Growth hormone decreases muscle glutamine production and
stimulates protein synthesis in hypercatabolic patients
Gianni
Biolo1,
Fulvio
Iscra2,
Alessandra
Bosutti1,
Gabriele
Toigo1,
Beniamino
Ciocchi1,
Onelio
Geatti3,
Antonino
Gullo2, and
Gianfranco
Guarnieri1
1 Istituto di Clinica Medica, 2 Istituto di
Anestesia, Rianimazione e Terapia Antalgica, and 3 Servizio
di Medicina Nucleare, University of Trieste, Trieste 34149, Italy
 |
ABSTRACT |
We determined
the effects of 24-h recombinant human growth hormone (rhGH) infusion
into a femoral artery on leg muscle protein kinetics, amino acid
transport, and glutamine metabolism in eight adult hypercatabolic
trauma patients. Metabolic pathways were assessed by leg arteriovenous
catheterization and muscle biopsies with the use of stable amino acid
isotopes. Muscle mRNA levels of selected enzymes were determined by
competitive PCR. rhGH infusion significantly accelerated the inward
transport rates of phenylalanine and leucine and protein synthesis,
whereas the muscle protein degradation rate and cathepsin B and UbB
polyubiquitin mRNA levels were not significantly modified by rhGH. rhGH
infusion decreased the rate of glutamine de novo synthesis and
glutamine precursor availability, total branched-chain amino acid
catabolism, and nonprotein glutamate utilization. Thus net glutamine
release from muscle into circulation significantly decreased after rhGH
administration (~50%), whereas glutamine synthetase mRNA levels
increased after rhGH infusion, possibly to compensate for reduced
glutamine precursor availability. We conclude that, after trauma, the
anticatabolic action of rhGH is associated with a potentially harmful
decrease in muscle glutamine production.
stable isotopes; competitive polymerase chain reaction; amino acid
transport; glutamine synthetase; trauma patients
 |
INTRODUCTION |
PATIENTS SUFFERING FROM
MAJOR INJURY have a rapid and progressive loss of skeletal muscle
protein that can be reduced only partly by nutritional support
(5, 14, 30). The administration of recombinant human growth hormone (rhGH) as adjunctive therapy has
been shown to be effective in slowing the loss of muscle mass in
patients (8, 16, 17,
28, 37, 38), despite the fact
that it may have some secondary harmful effects (31,
37). Changes in protein mass derive from a balance between
protein synthesis and degradation, which in turn can be influenced by the rate of transmembrane amino acid transport. Evidence indicates that
stimulation of protein synthesis is a primary mechanism of rhGH action
on muscle (12, 13). However, the
hormone effects on protein degradation and amino acid transport have
not been clarified.
Besides an anticatabolic effect, rhGH action on muscle also involves a
suppression of glutamine efflux, which is much greater than that
expected on the basis of the hormone's effects on protein metabolism
(4, 28). Such a decrease in glutamine efflux
can result from changes in glutamine de novo synthesis and/or outward transmembrane transport. Glutamine is the most abundant free amino acid
in the body. It is a precursor of many compounds (e.g., glucose, glutathione, and nucleic acids) and a major fuel for rapidly dividing cells (intestinal mucosa, immune system, and wound tissue). Glutamine is synthesized primarily in skeletal muscle and released into the
bloodstream to serve as an interorgan vehicle for carbon and nitrogen.
The key enzyme for glutamine synthesis is glutamine synthetase, whereas
precursor substrates are glutamate,
-ketoglutarate, free ammonia,
and amino-nitrogen derived from the catabolism of the
branched-chain amino acids (9). In skeletal muscle, there is a large intracellular pool of preformed free glutamine that serves
as a reservoir for any increased glutamine requirement in extramuscular
tissues. A decline in muscle free glutamine has consistently been
observed in trauma patients (5).
The aim of this study was to define the mechanisms of the rhGH-mediated
changes in muscle glutamine and protein kinetics in severely
traumatized patients during combined enteral and parenteral nutrition.
Leg arteriovenous catheterization, muscle biopsy, and stable isotopic
tracer of amino acids were used to determine the rates of muscle
protein synthesis, proteolysis, glutamine de novo synthesis, nonprotein
utilization of glutamate (mainly to form glutamine) and the
branched-chain amino acids (catabolism), as well as transmembrane
transport of glutamine and selected essential amino acids
(2-4, 6). Competitive RT-PCR was used to
determine muscle mRNA levels of key enzymes for glutamine synthesis
(glutamine synthetase) and for myofibrillar (ubiquitin) and
nonmyofibrillar (cathepsin B) protein degradation (7,
15, 27, 34, 35).
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METHODS |
Patients
Eight adult patients (6 males, 2 females; age 39 ± 5 yr,
weight 74 ± 4 kg, height 172 ± 3 cm) with multiple injuries
(APACHE II score 15 ± 1) were studied between days 7 and 12 after admission to the Intensive Care Unit of
the University Hospital of Cattinara, Trieste, Italy. Leg volume
(8,603 ± 520 ml) was estimated by use of an anthropometric
approach (21). Informed consent was obtained from the
patients' close relatives. The protocol was approved by the competent
hospital authority. All patients received continuous combined
intravenous (80% of total energy as amino acids, glucose, and lipids)
and enteral [20% of total energy: Nutrisond (Nutricia, Zoetermeer,
Netherlands) or Sondalis ISO (Clintec, Rome, Italy)] nutrition
providing 35 kcal · kg
1 · day
1 with 250 mg of nitrogen · kg
1 · day
1. Nutrient administration
was constant for
2 days before and during the study.
Experimental Protocol
Each patient was studied twice. Two leg muscle metabolic studies
(which included determination of protein and glutamine metabolism by
stable isotopes and competitive RT-PCR) were performed 24 h apart
during rhGH administration or saline infusion (control study). To
account for time-related changes of muscle metabolism after trauma and
for potential interference between two close stable isotope infusions,
the control study was performed either before (protocol 1)
or after (protocol 2) rhGH infusion. Patients were randomly
assigned to protocol 1 (n = 4, 3 males, 1 female, age 42 ± 6 yr, weight 74 ± 6 kg, height 171 ± 5 cm; APACHE II score 15 ± 2) or to protocol 2 (n = 4, 3 males, 1 female, age 36 ± 5 yr, weight
75 ± 6 kg, height 174 ± 4 cm; APACHE II score 15 ± 2). Seven to twelve days after trauma, patients were in relatively stable clinical and metabolic conditions. A continuous infusion of
either saline (protocol 1) or rhGH (0.10 µg · kg
1 · min
1; Genotropin, Pharmacia
and Upjohn Stockholm, Sweden; protocol 2) was started at 1 PM and continued for 24 h. rhGH was infused directly into the
femoral artery to reduce systemic hormone effects. The next day, a leg
muscle metabolic study was performed from 8 AM to 1 PM. Thereafter, in
the four patients previously infused with saline, rhGH infusion was
started into the femoral artery and was continued for 24 h. In the
other four patients previously treated with rhGH, a 24-h saline
infusion was commenced and then the second metabolic study was
performed during the last 5 h of either rhGH or saline administration.
Leg Metabolic Study
Indwelling catheters were placed in a central vein (subclavian
or internal jugular) for isotope infusion and in the femoral artery and
vein of one leg for blood sampling. The femoral artery catheter was
also used for continuous infusion of rhGH and for primed continuous
infusion of indocyanine green (Infracyanine, SERB, Paris, France) to
measure leg blood flow (2-4, 6,
22). Indocyanine green recycling was assessed by measuring
dye concentration in the left antecubital vein. Most of the catheters
were placed for clinical purposes. At hour 19 (8 AM) femoral
venous blood samples were obtained to measure background amino acid
enrichments and indocyanine green concentration. Then a primed
continuous infusion of
L-[5-15N]glutamine (MassTrace, Woburn, MA)
was started, followed at hour 21 by
L-[ring-2H5]phenylalanine
(MassTrace) and L-[1-13C]leucine (MassTrace).
Tracer infusions were maintained constant throughout the experiment.
The following tracer infusion rates(IR) and priming doses (PD)
were used:
L-[ring-2H5]phenylalanine:
IR = 0.05 µmol · kg
1 · min
1, PD = 2 µmol/kg;
L-[1-13C]leucine: IR = 0.08 µmol
· kg
1 · min
1, PD = 4.8 µmol/kg; L-[5-15N]glutamine: IR = 0.35 µmol · kg
1 · min
1, PD = 63 µmol/kg. Isotope infusions were not started simultaneously, because the equilibration period of each tracer varied
(2). L-[5-15N]glutamine required
5 h to reach steady state in muscle (2). L-[ring-2H5]phenylalanine
and L-[1-13C]leucine were infused for 3 h.
To measure leg blood flow, at hour 23 a primed
continuous infusion of indocyanine green dye (IR = 0.5 mg/min;
PD = 5 mg) into the femoral artery was started and maintained for
30 min. During the last 15 min of indocyanine green infusion, four
blood samples were taken every 5 min from the femoral and left
antecubital vein for spectrophotometric determination of steady-state
dye serum concentrations. The rate of leg plasma flow was calculated at steady state from the ratio between the dye infusion rate and the
difference between serum dye concentrations in femoral and antecubital
veins. Leg blood flow was calculated from the hematocrit. During the
last 30 min of the intra-arterial rhGH or saline infusion, four blood
samples were taken every 10 min from the femoral artery and vein to
determine amino acid enrichments and concentrations. To allow sampling
from the femoral artery, the rhGH or the saline infusion was stopped
for <10 s and then quickly resumed. Blood samples were also taken from
the femoral and left antecubital veins to determine local and systemic
growth hormone concentrations, respectively. Insulin and insulin-like
growth factor I (IGF-I) concentrations were measured in blood samples
taken from the left antecubital vein. At hour 24, after the
last blood sample was taken and before stopping the tracer and the rhGH
or saline infusions, a muscle biopsy was taken to measure enrichments
and concentrations of free amino acids and mRNA levels of cathepsin B,
ubiquitin, and the glutamine synthetase enzyme. The biopsy was taken
from the lateral portion of the vastus lateralis muscle ~20 cm above the knee with a Bergström biopsy needle (Stille, Stockholm,
Sweden) (2-4, 6). Approximately 80-100
mg of muscle tissue were obtained with each biopsy. This procedure
yields a sample of mixed skeletal muscle. Blood, visible fat, and
connective tissue were quickly removed from the specimen, and the
tissue was immediately frozen in liquid nitrogen and stored at
80°C for later analysis. Thereafter, in the four
patients previously infused with saline, an rhGH infusion was started
into the femoral artery and continued for 24 h. In the other four
patients previously treated with rhGH, a 24-h saline infusion was
commenced. During the last 5 h of either rhGH or saline
administration, tracer infusion was performed, leg blood flow was
measured, and blood samples and muscle biopsies were taken, as described.
Amino Acid Enrichments and Concentrations
Blood samples taken from the femoral artery and vein to
determine amino acid enrichments and concentrations were collected in
preweighed tubes containing 1% saponin. Simultaneously, a known amount
of a homoserine solution was added to the tube as internal standard and
thoroughly mixed. The blood was then precipitated with 15%
sulfosalicylic acid (SSA). The tubes were weighed again, and the
difference was recorded as blood volume after subtraction of the
internal standard and SSA volumes. The supernatant was frozen for later
analysis. To determine tracer enrichments, the t-butyldimethylsilyl derivatives were prepared as described
(2).
Each tissue sample was weighed, and muscle protein was precipitated
with 15% SSA. A known amount of a homoserine solution was added as
internal standard and thoroughly mixed. The tissue was then homogenized
and centrifuged, and the supernatant was collected. This procedure was
repeated twice more. The pooled supernatant was frozen for later
analysis. Muscle and whole blood SSA extracts were processed to
determine free amino acid enrichments and concentrations by gas
chromatography-mass spectrometry (Incos XL, Finnigan, Bremen, Germany)
and HPLC (Beckman, Berkeley, CA), respectively. Mass spectrometry
analysis was performed by electron impact ionization and selected ion
monitoring for the t-butyldimethylsilyl derivatives of
leucine ([mass-to-charge ratios (m/e) 302 and
303], phenylalanine (m/e 234 and 239)
and glutamine (m/e 431 and 432). Data were
expressed as a tracer-to-tracee ratio. Concentrations (nmol/ml) of free
amino acids in blood and total muscle water were calculated as
referenced (2). Measured values of enrichment and
concentrations relative to total tissue water were corrected (2, 18) to obtain intracellular values. These
corrections required the knowledge of amino acid concentration and
enrichment in the interstitial fluid, as well as knowledge of the
proportion between intra- and extracellular water in muscle. We assumed
that amino acid enrichment and concentrations in the interstitial fluid equaled blood values in the femoral vein and that the ratio between intra- and extracellular water in muscle was 0.16, as previously determined in humans by the chloride method (2).
Calculation of Kinetic Parameters
At the whole body level, amino acid rates of appearance
(Ra) were calculated by dividing isotope infusion rates by
arterial enrichments. Because amino acid intakes were identical in the control studies and during rhGH infusions, changes in the
Ra of the essential amino acids phenylalanine and leucine
can be considered as markers of changes in whole body proteolysis.
Whole body glutamine clearance was calculated by dividing the glutamine
Ra by the arterial glutamine concentration.
The net leg balance for amino acids was calculated from the Fick
principle
|
(1)
|
where CA and CV are whole blood amino
acid concentrations in femoral artery and vein, respectively; BF is leg
blood flow. A positive value indicates net uptake, whereas a negative
value indicates net release. Skeletal muscle is considered to account largely for amino acid metabolism in the whole leg (2). In the steady-state condition of muscle free amino acid concentrations, amino acid uptake or release across the leg reflects the balance between intracellular production and disposal for that particular amino
acid. Thus net phenylalanine, tyrosine, and lysine release from leg
muscle are markers of net protein catabolism, because these amino acids
are not synthesized or oxidized in muscle tissue (1,
26, 36). Furthermore, phenylalanine is not
hydroxylated to tyrosine in muscle (1, 26,
36), whereas the kidney and the splanchnic bed together
account for all of the whole body hydroxylation of this amino acid
(32). In contrast, skeletal muscle is the main site of
catabolism of the branched-chain amino acids leucine, valine, and
isoleucine, and of synthesis of alanine from pyruvate and of glutamine
from glutamate. We assumed that amino acids are released from
proteolysis in proportion to their relative content in muscle protein
(2-4, 6, 24). Thus the net
rates of release from protein catabolism of glutamate, glutamine, alanine, and the branched-chain amino acids can be calculated from the
net rate of phenylalanine release, corrected for the molar ratios
glutamate/phenylalanine (1.87), glutamine/phenylalanine (2.05),
alanine/phenylalanine (2.35), leucine/phenylalanine (3.10), isoleucine/phenylalanine (2.55), and valine/phenylalanine (1.77) determined in mixed human muscle protein (2,
24). Then, the rates of net alanine and glutamine
synthesis (i.e., the differences between the rates of synthesis and
nonprotein utilization of the amino acids) of net nonprotein disposal
of glutamate (mainly to form glutamine) and of branched-chain amino
acid catabolism can be calculated by subtracting from the total release
or uptake of these amino acids the component accounted for by protein
catabolism
|
(2)
|
|
(3)
|
|
(4)
|
|
(5)
|
|
(6)
|
|
(7)
|
|
(8)
|
This calculated rate of branched-chain amino acid catabolism
(i.e., nonprotein branched-chain amino acid disposal) is not equivalent
to the actual rate of branched-chain amino acid oxidation; in fact,
branched-chain amino acids are deaminated to the corresponding ketoacids before undergoing irreversible decarboxylation and oxidation. Some of these ketoacid molecules could escape intracellular metabolism and be released into the bloodstream. In our study, we did not measure
the arteriovenous balance of branched-chain ketoacids across the leg to
directly determine branched-chain amino acid oxidation. Nonetheless,
the difference between the rates of branched-chain amino acid
catabolism and oxidation is likely to be very small for two reasons.
First, the arteriovenous difference across the leg of the leucine
ketoacid
-ketoisocaproate is usually very small (33).
Second, during leucine tracer infusion, there is a net uptake across
the leg of the
-ketoisocaproate tracer derived from deamination of
the leucine tracer (33).
Inward and outward amino acid transports are calculated as the rates of
net amino acid movements from the femoral artery to muscle and from
muscle to the femoral vein (2, 3,
6), respectively
|
(9)
|
|
(10)
|
where EA, EV and EM were
amino acid enrichments in the femoral artery, femoral vein, and muscle, respectively.
Muscle protein synthesis and proteolysis were calculated as rates of
intracellular phenylalanine disposal and appearance (2, 3, 6), respectively
|
(11)
|
|
(12)
|
where (PHE) indicates the values of phenylalanine concentrations
and enrichments.
The absolute rate of glutamine de novo synthesis was calculated from
the rate of total intracellular glutamine appearance and glutamine
appearance from proteolysis. The latter is calculated from the rate of
intracellular phenylalanine appearance from proteolysis (Eq. 12) and the molar ratio glutamine/phenylalanine in mixed muscle proteins (2)
|
(13)
|
where (GLN) indicates the values of glutamine concentrations and enrichments.
Determination of Specific mRNA Levels
Muscle mRNA levels of cathepsin B (7), UbB
polyubiquitin (7, 35), and glutamine
synthetase (34) were determined by competitive RT-PCR
(15). Ubiquitin is encoded in the human genome as a
multigene family (35). Among the different ubiquitin genes, we assessed the UbB polyubiquitin gene, which codes for three
direct repeats of the ubiquitin sequence (7,
35). Total RNA was extracted from 20-30 mg of muscle
tissue by the guanidine thiocyanate procedure (7,
15). The quality and integrity of total RNA and its
approximate concentration were evaluated by denaturing agarose gel
electrophoresis and ethidium bromide staining. The presence of
contamination from genomic DNA in the extracted total RNA was checked
by amplification with primers for glyceraldehyde-3-phosphate
dehydrogenase (GAPDH) (see below). If contamination was detected, the
samples were treated with RNase-free DNase I (10 U/µl) (Boehringer
Mannheim, Mannheim, Germany) and extracted with
phenol-chloroform-isoamylic alcohol.
Approximately 5 µg of the extracted total RNA were incubated with 100 ng of oligonucleotide NOT I-(dT)18 (Pharmacia) for 10 min
at 68°C. Then reverse transcription was performed at 42°C for
1 h using murine myeloblastosis virus reverse transcriptase (GIBCO
BRL Life Technologies, Paisley, Scotland) in the presence of
ribonuclease inhibitor RNASE BLOCK (Stratagene, La Jolla, CA).
Amplification of GAPDH cDNA was performed using the primers 5'
CCATCACCATCTTCCAGGAGCG 3' (nucleotides 278-299 of file
M33197 in GenBank) for the sense strand and 5' ACGGAAGGCCATGCCAGTGA 3' (762-743) for the antisense strand. Amplification of
cathepsin B cDNA was performed using the primers 5'
GGGCACAACTTCTACAACGTGG 3' (nucleotides 307-328 of file L16510 in
GenBank) sense and 5' GTGTGGATGCAGATGCGGTCA 3'
(551-531) antisense. The sense strand primer for UbB
polyubiquitin cDNA amplification was 5' CAGTCATGGCATTCGCAGTGCC 3'
(nucleotides 1789-1810 of file X04803 in GenBank), whereas oligonucleotide NOT I-(dT)18 was used directly as antisense
primer. The sense strand primer for glutamine synthetase cDNA was 5'
GACTTTGGAGTGATAGCAACC 3' (nucleotides 810-830 of file Y00387 in
GenBank) and primer 5' GCAGTTGGCAGAGGGGCGACGA 3'
(1136-1157) for the antisense strand. Competitors for
cathepsin B, UbB polyubiquitin, and glutamine synthetase were
constructed by a recombinant PCR methodology, as already described for
cathepsin B and UbB polyubiquitin (7, 15).
Each competitor corresponding to the sequence amplified by the primers
reported above, plus an insertion of 26 bp in the middle, was
constructed by two separate PCR amplifications. The sequences of the
internal primers for the glutamine synthetase competitor were as
follows: 5' CTTCGGAGAGAATCCCCTCAGCGGGAGGAGGCCATTGAGAAAC 3' for the
sense strand and 5' CCGCTGAGGGGATTCTCTCCGAAGGATGTACTTCAGACCATTC 3' for
the antisense strand. The competitor for GAPDH quantification was the
plasmid pBluGAPDH 260 (kindly donated by Prof. F. E. Baralle, ICGEB, Trieste, Italy) obtained from the pBluescript II KS vector (Pharmacia) (7, 15).
Quantitative PCR amplifications were performed by mixing scalar amounts
of competitor DNA to the target cDNA followed by PCR amplification with
the appropriate primer pairs. All amplifications were conducted in PCR
buffer containing the two primers, the four dNTPs, 2.5 U of Taq DNA
polymerase (Boehringer), 1 µl of cDNA, and 1 µl of appropriately
diluted competitor DNA, using a DNA Thermo Cycler (Perkin-Elmer Cetus).
Samples were submitted to 38 cycles of amplification. After
amplification, PCR products were resolved on an 8% nondenaturing
polyacrylamide gel, visualized under ultraviolet light after ethidium
bromide staining, and photographed. Quantification of the amplification
products was obtained by densitometric scanning on photographs of the
ethidium bromide-stained gels. According to the principles of
competitive PCR, the ratio between the amount of the PCR products for
the target cDNA and that of the competitor DNA is linearly correlated
with the initial amount of cDNA in the reaction. To verify such a
linear correlation, we plotted the densitometric ratios between the PCR
products of scalar amounts of competitor DNA (0 pg, 0.1 pg, 0.5 pg, 1 pg, 2 pg) and fixed amounts of target glutamine synthetase cDNA with the initial amount of competitor DNA in the reaction. The correlation coefficient (r2) was 0.9989, and the equation
was y = 2.55 × +0.04.
Cathepsin B, UbB polyubiquitin, and glutamine synthetase mRNA levels
were calculated from the ratios between cathepsin B and GAPDH cDNA,
between UbB polyubiquitin and GAPDH cDNA, and between glutamine
synthetase and GAPDH cDNA quantities in the same sample. The units of
cathepsin B, UbB polyubiquitin, or glutamine synthetase mRNA content
are expressed as %GAPDH mRNA levels. It is assumed that rhGH
administration does not modify GAPDH mRNA levels in skeletal muscle of
trauma patients (19, 25). The variation coefficient for GAPDH mRNA content for four different samples from the
same muscle specimen was 3.2% of the mean when the samples were
processed on the same day and 4.3% of the mean when the samples were
processed on different days. The two muscle samples from a single
subject, whether studied in basal conditions or during rhGH infusion,
were always processed together.
Data Presentation and Statistics
All data are expressed as means ± SE. Because the values
of leg blood flow, amino acid concentrations, and kinetics were not significantly different in the control studies performed before rhGH
infusion (protocol 1) and 24 h after rhGH
discontinuation (protocol 2), the results obtained in the
two control studies were pooled together. Then, the values of leg blood
flow, amino acid concentrations, and kinetics obtained during rhGH
infusions (protocols 1 and 2) were compared with
those in the control studies (protocols 1 and 2)
by means of Student's t-test for paired samples. The
effects of rhGH infusion (protocol 1) and rhGH
discontinuation (protocol 2) on muscle levels of selected
mRNAs were assessed separately in the two protocols by means of
Student's t-test for paired samples. P < 0.05 was considered statistically significant.
 |
RESULTS |
Before rhGH infusion, plasma growth hormone concentration was
3 ± 1 ng/ml (protocol 1) and increased to 91 ± 9 ng/ml in the femoral vein and to 39 ± 3 ng/ml in the antecubital
vein during rhGH infusion (protocols 1 and 2).
After rhGH discontinuation, plasma growth hormone concentration
decreased to 3 ± 1 ng/ml (protocol 2). Plasma insulin
concentration increased (P < 0.05) during rhGH infusion (protocol 1) from 77 ± 22 to 141 ± 20 µU/ml. Insulin concentration did not change significantly after rhGH
discontinuation (protocol 2) (from 131 ± 22 to
122 ± 33 µU/ml). Mean values of insulin concentrations were not
significantly different in the pooled control studies (99 ± 20 µU/ml) and during rhGH infusions (136 ± 14 µU/ml;
protocols 1 and 2). Plasma IGF-I concentration tended to increase (P = 0.08) during rhGH infusion
(protocol 1) from 93 ± 14 to 153 ± 27 ng/ml.
IGF-I concentration did not significantly change after rhGH
discontinuation (protocol 2; from 188 ± 15 to 187 ± 17 ng/ml). Mean values of IGF-I concentrations were not significantly different in the pooled control studies (140 ± 21 ng/ml) and during rhGH infusions (170 ± 16 ng/ml; protocols
1 and 2).
Leg blood flow was similar in the control studies (5.25 ± 0.62 ml · min
1 · 100 ml leg vol
1)
and during rhGH infusions (5.01 ± 0.83 ml · min
1 · 100 ml leg vol
1).
The values of amino acid concentrations in femoral artery and vein were
not significantly different in the control studies and during rhGH
infusion (Table 1). In skeletal muscle,
intracellular concentrations of most amino acids tended to decrease
during rhGH infusion. Intramuscular glutamine concentrations decreased
(P = 0.07) by ~10%. Table
2 shows the values of amino acid balance across the leg in the control studies and during rhGH infusion. In the
basal studies, the net balance of most amino acids was negative and
significantly different from zero. Total amino acid release from leg
muscle decreased after rhGH infusion by ~55%. In particular, net
phenylalanine, tyrosine, and lysine release, which are markers of net
muscle protein catabolism, significantly decreased by ~45-55%
during rhGH infusion. Glutamine release also decreased significantly
after rhGH infusion.
Table 3 shows the effects of rhGH
infusion on selected parameters of muscle amino acid metabolism derived
from leg arteriovenous balance of unlabeled amino acids (see
Calculation of Kinetic Parameters). rhGH decreased the rates
of net nonprotein glutamate disposal (mainly to form glutamine) and of
net glutamine synthesis from glutamate, whereas the hormone infusion
did not change the rate of net alanine synthesis. rhGH infusion
decreased the rates of catabolism of the branched-chain amino acids
leucine (from 93 ± 18 to 49 ± 12 nmol · min
1 · 100 ml leg vol
1;
P < 0.05), valine (from 46 ± 18 to 18 ± 8 nmol · min
1 · 100 ml leg vol
1; P < 0.05), and isoleucine
(from 57 ± 11 to 31 ± 5 nmol · min
1 · 100 ml leg vol
1;
P < 0.05). The rate of catabolism of total
branched-chain amino acids decreased by ~50% during rhGH infusion
(Table 3).
Table 4 shows the values of amino acid
enrichments in femoral artery and vein and in muscle tissue. rhGH
infusion tended to increase amino acid enrichments in all sampled
compartments. rhGH administration significantly (P < 0.001) decreased the rate of whole body phenylalanine Ra
from 1.34 ± 0.07 to 1.17 ± 0.08 µmol · kg
1 · min
1 and
tended to decrease the rate of whole body leucine Ra from 2.92 ± 0.26 to 2.72 ± 0.22 µmol · kg
1 · min
1.
rhGH administration tended also to decrease the rates of whole body
glutamine Ra (from 8.01 ± 0.90 to 6.95 ± 0.73 µmol · kgq
1 · min
1) and
clearance (from 194 ± 24 to 167 ± 24 ml · kg
1 · min
1).
Figure 1 shows the rates of intracellular
phenylalanine disposal for protein synthesis and release from protein
degradation during saline and rhGH infusions. rhGH almost doubled the
rate of protein synthesis, whereas the hormone infusion did not
significantly change the rate of protein degradation. Also, the rate of
intracellular leucine release from protein degradation was not
significantly different during saline (220 ± 40 nmol · min
1 · 100 ml leg vol
1)
and rhGH (240 ± 42 nmol · min
1 · 100 ml leg vol
1)
infusions. In addition, muscle mRNA levels of cathepsin B and UbB
polyubiquitin did not significantly change after rhGH infusion (protocol 1; from 17 ± 2 to 19 ± 9% of GAPDH
mRNA and from 4.9 ± 1.8 to 4.0 ± 0.2% of GAPDH mRNA,
respectively) and after rhGH discontinuation (protocol 2;
from 24 ± 9 to 28 ± 9% of GAPDH mRNA and from 10.0 ± 1.8 to 8.7 ± 3.4% of GAPDH mRNA, respectively).

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Fig. 1.
Rates of protein synthesis and degradation in leg
skeletal muscle of trauma patients during saline or recombinant human
growth hormone (rhGH) infusion. * P < 0.05 rhGH vs.
saline.
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|
Table 5 shows the rates of inward and
outward transport of phenylalanine, leucine, and glutamine in skeletal
muscle during saline and rhGH infusions. rhGH significantly increased
the rates of inward transport of the essential amino acids leucine and
phenylalanine, whereas the rates of glutamine transport were decreased
by rhGH in both the outward and the inward directions. Figure
2 shows the isotopically derived rates of
intramuscular glutamine de novo synthesis during saline and rhGH
infusions. The absolute rate of glutamine de novo synthesis was 44 ± 11% lower during rhGH than during saline infusions (protocols
1 and 2). In contrast, rhGH infusion (protocol
1) significantly (P = 0.03) increased glutamine
synthetase mRNA levels from 26 ± 14 to 51 ± 12% of GAPDH mRNA. In the patients studied according to protocol 2,
glutamine synthetase mRNA did not change significantly after rhGH
discontinuation (from 22 ± 6 to 27 ± 12% of GAPDH mRNA).

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Fig. 2.
Absolute rate of de novo glutamine synthesis in leg
skeletal muscle of trauma patients during saline or rhGH infusion.
* P < 0.05 rhGH vs. saline.
|
|
 |
DISCUSSION |
In skeletal muscle of hypercatabolic traumatized patients, 24-h
rhGH infusion 1) increased muscle protein synthesis without significantly affecting protein degradation, 2) suppressed
the rates of branched-chain amino acid catabolism, 3)
increased the rates of transmembrane transport of the essential amino
acids leucine and phenylalanine, whereas it decreased membrane
transport of glutamine, 4) decreased the rates of de novo
synthesis of glutamine, and 5) decreased the rates of
release of total essential and nonessential amino acids from skeletal muscle.
Patients were studied during the second week after trauma in relatively
stable clinical and metabolic conditions. They were hypercatabolic, as
shown by the net efflux of essential amino acids from skeletal muscle
despite a continuous combined parenteral and enteral artificial
nutrition. We have shown that rhGH administration significantly reduces
the net muscle protein loss of patients. This anticatabolic effect of
rhGH is completely accounted for by an acceleration of protein
synthesis. Previous studies have demonstrated a selective increase in
muscle protein synthesis induced by rhGH administration in humans
(12, 13). It is possible, however, that many
of the observed anabolic effects of rhGH are mediated via the
stimulation of endogenous IGF-I synthesis, which may exert its effects
via endocrine and/or paracrine mechanisms. In humans, IGF-I
administration promoted protein anabolism both by stimulating protein
synthesis and by inhibiting protein degradation both in muscle and at
the whole body level (10, 11). In our study,
rhGH administration did not result in significant modifications of the
rates of muscle protein degradation. The effects of rhGH on protein
degradation were evaluated with different approaches. Stable isotopes
and mass spectrometry techniques were used to measure the rate of
protein degradation in skeletal muscle as the rate of appearance of the
essential amino acids phenylalanine and leucine in muscle cells.
With competitive PCR, we determined in skeletal muscle the mRNA levels
of cathepsin B and ubiquitin as markers of nonmyofibrillar and
myofibrillar protein degradation, respectively. It is known that growth
hormone has the potential to decrease proteolysis, possibly through
stimulation of IGF-I synthesis (10, 11). In
this study, we did not observe any decrease in muscle proteolysis: we
observed a substantial stimulation of protein synthesis and a tendency
toward decreasing intracellular concentration of total amino acids. It
is likely that the rate of degradation was maintained and possibly
increased to prevent a further decrease in amino acid concentrations.
In contrast to the rhGH effects on muscle proteolysis, at the whole
body level, the rate of phenylalanine appearance was significantly lower during the hormone infusion. Such a decrease of a marker of whole
body proteolysis clearly indicates a systemic effect of the
intra-arterial rhGH infusion. In fact, in the systemic circulation, not
only growth hormone but also insulin and IGF-I increased during the
local rhGH infusion. It is likely that such a decrease of whole body
proteolysis could have been mediated by insulin and/or IGF-I.
Furthermore, such systemic hormonal changes could also have contributed
to the stimulation of muscle protein synthesis observed in our study
(3, 6, 11).
In agreement with previous observations in animals (20,
23), this study shows that rhGH infusion in traumatized
patients accelerates the rates of transmembrane transport of the
essential amino acids leucine and phenylalanine. This effect was
independent of changes of leg blood flow and arterial amino acid
concentrations. This rhGH-mediated increased ability of transmembrane
systems to transport essential amino acids confirms previous
observations in vitro (20, 23) and represents
a novel observation in vivo. This acceleration of transport of
essential amino acids may have contributed to the anabolic effect of
rhGH by increasing intracellular amino acid availability for protein
synthesis. However, the slight decrease in intracellular amino acid
concentrations suggests that the stimulation of amino acid transport is
not the primary mechanism for stimulation of synthesis. In contrast,
transport acceleration may represent a compensatory mechanism to
prevent depletion of intracellular amino acid pools.
Besides stimulating protein synthesis, growth hormone suppressed the
rate of catabolism of the branched-chain amino acids leucine,
isoleucine, and valine. This effect has been reported by several other
authors using isotopic tracers of leucine at the whole body level
(8, 12).
In this study, the anticatabolic growth hormone effects on protein and
branched-chain amino acid metabolism were paralleled by a suppression
of the rate of de novo muscle glutamine production. De novo muscle
glutamine production was determined either as absolute intracellular
synthesis by stable isotopes or as net rate of synthesis appearing in
the circulation without stable isotopes. With both methods, muscle
glutamine production decreased by ~50% during rhGH infusion.
Immediate precursors of glutamine synthesis are glutamate and free
ammonia, whereas precursors for glutamate synthesis are
-ketoglutarate and amino-nitrogen from the branched-chain amino
acids (9). The rates of branched-chain amino acid
catabolism and of nonprotein utilization of glutamate, which is mainly
for glutamine synthesis, decreased after rhGH infusion. Thus a
decreased glutamine de novo synthesis could be secondary to a decreased glutamate availability from reduced transamination of the
branched-chain amino acids with
-ketoglutarate. The fact that mRNA
levels of glutamine synthetase (the key enzyme for glutamine synthesis) were found to be increased after rhGH infusion in the present study, as
well as in previous studies in animals (29), supports the
hypothesis that the growth hormone-mediated decrease in glutamine synthesis was determined by decreased precursor availability. Gene
expression of glutamine synthetase may increase after rhGH infusion to
compensate for reduced glutamine precursor availability.
In our study, besides a suppression of the rate of glutamine
production, we observed a decrease of glutamine transmembrane outward
transport from skeletal muscle, which prevented excessive depletion of
the intracellular glutamine pool in skeletal muscle. Previously,
Hammarqvist et al. (17) showed that rhGH administration prevented the decrease in muscle free glutamine typically observed after surgical trauma. We may speculate that, in their study, an
rhGH-mediated decrease of outward glutamine transport prevailed over
the effect on glutamine production. Consequently, glutamine concentration tended to increase during rhGH administration.
Glutamine and alanine constitute the major carriers of nitrogen among
body tissues (2). In skeletal muscle, these amino acids
are constantly being synthesized and released into the bloodstream (2). In severe trauma, alanine release from muscle is
greatly accelerated, whereas glutamine release was found to be
increased or unchanged (5). Our results indicate that rhGH
administration selectively decreases the rates of synthesis and release
of glutamine, whereas alanine synthesis did not change during the
hormone administration.
In our experimental protocol, the control study was performed either
before rhGH administration (protocol 1) or 24 h after the hormone discontinuation (protocol 2). This approach was
adopted to account for time-related changes of muscle metabolism after trauma and for potential interference between two close stable isotope
infusions. However, in the control studies of protocols 1 and 2, the whole body hormonal pattern tended to be
different. In protocol 1, local rhGH infusion tended to
increase systemic concentrations of both insulin and IGF-I; in the
control study of protocol 2, despite the fact that
circulating growth hormone returned to basal levels, both insulin and
IGF-I concentrations failed to decline after rhGH discontinuation.
However, this does not seem to constitute a problem for evaluating the
rhGH effects on muscle protein and glutamine kinetics. In fact, the
values of amino acid concentrations and kinetics were not significantly different in the control studies performed before rhGH infusion (protocol 1) and 24 h after rhGH discontinuation
(protocol 2). Furthermore the rhGH-mediated changes of amino
acid kinetics were similar in protocols 1 and 2.
Protein synthesis increased by 137 ± 0.68% after rhGH infusion
(protocol 1) and decreased by 113 ± 0.42% after rhGH
discontinuation (protocol 2). Branched-chain amino acid
catabolism decreased by 41 ± 7% after rhGH infusion (protocol 1) and increased by 42 ± 12% after rhGH
discontinuation (protocol 2). Glutamine de novo synthesis
decreased by 58 ± 14% after rhGH infusion (protocol
1) and increased by 31 ± 15% after rhGH discontinuation
(protocol 2). It is likely, therefore, that the observed
changes of protein and glutamine kinetics were directly mediated by an
increase of growth hormone concentration and not by a secondary
stimulation of insulin and IGF-I secretions.
In contrast to the rhGH-mediated changes in amino acid kinetics, muscle
mRNA levels of glutamine synthetase significantly increased after rhGH
infusion (protocol 1) but failed to decline after rhGH
discontinuation (protocol 2). Several potential explanations may account for such a discrepancy between protocols 1 and 2. 1) Growth hormone induction of the
glutamine synthetase gene may continue for many hours after the
infusion has been discontinued. 2) Glutamine synthetase mRNA
could be very stable. 3) Induction of the glutamine
synthetase gene might be mediated by the secondary stimulation of
insulin and/or IGF-I secretion, which also failed to decline in
protocol 2. Finally, 4) a hormone-mediated change of the housekeeping gene GAPDH is unlikely, because this change would
have also affected the determination of mRNA levels of cathepsin B and
of UbB polyubiquitin, which did not change after rhGH infusion in
either protocol 1 or protocol 2. Whatever the
reason for the discrepancy between protocols 1 and
2, the results of protocol 1 confirm the ability of the
growth hormone to induce in human skeletal muscle the glutamine
synthetase gene previously shown in different animal tissues
(29).
To calculate intracellular enrichments and concentrations in skeletal
muscle, we assumed a ratio of 0.16 between the extra- and intracellular
spaces in muscle samples. This value had been obtained previously in
normal human subjects (2). Nonetheless, despite the fact
that none of our patients showed clinical evidence of changes in fluid
retention during the study, we cannot exclude the possibility that
growth hormone could have slightly increased extracellular space volume
in muscle tissue. In fact, sodium and water retention are well
recognized side effects of rhGH administration (16). We
therefore evaluated the potential effects of changes in the ratio
between the extra- and intracellular compartments on the calculated
kinetic parameters of protein and glutamine metabolism. When such a
ratio was increased by 100% over the assumed initial value of 0.16, the changes in protein synthesis and degradation, glutamine de novo
synthesis, and amino acid transport were <5%. It is unlikely,
therefore, that growth hormone-mediated changes in extracellular fluid
in muscle could have affected the conclusions of our study.
Takala et al. (31) recently reported the results of two
large multicenter studies indicating an increased morbidity and mortality in critically ill patients treated with high doses of growth
hormone. Multiple organ failure and septic shock or uncontrolled infections were the main causes of death, suggesting that a modulation of the immune system or gut function was involved. However, the reason
for such a deleterious effect of growth hormone is unclear. Our study
describes a potential side effect of rhGH administration in critically
ill patients. In these patients, glutamine is an essential substrate
for rapidly dividing cells in the immune system and in the intestinal
mucosa, as well as for glutathione synthesis in the liver. Skeletal
muscle is the main tissue involved in glutamine de novo synthesis. In
our patients, whole body skeletal muscle released ~19 g of glutamine
per day into the bloodstream before rhGH administration [to
extrapolate the data to whole body skeletal muscle (33),
the data for one leg were multiplied by four]. After rhGH
administration, glutamine release from skeletal muscle decreased by
~50%, whereas at the whole body level, glutamine clearance tended to
decrease by ~15%, despite the fact that the highest growth in
hormone levels was achieved in only one leg. We may speculate,
therefore, that in a clinical setting, rhGH therapy could decrease
systemic glutamine availability and have negative effects on the immune
system and gut function. The obvious solution for this potential side
effect of growth hormone treatment in critically ill patients is to
simultaneously administer exogenous glutamine to offset the decreased
availability of the endogenous amino acid.
 |
ACKNOWLEDGEMENTS |
We thank Prof. F.E. Baralle and Prof. M. Giacca (International
Center for Genetic Engineering and Biotechnology, Trieste), and
Dr. R. Situlin (Istituto di Clinica Medica, University of Trieste) for
invaluable advice and collaboration. We thank Drs. L. Luzi and A. Battezzati (San Raffaele Scientific Institute, Milan) for the analysis
of IGF-I plasma levels. The human recombinant growth hormone and
additional support were kindly provided by Pharmacia and Upjohn (Milan,
Italy). We also wish to express our appreciation to Anna De Santis and
Mariella Sturma for excellent technical assistance.
 |
FOOTNOTES |
G. Biolo was awarded the European Society of Parenteral and Enteral
Nutrition (ESPEN)-Clintec Fellowship 1994 ("Effects of growth hormone
administration on skeletal muscle glutamine metabolism in severely
traumatized patients"). A. Bosutti was awarded the ESPEN Fellowship
1996 ("Molecular regulation of protein catabolism in trauma
patients").
Address for reprint requests and other correspondence:
G. Biolo, Istituto di Clinica Medica, Ospedale di Cattinara, Strada di
Fiume 447, Trieste 34149, Italy (E-mail:
gianni.biolo{at}clmed.univ.trieste.it).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Received 13 August 1999; accepted in final form 24 February 2000.
 |
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