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The Journal of Clinical Endocrinology & Metabolism Vol. 86, No. 11 5342-5347
Copyright © 2001 by The Endocrine Society

Endocrine Care

The Effects on Insulin Action in Adult Hypopituitarism of Recombinant Human GH Therapy Individually Titrated for Six Months

E. M. McConnell, A. B. Atkinson, C. Ennis, D. R. Hadden, D. R. McCance, B. Sheridan and P. M. Bell

Sir George E. Clark Metabolic Unit (E.M.M., A.B.A., C.E., D.R.H., D.R.M., P.M.B.) and Regional Endocrine Laboratory (B.S.), Royal Victoria Hospital, Belfast, BT12 6BA, Northern Ireland, United Kingdom

Address all correspondence and requests for reprints to: Dr. P. M. Bell, Sir George E. Clark Metabolic Unit, Royal Victoria Hospital, Belfast BT12 6BA, Northern Ireland, United Kingdom.

Abstract

There is controversy about the effect of replacement GH on insulin action in adult hypopituitary patients. GH replacement calculated from weight leads to unacceptable side effects in some patients. Recent studies suggest it should be individually titrated in adults using serum IGF-I levels. We have assessed the effect of titrated GH replacement on peripheral and hepatic insulin action in 13 adult-onset hypopituitary patients (8 males and 5 females; ages 47 ± 10 yr, mean duration of hypopituitarism 6 yr) with confirmed GH deficiency (GHD; maximum GH <5 mU/liter during insulin induced hypoglycemia), ACTH deficiency, and normal glucose tolerance. All patients were on stable hydrocortisone replacement (15 mg with breakfast, 5 mg with evening meal) for at least 2 months before the trial. Insulin action was assessed by the euglycemic hyperinsulinemic glucose clamp technique (1 mU/kg·min) before and after 6 months of GH therapy. GH was started at 0.8 IU sc daily and titrated monthly until the serum IGF-I increased to within 1–2 SD of the mean of normal age-matched controls. Body mass index did not change significantly during the 6 months of GH therapy. Fasting plasma glucose and HbA1c increased significantly after 6 months (5.2 ± 0.0 vs. 5.5 ± 0.0 mmol/liter, P < 0.0001, and 4.5 ± 0.1 vs. 4.7 ± 0.1%, P < 0.0005, respectively). There was no increase in fasting serum insulin (51.6 ± 10.2 vs. 60.0 ± 10.2 pmol/liter, P = 0.12). Exogenous glucose infusion rates required to maintain euglycemia were similar after GH (23.0 ± 0.4 vs. 21.1 ± 0.3 µmol/kg·min, P = 0.6). Endogenous glucose production in the fasting state was also unchanged following GH (11.8 ± 0.7 vs.12.3 ± 0.9 µmol/kg·min, P = 0.5) and suppressed to a similar extent following insulin (4.4 ± 0.8 vs. 5.5 ± 0.8 µmol/kg·min, P = 0.3). In summary, GH therapy for 6 months, with serum IGF-I maintained in the upper physiological range, increased fasting plasma glucose and HbA1c. There was no effect on peripheral or hepatic insulin sensitivity. Patients receiving GH therapy require long-term monitoring of glucose tolerance.

ADULTS WITH GH deficiency (GHD) have been shown to be insulin resistant compared with matched controls (1, 2, 3). GHD patients have decreased lean body mass and increased fat mass. The increased fat mass, which tends to be distributed in the truncal region (4, 5), may impair insulin sensitivity, with consequent adverse metabolic effects. The etiology of insulin resistance in hypopituitary patients may relate to abnormal body composition and the deficiency of GH or possibly to unphysiological replacement of other pituitary hormones. GH therapy reverses increased fat mass (4, 6, 7, 8), which might be expected to improve insulin sensitivity. It is also recognized, however, that GH in excess has substantial insulin antagonistic effects (9, 10, 11, 12).

Studies examining insulin resistance in GHD patients have used higher doses of GH than currently recommended (13, 14, 15, 16). The initial doses employed were established from the experience of treating GHD children. Invariably, studies using a GH dose based on body weight or surface area required dose reductions due to side effects and have resulted in a serum IGF-I higher than the normal reference range. Some of these studies demonstrated an adverse effect on insulin action (13), whereas others showed no change (14, 15), and one demonstrated an improvement (16). When individually titrated to the serum IGF-I level, lower doses of GH are required and are associated with fewer adverse effects (17, 18, 19, 20).

Because of the controversy surrounding effects of GH therapy on insulin action, we decided to examine the effect of 6 months of low-dose GH therapy, individually titrated to normalize the serum IGF-I concentration, on peripheral and hepatic insulin sensitivity in hypopituitary patients. All patients were receiving the lower, more physiological, total daily steroid replacement of 20 mg hydrocortisone (21, 22, 23, 24).

Subjects and Methods

Subjects

Thirteen patients with GHD and ACTH deficiency for at least 1 yr were recruited from the Royal Victoria Hospital Endocrine Clinic (Table 1Go). GHD and ACTH deficiency were defined as peak GH levels less than 5 mU/liter and peak cortisol levels less than 550 nmol/liter, respectively, during insulin-induced hypoglycemia. The oral dose of hydrocortisone had been changed to 15 mg at 0800 h and 5 mg at 1700 for at least 2 months before the study. Those patients with other concurrent pituitary deficiencies were on stable replacement therapy (13 T4, 12 sex hormone therapy, and 2 desmopressin) that had remained unchanged for at least 1 yr before study entry. No patient had received GH replacement during this time. Patients had acquired pituitary insufficiency (Table 1Go) as a result of surgery for pituitary or peripituitary tumors. Patients were excluded from the study if they exceeded 125% of ideal body weight (Metropolitan Life Insurance tables, 1959), had abnormal glucose tolerance (defined by a 2-h plasma glucose level >7.8 mmol/liter during a 75-g oral glucose tolerance test), were hypertensive, had hepatic or renal disease, or had a history of cardiac or cerebrovascular events.


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  		Table 1. Patient characteristics

 
Study design

Insulin action was assessed first at baseline and then after 6 months of GH therapy. All male patients were on im T replacement (250 mg monthly) except 1 patient, who used Andropatch (SmithKline Beecham, Hertfordshire, UK) 2 times daily. Studies were conducted 7–14 d after an injection of T to reduce the effect that relative T deficiency may have on insulin action. Five females were included in the study, 1 of whom was postmenopausal and not on any hormone replacement therapy. The remaining 4, who were on oral preparations, were studied in the first 10 d of the menstrual cycle. GH (Genotropin, Pharmacia & Upjohn, Inc., Stockholm, Sweden) treatment was commenced at 0.8 IU sc at 2200 h. The dose was then increased at monthly intervals to raise the serum IGF-I concentration into the upper part of the reference range for age-related normal controls.

Approval for the studies was obtained from the Research Ethical Committee of Queen’s University of Belfast (Belfast, UK).

Assessment of insulin action

Insulin action was assessed using the euglycemic glucose clamp technique as described previously by us (11, 25). Patients were admitted to the Metabolic Unit, Royal Victoria Hospital, at 0745 h, after a 12-h overnight fast, with hydrocortisone (15 mg) and other pituitary replacement therapies taken at 0700 h. A plastic cannula (18 gauge; Venflon Viggo, Helsingborg, Sweden) was placed in a left forearm antecubital vein and a blood sample obtained for subsequent use in analysis of plasma glucose specific activities. At 0745 h, a carrier infusion of 0.9% NaCl was connected at a rate of 50 ml per hour. All subsequent infusions were connected to this line.

A dorsal hand vein on the opposite side was cannulated retrogradely (21 gauge; Venflon Viggo), and the hand was placed in a temperature-controlled plexiglass box (Northern Ireland Technology Center, Automation Division, Queen’s University of Belfast) maintained at 55 C to allow intermittent sampling of arterialized venous blood.

Glucose turnover was assessed using a primed continuous infusion of HPLC-purified [3-3H]glucose [NEN Life Science Products Research Products Division, DuPont Ltd., Stevenage, UK (NET100C)] administered during a 2-h equilibration period (-120 min to zero time), and subsequent 2-h continuous (1 mU/kg·min) infusion of insulin (Humulin S; Eli Lilly & Co., Basingstoke, UK). Plasma glucose was maintained at 5.1 mmol/liter by an exogenous glucose infusion (20%). Exogenous glucose was prelabeled with [3-3H]glucose to match the predicted basal plasma glucose specific activity, with the primed continuous tracer infusion being reduced to 50% of the basal rate after 20 min and to 25% of basal the rate after 40 min (to maintain tracer steady state) and was maintained at this rate throughout the remainder of the hyperinsulinemic period.

Analytical techniques

Arterialized venous blood was used for all analyses. Blood samples for determination of plasma glucose specific activities were taken at 10-min intervals from -30 to 0, and from 90 to 120 min, relative to the start of the insulin infusion. Plasma for measurement of glucose specific activity was deproteinized with barium hydroxide and zinc sulfate by the method of Somogyi (26). Samples were counted in a liquid scintillation spectrometer (Tri-Carb 2000 CA, Canberra Packard, Pangbourne, UK). Aliquots of tracer infusate and labeled exogenous glucose infusion were spiked into nonradioactive plasma processed in parallel with plasma samples to allow calculation of [3-3H] glucose infusion rates.

Serum insulin concentration was measured by RIA with insulin antibody precipitate (27), using reagents supplied by Abbott Laboratories (Maidenhead, Berkshire, UK) on an IMX analyzer (Abbott Laboratories, Chicago, IL). The interassay coefficient of variation (CV) was 5.2% at a mean value of 7.3 mU/liter, 3.8% at a mean value of 16.7 mU/liter, and 4.1% at a mean value of 58.4 mU/liter. Serum C-peptide was measured using reagents supplied by Diagnostic Products Corp. (Los Angeles, CA), using an Immulite analyzer (Diagnostic Products). The interassay CV was 6.2% at a mean value of 2.7 µg/liter, 3.8% at a mean value of 6.4 µg/l. Serum cortisol was determined by RIA using reagents supplied by Diagnostic Products Corp. The interassay CV was 3.5% at a mean value of 234 nmol/liter, 3.8% at a mean value of 432 nmol/liter, and 3.6% at a mean value of 981 nmol/liter. Urinary free cortisol was measured using reagents supplied by Orion Diagnostica (Espoo, Finland). The interassay CV was 4.8% at a mean value of 115 nmol/liter, 5.2% at a mean value of 421 nmol/liter, and 3.4% at a mean value of 615 nmol/liter. Serum IGF-I was measured using reagents supplied by Immunodiagnostic Systems Ltd. (Boldon Business Park, Boldon, Tyne, and Wear, UK). Patient samples were incubated with a releasing agent to inactivate binding proteins and were then diluted for assay. This pretreated diluted sample was then incubated, together with horseradish peroxidase-labeled monoclonal anti-IGF-I, in purified sheep polyclonal anti-IGF-I-coated polystyrene microtiter wells for 2 h at room temperature. The wells were washed, and a single-component chromogenic substance was added to develop color. The adsorbance of the stopped reaction mixture was read in a microtiter plate reader, with the color intensity developed being directly proportional to the amount of IGF-I present in the sample. The interassay CV was 6.9% at a mean value of 11.9 nmol/liter and 7.1% at a mean value of 24.4 nmol/liter. Blood samples for lactate and pyruvate were collected in glass tubes containing an equal volume of aqueous perchloric acid solution (8% wt/vol) and immediately shaken. After centrifugation, extracts were separated and analyzed immediately or stored at -20 C until analysis (Sigma, Dorset, UK).

Calculations

Rates of glucose appearance and disappearance were determined during the periods from -30 min to zero time and from 90 to 120 min, using the non-steady state equations of Steele et al. (28) as modified by De Bodo et al. (29), assuming a pool fraction of 0.65 and an extracellular volume of 190 ml/kg. Infusion rates were then calculated as the sum of the tracer infused continuously and the tracer in the labeled exogenous glucose infusion. Rates of endogenous (hepatic) glucose production were then calculated by subtraction of the exogenous glucose infusion rates required to maintain euglycemia from the isotopically determined rates of glucose appearance.

Statistical methods

The power of the study, calculated from previous clamp data (11, 25, 30), gave a 90% chance of detecting a 10% change in insulin action at the 5% level of significance. Significance was assessed with a two-tailed t test for paired data. Significance was taken as P less than 0.05. The values given in the text are presented as means ± SEM. On the morning of the clamp, the area under the curve for serum cortisol was compared over the time interval 0800–1200 h.

Results

Clinical and biochemical parameters in the 13 patients are shown in Table 2Go. During GH treatment, serum IGF-I increased significantly from a baseline of 8.3 nmol/liter to 22.5nmol/liter [P < 0.0001; the laboratory reference range (mean ± SD) was 14 ± 5 nmol/liter for 30- to 40-yr-old adults, 13 ± 4 nmol/liter for 40- to 50-yr-olds, 16 ± 6 nmol/liter for 50- 60-yr-olds, and 14 ± 5 nmol/liter for 60- to 70-yr-old adults]. As can be seen in the age range of our patients, there was no significant change in serum IGF-I with age. Our mean value (22.5 ± 2.2 nmol/liter) was broadly 1–2 SD values above the normal mean concentrations. The mean GH dose employed was 1.4 ± 0.1 IU. The dose was reduced in 1 male patient because of carpal tunnel symptoms, despite the IGF-I level being in the normal range. Body mass index (BMI) did not change significantly over the 6 months (Table 2Go). Fasting arterialized venous plasma glucose levels increased significantly after 6 months of GH replacement (5.2 ± 0.0 vs. 5.5 ± 0.0 mmol/liter, P < 0.0001) as well as HbA1c (4.5 ± 0.1 vs. 4.7 ± 0.1%, P < 0.0005). There was no significant change in fasting serum insulin (51.6 ± 10.2 vs. 60.0 ± 10.2 pmol/liter) or C-peptide levels (0.7 ± 0.1 vs. 0.8± 0.1 nmol/liter) (Table 2Go).


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  		Table 2. Biochemical parameters at baseline and following 6 months of individually titrated GH replacement therapy (n = 13)

 
During the glucose clamp, plasma glucose levels remained constant in both studies (CV range, 2.8–8.1%). The insulin infusion of 1 mU/kg·min led to comparable steady state insulin concentrations before and after 6 months of GH replacement (522 ± 33 vs.508 ± 32 pmol/liter (Fig. 1Go). There was no statistically significant difference in exogenous glucose infusion rates following GH (23.0 ± 0.4 vs. 21.1± 0.3 µmol/kg·min, P = 0.6) (Fig. 1Go). Postabsorptive endogenous glucose production rates were similar in fasting state (11.8 ± 0.7 vs. 12.3 ± 0.9 µmol/kg·min, P = 0.5) and suppressed to the same degree during hyperinsulinemia (4.4 ± 0.8 vs. 5.5 ± 0.8 µmol/kg·min, P = 0.3) (Fig. 2Go). There was no difference in insulin-mediated glucose disposal determined isotopically following GH (27.6 ± 3.4 vs. 27.1 ± 4.2, P = 0.9). Postabsorptive levels of serum nonesterified fatty acids, ß-hydroxybutyrate, and glycerol were also similar in the fasting state and suppressed to the same degree during hyperinsulinemia (Fig. 3Go).



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  		Figure 1. Plasma glucose, insulin, and glucose infusion rates at baseline and following 6 months of individually titrated GH therapy in adult hypopituitary patients (mean ± SEM).

 


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  		Figure 2. Glucose turnover during hyperinsulinemic euglycemic clamps at baseline and following 6 months of individually titrated GH therapy in adult hypopituitary patients (mean ± SEM). No significant differences were observed. EGP, Endogenous glucose production.

 


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  		Figure 3. Plasma NEFA, ß-hydroxybutyrate, and glycerol before and during hyperinsulinemic clamps at baseline and following 6 months individually titrated GH therapy in adult hypopituitary patients (mean ± SEM). No significant differences were observed.

 
There was no significant difference in the area under the curve for serum cortisol levels during the morning of the clamp study, as well as no statistical difference in 24-h urinary free cortisol. Serum-free T4 or serum T3, pyruvate, and lactate were unchanged by 6 months of GH treatment (Table 2Go). There was a trend, however, toward a lower fasting basal lactate following GH, which did not achieve statistical significance (P = 0.07) (Table 2Go). T levels were measured at the initial clamp to ensure that levels were neither supra- nor subphysiological. These values ranged from 11.9 to 30.6 nmol/liter (normal range, 10–30 nmol/liter). Repeat studies were again performed within 7–14 d of injection.

Discussion

It is now accepted that GH doses in adults should be adjusted so as to maintain serum IGF-I levels in the upper range for age-matched controls. Until now, no study has examined in detail the effect of these lower doses on insulin resistance (both peripheral and hepatic). The mean GH dose employed in this study (1.4 IU/d) is similar to others in which this method of dose titration has been used (18, 19, 20). We noted, as have other studies (19), that females had lower serum IGF-I levels before commencing GH (mean 5.3 vs. 11.0 nmol/liter, P = 0.04), but there was no difference in mean GH doses used between females and males (mean 1.6 IU/d vs. 1.2 IU/d, P = 0.1). The small but significant increase in fasting glucose (0.3 mmol/liter) is of the same magnitude as that reported by Johannsson et al. (19) using a similar regimen. In addition, and in contrast to other studies (13, 14, 15, 16, 31), we demonstrated a significant increase in HbA1c. However, fasting insulin and C-peptide levels were not significantly increased after 6 months of GH therapy compared with our previous study (8) and similar studies (2, 4, 13, 14, 15, 16, 19, 32), which used a GH dose based on patients’ weight. Steady-state C-peptide levels did not change significantly over the 6 months (Table 2Go). It is unclear whether this is a reflection of the lower doses of GH used in this study or of the time period over which the study was conducted.

Our patients were insulin resistant before commencing GH with a reduced glucose infusion rate of 23.0 µmol/kg·min compared with approximately 40 µmol/kg·min seen in healthy controls of similar age (33, 34). This was despite the fact that our patients were on the lower doses of steroid replacement therapy currently recommended. In the present study, there was no increase in peripheral or hepatic insulin resistance after 6 months of carefully titrated GH replacement therapy. There was no change in fasting hepatic glucose production despite a significant increase in fasting plasma glucose, and this may possibly reflect a type-2 error. However, our study was powered (90% chance) to pick up a 10% difference at the 5% level of significance. Our results are in contrast to Christopher et al. (13), who demonstrated a suppression of hepatic glucose production with no change in peripheral insulin sensitivity (Table 3Go). However, they are similar to those of Fowelin et al. (14) and O’Neal et al. (15), who demonstrated a temporary state of insulin resistance shortly after commencing GH, which returned to baseline after 6 months of GH therapy (Table 3Go). Unfortunately, their methods did not allow the assessment of hepatic insulin sensitivity. Hwu et al. (16), in complete contrast, demonstrated an improvement in peripheral insulin sensitivity to levels similar to a control group, whereas Weaver et al. (4) demonstrated a deterioration in insulin sensitivity. All of the above groups employed larger doses of GH based on weight (ranging from 1.8 to 6.6 IU daily) than currently recommended (Table 3Go). The euglycemic hyperinsulinemic glucose clamp is accepted as the gold standard in assessing insulin sensitivity. When combined with isotope dilution methodology, it enables calculation of the rates of glucose production and an assessment of the contribution of peripheral and hepatic insulin sensitivity. Christopher et al. (13) are the only group to use this method in adult onset patients, albeit using weight-based GH doses.


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  		Table 3. The effects of GH on insulin action in hypopituitary patients in five trials

 
The insulin antagonistic effect of GH is thought to be partly due to its lipolytic effect leading to increased use of lipid intermediates and decreased peripheral glucose uptake (35). With the lower doses of GH used in our study, there was no change in nonesterified fatty acids (NEFA), ß-hydroxybutyrate, or glycerol, suggesting no major effect on lipoplysis, whereas GH doses based on weight resulted in raised NEFA (14).

Other standard pituitary replacement therapies may contribute to the development of insulin resistance in panhypopituitary patients. Euthyroidism was maintained throughout the study on the thyroid hormone replacement used and consequently should not have exerted any effect on insulin action (36, 37, 38). All males were studied between 7 and 14 d after an injection of T, avoiding relative T deficiency, another factor influencing insulin resistance (39). Likewise, females were studied between d 1 and 10 of the menstrual cycle, and all except one postmenopausal lady (patient 8, age 60 yr) were receiving stable low-dose oral cyclical hormone replacement therapy, which has no effect on insulin sensitivity (40). No females were receiving transdermal E. Recent evidence demonstrates that the routine steroid replacement therapy previously used may have been excessive (21, 22, 23). In addition, cortisol induces hyperinsulinemia (41, 42, 43) and insulin resistance and may contribute to increased insulin resistance in GHD patients. Previous studies examining insulin resistance have included some patients with multiple pituitary hormone deficiencies on larger doses of steroid replacement therapy (14, 15). We have demonstrated that hypopituitary patients are not more insulin resistant when receiving a total daily hydrocortisone dose of 20 mg, compared with an iv infusion simulating normal serum cortisol concentrations (24). Therefore, the dose of steroid replacement therapy in this study should not have caused insulin resistance before commencing GH. During the study, measurements of cortisol replacement were similar before and after GH therapy.

In conclusion, this study demonstrates that the use of low-dose individually titrated GH may prevent the development of peripheral and hepatic insulin resistance, as seen in other studies using weight-based doses. The study was conducted over 6 months, and it remains to be seen if longer-term low-dose GH, by promoting favorable changes in body composition, may offset or eventually reverse the insulin resistance of GHD. If insulin sensitivity does not improve in the long term following GH replacement, it may be that other standard replacement therapies should be re-examined to assess their impact. Certainly, in the first week following im T replacement, T levels can be supraphysiological and subphysiological before the next injection. T patches may provide some resolution to this problem, but problems with application have not led to a high take-up rate among our patients. There was, however, a small but significant rise in fasting glucose and hemoglobin A1c, and therefore, glucose tolerance should be monitored long term.

Acknowledgments

We are grateful to Dr. C. Patterson (Department of Community Medicine and Medical Statistics, Queen’s University of Belfast) for statistical advice.

Footnotes

During these studies, Dr. M. McConnell was a Royal Victoria Hospital research fellow. Pharmacia & Upjohn, Inc. also provided financial support.

Abbreviations: BMI, Body mass index; CV, coefficient of variation; GHD, GH deficiency; NEFA, nonesterified fatty acids.

Received May 16, 2000.

Accepted July 18, 2001.

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