Protein depletion and replenishment in mice: different roles of muscle and liver

Oscar A. Scornik, Scott K. Howell, and Violeta Botbol

Department of Biochemistry, Dartmouth Medical School, Hanover, New Hampshire 03755-3844

    ABSTRACT
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Abstract
Introduction
Results
Discussion
References

Fully grown male CD-1 mice, fed a protein-free diet for 3 days, received 1 g of starch with or without 300 mg casein by intragastric intubation. We surveyed the acute effects of these nutrients on protein synthesis in all tissues (by extrapolating to infinity the incorporation of radioactive leucine after its injection in massive doses) and protein degradation in skeletal muscle and liver (by the accumulation of bestatin-induced peptide intermediates). Muscle proteolysis was the major source of N during depletion. Compared with postabsorptive animals, starch suppressed muscle protein loss (synthesis +21%, degradation -24%, P < 0.01) and stimulated hepatic proteolysis (degradation +28%, P < 0.01). Addition of casein to the starch was anabolic in liver (synthesis +41%, degradation -33%, P < 0.01), gastrointestinal tract, pancreas, and skin (synthesis +38, +69 and +38%, respectively, P < 0.01) but had no effect on muscle. Protein turnover proved uniquely sensitive to the dietary supply of carbohydrates in muscle and to the endogenous or exogenous supply of amino acids in liver.

protein turnover; protein synthesis; protein degradation

    INTRODUCTION
Top
Abstract
Introduction
Results
Discussion
References

THE TURNOVER OF TISSUE PROTEINS plays a major role in the control of free amino acid pools. Net proteolysis supplies amino acids between meals, during periods of dietary protein deprivation, and even larger amounts for gluconeogenesis during starvation. After adequate food intake, protein deposition restores depleted tissue proteins (8, 19, 20, 26, 28). Not all tissues participate in these processes the same way. Liver (because of its high rate of protein turnover) and muscle (because of its large mass) are the most important ones. Other tissues are very active in protein synthesis, but the proteins they synthesize are destined for export (pancreas) or renewal (skin, intestinal epithelium, hemopoietic organs), not intracellular protein turnover.

Our understanding of the regulation of protein turnover in vivo is still fragmentary and incomplete. In the experiments presented here, we took advantage of procedures developed in our laboratory to estimate both protein synthesis (24, 27) and degradation (4, 5) in tissues of live mice. We measured protein synthesis in all tissues (by extrapolating to infinity the incorporation of radioactive leucine after its injection in massive doses) and protein degradation in skeletal muscle and liver (by the accumulation of bestatin-induced peptide intermediates). These experiments were coupled with measurements, under the same conditions, of nitrogen balance and tissue protein loss. We sought experimental conditions where replenishment of depleted tissue protein could be distinguished from protein-sparing effects of carbohydrates in the meal or resumption of growth. Also, we used mice, which, because of their small size, made these experiments affordable.

The focus of this study is the acute replenishment of tissue proteins following the supply of dietary carbohydrates and amino acids. We have already shown that, after an overnight fast, a single meal stimulates protein synthesis and inhibits protein breakdown in both liver (4) and muscle (5). The effects of a meal on protein turnover are, however, complex. They may reflect not only replenishment of depleted tissue protein but also the protein-sparing effects of carbohydrates in the food and, in young animals, resumption of growth (suspended during the fast). In the present study, we use fully grown mice after a short period of protein depletion, either in the postabsorptive state or fed by intragastric intubation starch, with or without casein. We can then distinguish the protein-sparing effects of carbohydrates from the additional anabolic effects of dietary amino acids (which permit net tissue protein increase). We demonstrate that muscle and liver protein turnover have very different roles in the regulation of amino acid pools. We show that muscle protein becomes the most important source of nitrogen during protein deprivation but that it plays little or no part in the rapid replenishment of tissue proteins after the ingestion of casein. The acute anabolic effect of dietary amino acids occurs primarily in liver and is also evident in other abdominal viscera and skin.

    EXPERIMENTAL PROCEDURE

Materials. Synthetic bestatin (Ubenimex), N-[(2S,3R)-3-amino-2-hydroxy-4-phenylbutanoyl]-L-leucine, was a generous gift of Nippon Kayaku (Tokyo, Japan). L-[4,5-3H(N)]leucine (5 mCi/µmol), L-[4,5-3H(N)]lysine (110 mCi/µmol), L-[guanidino-14C]arginine (51.5 µCi/µmol), and L-[1-14C]ornithine (47.7 µCi/µmol) were obtained from DuPont NEN (Boston, MA). Dulbecco's phosphate-buffered saline, in powder form, was purchased from GIBCO Laboratories (Grand Island, NY). Unless indicated otherwise in the text or the original publications, other chemicals were purchased from Sigma Chemical (St. Louis, MO).

Mice. Fully grown male CD-1 mice, 36-38 g, were purchased from Charles River Laboratories (Wilmington, MA). They were kept in a room illuminated from 6 AM to 6 PM and were fed ad libitum with standard laboratory chow. Where indicated, this was replaced by custom-made pellet diets (ICN Biochemicals). One, defined as "4% casein diet," contained 4% casein, 0.3% methionine, 55% corn starch, 27% dextrose, 5% corn oil, 5% cellulose fiber (alphacel), and standard supplements of vitamin and minerals. Another, defined as "protein-free diet," omitted the casein and methionine and replaced them with starch. In a third diet, starch was replaced by additional casein to make a "50% casein diet."

Intragastric intubation. For the experiments described in Figs. 2 and 3, starch and casein suspensions were administered intragastrically to protein-depleted mice through a no. 18 gavage needle (Pepper and Sons, New Hyde Park, NY). Food was removed from the cages at 7 AM. For the groups labeled S or S+C, the mice received 0.5 g of starch suspended in 1 ml water at 8 AM. Starting 2 h later, each mouse received an additional 0.5 g of starch, with or without 300 mg casein, suspended in 2.7 ml water. One drop of 0.5% phenol red was added first to the casein and water with continual stirring, followed by dropwise additions of 10 N NaOH until the indicator turned orange and the casein was free of clumps; the starch was mixed in last. The suspension of starch, or starch plus casein, was divided by weight into four equal portions administered at hourly intervals from 10 AM to 1 PM. Measurements of protein synthesis and degradation were performed at 2 PM. In other experiments (not shown), we observed that excess urea excretion peaked at this time, indicating maximum rates of absorption of the dietary amino acids. These experiments also included a third group of mice, labeled in the figures as "none," which received only water from 8 AM; their gastrointestinal tract at 2 PM was empty, and the mice were considered postabsorptive. For the experiments on 16-h liver protein regain (see Fig. 1D), food was removed from the cages at noon and the mice received 0.5 g of starch suspended in 1 ml water at 1 PM. Starting 2 h later, each mouse received an additional 0.5 g of starch, with or without different amounts of casein (as indicated in Fig. 1) divided into four hourly portions as before. For the largest amount, 450 mg, the volume of water in the suspension was increased to 4 ml. The animals received an additional 0.5 g starch at 10 PM. In other experiments (not shown), the timing of the feeding made no difference, at least as measured by excess urinary urea and ammonia after 300 mg casein.

Intravenous injections. For intravenous bolus injection, each mouse was placed in a 50-ml conical plastic restrainer. The protruding tail was immersed in water at 45°C for 1 min or more, until the lateral veins were clearly visible. The solutions were injected into one of these veins at a rate of 100 µl/s, with a disposable 1-ml insulin syringe through a 28-gauge 13-mm needle (Micro-fine-IV, Becton-Dickinson, Franklin Lakes, NJ).

Collection of tissues. Each mouse was stunned with a blow to the head and killed by decapitation with large scissors. In the protein degradation protocol, the liver and abdominal muscle were obtained immediately. In the other protocols the decapitated mouse was kept in ice until the dissection. The tissues obtained are described in the legends to Figs. 1 and 2. In these figures, "carcass" includes everything left after the skin, brain, gonads, and thoracic and abdominal viscera are removed; it includes all the bones and skeletal muscles, parotid and thyroid glands, eyes, and the upper respiratory and digestive tract included in the head.


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Fig. 1.   Tissue protein content during protein depletion and replenishment. Total protein was determined in all tissues gravimetrically as explained in EXPERIMENTAL PROCEDURE. All points represent averages ± SE. A and C: mice were kept on a 4% casein diet for 5 days; they were shifted to a protein-free diet (0% casein) for 3 days, and again to a 50% casein diet for 3 additional days, as indicated in the abcissa. Each point represents a group of 10 animals; results are plotted in a different scale for eviscerated carcass, skin, all viscera, and the sum of brain, lungs, heart, gonads, and spleen (A) and the kidneys, liver, and the sum of stomach, intestines, pancreas, and salivary glands (C). Except where indicated (ns, nonsignificant), all values after 3 days of depletion were significantly different from initial value (2-tailed t-test, P < 0.01). B: groups of 20 mice each were killed at time of shift from 4% casein to protein-free diet or 2, 3, 5, and 8 days afterward. Points represent total protein in liver and entire abdominal wall muscles (cut along the ribs, lumbar, and inguinal insertions). D: mice were fed by intragastric intubation starch, or starch plus indicated amounts of casein, as explained in EXPERIMENTAL PROCEDURE. Values show difference in total liver protein between each dose and the group receiving starch only, in 4 groups representing (from left to right) 21, 9, 9, and 20 mice, 16 h from the beginning of casein feeding.


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Fig. 2.   Incorporation of massive doses of [3H]leucine in tissues of mice under different conditions. Protein synthesis was measured by extrapolation, as described in EXPERIMENTAL PROCEDURE. A: groups of 10 mice each received 25, 50, or 100 µmol of leucine of constant specific radioactivity. Resulting incorporation in 4.75 min was measured in abdominal muscle (M), liver (L), pancreas (P), skin (SK), kidney (K), small intestine (SI), large intestine (LI), and stomach (ST) after feeding of starch (closed symbols) or starch and 300 mg of casein (open symbols). Curves were plotted in a reciprocal plot, and the reciprocal of saturation (1/extrapolated maximum, defined as 1 in the ordinate) was determined for each dose. B: incorporation with 50 µmol of leucine, in this and two other identical experiments, was pooled, multiplied by the reciprocal of the saturation calculated in A, and corrected for specific radioactivity of injected leucine to calculate total tissue leucine incorporation (right ordinate) and total protein synthesis (left ordinate), with the assumption that leucine is 10.5% of average tissue protein (24). Values represent 3 groups of mice receiving starch, starch and 300 mg casein, or water only. Values (average mg/day ± SE of no. mice in parentheses) represented, for 3 conditions, follow. Carcass, 128.4 ± 5.7 (9); 134.3 ± 5.6 (8); 134.6 ± 7.9 (8). Skeletal muscle, 59.7 ± 2.7 (17); 72.4 ± 3.7 (18); 75.6 ± 4.2 (17). Skin, 42.3 ± 3.0 (25); 58.2 ± 3.0 (26). Liver, 101.7 ± 3.3 (18); 97.8 ± 3.1 (25); 137.9 ± 3.3 (26). Gastrointestinal tract, 55.3 ± 2.4 (10); 57.8 ± 2.0 (19); 79.5 ± 2.2 (17). Pancreas, 43.7 ± 3.2 (19); 46.1 ± 1.9 (26); 78.0 ± 2.3 (24). Kidney, 18.8 ± 0.8 (19); 21.2 ± 0.6 (27); 26.5 ± 0.8 (26). Other viscera (submaxillary gland + spleen + heart + lung), 21.5 ± 0.7 (8); 21.4 ± 0.7 (9); 24.3 ± 1.2 (9). Combined values for "gastrointestinal tract" include separate measurements in stomach and small and large intestine, all of which showed significant increases after casein. Individual values for these separate tissues after starch or starch and casein were, respectively, (in mg/day), 8.7 ± 0.3 and 14.4 ± 0.6; 42.5 ± 1.9 and 54.4 ± 2.1; and 6.7 ± 0.2 and 10.5 ± 0.3. * Significant increase [2-tailed t-test (P < 0.01)] by addition of casein to starch. Postabsorptive values were significantly lower than starch-fed ones only for muscle, kidney, and heart (values for latter, included under "other viscera," were 1.8 ± 0.1, 2.4 ± 0.2, and 2.8 ± 0.2 mg/day).

Determination of tissue protein. In preliminary experiments, protein was determined in liver homogenates colorimetrically by the Lowry procedure, or gravimetrically by precipitation of the protein with hot 10% trichloroacetic acid and successive extractions of the precipitate with ethanol-ether-chloroform (2:2:1), acetone, and ether. In some experiments the extracted protein from all tissues was also dissolved at 90°C for 2 or more days in 10% N-lauroylsarcosine sodium salt in 1 M bicarbonate and measured by the Lowry procedure, with a correction factor for alkaline hydrolysis determined separately with bovine serum albumin or ovoalbumin standards. All of these procedures gave essentially the same results. The results presented here are from the gravimetric measurement. In the skin, the hair was not dissolved by the alkaline detergent solution; it was rinsed with water, dried, and its weight subtracted from the total. The carcass was ground in a mortar with dry ice, and a portion of it was used in the determination. The stomach and small and large intestines were opened up lengthwise and rinsed free of content. For liver and kidneys we used an aliquot of homogenates. The other tissues were used whole.

Protein radioactivity. For the determination of protein radioactivity, the protein from the tissues was extracted as described in the previous paragraph, except for three additional extractions with cold 10% trichloroacetic acid (to remove the radioactive precursor), and dissolved in the alkaline detergent at a concentration of up to 50 mg/ml; 0.2-ml aliquots were mixed with 0.8 ml 0.125 M HCl and 10 ml of scintillation fluid for counting.

Protein synthesis in all tissues: extrapolated incorporation of a massive amount of [3H]leucine. Protein synthesis was determined by the incorporation of radioactivity into protein after the intravenous injection of massive amounts of leucine, as first described for liver (24). The conceptual background and support for this approach, from experiments in isolated or cultured cells, are discussed elsewhere (27). Mice were injected with L-[4,5-3H(N)]leucine (25, 50, or 100 µmol per mouse, 0.2 µCi/µmol). Incorporation into protein was interrupted 5 min later by the intraperitoneal injection of cycloheximide (3 mg) and puromycin (1 mg) (3). The mice were killed 1 min after the second injection, and tissues were obtained as described above. For each condition, two controls were included, in which cycloheximide and puromycin were injected 15 s after [3H]leucine. Incorporation above controls was considered to have occurred in 4.75 min, the interval between the injection of inhibitors in the control and experimental animals. Protein synthesis was calculated from this value, the specific radioactivity of the injected leucine, the extent to which it saturates the muscle precursor pool (determined by extrapolation of reciprocal plots), and the average proportion of leucine in protein (10.5%). In this short interval, incorporation into protein is linear with time, and massive amounts of leucine have no effect on protein synthesis as measured by the incorporation of lysine (18, 24). We confirmed these observations for liver, muscle, intestine, kidney, and pancreas in two additional experiments. In one of them, three groups of 10 mice each received the protein synthesis inhibitors 15 s, 3 min, and 6 min after the injection of leucine; incorporation was linear in this interval in all tissues (not shown). In another experiment with two groups of 9 mice each, 5 min after the injection of [3H]lysine (50 µmol, 0.1 µCi/µmol), when 50 µmol nonradioactive leucine was included in the injection, incorporation was (relative to the control) for liver, 102 ± 4%; muscle, 108 ± 17%; intestine, 109 ± 5%; kidney, 101 ± 5%; and pancreas, 99 ± 4%.

Protein degradation in liver and muscle: accumulation of bestatin-induced peptides. The degradation of arginine-labeled long-lived proteins was studied separately in liver (4) 1 day after the intraperitoneal injection of 10 µCi of L-[1-14C]ornithine, and in abdominal wall muscles (5) 3 days after the subcutaneous injection of 20 µCi of L-[guanidino-14C]arginine. Accumulation of peptide intermediates was elicited by the intravenous injection of bestatin (1 mg, 15 min in liver; 5 mg, 10 min in muscle). The tissues were removed quickly and extracted with cold 10% trichloroacetic acid; the peptides were recovered from the extracts after the elimination of free amino acids by acid-ninhydrin treatment, ethyl-ether extraction, and fixation-elution in a cationic exchange resin (4). In muscle, the purification was continued further by acid hydrolysis of the peptides and enzymatic conversion of the guanidino-14C in the resulting free arginine to 14CO2 by arginase and urease (5). The radioactivity was corrected for the recovery of peptides in the purification procedure (5).

Estimation of balance in muscle and liver. The accumulation of bestatin-induced intermediates is proportional to total protein degradation but does not measure it directly. To calculate actual protein degradation we must calibrate the measured rates by balance whenever sustained conditions permit it (see Ref. 5).

For muscle, in nongrowing mice on the 4% casein diet, where balance is zero and there are no daily fluctuations in turnover, the actual rate of degradation was considered to equal the rate of synthesis; the rate of accumulation of bestatin-induced peptides was found to be 67% of this rate of turnover (5). Using this calibration factor for the 3-day protein-depleted mice, we estimated a protein loss of 3.3% per day (5), which agrees closely with the actual loss of 3.0% per day shown in Fig. 1B. This gave us a confirmed calibration for muscle and considerable confidence in the estimated balances.

For liver, the margin of error was larger. First, the rate of protein turnover in liver was an order of magnitude faster than in muscle, and the balance was calculated as a small difference between two large numbers. Second, the calculations were more indirect, because we took into account that only one-half of the proteins synthesized remained in the cells as stable components (6, 25). We calibrated the degradation in the state of protein depletion, where the rate of protein loss was 4.8% per day (Fig. 1B) or 9.0 mg per day, and the rate of synthesis was 99.7 mg per day, of which one-half, or 49.9 mg per day, was retained. We took then the rate of degradation in this condition to be 58.9 mg per day (9.0 + 49.9); the rate of accumulation of bestatin-induced peptides was 72% of this calculated value.

Scintillation counting. Samples were mixed with 10 ml of scintillation fluid (Liquiscint, National Diagnostics, Atlanta, GA) and counted in an LKB 1209 Rackbeta Scintillation Counter. In each run, internal standards were added to blanks under the same conditions to determine counting efficiency.

Determination of nitrogen, urea, and ammonia in urine or feces. For the collection of urine and feces, each mouse was placed in a square stainless steel metabolic cage (13 cm each side) in a room heated at 28°C. The protein-sparing effect of a starch meal was assessed by the decrease in urea excretion in eight mice fed intragastrically 1 g starch and 2 ml of water in two portions, with a 1-h interval, after 8 h of food deprivation. Urine was collected for 4 h up to the last portion of starch and for another 4 h afterward. To ensure an abundant urinary flow, the mice also received 1 ml of water intragastrically 1 h before the beginning of the first period. At the beginning of the first collection and at the end of each of the periods, the bladder was pressed gently to stimulate urination, the lower belly and genital regions were washed with distilled water, and the whole surface of the cage was rinsed by spraying it with a water mist. The urine and rinses were collected into a vial containing 1 drop of 1 M acetic acid brought to 20 ml with water, the urea in the solution was converted to ammonia with urease, and the combined urea and ammonia nitrogen (N) was determined colorimetrically (kit no. 640, Sigma Chemical). In a separate experiment, for the calculation of the obligatory N loss, eight mice were fed 0.5 g of starch and 1 ml of water at 8 AM, 12 AM, and 6 PM. Urine was collected for 16 h beginning at 8 AM, for the determination of urea and ammonia and total urinary N. In another experiment, eight mice were kept in the metabolic cages with the nonprotein diet and water ad libitum for 24 h, and the feces were collected for N determination. Total urinary and fecal N were determined for us by the Kjeldahl procedure by Galbraith Laboratories (Knoxville, TN).

    RESULTS
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Abstract
Introduction
Results
Discussion
References

Protein depletion and repletion. In fully grown animals, the supply of dietary amino acids is necessary for the maintenance of tissue mass and nitrogen balance, but excess supply does not lead by itself to protein gain. To study protein deposition after a meal, we must first cause a depletion, either by fasting (where depletion is faster, as amino acids are used for gluconeogenesis) or by selective protein deprivation (where depletion slows down to a minimum "obligatory loss") (8). Because the focus of this study is the maintenance of amino acid pools (rather than energy supply), we chose the latter. To facilitate the transition to the protein depletion condition, mice were first adapted for 5 days to a protein-restricted diet (4% casein, supplemented with 0.3% methionine), which in these nongrowing mice is sufficient to maintain balance and prevent weight loss for >= 2 wk. We then shifted these low protein-adapted mice to a nonprotein diet for 3 days, an interval sufficient to reduce urea excretion to its minimum (not shown).

The conditions chosen for this study are represented in Fig. 1, A and C, which summarizes the changes in tissue protein content in response to protein depletion and its subsequent repletion after the feeding of a 50%-casein diet. With minor differences, this resembles the classic experiment of Addis et al. (1) in rats. Liver and the gastrointestinal tract suffered the largest percentage of loss and the fastest subsequent gain. The protein loss in skin (without the hair) represented a smaller fraction of its total protein (7%), but the total loss (115 ± 64 mg, P < 0.05) was almost as large as that of all visceral organs combined (121 ± 16 mg, P < 0.01).

Much of the protein loss in renewable tissues may represent the failure to replenish shed cells (epidermis, intestinal epithelium) or depleted secretions (pancreas). In this study, we are primarily interested in the role of cellular protein turnover in the maintenance of amino acid pools, which occurs largely in muscle and liver. Whereas the hepatic changes are the most dramatic, muscle protein turnover is much slower than in liver, and the protein lost in 3 days by the total eviscerated carcass was not statistically significant. In a separate experiment, using larger groups of mice for greater accuracy, we took a closer look at the loss of both skeletal and hepatic protein over a longer period of protein depletion. The results are shown in Fig. 1B. This experiment showed that, between 2 and 8 days on a protein-free diet, the skeletal muscles of the abdominal wall lost exponentially 3% of their protein per day, and the liver 4.8% per day. Neither of these curves extrapolated to the 100% value at time 0. It appears that muscle protein starts to decrease after a 1-day lag, although the differences are too small to be certain. The liver, on the other hand, initially lost a larger portion of its protein (>20% in the first 2 days). A rapid initial loss in liver and intestine has been widely documented before in rats (1, 15), and it has been suggested that the abdominal viscera are the source of most of the "labile body protein," i.e., the additional N lost initially, before the minimum obligatory N loss is attained (28).

Contributions of muscle and liver protein to the obligatory N loss. Because of its large mass, changes in muscle protein are of great physiological interest. The measurements, summarized in Table 1, demonstrate that within 2 days muscle becomes the most important source of N in protein-deprived animals. In Table 1, the values for urinary and fecal N are direct determinations. The N lost by the skin is a maximum estimate based on the rates of skin protein synthesis and the assumption that most of this synthesis is used in the renewal of epidermal cells and hair lost to the outside. We estimate that the loss of 3% per day of muscle protein contributes 51% of the obligatory N loss, or 72% of the N lost in the urine and feces. The contribution by the liver is an order of magnitude lower.

                              
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Table 1.   Contribution of liver and skeletal muscle to nitrogen loss in protein-depleted mice

Replenishment of tissue proteins after intragastric administration of starch and casein. We next studied the replenishment of tissue proteins in mice depleted for 3 days, after the intragastric administration of a water suspension of starch and casein (supplemented with methionine, 5 mg per 95 mg of casein). The animals received 0.5 g of starch, followed 2 h later by another 0.5 g of starch with or without casein, in four portions at 1-h intervals.

Because its changes are the most pronounced, hepatic protein content was a useful indication of the successful repletion of some of the lost protein. In the experiment shown in Fig. 1D, five groups of mice received 0, 40, 110, 300, or 450 mg of casein, and total liver protein was determined 16 h after the beginning of the casein feeding. Compared with the control, a statistically significant increase was observed even with the smallest dose. With 300 mg of casein (see arrow in Fig. 1D), the difference from the control, 35 mg, represented ~10% of the dietary amino acids. This difference is comparable to the replenishment of 38 mg 16 h after refeeding a high-protein diet ad libitum (Fig. 1C). In the remaining experiments presented here, we measured the acute effects on tissue protein synthesis or degradation 1 h after the last portion of a meal containing either starch alone or starch with 300 mg of casein.

Effect of starch and casein on protein synthesis in all major tissues. We measured protein synthesis by extrapolation, as explained in EXPERIMENTAL PROCEDURE. Figure 2A shows the reciprocal plots for all major tissues after a meal containing starch alone (closed symbols) or starch and casein (open symbols). In these protein-depleted mice, the influx of nonradioactive leucine after the meal containing casein made a difference in the ability of the massive dose of radioactive precursor to saturate the leucine pools; this difference was taken into account in the calculations (see Fig. 2 legend). Figure 2B summarizes the calculated rates of protein synthesis after either meal. We also included a third group of postabsorptive mice, which received neither starch nor casein but water alone for 6 h (labeled as "none").

All major tissues are represented in Fig. 2. The eviscerated, skinless carcass (first set of columns) includes all skeletal muscle, calculated separately in the second set. Although muscle comprises ~90% of the carcass protein, because of its low fractional rate of synthesis it represents only one-half of the carcass protein synthesis. The rest must represent less abundant tissues with much higher fractional rates of synthesis, such as the lining of the mouth and the upper respiratory tract, the bone marrow, and the thyroid and parotid glands. These tissues were not removed, but the effects of starch and casein can be explained by those on muscle alone. The whole carcass is included to provide a complete catalog of rates of protein synthesis in all tissues. Total body protein synthesis, i.e., the sums of the values for carcass, skin, and viscera, were not shown in Fig. 2. They were 421 ± 7.6 mg per day after starch and 539 ± 9.7 mg per day after starch and casein. This increment, if sustained for 12 h, would by itself lead to deposition of 60 mg of tissue protein, or 20% of the protein in the meal (not counting inhibition of protein degradation, which will be discussed).

In these protein-depleted mice, the liver contains only 3% of total body protein but accounts for 25% of the total body protein synthesis. The increment in hepatic protein synthesis caused by inclusion of casein in the meal represents one-third of the total increment. We have shown before, in this and other conditions, that of the newly synthesized proteins, 30% is exported as plasma proteins, 20% turns over within 3 h, and 50% represents stable liver components (6, 25).

As we have seen in Fig. 1, other than liver, depletion and replenishment of tissue protein was most evident in the gastrointestinal tract and pancreas. This was also evident by the stimulation of protein synthesis in these tissues. The combined values for "gastrointestinal tract" include separate measurements in stomach and small and large intestine, all of which showed significant increases after casein (see legend to Fig. 2). We do not know how much of this synthesis was in the renewable lining of the gastrointestinal tract. The increment of synthesis in pancreas was as large as that of liver, and most likely it was directed toward its secretory function.

Casein feeding also stimulated protein synthesis in skin and kidneys. The one major exception to the effect of casein was muscle. Total muscle synthesis was calculated from total muscle protein (45% of body weight × 170 mg protein/g) and the average fractional rates of the combined abdominal wall and leg muscles, which were indistinguishable. Muscle protein synthesis showed the most vigorous response to the starch meal but very little additional effect of casein. The response of heart muscle was very similar to that of skeletal muscle (see legend to Fig. 2).

Protein turnover: contrasting effects of starch and casein in muscle and liver. In addition to protein synthesis, we also measured instant rates of protein breakdown in liver and muscle by the accumulation of bestatin-induced peptide intermediates during the degradation of arginine-labeled long-lived proteins (see EXPERIMENTAL PROCEDURE). Figure 3 compares the relative rates of synthesis and degradation in liver and muscle in postabsorptive animals (receiving no food for 6 h, labeled "none") or after a meal containing only starch (S) or starch and casein (S+C). The actual rates of protein synthesis are those in Fig. 2. The actual values for protein degradation are given in the legend to Fig. 3. The statistical significance of the differences is also noted in Fig. 3. To facilitate the discussion we present them in two ways. In Fig. 3A we compare both meals with the postabsorptive state (none = 100). In this comparison we observe that the complete meal (S+C) had a similar effect in both tissues: protein synthesis was 36% higher in liver and 26% higher in muscle. Protein breakdown was 15% lower in liver and 30% lower in muscle.


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Fig. 3.   Protein turnover in liver and muscle. A and C: values for liver and muscle protein synthesis and degradation are percentages of those in postabsorptive state (A) or of those after starch meal (B). Values for synthesis were the same as those shown in Fig. 2 legend. For purposes of balance, values for degradation were calculated from rate of accumulation of bestatin-induced peptides, after a correction for purification recovery (88% for liver and 47% for muscle) and calibration against synthesis (62% for liver and 67% for muscle), as explained EXPERIMENTAL PROCEDURE. Calibrated degradation values, average mg/day ± SE for nos. of mice in parentheses, follow. Muscle, 159.6 ± 17.3 (6); 121.1 ± 14.8 (6); 111.2 ± 14.8 (6). Liver, 59.8 (6), 76.6 ± 5.0 (7); 51.1 ± 1.0 (7). B: balances are calculated from these values for each condition as the difference between synthesis and degradation. Values were the same, except that for liver, only 50% of rates of synthesis were computed, because the other 50% represent rapidly turning over proteins and exported plasma proteins (6, 25). Data also include urinary elimination of urea and ammonia, as explained in text. All values are mg protein (catabolized to urea, gained or lost by tissues) per h. S, starch fed; S + C, starch + casein fed.

When we include in the comparison animals receiving only starch, however, a very different picture emerges. Starch had a protein-sparing effect, detectable directly by a decrease in urea excretion (see next section). This anticatabolic effect of carbohydrates was not the same, however, as protein deposition, which in these protein-depleted animals could only happen with a net influx of dietary amino acids. In muscle, but not in liver, most of the effects of the meal were attained by starch alone; compared with postabsorptive mice (Fig. 3A), starch produced a 21% increase in synthesis and a 24% decrease in degradation. Contrary to that in muscle, starch alone had a catabolic effect in liver. Compared with the postabsorptive animals, starch had no effect on protein synthesis but increased protein breakdown by 28%.

Whereas starch decreased N loss, replenishment of tissue proteins in the protein-depleted mice could only occur when casein was added to the starch. When the same data are replotted to compare the effects of dietary amino acids with carbohydrates alone (Fig. 3C, S = 100%), it is shown that casein stimulated hepatic protein synthesis by 41% and inhibited degradation by 33%. In muscle, however, the effects were marginal and statistically insignificant.

Effect of starch on urea excretion. The protein-sparing effects of starch feeding could be observed by direct measurement of urea excretion. In one experiment with seven postabsorptive mice, intragastric administration of 1 g of starch caused a 59% decrease in the urinary excretion of combined urea and ammonia N, from 2.51 ± 0.16 mg in the 4 h preceding the starch meal to 1.04 ± 0.01 mg in the 4 h that followed it (P < 0.001). When a conversion factor of 6.25 mg protein per milligram N is used, these values translate, respectively, to the catabolism of 3.9 and 1.6 mg of endogenous protein per hour (Fig. 3B).

Estimation of balance in muscle and liver. To better visualize some of the points discussed in DISCUSSION, we estimated the balance between protein synthesis and degradation in muscle and liver, as described under EXPERIMENTAL PROCEDURE (Fig. 3B). In the postabsorptive mice, fasted from 7 AM to 1 PM, muscle lost 4.1 ± 0.7 mg protein/h. The starch meal reduced this loss significantly to 2.1 ± 0.6 mg/h. As we have seen, casein in the meal had little additional effect; despite the dietary influx of amino acids, muscle still lost 1.5 ± 0.6 mg/h. In liver, the starch meal had a catabolic effect on both synthesis and degradation (Fig. 3, A and B), and this resulted in an estimated increase in the protein loss to 0.7 ± 0.2 mg/h. Addition of casein to the meal had a strong anabolic effect in both synthesis and degradation and resulted in an estimated gain of 1.0 ± 0.1 mg/h.

    DISCUSSION
Top
Abstract
Introduction
Results
Discussion
References

Research in protein turnover in live animals has been limited in the past by difficulties in the measurement of tissue protein synthesis or degradation. The information available up to now has been fragmentary and incomplete. The procedures developed in our laboratory, utilized here, have enabled us to provide, in a single study, a comprehensive survey of the effects of a meal on instant rates of protein synthesis in all the major tissues, as well as degradation in the two major turning over tissues, muscle and liver.

Protein depletion and replenishment. In designing this study, we chose to deplete tissue proteins by selective deprivation of dietary amino acids for 3 days, the shortest interval necessary to reduce N loss to its minimum. In the experiments shown in Figs. 2 and 3, we assessed separately the effects of carbohydrates and protein in the food. It should be noted that the addition of 300 mg of casein to 1 g of starch increased by 30% the caloric content of the intragastric meal. Nevertheless, these mice were depleted of N, not energy; furthermore, 1 g of starch between 8 AM and 1 PM represented as much carbohydrate as these animals would normally eat during the daytime. For these reasons, in the following discussion we assume that the effects of intragastric casein on protein turnover are due to the dietary supply of amino acids, not excess energy. We also used fully grown mice to distinguish the replenishment of lost tissue protein from the resumption of growth. From the results presented in this paper, we can draw the following conclusions.

1) Muscle protein was the major source of N in the protein-depleted state. Although the importance of muscle in long-term protein deprivation (Table 1) has been well recognized before (2, 8, 28), the time required for the establishment of net muscle loss has been inconsistent. In one experiment, with very young rats fed a very low protein diet (7), liver lost 40% of its protein in 1 wk but muscle did not lose any until after 2 wk. In another experiment with young rats on a protein-free diet, there was no change in muscle protein content until after 3 days (13). In our study, the loss of muscle protein was well established in the 2nd day of depletion (Fig. 1B). This earlier response may have been due to our use of fully grown animals (reversal of growth may take longer). It is also possible that the adaptation to a 4% casein diet preceding depletion, although sufficient for balance, was restricted enough to prime the effect and shorten the lag.

2) Muscle was also the major additional source of amino acids for gluconeogenesis. Compared with the effect in postabsorptive mice, starch produced a profound redistribution in the N economy. It suppressed muscle protein loss by one-half, from 4.1 to 2.1 mg/h (a decreased loss of 2.0 mg/h) (Fig. 3B). There was also a significant increase in synthesis in kidney and heart (see Fig. 2 legend). If, as we have shown, muscle was the major source of N in these protein-depleted mice (Table 1), what are the consequences of the suppressed muscle catabolism in the total body N balance? Some of the difference was contributed by the liver, the only tissue that showed a catabolic response to starch, resulting in an estimated increase in hepatic protein loss of 0.7 mg/h (Fig. 3B). The largest effect of starch, however, was in suppressing amino acid catabolism. As shown in Fig. 3B, there was a drop in the urinary elimination of urea and ammonia equivalent to 2.3 mg protein/h. The additional excretion of urinary urea and ammonia in the postabsorptive state must have represented increased gluconeogenesis from amino acids. Because the excess supply of amino acids in this condition was largely from muscle, it follows that muscle protein was not only the major source of N in the protein-depleted state but also the major additional source of amino acids for gluconeogenesis.

3) Muscle failed to replenish its protein after a single meal. Whereas starch suppressed muscle catabolism, it could not, in these depleted mice, lead to protein deposition without an exogenous supply of amino acids. Addition of casein to the meal had little effect on muscle protein synthesis or degradation (Fig. 3). This does not mean that muscle protein cannot be replenished (it must at some point), but that this is a sluggish process, which is not immediately regulated by the dietary influx of amino acids.

4) Liver protein turnover responded primarily to the supply of amino acids. When casein was added to the meal, the liver exhibited a strong anabolic response (Fig. 3). The unique hepatic responsiveness to the supply of amino acids was most clearly shown after starch alone, when net muscle proteolysis was drastically decreased. The liver exhibited a strong catabolic response (Fig. 3) despite the anabolic hormonal signals (increased insulin and decreased glucagon) that must have resulted from the abundant influx of dietary glucose.

5) The gastrointestinal tract, pancreas, and skin were as quick to lose and regain protein as the liver. These tissues (Fig. 1) showed a vigorous stimulation of protein synthesis by the addition of casein to the meal (Fig. 2). As a fraction of its protein, skin losses were smaller, but much of the skin protein is metabolically inert connective tissue. After 3 days of depletion, the skin lost as much protein as all viscera combined (Fig. 1). We do not know what proportion of the changes represents loss and replenishment of secretions (pancreas) or renewable cells (epidermis, hair follicles, gastrointestinal epithelium) rather than turnover of cellular components. The difference is important, because it is the latter that contributes most directly to the regulation of amino acid pools.

Mechanistic implications. The mechanistic questions posed by this survey are very different for muscle and liver. The abundant influx of glucose (the digestion product of starch) must have produced a strong stimulation of insulin secretion and inhibition of the secretion of glucagon. The suppression by starch of muscle proteolysis and amino acid catabolism, shown here, was consistent with these changes in the pancreatic hormones. Insulin has been shown to stimulate the synthesis of muscle proteins or inhibit their degradation (9, 16, 19, 20). Increased insulin and decreased glucagon also directly inhibit the hepatic gluconeogenic pathway (23).

The responsiveness of the liver to the supply of amino acids is shown not only by its anabolic response to dietary amino acids but also by its unique catabolic response to starch feeding (and the resultant suppression of amino acid supply from muscle). Experiments in perfused livers have clearly established the anabolic effects of amino acids and insulin and the catabolic effects of glucagon (21, 22, 29, 30). The catabolic response to starch feeding in the present study suggests that, under these conditions, the decreased supply of amino acids overshadowed the anabolic changes in the secretion of pancreatic hormones.

The vigorous responses to dietary protein in intestine and liver could be due to the exposure of these tissues to the highest concentrations of amino acids (in intestine during absorption, in liver in the portal circulation). That would not explain, however, the even larger responses of protein synthesis to casein feeding in large intestine and pancreas, which are not exposed to the absorbed amino acids except through the systemic circulation.

The explanation for the lack of acute effects of dietary amino acids in muscle is less obvious. Amino acids, particularly leucine, have been found to stimulate protein synthesis and/or inhibit protein degradation in the muscle of young rats (11), as well as in isolated diaphragm (10) and perfused legs (14). The effect is not consistent (17) and may depend on the metabolic conditions of the tissue. Contrary to effects on liver, the effects on muscle may be indirect, increasing the sensitivity of the tissue to insulin (12). In our experiments, starch (through insulin) may already have elicited the maximum possible anabolic response, and that is why amino acids had no additional effects. Also, muscle may have failed to achieve net protein deposition, because the depleted tissue lost some of its protein-synthesizing machinery and, hence, its capacity for protein synthesis. Muscle can lose its RNA rapidly during protein depletion, at least in young rats (13). The fractional rate of synthesis after starch and casein in our mice (3.0% per day) was 30% lower than before the depletion (4.4% per day) (5).

The slow recovery of muscle mass may be a necessary consequence of its specialized nature. Its highly organized structure may not favor the random addition of cytoplasmic components; rather, muscle growth involves remodeling and increased protein turnover (28). A coordinated regrowth may be elicited by slower effects of an adequate supply of nutrients, for instance through delayed transcriptional effects of insulin, or changes in insulin-like growth factor I levels (9). A delayed response of muscle to a protein-containing meal may be of physiological advantage to the animal, if it facilitates the preferential utilization of a limited influx of dietary amino acids to replenish the visceral organs.

Conclusions. The present study in protein-depleted mice shows contrasting roles of liver and muscle in the maintenance of amino acid pools. Protein turnover proves uniquely sensitive to the dietary supply of carbohydrates in muscle and to the endogenous or exogenous supply of amino acids in liver. The results illustrate the importance of liver protein turnover in the minute-by-minute regulation of amino acid pools and the predominant role of muscle in the long-term supply of amino acids during protein and energy deprivation. They indicate that muscle protein loss can be acutely suppressed by food intake but that, once depletion is established, muscle replenishment is a delayed process.

    ACKNOWLEDGEMENTS

We are grateful to Nippon Kayaku (Tokyo, Japan) for their generous gift of bestatin.

    FOOTNOTES

This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant 5RO1 DK-43833.

Address for reprint requests: O. Scornik, Dept. of Biochemistry, Dartmouth Medical School, 7200 Vail Bldg., Rm. 402, Hanover, NH 03755-3844.

Received 13 February 1997; accepted in final form 15 August 1997.

    REFERENCES
Top
Abstract
Introduction
Results
Discussion
References

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