Endocrinology Vol. 139, No. 11 4466-4469
Copyright © 1998 by The Endocrine Society
Plasma Leptin Turnover Rates in Lean and Obese Zucker Rats1
R. Vilà,
C. Adán,
I. Rafecas,
J. A. Fernández-López,
X. Remesar and
M. Alemany
Departament de Bioquímica i Biologia Molecular, Facultat de
Biologia, Universitat de Barcelona, 08028 Barcelona, Spain
Address all correspondence and requests for reprints to: Prof. Dr. Marià Alemany, Departament de Bioquímica i Biologia Molecular, Facultat de Biologia, Universitat de Barcelona, Avinguda Diagonal 645, 08028 Barcelona, Spain. E-mail:
alemany{at}porthos.bio.ub.es
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Abstract
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Conscious female adult lean and obese Zucker rats were injected through
the jugular vein with radioactive iodine-labeled murine leptin; in the
ensuing 8 min, four blood samples were sequentially extracted from the
carotid artery. The samples were used in a modified RIA for leptin, in
which paired tubes received the same amount of either labeled or
unlabeled leptin, thus allowing us to estimate both leptin levels and
specific radioactivity. The data were used to determine the decay curve
parameters from which the half-life of leptin (5.46 ± 0.23 min
for lean rats and 6.99 ± 0.75 min for obese rats) as well as the
size of its circulating pool (32 pmol/kg for lean rats and 267 pmol/kg
for obese rats) and the overall degradation rate (96 fkat/kg for
lean rats and 645 fkat/kg for obese rats) were estimated. These
values are consistent with the hormonal role of leptin and the need for
speedy changes in its levels in response to metabolic challenge.
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Introduction
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LEPTIN, the product of the ob
gene (1), is a key factor in the regulation of body weight. Its precise
role, however, has been subject to open controversy and discussion (2)
despite earlier claims that its main role was that of a ponderostat
signal, able in itself to modify the energy partition in some strains
of rodents (3). A large amount of research has been carried out to
establish the functions of leptin (4, 5) since its discovery in 1994
(1), but our knowledge of its in vivo dynamics is sketchy,
in part because of methodological difficulty and in part because of the
priority given to eventual pharmacological use and the need to
characterize the mechanisms regulating its expression and signaling
role.
The kinetic analysis of circulating hormones is a good source of
information about the theoretical speed of response to change as well
as on the demand for synthesis and/or degradation that the living
organism devotes to the maintenance of a fully functional and
responsive system. Short half-lives represent a higher energy
expenditure but allow for faster responses and more immediate
adaptation; longer turnover rates are usually correlated with longer
term regulatory activity. Leptin turnover has been estimated in humans
(half-life of about 25 min) (6) using arterio-venous differences and in
mice (half-lives of 1.5–3 h) (7, 8) by measuring the decay of
circulating label after the injection of labeled leptin.
Our method for the estimation of circulating peptide hormone turnover
rates in vivo, which we have applied to insulin (9), has the
advantages that only the label in the immunoreactive peptide hormone is
taken into account, and all the measurements are performed in a single
animal.
We chose the hyperleptinemic Zucker fa/fa rat, because of
overexpression of the ob gene (10), to test whether this
overexpression affects the turnover rate of leptin. The main objective,
however, was to determine the range of the half-life of circulating
leptin to elucidate the predominance of its short or long term
metabolic actions.
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Materials and Methods
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Materials and animals
Pure recombinant murine leptin (Biotrend, Köln, Germany)
and 125I-labeled murine leptin (specific radioactivity,
69.2 GBq/µmol; Anawa, Zurich, Switzerland) were used. The labeled
leptin used has a chromatographic purity of 98.6%.
Zucker lean (Fa/?) and obese (fa/fa) female adult rats,
weighing 224 ± 4 and 398 ± 19 g, respectively, bred at
the Animal Service of the University of Barcelona (Barcelona, Spain)
from heterozygous stock obtained from Charles River Laboratories,
Inc. (Wilmington, MA), were used. The animals were housed in
individual polypropylene-bottomed cages under standard conditions
(lights on from 0800–2000 h; 22–23 C; 70–75% relative humidity),
and were fed standard chow pellets (type A04, Panlab, Barcelona,
Spain). A series of four rats of each phenotype was cannulated, under
ethyl ether anesthesia, in the left carotid artery (bringing the tip of
the cannula just to the heart) with P-50 and in the right jugular vein
with P-10 polyethylene tubes (Clay-Adams, Parsippany, NJ). The cannulas
were filled with heparinized saline and sealed; they were threaded sc,
exiting the rat through the back, where they were coiled and held in
place with surgical tape. The rats were used for the experiments 5
h later. At the end of the experiment, the rats were killed by
decapitation. This study was conducted in accordance with European
Community principles, guidelines, and procedures for animal
experimentation.
Leptin turnover measurement
The experiment was begun (6 h after the beginning of the light
cycle) by injecting each rat (within 5–8 sec) through the jugular
venous cannula with 80 kBq (1.16 pmol) labeled leptin in 0.2 ml
isotonic saline solution. The radioactivity initially present (and that
remaining after the injection) in the syringe was measured with a
-counter. At timed intervals of 1, 2, 5, and 8 min, aliquots of 0.4
ml blood were extracted through the carotid cannula and stored at 4 C
in heparinized plastic vials. The blood samples were immediately
centrifuged at 4 C to separate the plasma samples, which were used
directly for labeled leptin estimation and leptin turnover according to
our method for insulin turnover (9) modified for use with leptin. The
rat was maintained conscious and unaware of the manipulations (except
for the uncoiling of the cannulas), as it remained in its cage out of
sight of the researchers throughout the experiment.
Plasma leptin levels were estimated using a standard RIA procedure
(Linco Research, Inc., St. Charles, MO) with some
modifications: each plasma sample was distributed in two tubes; in the
first, in addition to the plasma sample [100 µl (lean) or 40 µl
(obese)], 100–160 µl buffer, 100 µl 125I-labeled
leptin solution containing 0.43 kBq (i.e. 6.25 fmol), and
100 µl diluted specific leptin antibody (Linco Research)
were added. In the remaining tube, the plasma samples [100 µl (lean)
or 40 µl (obese)] received 100–160 µl buffer, 100 µl unlabeled
murine leptin (6.25 fmol), and 100 µl specific leptin antibody. Thus,
the second tube finally contained the same amount of leptin as the
first (the amount initially present and that added were the same as
those in the first tube), but the amount of labeled leptin present in
either was different; the second tube lacked the added labeled leptin
used for the standard RIA procedure. Since the total amount of leptin
was the same in both series of tubes, the labeled leptin initially
present in plasma bound in the same proportion to the antibody
preparation; thus, the second tube could be used as a blank for the
first as in a standard RIA procedure. This allowed estimation of the
apparent leptin concentration regardless of the amount and distribution
in molecular species of radioactivity initially present in the plasma.
This approach circumvented the problems posed by the presence of
radioactive sources (leptin and other) in the samples. The RIA was
completed with a series of standards of murine leptin, blanks, and
several tubes for the estimation of nonspecific binding. All
measurements were carried out in duplicate for each animal and time
point.
Calculations
The leptin label present in a given sample of plasma was
estimated assuming that labeled leptin was bound by the antibody in the
same proportion as unlabeled leptin from the same source. From the RIA
data, a plot of leptin bound to the antibody vs. the
concentration of leptin in the tube was drawn using murine leptin as a
standard. The data were fitted to an asymmetric sigmoid curve using the
FiG-P program (Biosoft, Cambridge, UK); the calculated
parameters of the curve were used to estimate the percentage of leptin
bound to each of the blood samples obtained in the experiment. This
percentage also reflected the proportion of leptin radioactivity bound
to the antibody; thus, the total amount of leptin radioactivity,
rt, per ml blood at a given time, t, was established for
each sample. The rt values were plotted against time t and
fitted to a standard decay graph using the FiG-P program:
rt = r0 x e-K x dt, from
which, K, the decay constant, and r0, the initial
radioactivity per ml blood, were obtained. The half-life,
t1/2, of leptin was calculated from K, since
t1/2 = 1/K. The ratio of total radioactivity injected,
R0/r0, was used to establish the volume, V, of
distribution of the injected label (i.e. the virtual or
practical leptin space): V = R0/r0. The
content of leptin was calculated from total leptin radioactivity and
the specific activity of the labeled leptin injected. Injected leptin
was a maximum of 16% (lean) to 1% (obese) of the total body leptin.
As this proportion was small, its influence on circulating leptin was
minimal. The rat leptin concentration in plasma did not vary during the
experiment. The concentration vs. time graphs were used to
obtain a mean leptin concentration value, lM,
and to check whether there were significant variations in leptin
concentration. As we knew both the virtual distribution volume and the
concentrations, we could derive the whole mass of circulating leptin
L0 at time zero: L0 = lM
x V. The rate of loss of leptin (rate of degradation, 
) from this
circulating pool could be derived from the decay curve and the mass of
leptin: = K x L0. Indeed, as the virtual
distribution volume, V, did not change, the leptin mass at a given
time, Lt, can be estimated from the plasma concentrations,
lt, and the degradation rates for different
times, t, may be calculated. The values obtained in all
cases were very similar, because the changes in leptin concentration
during the 8-min analysis were insignificant.
The loss of radioactivity from the labeled leptin pool was studied by
establishing the total leptin label values, Rt, at a given
time from the radioactivity per ml plasma and the virtual volume of
distribution: Rt = rt x V.
Statistical comparison between groups was established with standard
ANOVA programs and Student’s t test.
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Results
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Plasma leptin levels were maintained within a maximal range of
variation of 6 ± 2% (lean) and 20 ± 7% (obese) of the
initial values in the time elapsed between the injection and the last
blood extraction. The significances of the effect of time on leptin
concentrations were P = 0.968 (lean) and
P = 0.250 (obese; by ANOVA).
Figure 1
presents the decay curves for
plasma leptin radioactivity vs. time. Lean and obese rats
showed a similar pattern over time. The differences between both series
of animals were significant, as was the effect of time. These decay
curves were used for the calculation of leptin space, turnover rates,
and cleavage, shown in Table 1. Leptin
levels were higher in the obese rats than in the lean rats. The
distribution space of leptin was larger for the larger obese rats, but
was comparable for both groups when the data were corrected for body
size despite the smaller relative lean body mass of the obese rats.
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Figure 1. Leptin radioactivity decay with time in the plasma
of Zucker lean and obese rats injected with labeled leptin. The data
are the mean ± SEM of four or five different animals
and are expressed as the fraction of the injected radioactivity
remaining in the whole circulating leptin pool at a given time.
Statistical analysis of the differences between groups (by ANOVA): lean
vs. obese, P = 0.004; effect of time
on lean, P = 0.000; effect of time on obese,
P = 0.000.
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Obese rats had a higher leptin mass than lean controls; this was
maintained even after correcting for body size. The half-life of leptin
was in the same range for lean and obese rats; the latter showed
somewhat longer (
28%) half-lives. The calculated leptin degradation
rates were almost 7-fold higher in obese rats than in the lean (per U
body mass).
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Discussion
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The method used here has been previously successfully used for the
analysis of insulin turnover in lean and obese rats (9). It is
conceptually simple, but requires a careful development, especially at
the critical point of evaluation of leptin radioactivity in the
samples. The injection of labeled leptin did not significantly affect
the mass of circulating leptin. The levels of leptin did not change as
a consequence of the injection of labeled leptin and remained uniform
in all of the blood extractions. An additional advantage of this method
was its use of relatively undisturbed animals.
A critical point in the investigation of leptin cleavage is the
assignment of radioactivity measurements to intact (i.e.
functional) leptin, without interference by free iodine or other
labeled peptide fragments eventually freed by the cleavage of leptin.
Leptin turnover estimations calculated from the decay of label in the
blood (7, 8) tend to give longer half-lives. The method used here is
not affected by this interference, as only the label bound to leptin is
measured; free iodine and labeled peptides are removed during the RIA
procedure. Only labeled (complete) leptin is bound to the antibody, and
thus only this radioactive molecular species is taken into account.
The main difficulty that may arise from a study based on calculated
constants taken from calculated values and used to derive the final
results is a cumulative effect of residuals in calculations, which may
lead to errors. This study has been designed to minimize this effect.
The adequacy of decay curve fitting is apparent in Fig. 1
, which shows
the loss of radioactive leptin in plasma. The low dispersion of data
suggests an acceptable degree of precision in the derivation of the
virtual volume of diffusion and decay rates shown in Table 1.
The virtual volume of distribution of leptin derived from data of
arterial blood may not be real, because we do not know whether the
leptin levels are representative; nevertheless, the data are useful and
admit comparison between different animals and situations. The leptin
pool size, Lt, is more reliable, because it is derived
directly from decay curves and leptin levels. Leptin pool size was much
higher in obese than in lean rats both in absolute terms and in
relation to body weight; in the latter case, the differences
diminished, probably because of the dilution effect of the large fat
mass in obese rats. The relative uniformity of the virtual volume of
distribution vs. body weight in lean and obese rats
suggested that the distributions of leptin are comparable in lean and
fat tissues. As a consequence, the comparisons between both groups were
directly referred to body size to establish comparisons despite their
different body weights.
The turnover rates found here are much shorter than those found in
normal mice (3 h) (7) or in db/db and ob/ob mice
(1.5 h) (8) calculated from label decay curves, and they are also
shorter than those estimated in humans from arterio-venous differences
(25 min) (6). These differences may be a consequence of the methodology
applied; in our case, the circulating levels of leptin were not
modified, and only leptin label was taken into account. The timing of
sample extraction was also in the range of the measured turnover, which
gave higher precision to our estimations.
The Zucker fa/fa rats are hyperleptinemic, but this leptin
is inoperative because they lack a functional hypothalamic leptin
receptor (10, 11). This deficiency has been postulated as the cause of
their obesity, because it deprives the rat of a key element in the
control of energy partition (12). The white adipose tissue of
fa/fa rats overexpresses the ob gene, thus
inducing massive synthesis of leptin (10). The synthesis of leptin in
Zucker obese rats is further enhanced by their large fat mass, as
leptin production has been found to be related to adipose tissue mass
in humans and rodents (13, 14). However, the half-life of leptin is in
a similar range in lean and obese rats; the latter show a mere 28%
higher mean rate. As the leptin pool size is much larger in the obese
rats, the maintenance of similar turnover rates implies a more active
degrading process in these animals than in the controls. We have no
clear idea where this leptin is degraded. Some reports suggest that the
kidney plays a significant role in the clearance of excess leptin from
the blood (15, 16). Lean and obese Zucker rats do not show different
kidney blood flows (our unpublished results), but the much
higher concentration of leptin in the blood of the obese rats may be a
key factor facilitating its removal; the mean ratio of plasma
concentrations (obese/lean) is 4.12, and that of the pool removal rates
is 6.17 (3.49 when corrected for body size).
As the levels of leptin in undisturbed animals are uniform, it may be
assumed that the removal rate is essentially identical to the synthesis
rate of leptin. This means that a 225-g lean rat synthesizes (and
degrades) about 30 µg leptin/day (i.e. 260-fold the whole
body leptin mass), much less (a mere 8%) than that of a 400-g obese
rat (358 µg/day, i.e. 210-fold the leptin mass).
The differences between lean and obese rats found for leptin were
fairly similar to those for insulin in this same animal model of
obesity. Zucker fa/fa rats are hyperinsulinemic (17) and
show insulin turnover rates similar to those of controls (9). The range
of t1/2 was also similar between leptin and insulin (only a
few minutes). This value is consistent with the hormonal role of leptin
and the eventual need for speedy changes in its levels as a response to
metabolic challenge, such as that found under starvation (18), diet
(19, 20), or hormonal manipulation (21), and may help explain the
ultradian rhythms of this hormone (22).
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Acknowledgments
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Thanks are given to Robin Rycroft from the Language Advisory
Service at the University of Barcelona for correction of the text.
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Footnotes
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1 This work was supported by Laboratoris SALVAT, SA, as well as by
grants from the CIRIT of the Government of Catalonia and the Plan
Nacional de Ciencia de los Alimentos (ALI96–1094) of the Government of
Spain. Work was carried out within the frame of the EC Network on
Metabolic Integration and Energy Control ERBCHRX-CT94–0490. 
Received March 18, 1998.
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Top
Abstract
Introduction
