipoprotein lipase and hepatic lipase. In: Converse C, Skinner ER, editors. Lipoprotein Analysis. Oxford: Oxford University Press, 1991:169 – 185. [5] Bengtsson-Olivecrona G, Sletten K. Primary structure of the bovine analogues to human apolipoproteins CII and CIII. Studies on isoforms and evidence for proteolytic processing. Eur J Biochem 1990;192:515–21. [6] Bonnet M, Faulconnier Y, Fle´chet J, Hocquette JF, 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. [7] Borensztajn J. Heart and skeletal muscle lipoprotein lipase. In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:133 – 148. [8] Chajek-Shaul T, Bengtsson-Olivecrona G, Peterson J, Olivecrona T. Metabolic fate of rat heart endothelial lipoprotein lipase. Am J Physiol 1988;255:E247–54. [9] Chilliard Y. Dietary fat and adipose tissue metabolism in ruminants, pigs, and rodents: a review. J Dairy Sci 1993;76:3897 – 931. [10] Chilliard Y, Robelin J. Activite´ lipoprote´ine-lipasique de diffe´rents de´poˆts adipeux et ses relations avec la taille des adipocytes chez le vache tarie en cours d’engraissement ou en de´but de lactation. Reprod Nutr Dev 1985;25:287–93. [11] Cryer A. Comparative biochemistry and physiology of lipoprotein lipase In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:277–327. [12] Cryer A, Jones HM. The distribution of lipoprotein lipase (clearing factor lipase) activity in the adipose, muscular and lung tissues of ten animal species. Comp Biochem Physiol 1979;63B:501 – 5. [13] Doolittle MH, Ben-Zeev O, Elovson J, Martin D, Kirchgessner TG. The response of lipoprotein lipase to feeding and fasting. J Biol Chem 1990;265:4570–7. [14] Eckel RH. Adipose tissue lipoprotein lipase. In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:79–132. [15] Faulconnier Y, The´venet M, Fle´chet J, Chilliard Y. Lipoprotein lipase and metabolic activities in incubated bovine adipose tissue explants: effects of insulin, dexamethasone, and foetal bovine serum. J Anim Sci 1994;72:184–91. [16] Galan X, Llobera M, Ramirez I. Lipoprotein lipase and hepatic lipase in Wistar and Sprague-Dawley rat tissues. Difference in the effects of gender and fasting. Lipids 1994;29:333 – 6. [17] Garfinkel AS, Schotz MC. Lipoprotein lipase. In: Gotto AM Jr, editor. Plasma Lipoproteins. Amsterdam: Elsevier, 1987:335 – 357. [18] Hocquette JF, Castiglia-Delavaud C, Graulet B, Ferre´ P, Picard B, Vermorel M. Weaning marginally affects glucose transporter (GLUT4) expression in calf muscles and adipose tissues. Br J Nutr 1997;78:251–71. [19] Hocquette JF, Graulet B, Castiglia-Delavaud C, Bornes F, Lepetit N, Ferre´ P. Insulin-sensitive glucose transporter tran-
[20]
[21]
[22]
[23]
[24]
[25]
[26]
[27]
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35] [36]
[37]
211
script levels in calf muscles assessed with a bovine GLUT4 cDNA fragment. Int J Biochem Cell Biol 1996;28:795 –806. Iverius P-H, Brunzell JD. Human adipose tissue and lipoprotein lipase: changes with feeding and relation to postheparin plasma enzyme. Am J Physiol 1985;249:E107– 14. Kirchgessner TG, LeBoeuf RC, Langner CA, Zollman S, Chang CH, Taylor BA, Schotz MC, Gordon JI, Lusis AJ. Genetic and developmental regulation of the lipoprotein lipase gene: loci both distal and proximal to the lipoprotein lipase structural gene control enzyme expression. J Biol Chem 1989;264:1473– 82. Kirchgessner TG, Svenson KL, Lusis AD, Schotz MC. The sequence of cDNA encoding lipoprotein lipase. J Biol Chem 1987;262:8463– 6. LaDu MJ, Kapsas H, Palmer WK. Regulation of lipoprotein lipase in adipose and muscle tissues during fasting. Am J Physiol 1991;260:R953– 9. LaDu MJ, Schultz CJ, Essig DA, Palmer WK. Characterization of serum-stimulated lipoprotein lipase from bovine heart. Int J Biochem 1991;23:405 – 11. Lindberg A, Olivecrona G. Lipase evolution: trout, Xenopus and chicken have lipoprotein lipase and apolipoprotein C-IIlike activity but lack hepatic lipase-like activity. Biochim Biophys Acta 1995;1255:205 – 11. Lithell H, Boberg J. Determination of lipoprotein-lipase activity in human skeletal muscle tissue. Biochim Biophys Acta 1978;528:58 – 68. Nilsson-Ehle P. Measurements of lipoprotein lipase activity. In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:59 – 77. Olivecrona T, Bengtsson G. Lipases in milk. In: Borgstro¨m B, Brockman HL, editors. Lipases. Amsterdam: Elsevier, 1984:205 – 261. Olivecrona T, Bengtsson-Olivecrona G, Chajek-Shaul T, Carpentier Y, Deckelbaum R, Hultin M, Peterson J, Patsch J, Vilaro S. Lipoprotein lipase. Sites of synthesis and sites of action. Atheroscler Rev 1991;22:21 – 5. Olivecrona T, Bengtsson-Olivecrona G, Hultin M, Peterson J, Vilaro S, Deckelbaum RJ, Carpentier YA, Patsch J. What factors regulate the action of lipoprotein lipase?. In: Malmendier CL et al, editors. Hypercholesterolemia, Hypocholesterolemia, Hypertriglyceridemia. New York: Plenum Press, 1990:335 – 339. Olivecrona T, Vilaro S, Bengtsson-Olivecrona G. Lipases in milk. In: Fox PF, editor. Advanced Dairy Chemistry. Amsterdam: Elsevier, 1992:292 – 309. Peterson J, Bihain BE, Bengtsson-Olivecrona G, Deckelbaum RJ, Carpentier YA, Olivecrona T. Fatty acid control of lipoprotein lipase: a link between energy metabolism and lipid transport. Proc Natl Acad Sci USA 1990;87:909 – 13. Pethick DW, Dunshea FR. Fat metabolism and turnover. In: Forbes JM, France J, editors. Quantitative Aspects of Ruminant Digestion and Metabolism. Wallingford: CAB International, 1993:291 – 311. Planche E, Boulange A, DeGasquet P, Tonnu NT. Importance of muscle lipoprotein lipase in rats during suckling. Am J Physiol 1980;238:E511– 7. Rao DR, Hawkins GE. Activity of lipoprotein lipase in adipose tissues from steers. J Dairy Sci 1976;59:161 – 3. Rule DC, Andersen MK, Bailey JW, Swain L, Ficek SJ, Thomas DP. Frozen storage of ovine and rat tissues adversely affects lipoprotein lipase activity. J Nutr Biochem 1996;7:577– 81. Senda M, Oka K, Brown WV, Qasba PK. Molecular cloning and sequence of a cDNA coding for bovine lipoprotein lipase. Proc Natl Acad Sci USA 1987;84:4369 – 73.
212
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[38] Soteriou A, Cryer A. Distinct immunoreactivities suggest the existence of potential tissue variants in rat lipoprotein lipase. Biochem J 1994;299:417–23. [39] Tornoci L, Scheraldi CA, Li XZ, Ide H, Goldberg IJ, Le NA, Tornoci L. Abnormal activation of lipoprotein lipase by non-equilibrating apo C-II: further evidence for the presence of non-equilibrating pools of apolipoproteins C-II and C-III in
.
plasma lipoproteins. J Lipid Res 1993;34:1793 – 803. [40] Tume RK, Thornton RF, Johnson GW. Lipoprotein lipase of sheep and rat adipose tissues. Aust J Biol Sci 1983;36: 41 – 8. [41] Vitic J, Stevanovic J. Comparative studies of the serum lipoproteins and lipids in some domestic, laboratory and wild animals. Comp Biochem Physiol 1993;106B:223– 9.
Lipoprotein lipase activity and mRNA levels in bovine tissues Jean-Franc¸ois Hocquette a,*, Benoıˆt Graulet a, Thomas Olivecrona b a
INRA, Laboratoire Croissance et Me´tabolismes des Herbi6ores, Theix, 63122 Saint-Gene`s Champanelle, France b Department of Physiological Chemistry, Uni6ersity of Umea˚, S-901 87 Umea˚, Sweden Received 20 March 1998; received in revised form 3 July 1998; accepted 13 August 1998
Abstract Lipoprotein lipase (LPL) in cattle has been extensively studied in adipose tissue, milk and mammary gland, but only to a limited extent in muscles. Therefore, we have adapted our in vitro LPL assay method for the measurement of LPL activity and describe, for the first time, sensitive procedures to quantify LPL activity and mRNA levels in bovine muscles. In vitro activation of bovine LPL activity is approximately 5-fold greater with rat than with bovine sera for heart and muscles, but not for adipose tissues. Values of LPL activity are in the upper range of those previously reported for rat or bovine tissues. With rat serum as activator, LPL activity in the heart of seven calves (662–832 mU g − 1) is at least 3-fold lower than in the rat heart (2150 – 2950 mU g − 1, PB 0.05). LPL activity is higher in bovine heart and oxidative muscles (412 – 972 mU g − 1), except the diaphragm, than in mixed or glycolytic muscles (33–154 mU g − 1, P B0.05). The levels of LPL transcripts are positively related to LPL activity in bovine tissues, including muscles and adipose tissues. © 1998 Elsevier Science Inc. All rights reserved. Keywords: Lipoprotein lipase activity; Gene expression; LPL; Muscles; Adipose tissues; Bovine
1. Introduction Triacylglycerols (TG) are unloaded from chylomicrons and very low density lipoproteins through hydrolysis by lipoprotein lipase (LPL) on capillary endothelial surfaces, where circulating lipoproteins bind briefly. Fatty acids liberated by LPL are available for tissues as energy sources especially in muscles, or for storage in the form of TG in adipose tissues. It is generally assumed that the rate-limiting factor for energy delivery from lipoproteins is the amount of active LPL available at the endothelium which is correlated to LPL activity in tissue cells (for review, see [7,14,29]). This may be true in most physiological conditions [30] except in case of fat overload [32]. Therefore, tissue LPL and its regulation became the subject of active investigation in Abbre6iations: LPL, lipoprotein lipase; TG, triacylglycerol; ICDH, isocitrate dehydrogenase. * Corresponding author. Tel.: + 33 473 624253; fax: + 33 473 624639; e-mail: [email protected]
rodents and humans (for review, see [29]) but also in ruminants (for review, see [9]). The study of LPL is indeed of particular interest in tissues of meat-producing ruminants, since LPL controls TG partitioning between adipose tissues and muscles, thereby increasing fattening or providing energy in the form of fatty acids for muscle growth. Some quantitative studies at the whole animal level have suggested an important function of LPL activity in muscle tissues with regard to the total TG removal capacity of the body in both single-stomached (for review, see [7]) and ruminant species (for review, see [33]). Thus, in the adult sheep at maintenance, 55–60% of the total amount of free fatty acids originate from hydrolysis of circulating TG by LPL, and the skeletal muscle mass and the heart together could utilize approximately 40% of the non-esterified fatty acid entry rate (for review, see [33]). For this reason, it seems to us very important to study LPL activity, not only in adipose tissues, but also in muscles and heart, to get a better knowledge of the control of TG partitioning between these tissues in meat-producing cattle.
0305-0491/98/$ - see front matter © 1998 Elsevier Science Inc. All rights reserved. PII S0305-0491(98)10090-1
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To our knowledge, LPL has been little studied in muscles of ruminant species, despite numerous studies in adipose tissues ([6]; for review, see [9]) or milk (for review, see [31]). However, the presence of LPL activity 8 – 17-fold lower in beef heart compared to that in rat heart has been previously reported [12,24]. As in the rat, bovine LPL activity is expected to be lower in skeletal muscles than in heart [23]. Therefore, the first objective of our work was to develop a more sensitive assay of LPL activity in bovine skeletal muscles using an improved protocol [4]. In addition, the expression of the LPL gene in bovines has been only studied in mammary gland [22,37] and adipose tissue [6] and, LPL transcripts were barely detectable in beef heart [24]. Therefore, the second objective of this work was to characterize and quantify the mRNAs coding for LPL in various bovine tissues, especially in muscles.
2. Material and methods
seter (MA), diaphragm (D), rectus abdominis (RA), longissimus thoracis (LT) at the level of the 6th rib, semitendinosus (ST) from the medial portion, tensor fasciae latae (TFL), cutaneus trunci (CT) from the thick part, perirenal adipose tissue (PAT), omental adipose tissue (OAT) and subcutaneous adipose tissue (SCAT). Tissue samples were frozen in liquid nitrogen within 10 min of exsanguination and stored at − 80°C for subsequent analysis. Frozen samples were pulverized in liquid nitrogen. Others tissues (heart, lungs, kidney, spleen, small intestine and liver) as well as PAT were taken from five-week-old preruminant crossbred Friesian-Holstein male calves. Brain, adrenal glands, ovaries and mammary gland were taken from non-lactating pregnant cows. Samples of PAT and masseter muscle were taken from 230–260-day-old foetus. Muscle tissues (heart, soleus, extensor digitorum longus, epithroclearis) were taken after an overnight fast from 2–4-month-old male Sprague-Dawley rats (Iffa-Credo, L’Arbresle, France) and were treated as bovine tissues.
2.1. Reagents 2.3. Analytical techniques [3H]triolein (1.85– 2.96 GBq mmol − 1) and Hyperfilms MP were supplied by Amersham (Bucks, UK). [a-32P]dCTP ( \ 111 TBq mmol − 1) and [g-32P]dATP ( \ 111 TBq mmol − 1) were purchased from ICN Biochemicals (Irvine, CA). Guanidium thiocyanate was obtained from Fluka (Ronkonkoma, NY). Genescreen membranes were from Du Pont-NEN (Boston, MA). T4 polynucleotide kinase was supplied by New England Biolabs (Beverly, MA). Saturated phenol, chloroform/ isoamyl alcohol (49:1 v/v), agarose and nonaprimer labeling kit were purchased from Applige`ne (Illkirch, France). RNA molecular weight markers were from BRL (Bethesda, MD). Other reagents were from Sigma (St. Louis, MO). Antibodies to LPL purified from bovine milk were raised in rabbits [8].
2.2. Animals and experimental design Seven Montbe´liard male calves, tied and housed in individual stalls were used. Animal characteristics were described previously [19]. Briefly, from weaning (107– 128 days of age) to slaughter (170 days of age), calves were fed individually concentrate (80%) and hay (20%) according to a feeding pattern designed to allow an average daily gain of 1300 g. Composition of the concentrate was dehydrated alfalfa (30%), sugar beet pulp (40%), barley (14%) soybean meal (11%) and mineral compound (5%). The concentrate contained 16.2% of protein and 1.8% of fat. Slaughtering was done at 09:00 h after an overnight fast by stunning (captive-bolt pistol) and exsanguination. Tissue samples (50 – 100 g) from the following tissues were taken as described [18,19]: heart (H), mas-
Tissue protein or DNA contents and isocitrate dehydrogenase (ICDH) activity were measured as previously described [18,19].
2.4. Assay of LPL acti6ity LPL activity was assayed after detergent extraction as previously described [4]. Fresh tissue (six to seven tissue samples corresponding to approximately 4 g) or frozen tissue powder were homogenised at 4°C in 9 ml per g of the following buffer: ammonia–HCl (25 mM), pH 8.2 containing EDTA (5 mM), Triton X-100 (0.8% w/v), SDS (0.01% w/v), heparin (5000 IU l − 1) and peptidase inhibitors [pepstatin A (1 mg ml − 1), leupeptin (10 mg ml − 1), and aprotinin (0.017 TIU ml − 1)]. The inclusion of protease inhibitors and/or elimination of Ca2 + inhibit loss of LPL activity induced by frozen storage [36]. Insoluble material was removed by centrifugation (20000× g for 20 min at 4°C). LPL assay was performed as below. The substrate emulsion was Intralipid 10% (120 mM of TG) into which a trace amount of [3H]triolein (approximately 14 MBq) had been incorporated by sonication (75 W, 10 min). The emulsion was stored at 4°C and used for approximately two weeks since blank values increase beyond 10–15 days. Incubation medium was prepared from 10 ml emulsion, 10 ml heat-inactivated serum as a source of apolipoprotein CII, 60 ml of deionized water and 100 ml of the incubation buffer which contained 12% fatty acid free bovine serum albumin, 0.02% standard heparin, 0.2 M NaCl and 0.3 M Tris–HCl, pH 8.5. The total volume was adjusted to 200 ml with the
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sample volume diluted 5-fold in water (20 ml, i.e. 4 ml of tissue homogenate). The final substrate concentration was 6 mM. Incubation was carried out for 60 min at 25°C in a shaking water bath. The reaction was stopped by addition of distilled water (0.5 ml) and 2 ml of isopropanol/heptane/H2SO4 (48:48.3:1 v/v/v). Total lipids were extracted and fatty acids separated from TG as follows: after centrifugation (2000× g for 3 min, 4°C), a sample of the upper phase (800 ml) containing total lipids was transferred to new tubes into which 1 ml alkaline ethanol (ethanol 95%/water/2 M NaOH, 500:475:25 v/v/v) and 3 ml heptane were added. After a second centrifugation, the upper heptane phase containing unhydrolysed TG was discarded. A new extraction was performed with 3 ml of heptane. Finally, an aliquot (800 ml) of the remaining alkaline ethanol phase containing fatty acids was counted. Extraction efficiency of fatty acids was determined to be 41.89 0.53% (mean 9S.E., n = 20) and was similar to that observed for rat tissues [4]. All incubations were performed in triplicate. LPL activity in mU per g tissue wet weight (1 mU =1 nmol of fatty acids released per min) was calculated taking into account dilution of sample, extraction efficiency of fatty acids, incubation time, specific activity of the substrate and tissue/volume ratios in homogenates.
2.5. Quantification of mRNA coding for LPL Total RNA was isolated and analysed as previously described [18,19]. For Northern analysis, RNA aliquots (40 mg) were denatured in a solution containing 2.2 M formaldehyde and 50% formamide (v/v) by heating at 65°C for 10 min, size-fractionated by 1.5% agarose gel electrophoresis, and electrophoretically transferred to Genescreen membranes. The bovine LPL probe was labeled by random priming with 32P using the nonaprimer labeling kit. Prehybridization and hybridization to the LPL probe were performed at 42°C for 2–4 h and 16–20 h, respectively, in solutions containing 45% deionized formamide [19]. Then, the membranes were washed twice for 5 min in 2X SSPE at room temperature (1X SSPE=0.15 M NaCl, 10 mM NaH2PO4, 1 mM EDTA, pH 7.4), once for 30 min in 2X SSPE/2% SDS at 55°C, once for 15 min in 1X SSPE/2% SDS at 55°C and once for 15 min in 0.1X SSPE/2% SDS at 55°C. Hyperfilm MP were exposed to the membranes for 2 – 15 days at − 80°C with two intensifying screens. Quantification was performed with scanning densitometry (Hoeffer, San Francisco, CA). Preliminary Northern blot experiments were performed with various levels of RNA loaded on the gel from 10 to 50 mg. Membranes were hybridized with the LPL probe and also with a probe for the rat 18S ribosomal RNA [19]. Levels of LPL mRNA were positively correlated with levels of 18S rRNA (r = 0.895,
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n = 7, PB 0.01, data not shown). Therefore, under standard conditions (40 mg of total RNA), quantification of LPL mRNA was corrected for variations of the amount of RNA loaded on each lane by using values of hybridization to the 18S rRNA probe [19]. Results were expressed in arbitrary densitometric units per mg of total RNA loaded on the gel.
2.6. Statistical analysis Results were expressed as means 9 S.E. of four to seven animals. Significant differences between enzyme activities or mRNA levels in two different tissues from the same animals were evaluated by paired Student’s t-test. 3. Results
3.1. De6elopment of the LPL assay procedure LPL activity in bovine heart was at maximum at pH 8.5 as in rat samples [4]. A minimum of 10% of albumin and of 0.02% of heparin in the incubation buffer was necessary to observe maximum LPL activity. The amount of substrate (6 mM in the incubation medium) was saturating since LPL activity was at a maximum between 6 and 9.3 mM for both heart and adipose tissue. LPL activity was linear with time and similar between assays at 25°C but not at 37°C (Fig. 1A) as noted previously [4]. Similar results were obtained with adipose tissues (results not shown). The assay was linear between 0.2 and 4 ml of tissue homogenate, equivalent to 20–400 mg of fresh tissue (Fig. 1B) independent of tissue (heart, muscles or adipose tissue). Beyond this point, a partial inhibition of LPL activity occurred. Therefore, LPL activity was routinely assayed at 25°C for 60 min with 4 ml of homogenate in order to measure the highest level of activity in the linear range [27]. Under these standard conditions, hydrolysis did not exceed 10% of total amount of substrate for any sample. Finally, the intra-assay variation was low (9 5–10%), whereas inter-assay variations were higher (13–30% for the samples with the lowest activity and less than 10% for tissues which exhibit the highest activity).
3.2. Specificity of the measured LPL acti6ity As in human adipose tissue [20], LPL activity was completely inhibited in presence of a high concentration (100 mM) of sodium deoxycholate because this detergent desorbs the enzyme from lipid droplets [28]. In contrast, only a partial inhibition of LPL was observed in the presence of 0.5–1 M NaCl: − 18 to − 21% for adipose tissue and − 31 to −39% for heart and skeletal muscles.
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Fig. 1. Biochemical characteristics of the LPL assay in bovine heart. (A) Effect of incubation time on the velocity of the enzyme reaction. Results are expressed in mmol of fatty acids released for the indicated time. Four assays were done at two different temperatures demonstrating the absence of replication at 37°C but showing the linearity of the assay at 25°C with respect to time. (B) Effect of the amount of tissue homogenate on the measurement of LPL activity. Results are expressed in nmol of fatty acids released per min for the indicated amounts of homogenate. In all experiments, 1 g of fresh tissue was homogenized in 9 ml of buffer. The incubation medium (200 ml) was prepared from 10 ml emulsion, 10 ml serum, 60 ml deionized water, 100 ml of the incubation buffer and 20 ml of diluted sample.
Since LPL binds to heparin, tissue homogenates of bovine muscles were passed through a column of heparin-sepharose. After elution with a NaCl gradient, only one peak of LPL activity was detected at the expected concentration of NaCl suggesting the existence of only one lipase assayed by our protocol. Finally, lipase activity was completely abolished in presence of polyclonal antibodies against bovine LPL, which may be considered as the best test of specificity [8].
than with bovine serum (Fig. 2). In contrast, LPL activity in adipose tissues was similar with the two sources of sera (ratio of 1.09 0.02; mean9 S.E. of 16 samples; Fig. 2). Nevertheless, levels of LPL activity assayed with rat serum were positively correlated with levels of LPL activity measured with bovine serum for both adipose tissues (r= 0.99, n = 24, PB 0.001) and muscles (r= 0.76, n= 28, P B 0.01) as shown in Fig. 2. From all these experiments, we decided to use the best batch of rat serum for LPL assay in standard conditions.
3.3. Effect of serum acti6ator on LPL acti6ity 3.4. Effect of tissue processing on LPL acti6ity For LPL assay, one must also add serum to the incubation medium to provide an activator (apo CII) to obtain maximum activity. Basal LPL activity (i.e. without serum) was between 19 and 52% of maximum activity for heart, skeletal muscles or adipose tissues. This relatively high and variable basal LPL activity may be explained by the presence of small amounts of blood serum in the tissue which might differ among samples. To activate LPL, the optimum amount of serum was found to be 5% by volume independent of the source of serum (rats or bovines). But great variations in LPL activity were observed depending on the source of serum in interaction with the source of tissue extract (Table 1). Therefore, we pooled the best rat sera and the best bovine sera to obtain two homogeneous sources of activators [4,27]. When these two batches of sera were tested in the same assay, rat serum was more effective than bovine serum to activate LPL in heart and skeletal muscles. In one representative experiment, LPL activity was 5.59 0.27-fold higher (mean9S.E. of 28 samples) with rat
At the time of slaughter of the animals, it is often impossible to perform LPL assays immediately. Therefore, we tested different conditions of processing and storage of samples. In a first experiment, two procedures were studied: the first batch of tissues was frozen in liquid nitrogen, stored at − 80°C, pulverized in liquid nitrogen and homogenized in the appropriate buffer on the day of assay; the second batch of tissues was frozen into flasks with 9 ml of homogenisation buffer for LPL assay (containing detergents and heparin to protect LPL against any degradation) per g tissue wet weight and stored at − 80°C until subsequent homogenisation after complete thawing on the day of assay. LPL activity in adipose tissues was 1.49 0.13-fold higher for samples frozen within homogenisation buffer than for tissue powder (mean9 S.E., n= 6, PB 0.07). In contrast, LPL activity in heart and masseter muscle were 1.49 0.08- and 2.7 90.38-fold higher, respectively, for tissue powder (mean9S.E., n=8, PB 0.05).
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Table 1 LPL activity in the presence of bovine or rat sera from different donors as source of activator Source of rat sera
Donor Donor Donor Donor Donor
1 2 3 4 5
LPL activity in assay 1 (%) Heart
Omental adipose tissue
100 68.4 84.8 77.2 60.8
100 90.4 110.0 90.4 81.9
Source of bovine sera
Donor Donor Donor Donor
A B C D
LPL activity in assay 2 (%) Heart
Omental adipose tissue
100 54.8 51.6 80.0
100 54.6 62.2 63.2
LPL activity in two samples was measured in triplicate with rat sera from five different donors (assay 1) or with bovine sera from four different donors (assay 2) to compare the effects of the source of serum. Results of LPL activity were first calculated in mU per g tissue wet weight as usual and then expressed within the same column relative to the values obtained for donors 1 and A for assays 1 and 2, respectively.
In a second experiment, we showed that, in heart, one or two of freeze/thaw cycles induced a 10–20% decrease in LPL activity. After six cycles, a 30% decrease was observed. In adipose tissues, a 20 –40% decrease in LPL activity was detected after only one cycle and a 40–60% decrease was observed after two to six cycles of freezing/thawing suggesting that LPL is less stable in adipose tissue than in heart. Thus, we recommend that LPL should be assayed on fresh samples, and, if this is not possible, that samples or tissue homogenates should be frozen only once.
3.5. LPL acti6ity and gene expression in 6arious bo6ine tissues LPL activity was barely detectable in small intestine, kidney, spleen, adrenal glands and lung from young calves. The LPL activity in liver was very low and even undetectable in some assays (Table 2). For these tissues, blank values might represent more than 50% of total
values. Thus, absolute values for LPL activity were considered unreliable. In contrast, LPL activity was higher in heart and adipose tissues (Table 2). Northern blot analysis revealed the presence of three different-sized species of LPL mRNAs in bovine tissues (Fig. 3). The sizes of the two major mRNAs were approximately 3.5–3.8 and 3.2–3.4 kb. The 3.5–3.8 kb species seemed to be the most abundant transcript in many, but not all, experiments. The minor mRNA species (1.7 kb), was detected at a low level in tissues which expressed the highest level of LPL transcripts. Only the two major mRNA species were quantified by scanning densitometry since the third mRNA species was minor and not always detected. In agreement with the results for activity, levels of LPL mRNAs were high in heart and adipose tissues from calves even during foetal life (Fig. 3). In contrast, LPL mRNAs occurred in very low levels in spleen, adrenal glands and ovaries, barely detectable in lung, small intestine and kidney and undetectable in liver and brain.
3.6. LPL acti6ity and gene expression in eight bo6ine muscles
Fig. 2. Effect of serum activator on LPL activity in bovine tissues. Tissue LPL activity was measured with either bovine serum or rat serum in the same assay in heart, masseter (MA), omental adipose tissue (OAT) and perirenal adipose tissue (PAT). LPL activity measured with rat serum was plotted against LPL activity measured with bovine serum.
Eight different muscles from an homogeneous group of calves were characterized using ICDH activity, characteristic of oxidative metabolism [18,19] (Fig. 4). LPL activity assayed with rat serum was higher in heart and masseter (the most oxidative muscles) than in other skeletal muscles (Fig. 4). Similar results were observed with bovine serum as activator although absolute values were lower than those with rat serum (449922, 2569 39 and 29–77 mU g − 1 tissue wet weight for heart, masseter and other skeletal muscles, respectively). Surprisingly, LPL activity in the diaphragm was as low as in glycolytic muscles although ICDH activity was higher than in other skeletal muscles of the carcass. LPL activity was also expressed in mU per mg of DNA and per mg of protein. Similar results and similar differences among muscle tissues were observed whatever the method of expression of the results and the source of serum activator used in the assay.
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Table 2 Range of LPL activity in various bovine tissues Liver
Small intestine
Kidney
Spleen
Adrenal glands
Lung
Heart
Perirenal adipose tissue
0–58
137–340
50–250
58–184
140–202
80–208
473–1465
580–3560
Hearts and adipose tissues were obtained from 12 1–6-month-old calves. No significant effect of age was found. Other tissues were taken from five 5-week-old preruminant calves. Adrenal glands were taken from cows. LPL activity was measured in triplicate with different sera (5% v/v) as source of activator in many different assays for heart and adipose tissue and in two different assays for other tissues. The highest and the lowest values in mU per g tissue wet weight obtained in the different assays for the different animals are indicated.
LPL activity levels in rat heart and soleus (2150– 2950 and 1160–1350 mU g − 1 wet tissue, respectively) were higher (PB0.05) than in bovine heart and masseter (662–832 and 412 – 972 mU g − 1 wet tissue, respectively). LPL activity was also higher in rat extensor digitorum longus and epithroclearis, the most glycolytic muscles in the rat (130 – 190 mU g − 1 wet tissue) than in RA, TFL, ST, LT and CT (33 – 154 mU g − 1 wet tissue), the most glycolytic muscles in the calf [18,19]. LPL mRNA levels expressed per mg of total RNA were highest in oxidative muscles (Fig. 5). However, the level of LPL mRNA in masseter was approximately 2-fold lower than that in heart, despite similar LPL activity levels (Figs. 4 and 5). When results of LPL mRNA levels were expressed per g wet tissue weight, differences between muscles were amplified (60.69 23.4, 21.794.1, 0.5– 8.4 arbitrary units for heart, masseter and other skeletal muscles, respectively) since yields of extracted RNA were higher in oxidative muscles, especially in heart as in rats [23] and in bovines [18,19].
3.7. LPL acti6ity and gene expression in three bo6ine adipose tissues LPL activity was higher in internal adipose tissues (OAT and PAT) than in SCAT (Fig. 4). Similar results were observed with bovine serum as activator (971 9 218, 2589 121 and 12469180 mU g − 1 tissue wet weight for OAT, SCAT and PAT, respectively). However, since protein and DNA contents were approximately 2- and 1.5-fold higher in SCAT, respectively, than in internal adipose tissues, differences between SCAT and internal adipose tissues were amplified when results of LPL activity were expressed per mg of protein or DNA [27]. LPL mRNA levels were higher in PAT and OAT than in SCAT (Fig. 6). LPL mRNAs in SCAT were barely detectable in some experiments (Fig. 6A), but quantifiable in most. However, the level of LPL mRNA in PAT was approximately 1.5-fold higher (P B 0.05) than in OAT, despite similar LPL activity (Figs. 4 and 6). Similar results were observed when data of LPL mRNA levels were expressed per g wet tissue weight, since yields of extracted RNA were similar in the three adipose tissues.
4. Discussion
4.1. Extraction and assay of LPL in bo6ine tissues It might seem a trivial task to develop assays of LPL in bovine tissues since many methods have been reported in the literature for LPL from milk or adipose tissues of ruminants [15,36,40]. However, by studying LPL in bovine muscles, we demonstrated that conditions for assay of LPL have to be set up and critically evaluated for each new application. Depending on the methods used to extract and to assay the enzyme, the activity recorded for muscles could vary by an order of magnitude. Factors needing special attention for the bovine muscles were sample storage, extraction procedures and the source of serum to provide activator. Tissue extracts of acetone-ether powders were widely used in the earlier literature [27], but have been found to give low recovery. Most investigators now favour homogenates in buffers with heparin and detergents which help to stabilize the enzyme, to extract it from intracellular vesicles and to dissociate it from heparin sulfate proteoglycans [4]. The earlier recommendation was to store tissue pieces frozen in this buffer until they were homogenized for assay [4]. In bovine, this worked well for adipose tissue, but not for muscles since the muscle pieces need to be pulverized in liquid nitrogen. As substrate, we used radioactive triolein that was incorporated into a commercial phospholipid-stabilized TG emulsion. This provides high sensitivity, but substantial between-assay variation, especially when different emulsion preparations are used [27]. However, the intra- and inter-assay variations of our assay were in agreement with other data in rodents with the same assay procedure [4] or in cow adipose tissues with another procedure [15]. Tissue samples may contain other lipase activities. In the present system, the substrate with long-chain TG in a lipoprotein-like emulsion virtually eliminates the contribution of non-specific esterases with water-soluble esters as substrate, and the high pH minimizes the contribution from lysosomal enzymes [27]. The specificity of our assay was demonstrated by the fact that, (i) heparin-agarose columns bound the lipase activity which then eluted as a single peak in the position
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Fig. 3. Northern blot analysis of the tissue distribution of LPL mRNAs in bovine tissues. Total RNA was extracted from perirenal adipose tissue (PAT), foetal perirenal adipose tissue (fPAT), liver (Li), small intestine (SI), kidney (Ki), spleen (Sp), adrenal glands (AG), brain (Br), lung (Lu), ovaries (Ov), heart, (H), foetal heart (fH) and masseter muscle (MA). RNAs (40 mg) was then analyzed by Northern blot using the bovine LPL cDNA. Films were exposed one (for adipose tissues) or seven days (for the other tissues) to the membranes.
characteristic for LPL [17], and (ii) polyclonal antibodies against bovine milk LPL suppressed the lipase activity almost completely. The commonly used criterion to differentiate LPL from other lipases, i.e. its inhibition by NaCl, is not a very reliable tool since it is very dependent on the composition of buffers (for review, see [11]) or on the conditions of incubation [3]. At 37°C, the inactivation by NaCl is rapid and almost complete. At 4 or 10°C, and with albumin, the enzyme works well at 1.5 M NaCl (for review, see [28]). In our assay at 25°C, 1 M NaCl caused only partial loss of activity and this differed with the source of tissue extract. The pH-dependency of the reaction depends on an interplay of factors [4] such as pH effects on the binding of LPL to lipid droplets and on the active site reaction (for review, see [28]). However, LPL extracted from human muscle showed about the same activity from pH 7.9–8.7 [26], whereas the highest level of activity for LPL from bovine muscles was obtained between pH 9 and 9.5 [24]. For the present assay system, small variations in pH of the incubation medium might affect the activity and hence be a source of interassay variability. This is the reason why in our laboratory, a large batch of incubation medium is prepared, and used in all assays to minimize inter-assay variability.
4.2. Differential effect of bo6ine or rat sera on LPL acti6ation Apo CII provided by the serum binds to the lipid– water interface and to the enzyme. With soluble model substrates, apo CII has little or no effect (for review, see [17]). With emulsified substrates and in monolayer systems, the effect of apo CII varies widely, and this appears to relate to the ‘quality of the interface’ [3]. There are examples of systems where LPL exerts full activity without apo CII and others where the enzyme has little or no activity without apo CII. Hence, it appears that the role of apo CII is to restore the activity in situations where other factors suppress it [3]. Particu-
larly important is the presence of other lipid-binding proteins, for instance, apolipoproteins such as apo CIII or fragments of these apolipoproteins, and also unrelated proteins [4]. A striking finding here was the 5-fold higher activation of the lipase activity with rat compared to bovine serum for heart and muscles, but not for adipose tissue. We tentatively ascribe this to the presence of much more protein in the muscle extracts, and note that haemoglobin strongly suppresses LPL activity [3]. Another possibility is that there are differences between the lipase in muscle and in adipose tissue as suggested by kinetic [26] or immunological studies [38]. There is, however, only one gene for LPL [21] but LPL undergoes a wide range of posttranscriptional [21] or posttranslational [13] modifications that may occur in a tissue related-fashion. It may seem paradoxical that rat serum was more efficient than bovine serum for LPL activation. Although species-specific immunological or enzymatic properties of the LPL molecule alone have been reported [8,40], there is no evidence for species specificity in the LPL–apo CII interaction. In model systems, a similar activation of bovine LPL was obtained using apo CII or analogues from as widely different animals as cows, humans, rats, guinea pigs, chicken and salmon. Conversely, human apo CII has been shown to activate LPL from many different animals including cows and rats [2,25]. In agreement with this, the region in apo CII that interacts with the enzyme is quite strictly maintained across bovine and other species in the C-terminal third of the molecule required for LPL activation [5]. Hence it appears unlikely that our observations relate to a particularly good fit between the bovine enzyme and rat apo CII. These differential effects of bovine and rat sera on LPL activation may be linked to the absolute and relative concentrations of apo CII and other apolipoproteins (for instance, apo CIII) which might differ between rats and cows. In addition, the shift of apolipoproteins from plasma lipoproteins of the added activator to the lipid–water interface depends on sev-
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Fig. 4. Metabolic activity of various muscles and adipose tissues from seven 170-day-old ruminant calves. Values are means 9 S.E. of four to seven observations. Isocitrate dehydrogenase (ICDH) activity was measured spectrophotometrically in muscle tissue homogenates and expressed in U g − 1 tissue wet tissue (1 U =1 mmol of substrate used per min). LPL activity was measured in tissue homogenates with rat serum as source of activator and expressed in mU g − 1 tissue wet weight (1 U = 1 mmol of fatty acid released per min). A value followed by a superscript differs significantly (PB 0.05) from all other values not followed by the same superscript. H, heart; MA, masseter; D, diaphragm; RA, rectus abdominis; TFL, tensor fasciae latae; LT, longissimus thoracis; ST, semitendinosus; CT, cutaneus trunci; OAT, omental adipose tissue; SCAT, subcutaneous adipose tissue; PAT, perirenal adipose tissue.
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Fig. 5. Northern blot analysis of the LPL mRNAs distribution in muscles from seven 170-day-old ruminant calves. Total RNA was extracted from bovine muscles and then analyzed by Northern blot using the bovine LPL cDNA (A) and the 18S rRNA probe (B) to verify that equivalent amounts of total RNA were loaded on each lane. Quantification of LPL mRNA and of 18S rRNA levels was performed by scanning densitometry. Different films exposed 1–8 days to the same membrane were used for quantification to be sure that the signals were within the linear range of the films depending on the samples and the time of exposure. Results of LPL mRNA levels were corrected for variations in 18S RNA levels and were then expressed in densitometric arbitrary units per mg of total RNA loaded on the gel relative to the level in longissimus thoracis (C). Values are means 9S.E. of five to seven observations (C). A value followed by a superscript differs significantly (P B0.05) from all other values not followed by the same superscript. H, heart; MA, masseter; D, diaphragma; RA, rectus abdominis; TFL, tensor fasciae latae; LT, longissimus thoracis; ST, semitendinosus; CT, cutaneus trunci.
eral factors including the relative concentrations of HDL and VLDL and the level of TG in the serum activator [39]. Therefore, the fact that the proportions of circulating VLDL and HDL differ markedly between rats and ruminants [41] might also induce differences in activation of LPL by the two types of sera. There are also differences in activation of LPL by apo CII from different species that relate to the interaction with the lipid interface rather than with the enzyme [2]. In support of this, there are rather large differences among apo CII from different species in the N-terminal twothirds of the molecule which is responsible for interaction with the lipid– water interface [5]. It may be this type of difference that made rat serum a much better source of activator for the LPL in bovine muscle samples. Whatever the reasons for the observed differences, we used rat serum in our assay to obtain high values for LPL activity in samples from muscle, and to reduce interassay variability. To enhance reproducibility, it is advisable to prepare a large batch and freeze this in aliquots.
4.3. Comparison of LPL acti6ity in similar tissues by different procedures When we applied the present method to rat heart, we obtained values (2150 – 2950 mU g − 1) similar to those reported using a similar procedure for LPL assay [8]. These values are higher than those reported by most investigators (150 [16], 500 [23], 950 – 1250 [12] mU g − 1, but similar to those reported by others (1300 – 6500 mU
g − 1 [34]). The values we obtained for LPL activity in bovine heart with rat (749 mU g − 1) or bovine (449 mU g − 1) serum as activator were much higher than those previously reported (90 mU g − 1 [24]). Our values for LPL in calf adipose tissues (372–923 mU g − 1) are similar to those in adipose tissues of steers (500–833 mU g − 1 [35]), but higher than those of cows (133 mU g − 1 [15]) or sheep (85–109 mU g − 1 [40]). The values for LPL activity in skeletal muscles from our calves (33–154 mU g − 1) are higher than those for sheep (6–14 mU g − 1 [40]; 15–55 mU g − 1 [36]) or human muscles (5–32 mU g − 1 [26]). Thus, LPL activity levels registered in this study are in the upper range of values reported in the literature for LPL activity in rat or ruminant tissues. Some of these differences may be explained by physiological parameters of the animals such as age or feeding [8,34,36], but the sensitivity and the quality of the assay procedure is of prime importance especially with tissues which exhibit a low activity such as bovine muscles. The LPL activity in bovine heart was 3.4–4.8-fold lower than that in rat heart depending on the serum used. This difference is not unexpected due to differences in sizes and, hence, in metabolic rate between the two studied species. However, a much higher difference (8–17-fold) was previously reported [12,24].
4.4. Detection of LPL transcripts in bo6ine tissues The detection of three different-sized species of LPL mRNAs in bovine tissues (the two larger species being present in significant amounts for quantification) was in
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Fig. 6. Northern blot analysis of the LPL mRNAs distribution in adipose tissues from seven 170-day-old ruminant calves. Total RNA was extracted from bovine adipose tissues and then analyzed by Northern blot using the bovine LPL cDNA (A) and the 18S rRNA probe (B) as described in Fig. 5. Results of LPL mRNA levels were corrected for variations in 18S RNA levels and were then expressed in densitometric arbitrary units per mg of total RNA loaded on the gel relative to the level in omental adipose tissue (C). Values are means9S.E. of four to seven observations (C). A value followed by a superscript differs significantly (P B0.05) from all other values not followed by the same superscript. OAT, omental adipose tissue; SCAT, subcutaneous adipose tissue; PAT, perirenal adipose tissue.
agreement with the previous observations in bovine mammary gland [22,37] and adipose tissue [6]. In contrast, it was reported that the primary LPL mRNA detected in beef heart was in amounts too low for accurate quantification [24]. This may be explained by methodological differences between the two studies: (i) we used an homologous probe instead of a mouse LPL cDNA, (ii) labelling was performed by random priming rather than by the nick translation method, (iii) total RNA were denaturated with 6.6% formaldehyde instead of 3.2%. Finally, the small differences reported for the size of the LPL transcripts in bovine tissues among laboratories [6,22,24,37]; and this study) could be explained by the different standards used for calculation (DNA or RNA kb ladders, ribosomal RNAs). However, polymorphisms affecting mRNA sizes among tissues and/or species or strains [21] cannot be excluded. Except in liver and brain, LPL mRNA transcripts were observed in all tissues examined albeit at very different levels. In addition, LPL mRNA levels were higher in growing calves than in foetuses as described in the rat [21]. As in mouse, human [22] and rat tissues [21,23], levels of LPL transcripts were higher in bovine heart and oxidative skeletal muscles which use fatty acids as energy source than in muscles composed of fast-twitch white fibers which use glucose as fuel [23]. Great differences in metabolism among adipose sites have been previously reported in pigs [1] and calves [18]. In the cow, LPL activity may also differ among adipose tissues from various anatomical sites [10]. These differences are mainly related to differences in LPL mRNA levels among adipose sites (Fig. 6). Our results also suggest that the higher the LPL
activity, the higher the adipocyte size as previously described in rats and humans [14]. The LPL system is subject to regulation at many levels involving gene expression, synthesis/degradation, intra- and intercellular translocation, activation/inactivation by serum factors. However, LPL activity depends primarily on a balance between rates of synthesis and degradation (for review, see [11]). Our results support this last idea since differences in LPL activity among a variety of bovine tissues are mainly reflected by differences in LPL mRNA levels, as in the rat [23]. However, other mechanisms of regulation may also be involved in the fasted rat [23] and in the bovine since LPL activity did not always parallel mRNA levels among tissues in rats [23] and in cattle (this study). In conclusion, although LPL specific activity and gene expression in bovine muscles are significantly lower than that found in rat muscles, we have developed sensitive procedures to assay LPL and to quantify LPL mRNA levels in bovine tissues. Acknowledgements We thank Dr D. Bauchart (INRA, Laboratoire Croissance et Me´tabolismes des Herbivores, France) for helpful discussions and Dr Y. Furuichi (Nippon Research Center, Kamakura, Japan) for kindly providing us the LPL bovine cDNA. We also thank Nicole Guivier and Samia Brazi for their expertise and contributions, Robert Jailler and his group for the management of the animals and Gilbert Cuylle and his group for the management of the slaughterhouse. This study was supported in part by grant 13x-727 from the Swedish Medical Research Council.
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References [1] Anderson DB, Kauffman RG, Kastenschmidt LL. Lipogenic enzyme activities and cellularity of porcine adipose tissue from various anatomical locations. J Lipid Res 1972;13:593–9. [2] Andersson Y, Lookene A, Shen Y, Nilsson S, Thelander L, Olivecrona G. Guinea pig apolipoprotein CII: expression in E. coli, functional studies of recombinant wild-type and mutated variants and distribution on plasma lipoproteins. J Lipid Res 1997;38:2111 – 24. [3] Bengtsson G, Olivecrona T. Lipoprotein lipase: some effects of activator proteins. Eur J Biochem 1980;106:549–55. [4] Bengtsson-Olivecrona G, Olivecrona T. Assay of lipoprotein lipase and hepatic lipase. In: Converse C, Skinner ER, editors. Lipoprotein Analysis. Oxford: Oxford University Press, 1991:169 – 185. [5] Bengtsson-Olivecrona G, Sletten K. Primary structure of the bovine analogues to human apolipoproteins CII and CIII. Studies on isoforms and evidence for proteolytic processing. Eur J Biochem 1990;192:515–21. [6] Bonnet M, Faulconnier Y, Fle´chet J, Hocquette JF, 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. [7] Borensztajn J. Heart and skeletal muscle lipoprotein lipase. In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:133 – 148. [8] Chajek-Shaul T, Bengtsson-Olivecrona G, Peterson J, Olivecrona T. Metabolic fate of rat heart endothelial lipoprotein lipase. Am J Physiol 1988;255:E247–54. [9] Chilliard Y. Dietary fat and adipose tissue metabolism in ruminants, pigs, and rodents: a review. J Dairy Sci 1993;76:3897 – 931. [10] Chilliard Y, Robelin J. Activite´ lipoprote´ine-lipasique de diffe´rents de´poˆts adipeux et ses relations avec la taille des adipocytes chez le vache tarie en cours d’engraissement ou en de´but de lactation. Reprod Nutr Dev 1985;25:287–93. [11] Cryer A. Comparative biochemistry and physiology of lipoprotein lipase In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:277–327. [12] Cryer A, Jones HM. The distribution of lipoprotein lipase (clearing factor lipase) activity in the adipose, muscular and lung tissues of ten animal species. Comp Biochem Physiol 1979;63B:501 – 5. [13] Doolittle MH, Ben-Zeev O, Elovson J, Martin D, Kirchgessner TG. The response of lipoprotein lipase to feeding and fasting. J Biol Chem 1990;265:4570–7. [14] Eckel RH. Adipose tissue lipoprotein lipase. In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:79–132. [15] Faulconnier Y, The´venet M, Fle´chet J, Chilliard Y. Lipoprotein lipase and metabolic activities in incubated bovine adipose tissue explants: effects of insulin, dexamethasone, and foetal bovine serum. J Anim Sci 1994;72:184–91. [16] Galan X, Llobera M, Ramirez I. Lipoprotein lipase and hepatic lipase in Wistar and Sprague-Dawley rat tissues. Difference in the effects of gender and fasting. Lipids 1994;29:333 – 6. [17] Garfinkel AS, Schotz MC. Lipoprotein lipase. In: Gotto AM Jr, editor. Plasma Lipoproteins. Amsterdam: Elsevier, 1987:335 – 357. [18] Hocquette JF, Castiglia-Delavaud C, Graulet B, Ferre´ P, Picard B, Vermorel M. Weaning marginally affects glucose transporter (GLUT4) expression in calf muscles and adipose tissues. Br J Nutr 1997;78:251–71. [19] Hocquette JF, Graulet B, Castiglia-Delavaud C, Bornes F, Lepetit N, Ferre´ P. Insulin-sensitive glucose transporter tran-
[20]
[21]
[22]
[23]
[24]
[25]
[26]
[27]
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35] [36]
[37]
211
script levels in calf muscles assessed with a bovine GLUT4 cDNA fragment. Int J Biochem Cell Biol 1996;28:795 –806. Iverius P-H, Brunzell JD. Human adipose tissue and lipoprotein lipase: changes with feeding and relation to postheparin plasma enzyme. Am J Physiol 1985;249:E107– 14. Kirchgessner TG, LeBoeuf RC, Langner CA, Zollman S, Chang CH, Taylor BA, Schotz MC, Gordon JI, Lusis AJ. Genetic and developmental regulation of the lipoprotein lipase gene: loci both distal and proximal to the lipoprotein lipase structural gene control enzyme expression. J Biol Chem 1989;264:1473– 82. Kirchgessner TG, Svenson KL, Lusis AD, Schotz MC. The sequence of cDNA encoding lipoprotein lipase. J Biol Chem 1987;262:8463– 6. LaDu MJ, Kapsas H, Palmer WK. Regulation of lipoprotein lipase in adipose and muscle tissues during fasting. Am J Physiol 1991;260:R953– 9. LaDu MJ, Schultz CJ, Essig DA, Palmer WK. Characterization of serum-stimulated lipoprotein lipase from bovine heart. Int J Biochem 1991;23:405 – 11. Lindberg A, Olivecrona G. Lipase evolution: trout, Xenopus and chicken have lipoprotein lipase and apolipoprotein C-IIlike activity but lack hepatic lipase-like activity. Biochim Biophys Acta 1995;1255:205 – 11. Lithell H, Boberg J. Determination of lipoprotein-lipase activity in human skeletal muscle tissue. Biochim Biophys Acta 1978;528:58 – 68. Nilsson-Ehle P. Measurements of lipoprotein lipase activity. In: Borensztajn J, editor. Lipoprotein Lipase. Chicago: Evener, 1987:59 – 77. Olivecrona T, Bengtsson G. Lipases in milk. In: Borgstro¨m B, Brockman HL, editors. Lipases. Amsterdam: Elsevier, 1984:205 – 261. Olivecrona T, Bengtsson-Olivecrona G, Chajek-Shaul T, Carpentier Y, Deckelbaum R, Hultin M, Peterson J, Patsch J, Vilaro S. Lipoprotein lipase. Sites of synthesis and sites of action. Atheroscler Rev 1991;22:21 – 5. Olivecrona T, Bengtsson-Olivecrona G, Hultin M, Peterson J, Vilaro S, Deckelbaum RJ, Carpentier YA, Patsch J. What factors regulate the action of lipoprotein lipase?. In: Malmendier CL et al, editors. Hypercholesterolemia, Hypocholesterolemia, Hypertriglyceridemia. New York: Plenum Press, 1990:335 – 339. Olivecrona T, Vilaro S, Bengtsson-Olivecrona G. Lipases in milk. In: Fox PF, editor. Advanced Dairy Chemistry. Amsterdam: Elsevier, 1992:292 – 309. Peterson J, Bihain BE, Bengtsson-Olivecrona G, Deckelbaum RJ, Carpentier YA, Olivecrona T. Fatty acid control of lipoprotein lipase: a link between energy metabolism and lipid transport. Proc Natl Acad Sci USA 1990;87:909 – 13. Pethick DW, Dunshea FR. Fat metabolism and turnover. In: Forbes JM, France J, editors. Quantitative Aspects of Ruminant Digestion and Metabolism. Wallingford: CAB International, 1993:291 – 311. Planche E, Boulange A, DeGasquet P, Tonnu NT. Importance of muscle lipoprotein lipase in rats during suckling. Am J Physiol 1980;238:E511– 7. Rao DR, Hawkins GE. Activity of lipoprotein lipase in adipose tissues from steers. J Dairy Sci 1976;59:161 – 3. Rule DC, Andersen MK, Bailey JW, Swain L, Ficek SJ, Thomas DP. Frozen storage of ovine and rat tissues adversely affects lipoprotein lipase activity. J Nutr Biochem 1996;7:577– 81. Senda M, Oka K, Brown WV, Qasba PK. Molecular cloning and sequence of a cDNA coding for bovine lipoprotein lipase. Proc Natl Acad Sci USA 1987;84:4369 – 73.
212
J.-F. Hocquette et al. / Comparati6e Biochemistry and Physiology, Part B 121 (1998) 201–212
[38] Soteriou A, Cryer A. Distinct immunoreactivities suggest the existence of potential tissue variants in rat lipoprotein lipase. Biochem J 1994;299:417–23. [39] Tornoci L, Scheraldi CA, Li XZ, Ide H, Goldberg IJ, Le NA, Tornoci L. Abnormal activation of lipoprotein lipase by non-equilibrating apo C-II: further evidence for the presence of non-equilibrating pools of apolipoproteins C-II and C-III in
.
plasma lipoproteins. J Lipid Res 1993;34:1793 – 803. [40] Tume RK, Thornton RF, Johnson GW. Lipoprotein lipase of sheep and rat adipose tissues. Aust J Biol Sci 1983;36: 41 – 8. [41] Vitic J, Stevanovic J. Comparative studies of the serum lipoproteins and lipids in some domestic, laboratory and wild animals. Comp Biochem Physiol 1993;106B:223– 9.