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Department of Intensive Care Medicine, Burn Unit and Center for Experimental Surgery and Anesthesiology (F.W., M.M., G.V.d.B.), Laboratory for Experimental Medicine and Endocrinology (E.V.H., W.C.), and Laboratory for Comparative Endocrinology (V.M.D.), Catholic University of Leuven, Leuven, Belgium; and Department of Medicine, Mayo Graduate and Medical Schools (J.D.V.), Mayo Clinic, Rochester, Minnesota 55905
Address all correspondence and requests for reprints to: Frank Weekers, M.D., Department Intensive Care Medicine, University of Leuven, B-3000 Leuven, Belgium. E-mail: frank.weekers{at}uz.kuleuven.ac.be.
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Abstract |
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Introduction |
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There appears to be a biphasic neuroendocrine response to critical illness. During the first few hours and days after an insult, the adaptive neuroendocrine response to critical illness consists primarily of activated anterior pituitary function with peripheral tissue resistance. In the later phase of critical illness, from 7 d onward, pulsatile secretion of anterior pituitary hormones is reduced, and organ function is further impaired. Reduced availability of hypothalamic TRH and the endogenous ligand of the GH-releasing peptide (GHRP) receptor (ghrelin) are postulated factors in reduced TSH and GH secretion. Pulsatile secretion of TSH and GH in critically ill patients can be reamplified by combined infusions of the releasing factors TRH and GHRP-2, which increase circulating levels of T4, T3, IGF-I, and GH-dependent IGF-binding proteins (8).
Anabolism is achieved when GHRP-2 and TRH are administered together, but not individually. Nonetheless, both GHRP and rhGH infusions in the critically ill patient exacerbate the stress-induced insulin resistance and hyperglycemia (2, 9). Indirect data suggest that adequate availability of thyroid hormone may be crucial for GHRP-2 or rhGH to evoke anabolic responses (10) and prevention of hyperglycemia may be essential to improve survival (11).
In view of lethal side-effects, studies with rhGH are now impossible in patients. We described an animal model marked by typical endocrine and metabolic characteristics of prolonged critical illness (7). This model allows possible investigation of the primary mechanisms of toxicity of rhGH in critical illness.
We hypothesized that the toxicity of high dose rhGH treatment during critical illness is explained by the induction of insulin resistance and hyperglycemia and that impaired anabolism reflects low thyroid hormone availability. According to this postulate, infusion of GHRP-2 in combination with TRH and insulin may provide a superior anabolic strategy.
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Materials and Methods |
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Two days later, animals were reweighed, and blood samples for biochemical and blood gas analyses were taken via the arterial line. Then, after shaving both flanks and under deep general anesthesia supplemented with a local paravertebral block, a 15% full thickness burn injury was imposed (Fig. 1
). After a 1-h recovery period with supplemented oxygen, the animals returned to their cages, and fluid resuscitation was started overnight with a continuous infusion of Ringer lactate at six drops per minute (±12 ml/h) through a volumetric infusion pomp. To assure free movement of the rabbits without twisting of the catheter and infusion lines, a swivel device was incorporated. In the evening, a supplemental dose of a major analgesic (Dipidolor, Janssen-Cilag, Beerse, Belgium) was given.
The next morning at 0800 h ± 30 min, parenteral nutrition was initiated in all rabbits at four drops per minute (±12 ml/h). Blood glucose levels were kept within narrow limits (<180 mg/dl) by frequent blood glucose monitoring and titration of insulin infusion (100 IU/ml Actrapid, Novo Nordisk, Bagsvaerd, Denmark; via an SE200B infusion pump, Vial Medical, Brezins, France) when necessary. All iv infusions were weighed before and after administration, allowing accurate determination of the infused quantity. Parenteral nutrition was prepared daily in the hospital pharmacy under laminar air-flow conditions. The infusion bags contained 103 ml 20% glucose, 37.5 ml Aminoplasmal L10 (Braun, Melsungen, Germany), 34 ml 20% Intralipid (Fresenius Kabi, Stockholm, Sweden), and 150 ml sterile water. Thus, bags with 325 ml solution contained 150 kcal nonprotein calories and 0.6 g nitrogen. Of all calories, 56% were delivered as carbohydrates, and 44% as fat. Protein intake equaled 1.25 g amino acids/kg·d. No additional vitamins or trace elements were added. Total parenteral nutrition was continuously administered. Animals had free access to water, but oral food intake was denied. Parenteral nutrition bags were changed daily.
On d 1 and 3 blood was sampled for later analysis. On the morning of d 4, surviving rabbits were randomly (by sealed envelopes) allocated to one of the following treatment groups. Group 1 (saline group) received a 4-d continuous infusion of 0.9% NaCl. Goup 2 (rhGH group) was injected sc on 4 consecutive d with 3.5 mg/kg·d of rhGH (Genotropin, Amersham Pharmacia Biotech, Stockholm, Sweden) in two divided injections (one at 0800 h ± 30 min and one at 2000 h ± 30 min), and insulin was kept at the same infusion rate as on the morning of d 4 regardless of glucose levels during further treatment. Group 3 (from hereon referred to as the GHRP-2+TRH group) received an initial bolus of 60 µg/kg GHRP-2 (Kaken Pharmaceutical Co. Ltd., Tokyo, Japan) plus 60 µg/kg TRH (200 µg/ml 0.9%NaCl; Ferring, Kiel, Germany), followed by a continuous infusion of 60 µg/kg·h GHRP-2 and 60 µg/kg·h TRH. Infusion syringes and infusion lines were protected from the light. The dose of GHRP-2 and TRH was defined in a previous study (7). In groups 1 and 3, insulin was titrated to maintain blood glucose levels below 180 mg/dl (9.9 mM/liter) throughout the study.
On d 4, between 0830 h ± 30 min and 1530 h ± 30 min, 1 ml blood was sampled every 15 min for determination of plasma rabbit GH (rGH) and the TSH concentration-time series over 7 h. Blood was drawn from the arterial line using a Vamp system (Baxter Healthcare Corp., Irvine, CA), which allows undiluted blood sampling without undue blood loss. Blood was collected into glass tubes containing 25 U heparin and then stored at 4 C until centrifugation. All samples were centrifuged, and plasma was stored at -70 C until assay. In all groups, study drugs were only injected after two baseline samples (i.e. the first two samples of the profiles) had been withdrawn. Blood for blood gas and biochemical analyses was taken before and after the profile. On d 5, 6, and 7, parenteral nutrition bags were changed daily, and additional blood samples were taken on d 5.
On d 8, again during TPN infusion and continuation of the study drugs, another 7-h sampling time series (sampling every 15 min) was performed to evaluate rGH and TSH secretion. After the last blood sample, animals were weighed and euthanized with sodium-pentobarbital (60 mg/ml Nembutal, Sanofi-Winthrop, New York, NY). Organs were harvested, and heart, lungs, liver, spleen, and kidney were weighed. Subsequently, tissue samples were taken from left and right ventricles, lung, liver, spleen, kidney, ileum, muscle, intact skin, and burned skin. In each of these tissue samples, a part was used to determine dry weight and water content. Water content was determined by weighing the samples before and after 24 h of lyophilization (Alfa I-5 Chriss, Aichach, Germany). Part of the liver was snap-frozen in liquid nitrogen-chilled isopentane (Sigma-Aldrich Corp., Bornem, Belgium) and stored at -70 C until further analysis of deiodinase activity.
All blood gas analyses were performed immediately after sampling. Blood for biochemical analyses was sampled in lithium citrate tubes (BD Biosciences, Temse, Belgium), immediately centrifuged, and then stored at -70 C until assay.
Assays
Arterial blood was analyzed on an ABL 700 analyzer (Radiometer Medical A/S, Copenhagen, Denmark) to quantitate pH, pO2, pCO2, hemoglobin, lactate, and glucose. Furthermore plasma concentrations of sodium, potassium, chloride, bicarbonate, ureum, creatinine, total calcium, phosphorus, total protein content, alkaline phosphatase, transaminases (aspartate aminotransferase and alanine aminotransferase), 
glutamyl transpeptidase, total bilirubin, uric acid, creatine kinase, lactate dehydrogenase, amylase, lipase, cholesterol, and triglycerides were determined using the Roche Modular with Roche reagents (Roche, Mannheim, Germany).
Plasma rGH concentrations were measured with a specific RIA (reagents provided by Dr. A. Parlow, National Pituitary Agency). The detection limit was 1 µg/liter, and the within-assay coefficient of variation was 2.3%. All samples were analyzed in duplicate within a single assay run. All samples contained detectable plasma concentrations.
Plasma IGF-I concentrations were measured in duplicate by RIA, using a slightly modified version of that described by Verhaeghe et al. (12). After acidification of the plasma samples with formic acid, binding proteins were separated by acid gel filtration on an Econo-Pac column (Bio-Rad Laboratories, Richmond, CA). Des-IGF-I was used as tracer to avoid binding to residual small IGF-binding proteins. The intraassay coefficient of variation was 4.6%. The detection limit was 20 µg/liter.
Plasma concentrations of rabbit TSH were measured by a specific RIA (reagents provided by Dr. A. Parlow, National Pituitary Agency). The detection limit was 1.2 mU/liter. All samples were analyzed in duplicate within a single assay run. For samples with undetectable levels, a value representing half the detection limit was entered.
Total plasma T4 and T3 concentrations were determined by a specific RIA (Immunotech, Marseilles, France). The sensitivity for T3 was 0.2 nmol/liter, and that for T4 was 6 nmol/liter. All samples were assayed in duplicate.
Plasma insulin concentrations were determined by immunoradiometric assay (Medgenix INS-IRMA, BioSource, Fleurus, Belgium).
Hepatic deiodinase activity
For determination of hepatic type 1 (D1) and type 3 (D3) iodothyronine deiodinase activities, liver microsomal fractions were prepared and assayed as described by Darras et al. (13). Final incubations were in a total volume of 200 µl. For D1 activity, final incubation mixtures contained 1 µM rT3, 50,000 cpm/tube [3',5'-125I]rT3 (labeled using 17 Ci/mg carrier-free 125I from NEN Life Science Products, Boston, MA), 250 µg/ml microsomal liver protein, 2 mM EDTA, and 5 mM dithiothreitol and were incubated for 30 min at 37 C. For D3 activity, incubations contained 1 nM T3, 200,000 cpm/tube [3'-125I]T3 (labeled as described above), 250 µg/ml liver microsomal protein, 1 µM rT3, 0,1 mM propylthiouracil, 2 mM EDTA, and 50 mM dithiothreitol and were incubated for 120 min at 37 C.
Analytical procedures
The time series of sequential plasma rGH concentrations measured every 15 min for 7 h on d 4 and 8 was transformed into pituitary secretion profiles using multiple parameter deconvolution analysis (14). This method is designed to compute hormonal half-life and the number and mass of underlying pituitary secretory bursts and to estimate basal secretion. The following parameters were calculated: half-life, temporal position of the secretory bursts, basal rGH secretion rate (amount of rGH released from pituitary to achieve serum concentrations approximating the mean of the lowest 5% of all values observed [micrograms per liter of distribution volume (Lv) per minute]), mass [area of the resolved secretion burst (micrograms per Lv)], mean pulsatile production [product of the number of secretory bursts and the mean secretory burst mass over the time interval considered (micrograms per Lv over 7 h)], and total rGH secretion (sum of basal and pulsatile released rGH) were calculated. Deconvolution analysis of the plasma TSH concentration profiles was not feasible because too many samples had undetectable TSH concentrations. Thus, TSH concentrations are given as the mean and the area under the curve (AUC) calculated by trapezoidal rule.
Statistics
Comparisons of normally distributed data among the three groups were made by ANOVA. Within-group changes were analyzed by t test with Bonferroni correction for multiple comparisons when indicated. Nonnormally distributed data were analyzed by Friedman test or Kruskal-Wallis test as appropriate. All data are expressed as the mean ± SD unless specified otherwise. P < 0.05 was considered significant.
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Results |
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Body weight
The mean body weight of the rabbits at the start of the study was 3004 ± 266 g in the saline group (n = 8), 3050 ± 157 g in the GHRP-2+TRH group (n = 9), and 3136 ± 211 g in the rhGH group (n = 7). All rabbits lost weight 4 d after burn injury: to 2828 ± 196 g in the saline group (P = 0.04 vs. baseline), to 2891 ± 134 g in the GHRP-2+TRH group (P = 0.01 vs. baseline), and to 2921 ± 214 g in the rhGH group (P = 0.007 vs. baseline). Loss of body weight was similar in the three groups. Thereafter there was no further change in body weight toward d 8. Equal amounts of parenteral feeding were administered in the three groups.
GH axis
Mean baseline plasma rGH concentrations were similar in the three study groups (4.0 ± 2.8 µg/liter in the saline group, 4.7 ± 4.2 µg/liter in the GHRP-2+TRH group, and 3.16 ± 2.03 µg/liter in the rhGH group; P = NS). Within 1 d of treatment, exogenous high doses of rhGH suppressed and GHRP-2+TRH infusion increased pulsatile rGH secretion compared with saline (Figs. 2 and 3 and Table 1). Between d 4 and 8, pulsatile rGH secretion decreased further in the saline and rhGH groups, whereas it remained elevated in the GHRP-2+TRH group. The rhGH-induced suppression was mainly caused by reduced rGH burst mass. The initial GHRP-2+TRH-induced amplification of rGH secretion was caused by an increase in rGH burst mass, whereas elevated basal secretion accounted for most of the increase on d 8 (Table 1).
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) in all three groups. The plasma IGF-I concentration remained stable between d 3 and 8 in the saline group, whereas rhGH administration significantly increased and GHRP-2+TRH tended to increase IGF-I by d 8. Serum IGF-I levels on d 8 were significantly higher in the rhGH group compared with the GHRP-2+TRH group (Fig. 4).
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and 6).
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) in all three groups. In the saline and rhGH groups, both TT4 and TT3 remained low between d 3 and 8, whereas in the GHRP-2+TRH group, there was a progressive increase in TT4 and TT3.
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Glucose control
At baseline, the mean blood glucose concentration was 156 ± 38 mg/dl (Fig. 8, upper left panel). On d 3, the insulin titration protocol to keep blood glucose levels below 180 mg/dl resulted in a mean blood glucose concentration of 130 ± 19 mg/dl (P = 0.02 vs. baseline and no difference among the three groups). At the start of treatment with either rhGH or GHRP-2+TRH, blood glucose levels transiently increased on d 5 compared with the saline group, with equal blood glucose levels on d 8. The total amount of insulin infused in the three groups was equal (Fig. 8, upper right panel). Plasma insulin concentrations increased immediately after trauma equally in the three study groups (Fig. 8, lower panel), with a trend for a further increase on d 5 (P = 0.06).
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Baseline plasma levels of aspartate aminotransferase (19 ± 10 U/liter) and alanine aminotransferase (32 ± 8 U/liter) transiently increased after burn injury on d 3 equally in the three groups and returned to baseline on d 8. Plasma bilirubin was below the detection limit of the assay in all animals (<0.1 mg/dl). Baseline plasma levels of alkaline phosphatase (124 ± 42 U/liter) tended to progressively decline toward d 8 in the saline (68 ± 79 U/liter) and GHRP-2+TRH (46 ± 16 U/liter) groups (both P = 0.06 vs. d 0), but not in the rhGH group. The plasma amylase concentration was 277 ± 56 U/liter at baseline and rose to 398 ± 148 U/liter on d 3 (P = 0.004). On d 8, the plasma amylase concentration remained higher in the GHRP-2+TRH group (316 ± 29 U/liter; P = 0.01 vs. saline), whereas it normalized in the two other groups. Levels of glutamyl transpeptidase, lipase, and lactate dehydrogenase remained unaltered.
Baseline plasma creatine kinase (2248 ± 3133 IU/liter) levels were similar in the three groups and transiently increased on d 3 postburn (3990 ± 3047 IU/liter; P = 0.0029 vs. d 0), returning to baseline on d 5.
The mean plasma triglyceride concentration was 65 ± 29 mg/dl at baseline. After injury and under parenteral nutrition, all animals developed hypertriglyceridemia (135 ± 76 mg/dl in the saline group, 363 ± 507 mg/dl in the GHRP-2+TRH group, and 190 ± 57 mg/dl in the rhGH group), which persisted throughout the experiment. There was no difference among the three groups.
Organ weight
There was no significant difference among the three groups in wet weights of heart, lungs, and spleen. The liver tended to be heavier in rhGH-treated animals compared with the saline group (P = 0.07).
Water contents of left and right ventricle, lungs, liver, spleen, small intestine, kidney, muscle, and burned and normal skin were identical in the three groups.
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Discussion |
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We hypothesized that the infusion of GHRP-2 and TRH combined with insulin-titrated maintenance of normoglycemia would be less toxic and more favorable to survival than administration of high doses of rhGH in an animal model of prolonged critical illness. Indeed, exacerbation of insulin resistance and hyperglycemia by high doses of rhGH and reduced systemic availability of thyroid hormones in critical illness are metabolic alterations that may have contributed to increased morbidity and mortality in a clinical study (2).
The high dose rhGH and GH secretion driven by GHRP-2 and TRH infusion did not substantially accentuate burn injury-induced insulin resistance and hyperglycemia. rhGH elevated circulating IGF-I concentrations and suppressed endogenous pulsatile rGH secretion, demonstrating efficacy in the rabbit model. It seems, therefore, that rabbits have reduced sensitivity to insulin antagonism by rhGH. This conjecture is in line with the results of a study in transgenic rabbits overexpressing the bovine GH gene (15). Whether the absence of an adverse rhGH effect on stress-induced insulin resistance and hyperglycemia explains the apparent absence of increased mortality is speculative. However, strict glycemic control with insulin in critically ill patients limits excessive inflammation, sepsis, organ failure, and mortality (11), whereas high dose rhGH treatment predicts insulin resistance, hyperglycemia, inflammation, sepsis, multiple organ failure, and doubling of mortality (2).
The present outcomes in the rabbit contrast with the findings of Angelin et al. in rats (16). Continuous infusion of rhGH, followed by endotoxin injection in the latter model, heightened the risk of cardiovascular shock, hypertriglyceridemia, and biochemical signs of multiple organ failure. Possible distinctions of relevance in the two studies include interventional sequence (17, 18), mode of rhGH administration (19, 20, 21), type of injury imposed, feeding status, and species. Although of major importance, the precise effect of rhGH on immune function is complex. In parenterally fed trauma patients, rhGH (0.15 mg/kg·d) for 7 d increased TNF-
and IL-6 (22), whereas in patients undergoing abdominal aneurysm surgery, preoperative injection of rhGH for 6 d did not alter IL-1a receptor antagonist, IL-6, or acute phase protein concentrations (23). In vitro, pharmacological concentrations of rhGH affected the release of cytokines (24). Finally, endotoxin evokes profound, but short-lasting, cytokine release, unlike sustained cytokine production in illness in humans or evolving burn injury (25).
The current study outcome is consistent with human (26) and other animal (7) studies in which prolonged critical illness depresses hypothalamic-pituitary secretion. In the control intervention, rabbits with major burns exhibited a fall in pulsatile rGH secretion and serum IGF-I concentrations between d 4 and d 8 of the experiment. As expected, sc injection of rhGH suppressed endogenous rGH secretion on d 4 and even more so on d 8. Mechanistically, negative feedback was attributable to a reduction in the amount of rGH released per burst. These data indicate that feedback inhibition is detectable during severe illness in the lagomorph, akin to inferences in the human (9, 27). Infusion of GHRP-2 and TRH stimulated pulsatile GH secretion from d 4 to 8 by the converse mechanism of augmenting GH pulse size and elevating nonpulsatile rGH release. The foregoing dynamics also operate in critically ill patients (27, 28). Reactivation of pulsatile rGH secretion by constant infusion of GHRP-2 and TRH extends the idea that blunted GH secretion in prolonged illness is not due to impaired GH synthesis by somatotropes but, rather, to deficient stimulation by hypothalamic and/or peripheral releasing factors. The significant IGF-I response to rhGH injection and the analogous trend after infusion of GHRP-2 and TRH indicates that target tissues such as the liver regain responsiveness to GH drive in prolonged critical illness, as distinguished from acute critical illness (9, 29). Pulsatile rGH secretion remained elevated after 4 d of GHRP-2+TRH infusion. The latter outcome may be explained by the limited initial IGF-I response with correspondingly lesser feedback inhibition (30, 31). Alternatively, continued hypothalamo-pituitary responsiveness may be relevant as inferred under low dose continuous GHRP stimulation (32).
The constellation of suppressed pulsatile TSH secretion with low serum T4 and T3 and elevated rT3 concentrations characterizes the low T3 syndrome of prolonged critical illness (5). Suppressed TSH secretion contributes to low T4 production and altered peripheral thyroid hormone metabolism depletes T3 availability. In humans, peripheral thyroid hormone metabolism is mediated by three iodothyronine deiodinases: D1, D2, and D3 (33). D1 is present in liver, kidney, and thyroid and plays a key role in the conversion of T4 to T3 and in the catabolism of rT3. D2 is present in brain, pituitary, thyroid, and skeletal muscle and also converts T4 by outer ring deiodination to T3. D3 is present in brain, skin, placenta, pregnant uterus, and fetal tissues and inactivates T4 and T3 by inner ring deiodination to rT3 and 3,3'-diiodothyronine, respectively. We previously showed that the reduced T3 and elevated rT3 concentrations in prolonged critically ill patients are in part due to decreased D1 activity and, hence, suppressed peripheral conversion of T4 to T3 (34). In contrast, D3 activity was higher in critically ill patients, thereby suppressing T3 and augmenting rT3 availability. Some alterations in deiodinase activity may be cytokine induced (35, 36).
In the rabbit model of critical illness with signs of the low T3 syndrome, similar changes in deiodinase activity could be inferred, although we did not include healthy controls in this study. Coinfusion of GHRP-2 and TRH activated the thyroid axis by stimulating TSH secretion, elevating serum total T4 and total T3 concentrations. TSH-dependent secretion of T4 and T3 may have contributed to the latter responses. On the other hand, combined GHRP-2 and TRH stimulation, respectively, augmented and depressed the catalytic activity of hepatic D1 and D3. The observed changes in deiodinase activities may have contributed to enhanced peripheral T4 to T3 conversion in the sick rabbits. Suppression of D3, but not stimulation of D1, also occurred in response to administration of rhGH. This may point to a role for GH, either endogenously released or exogenously administered, in mediating the effect on D3 (37). Such an effect of GH on D3 activity, also shown in healthy chickens (38), would explain the increase in T3 and decrease in rT3 previously observed after treatment with 4 mg/d rhGH for 2 wk in healthy humans (39). Reactivation of hepatic D1 activity, on the contrary, was absent with exogenous GH and thus could be due to GHRP-2, TRH, or the rise in T4 and T3 concentrations (33, 40). The present data do not distinguish among these options. Although an effect of insulin on liver deiodinase activity has been suggested (41), our observation cannot be explained by such an effect, as blood glucose and insulin concentrations were identical in all intervention groups.
In summary, in a rabbit model of critical illness, combined infusion of GHRP-2 and TRH stimulates GH and TSH secretion and elevates T4 and T3 concentrations inferentially by increasing T4 secretion and favoring the hepatic interconversion of T4 to T3. In contradistinction to clinical data, a high dose of rhGH in the lagomorph did not increase short-term mortality after a major burn. Whether the absence of rhGH-induced insulin resistance and hyperglycemia in this species explains the latter outcome remains to be elucidated.
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Acknowledgments |
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Footnotes |
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Abbreviations: AUC, Area under the curve; D1, type 1 deiodinase; D3, type 3 deiodinase; GHRP, GH-releasing peptide; rGH, rabbit GH; rhGH, recombinant human GH; Lv, distribution volume; TT3, total T3; TT4, total T4.
Received August 5, 2003.
Accepted for publication September 30, 2003.
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| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
| Molecular Endocrinology | Recent Prog. Horm. Res. | All Endocrine Journals |
), GHRP-2+TRH (
), and rhGH (
). Day 4 is the first day, and d 8 is the fourth day of intervention. Data are the mean ± SEM.
, Pooled IGF-I concentrations before randomization; 
, P = 0.07 (vs. d 3).
), GHRP-2+TRH (
), and rhGH (
) after 1 (d 4) or 4 (d 8) d of intervention. Data are the mean ± SEM. P < 0.007 denotes the difference between the effects of GHRP-2+TRH compared with those of saline and rhGH.
