Section of Islet Transplantation and Cell Biology, Research Division, Joslin Diabetes Center, Boston, Massachusetts 02215
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ABSTRACT |
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Although type 2 diabetes mellitus
is associated with insulin resistance, many individuals compensate by
increasing insulin secretion. Putative mechanisms underlying this
compensation were assessed in the present study by use of 4-day glucose
(GLC; 35% Glc, 2 ml/h) and lipid (LIH; 10% Intralipid + 20 U/ml
heparin; 2 ml/h) infusions to rats. Within 2 days of beginning the
infusion of either lipid or glucose, plasma glucose profiles were
normalized (relative to saline-infused control rats; SAL; 0.45% 2 ml/h). During glucose infusion, plasma glucose was maintained in the normal range by an approximately twofold increase in plasma insulin and
an ~80% increase in -cell mass. During LIH infusion, glucose profiles were also maintained in the normal range. Plasma insulin responses during feeding were doubled, and
-cell mass increased 54%. For both groups, the increase in
-cell mass was associated with increased
-cell proliferation (98% increase during GLC and 125% increase during LIH). At the end of the 4-day infusions, no
significant changes were observed in islet-specific gene transcription (i.e., the expression of islet hormone genes, glucose metabolism genes,
and insulin transcription factors were unaffected). Two days after
termination of the infusions, the glucose-stimulated plasma insulin
response was increased ~67% in glucose-infused animals. No sustained
effect on insulin secretory capacity was observed in the LIH animals.
The increase in plasma insulin response after glucose infusion was
achieved in the absence of any change in insulin clearance. We conclude
that, in rats, an increase in insulin demand after an increase in
glucose appearance or free fatty acid leads to an increase in
-cell
mass, mediated in part by an increase in
-cell proliferation, and
that these compensatory changes lead to increased insulin secretion,
normal plasma glucose levels, and the maintenance of normal islet gene expression.
insulin resistance; insulin secretion; -cell; mitosis; maturity
onset diabetes of the young
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INTRODUCTION |
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ALTHOUGH TYPE
2 DIABETES MELLITUS is associated with insulin
resistance (11), not all insulin-resistant subjects
develop diabetes; rather, many individuals compensate for the
resistance by increasing insulin secretion (22, 23,
36). Although the increased insulin response protects
individuals from developing diabetes, very little is known about the
cellular or molecular mechanisms underlying the adaptive change. An
increase in insulin secretory capacity can result from either an
increase in the ability of existing -cell mass to manufacture and
secrete insulin or from an increase in the
-cell mass per se
(6). An increase in the secretory capacity of the
existing
-cell mass most likely involves changes in the rates of key
steps in the
-cell stimulus/secretion pathway. An increase in
-cell mass can result from either hypertrophy or hyperplasia, with
hyperplasia resulting from an increase in
-cell replication
(mitosis), a decrease in
-cell death (apoptosis), or the
formation of new
-cells from precursor cells (neogenesis) that
reside in pancreatic ducts (14).
One reason for our lack of understanding of the adaptive -cell
response is the lack of an established animal model in which to
investigate in vivo the temporal relationships between changes in
insulin secretion, expansion of
-cell mass, and alterations in
islet-specific gene expression. Possible animal models include the
glucose- or lipid-infused rat. It has long been known that
-cell
mass increases during glucose infusion (49) and that the
increase is due to both hypertrophy and hyperplasia (7). However, although glucose infusion increases
-cell mass, it is unclear whether the increase leads to an increase in insulin secretory capacity. During glucose infusion, plasma insulin levels increase within 1-2 days to reestablish euglycemia (7). The
increase in insulin secretion at the renormalized glucose level is
thought to be due to an increase in the catalytic rate of glucokinase (GK) (9), GK being rate limiting for glucose entry into
the
-cell (38). However, despite the increase in both
-cell mass and GK activity, studies in the in vitro perfused
pancreas have consistently indicated that the glucose-stimulated
insulin response is decreased, rather than increased, after glucose
infusion (28), leading to the belief that the model
represents
-cell exhaustion or glucotoxicity. Nonetheless, although
the in vitro insulin response is reduced, one study has reported an
increase in insulin secretion when the response is assessed in vivo
(25), suggesting that the model may be appropriate for the
study of
-cell adaptation.
Free fatty acids (FFA) have also been implicated in altering -cell
mass and insulin secretion. In vitro, short-term elevations in FFA
increase
-cell proliferation in islets isolated from Wistar rats
(39) but not in islets isolated from Zucker diabetic fatty rats (19). However, whereas an elevation in FFA may
initiate a hyperplasia response, prolonged exposure has been shown to
increase
-cell apoptosis (43). Thus the net
effect on in vivo
-cell mass, which is determined by a balance of
mitosis, apoptosis, and islet neogenesis (14), is
unclear. Functionally, short-term increases in FFA augment in vivo
glucose-stimulated insulin secretion (12, 44), whereas,
with one exception (34), longer-term elevations decrease
the response (33, 37). Thus it is also unclear whether
elevations in FFA lead to a compensatory increase in
-cell mass or
function (adaptation) or to a failure of the existing
-cells (lipotoxicity).
Ultimately, -cell adaptation, be it a change in
-cell mass or
secretory capacity per unit mass, is likely to be under the control of
key regulatory genes, of which there are many candidates. All known
genes leading to maturity-onset diabetes of the young [the "MODY"
genes: hepatic nuclear factors (HNF)4-
, HNF1-
, and HNF1-
,
pancreatic duodenum homeobox protein (PDX)-1, and GK] have been
implicated in defective insulin secretion, and at least two of these
(HNF4-
and PDX-1) have been reported to be affected by FFA
(16, 18). Knockout studies in mice have shown that PDX-1
is essential for pancreas development (21) and that the basic helix-loop-helix factor NeuroD/BETA2 is required for
-cell development (41). PDX-1, NeuroD/BETA2, and NeuroD/BETA2's
dimeric partner pan1 all play a postdevelopmental role in regulating
insulin gene transcription (17). The link between these
genes and
-cell development and/or function suggests that any or all
of them may be instrumental during
-cell adaptation.
The present study was conducted to determine whether rat -cells can
adapt to the increase in insulin demand during glucose or lipid
infusion, and if so, to determine what physiological and molecular
mechanisms underlie the adaptation. Results indicated that within
1-2 days of either glucose or lipid infusion, rats renormalize
their plasma glucose profiles by dramatically increasing their plasma
insulin responses. The increased insulin responses were associated with
increases in
-cell mass, which were paralleled by increased rates of
-cell proliferation. At the end of the 4-day substrate infusions,
islet gene expression was completely normal. Two days after the
infusions were concluded, insulin secretory capacity remained elevated
in glucose-infused animals but returned to normal in lipid-infused animals.
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METHODS |
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Substrate infusions.
Male Sprague-Dawley rats (~250 g) with indwelling jugular vein and
carotid artery catheters were obtained from Taconic Farms (Germantown,
NY). After arrival at the animal facility, rats were observed for 3
days to ensure general well-being as assessed by weight gain and
appearance. Rats were then randomly divided into four groups: a glucose
infusion group (GLC; 35% hydrated dextrose; McGaw, Irvine, CA), a
lipid infusion group (LIH; 10% Intralipid + 20 U/ml heparin;
Baxter iv), a saline-infused control group (SAL; 0.45%), and a
noninfused nonrestrained control group (NON). Substrates were infused
into the carotid artery at 2 ml/h for 96 h beginning at ~9:30
AM. The infusion system consisted of a swivel and saddle (Lomir
Biomedical, Malone, NY) designed to allow the animal unrestricted
mobility. Blood samples (~0.1 ml) were drawn from the jugular vein at
0.5, 0 (before start of infusions), 0.1, 0.5, 1, 4, 6, 12, 24, 36, 48, 60, 72, 84, and 96 h, placed in heparin-coated tubes, and
immediately centrifuged. The samples at 12, 36, 60, and 84 h were
obtained during the time rats normally eat (~10 o'clock in the
evening). For all samples, a portion of the supernatant (10 µl) was
immediately assessed for glucose (Beckman Glucose Analyzer II; Beckman,
Brea, CA), and the remainder (~40 µl) was frozen and stored at
20°C for later determination of plasma insulin concentration.
Additional samples (~0.1 ml) were taken at
0.5, 1, 24, 48, 72, and
96 h for the determination of plasma FFA levels; these samples
were taken in tubes coated with EDTA (diethyl p-nitrophenyl
phosphate) and paraoxon to prevent lipoprotein lipase-stimulated
breakdown of triglycerides (50). Rats were allowed ad
libitum food and water during the 4-day infusions, and a standard
(12:12-h light-dark) cycle was maintained. At the end of the infusion
period, animals were removed from the restraint system, returned to
their cages, and eventually euthanized for morphological assessment of
-cell mass and proliferation or for islet isolation and
semiquantitative RT-PCR analysis of islet gene expression. After return
to their cages, animals were given 2 days to renormalize plasma glucose and insulin levels; then the animals were used to assess postinfusion glucose-stimulated plasma insulin responses (hyperglycemic clamp) or
peripheral insulin clearance (euglycemic-hyperinsulinemic clamp). Infusions for each of the four measures (
-cell mass, islet gene expression, insulin response, and insulin clearance) were randomized by
group (NON, SAL, GLC, LIH). Rats removed from the study due to
technical problems were replaced, and the infusion process was
rerandomized; the rerandomization resulted in some groups having a
higher number of experiments. Because the number of infusions was four
times greater than the number of postinfusion tests (assessment of
-cell mass, islet gene expression, insulin secretion, and insulin
clearance each requires a separately infused animal), the insulin and
FFA levels during the infusions were not assessed in all animals (the
total number of infused animals in each group was n = 32, SAL; n = 37, GLC, and n = 33, LIH)
but rather in a random subset of 6-9 animals, as reported in the
text and Figs. 1-4 of RESULTS.
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Hyperglycemic clamps.
Hyperglycemic clamps (n = 7 NON; n = 6 SAL; n = 9 GLC; n = 6 LIH) were
performed in conscious unrestrained fed animals. For each clamp, two
basal blood samples (~0.1 ml each; at 30 and
5 min) were taken in
heparin-coated tubes and immediately centrifuged. As described in
Substrate infusions, 10 µl of supernatant were assessed
for plasma glucose, with the remainder immediately frozen and stored at
20°C for later determination of insulin and C-peptide concentrations. A priming bolus of 20% hydrated glucose was then calculated on the basis of the measured basal glucose level, an assumed
glucose distribution space of 25% body weight, and a desired clamp set
point of 250 mg/dl. The priming bolus was given at 0 min into the
carotid artery, together with an initial exogenous glucose infusion of
6 mg/min. Subsequent blood samples (each ~0.1 ml) were then taken at
1, 3, 5, 10, 15, 20, 25, 30, 40, 50, 60, 70, 80, and 90 min, and the
exogenous glucose infusion was adjusted in proportion to the difference
between the measured glucose and the set point and the rate of change
of this difference (integrated proportional-derivative control).
Insulin secretory capacity was assessed as the average plasma insulin
concentration during the final 0.5 h of the clamp (60-90
min). Hematocrit was assessed at the beginning and end of clamps in a
subset of animals and typically decreased from ~35 to ~32%.
Hyperinsulinemic clamps.
Increased plasma insulin levels can be due to either an increase in
insulin secretion or a decrease in insulin clearance. Both glucose
(26) and lipid infusions (48) have been shown to decrease insulin clearance. Although C-peptide might be considered a
surrogate of insulin secretion, the clearance of this peptide has not
been shown to be constant during chronic infusion studies. As well, the
rat has two insulin/C-peptide genes, preventing the assessment of
prehepatic insulin secretion with traditional C-peptide deconvolution.
Because C-peptide could not be used to correct for changes in insulin
clearance, the clearance of insulin was directly assessed with
euglycemic-hyperinsulinemic clamps. These clamps (n = 6 for all groups) were performed in an identical manner to the
hyperglycemic clamps, with the exception that glucose was clamped at
basal with a variable glucose (20% hydrated glucose) infusion that was
co-infused with human insulin (Eli Lilly, Indianapolis, IN; 4.2 mU · min1 · kg
1, 0.032 ml/min) through a small Y-connector. The variable glucose infusion was
also calculated on the difference, and rate of change of the
difference, between plasma glucose and the set point, in this case
basal. Peripheral insulin clearance was calculated as the insulin
infusion rate
(µU · min
1 · kg
1) divided
by the average insulin concentration during the final 0.5 h of the
clamp (60-90 min). Changes in hematocrit were similar to those
observed during hyperglycemic clamps.
Quantification of -cell mass and mitotic rate.
At ~6 AM on the 4th day of substrate infusion, rats
(n = 6 for all groups) were lightly anesthetized with
pentobarbital sodium (Nembutal; ~0.05 ml, 50 mg/ml into the jugular
vein) and given an intraperitoneal injection of BrdU (100 mg/kg
crystalline grade 5-bromo-2'-deoxyuridine, diluted in PBS to 10 mg/ml).
Rats typically recovered from the anesthesia/BrdU injection within
5-10 min. Heparin was discontinued for ~8 h before the BrdU
injection to prevent internal bleeding at the injection site and was
restarted immediately after the injection. Exactly 6 h after the
BrdU injection, a second nonlethal dose of pentobarbital sodium
(0.1-0.2 ml, 50 mg/ml into the jugular vein) was administered, and
the pancreas was rapidly removed. The animal was then killed by
pentobarbital sodium overdose. The pancreas was cleaned of lymph nodes
and fat, divided into head and tail portions, weighed, and fixed in
Bouin's solution. The head and tail portions of the pancreas were
embedded in paraffin, sectioned, and double-stained for BrdU (cell
proliferation kit; Amersham; Arlington Heights, IL) and
non-
-endocrine cells (by use of antibodies against glucagon,
somatostatin, and pancreatic polypeptide as described in Ref.
40). Total
-cell mass and percent
-cell
proliferation were then quantified on systematically chosen fields
within a single section spanning a complete cross section of the head
and tail (one section each). Mass was quantified by point-counting, as
originally described by Weibel (47) and applied by
Bonner-Weir et al. (7) to the endocrine pancreas. To
minimize variance and prevent bias, all quantification was performed by
one observer (N. Trivedi), who was unaware of the infusion group.
Quantification of mRNA levels (gene expression). Islet mRNA level was measured by semiquantitative radioactive multiplex RT-PCR. On the morning of the 4th day of infusion, rats were removed from the restraint system under Nembutal anesthesia, and the pancreas was digested with collagenase (n = 7 NON; n = 5 SAL, n = 7 GLC; n = 5 LIH). After digestion, islets were isolated and hand picked as previously described (20). The mRNA oligonucleotide primers, appropriate control genes, multiplex RT-PCR conditions, and validation and test of linearity have also previously been reported in detail (20).
Plasma insulin and FFA assays. Rat insulin was assayed using radioimmunoassay kits (Linco Research, St. Charles, MO). Kits were modified for small blood samples by reducing the manufacturer's recommended sample and diluent volumes by two (from 100 µl/sample to 50 µl/sample). Samples for insulin were diluted 1:5 (basal) or 1:10 (stimulated) such that determinations were performed in duplicate with only 5-10 µl of plasma. For the euglycemic-hyperinsulinemic clamps in which human insulin was infused, the plasma immunoreactive insulin level was assayed using a porcine standard. Plasma FFAs were measured using a colorimetric assay (Wako Chemicals, Neuss, Germany) modified for small samples by reducing the manufacturer's recommended sample and diluent volumes (factor of 2.5).
Data analysis and statistics. Results are presented as means ± SE. Differences among groups (SAL, GLC, LIH, and NON) were assessed with one-way ANOVA followed by Dunnett's test, with saline as the control group (P < 0.05 was considered significant). Differences between the saline-infused controls and noninfused animals (SAL and NON) were evaluated by unpaired t-tests [this was considered an a priori or planned test (24)]. Differences between time points within an animal (for example, pre- vs. postinfusion body wt) were evaluated with paired t-tests. In one case, Grubb's outlier test (2) was used to remove a data point (P < 0.05). Statistical tests were performed using Graphpad software (Graphpad, Irvine, CA). All animal procedures were approved by the Joslin Diabetes Center Institutional Animal Care Committee.
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RESULTS |
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Substrate infusions. During saline infusion, the morning fed glucose level fell from 125 ± 2 mg/dl on day 1 to 117 ± 3 mg/dl on day 4 (Fig. 1; P < 0.05; paired t-test, n = 32), whereas FFA [0.32 ± 0.06 vs. 0.35 ± 0.13 mM; not significant (NS); n = 6] and insulin (1.7 ± 0.2 vs. 1.6 ± 0.2 ng/ml; NS; n = 13) levels were unchanged. Animals maintained, but did not gain, body weight during the infusion (278 ± 8 vs. 278 ± 7 g; n = 32; NS).
During glucose infusion, plasma glucose increased from an initial basal level of 129 ± 2 mg/dl to a peak level of 214 ± 7.6 mg/dl and then slowly returned to near basal levels by ~36 h (Fig. 1A). The morning glucose level was slightly lower on day 4 compared with day 1 (121 ± 3 vs. 129 ± 2 mg/dl; P < 0.01; n = 37). During days 3 and 4, glucose levels were maintained near basal and were similar to the values observed in saline-infused controls (Fig. 1B). FFA levels tended to be lowest in the glucose-infused rats (Fig. 1C), but no significant differences were observed between saline- and glucose-infused rats during feeding on days 3 and 4 (60 and 84 h) or on the morning of the 4th day (Fig. 1D, ANOVA; n = 6). Morning FFA levels on days 1 and 4 were also similar (0.23 ± 0.04 vs. 0.19 ± 0.12 mM; NS; n = 6). By day 4, an approximately twofold increase in basal insulin was needed to maintain morning euglycemia (1.6 ± 0.1 to 3.0 ± 0.3 ng/ml; P < 0.05 by paired t-test; n = 21). Compared with saline-infused controls, insulin levels were doubled during feeding on days 3 and 4 (Fig. 1, E and F; ANOVA; P < 0.01). As with saline infusion, animals maintained, but did not gain, body weight over the 4-day infusions (271 ± 9 vs. 267 ± 11 g; n = 37; NS). During lipid infusion, plasma glucose levels were slightly suppressed at all time points measured. The morning glucose level on day 4 (99 ± 2 mg/dl) was significantly lower than the value on day 1 (120 ± 2 mg/dl; P < 0.05; paired t-test, n = 33) and lower than that observed in saline-infused controls (Fig. 1B; ANOVA; P < 0.01). FFA levels were elevated ~10-fold for the entire infusion period (0.32 ± 0.05 vs. 3.0 ± 0.97 mM, basal vs. 96 h; Fig. 1, C and D; n = 6; P < 0.01). During day 2, plasma insulin levels were similar in the saline- and lipid-infused groups (1.4 ± 0.3 vs. 1.6 ± 0.2 ng/ml at 24 h and 2.0 ± 0.4 vs. 2.2 ± 0.3 ng/ml at 36 h; NS) but were significantly elevated during feeding on days 3 and 4 (Fig. 1, E and F, P < 0.05 by ANOVA), suggesting that the insulin resistance induced by the high FFA level was being compensated for by an increase in insulin secretion in the absence of any change in plasma glucose (glucose levels were similar during days 3 and 4; Fig. 1, A and B; NS, ANOVA). By day 4, a small but significant increase in basal insulin was needed to maintain morning euglycemia (2.0 ± 0.5 vs. 1.7 ± 0.3 ng/ml; P < 0.05 by paired t-test; n = 10). Again, animals maintained, but did not gain, body weight during the infusion (292 ± 5 vs. 289 ± 5 g; n = 33; NS).Islet gene expression.
With the exception of a small change in islet amyloid polypeptide
(IAPP), islet gene expression assessed at the end of the 4-day
infusions was identical among the groups (Table
1). Although IAPP significantly increased
compared with saline-infused controls, this increase was not
significant compared with that of noninfused animals.
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Quantification of -cell mass and mitotic rate.
Once sectioning and staining were completed, sections were evaluated
for BrdU incorporation; sections in which BrdU staining could not be
adequately detected were excluded (two noninfused animals and one
glucose-infused animal could not be quantified because of poor BrdU
staining). As well, at the completion of the
-cell mass
calculations, one saline-infused animal was observed to have a mass
that was >4 SD from the mean; following outlier analysis
(P < 0.05, Grubb's outlier test), this animal was
also excluded. Thus the final number of animals used for assessment of
-cell mass was 5 in the SAL group and 6 in all of the other groups,
and the final number of animals used for quantification of
-cell
proliferation was 4 in the NON group, 5 in the SAL and GLC groups, and
6 in the LIH group. In these animals,
-cell mass was increased 80%
by glucose infusion (Fig. 2A;
P < 0.01; compared with SAL) and 54% by lipid
infusion (Fig. 2A; P < 0.05; compared with
SAL), and both glucose and lipid increased
-cell proliferation (Fig.
2B; P < 0.01; both comparisons relative to
SAL). However, compared with noninfused controls, saline infusion
lowered
-cell proliferation (P < 0.05, t-test), and the animals tended to end the infusion with
lower
-cell mass (P = 0.23; t-test),
suggesting that part of the differences observed between the saline-
and substrate-infused animals may have been due to an effect of the restraint or fluid loading (saline infusion controls for these effects,
because all infused animals have the same restraint and the same level
of fluid loading).
Assessment of postinfusion -cell function.
During hyperglycemic clamps, plasma glucose was rapidly elevated to 250 mg/dl (Fig. 3A). The glucose
infusion rate required to maintain this level of glycemia was similar
in NON, SAL, and LIH animals but was significantly elevated in the GLC
animals (Fig. 3B; P < 0.01). The high
glucose infusion rate was associated with a significantly higher plasma
insulin response (Fig. 3C; P < 0.05).
Assessment of postinfusion peripheral insulin clearance. To assess whether the increase in plasma insulin observed during the postglucose infused hyperglycemic clamp (Fig. 3C) was due to a decrease in peripheral insulin clearance, euglycemic hyperinsulinemic clamps were performed in a separate set of rats undergoing similar infusions. For these clamps, the glucose infusion rate was similar in NON, SAL, and LIH animals but was increased ~20% in the GLC group (P < 0.05; Fig. 4B). Steady-state plasma insulin levels tended to be lower in the LIH-infused animals (Fig. 4C), suggesting that insulin clearance may have been elevated in this group (22% relative to saline-infused controls, Fig. 4D); however, this difference did not achieve statistical significance (P = 0.17).
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DISCUSSION |
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The present study demonstrated that, during chronic substrate
oversupply, rats undergo compensatory -cell adaptations leading to
increased plasma insulin responses and normal glucose levels. During
glucose infusion, euglycemia was maintained by an approximately twofold
increase in plasma insulin (Fig. 1), and after 4 days,
-cell mass
was increased ~80% relative to saline-infused animals (Fig.
2A). After these changes, gene expression was unchanged for
islet hormones, genes involved in
-cell glucose uptake and metabolism, and transcription factors linked to insulin gene expression and
-cell development (Table 1). Compensatory increases in plasma insulin with normal glucose levels were also seen with lipid infusion, particularly during feeding (Fig. 1F). After 4 days of lipid
infusion,
-cell mass increased ~54% (Fig. 2A) and,
with the exception of a small change in IAPP, islet gene expression was
again unchanged (Table 1). Saline infusion resulted in the lowest
plasma insulin levels of any infused group and the lowest rate of
-cell proliferation (Fig. 2B). At the end of the saline
infusion,
-cell mass tended to be lower in saline-infused animals
compared with noninfused, nonrestrained animals. Of all the changes in
-cell mass, only the increase with glucose infusion yielded a
sustained (2-day) increase in the plasma insulin response to glucose
(Fig. 3C). This increase in plasma response was not due to
any change in peripheral insulin clearance (Fig. 4D),
indicating that the insulin secretory capacity per se was increased.
The lack of significant changes in islet gene expression after
prolonged glucose or lipid oversupply is surprising. During glucose
infusion, plasma insulin levels were doubled, but there was no change
in the expression of the insulin gene per se or of genes regulating the
entry into and metabolism of glucose in the -cell (GLUT-2, GK,
hexokinase and mitochondrial glycerol-3-phosphate dehydrogenase, and lactic dehydrogenase; Table 1). However, at the time gene expression was assessed (end of the 4-day infusions), glucose had been normal for >2 days (Fig. 1A) and
-cell
mass was significantly increased (Fig. 2A). The increase in
-cell mass may have allowed individual
-cells to secrete at a
normal basal level, obviating the need for an increase in the insulin gene transcription or genes required for glucose metabolism in the
-cell.
Genes known to be affected by FFA in vitro were also unchanged in the
present study despite an ~10-fold increase in plasma FFA levels
(Table 1). Elevated FFA has been shown to decrease PDX-1, GLUT-2, GK,
and insulin expression in vitro (16). The difference in
the results presented here and in Ref. 16 can likely be
attributed to different antecedent glucose levels. In the present
study, glucose levels were normal or elevated for the 4 days preceding
the islet isolation, whereas in Ref. 16, the
glucose level was deceased to 2.8 mM and then increased to 18 mM.
Antecedent hypoglycemia has been shown to alter -cell gene
expression (15, 42). In addition to PDX-1, FFA has also been proposed to act as a natural ligand of HNF4-
(18)
and potentially regulate its transcriptional activity on target genes. However, GLUT-2, which is a target of HNF4-
(45), was
not affected in the present study. Finally, high FFA levels have been
reported to reduce insulin gene translation in vitro (5).
Although we cannot rule out an effect of FFA on the translation of
insulin mRNA, we can rule out an acute consequence of such an effect, in that, in the present study, the
-cell was able to maintain normal
glucose levels and to dramatically increase insulin secretion during
feeding despite the increase in FFA. One possible reason for the
discrepancies between the in vivo results reported here and many of the
in vitro results may be related to the high FFA-to-BSA ratios used in
vitro; high FFA-to-BSA ratios result in unphysiological levels of
unbound FFA (46). The lack of a detrimental effect of FFA
on gene expression or insulin secretory capacity in vivo makes many of
the reported in vitro effects difficult to interpret.
Although neither glucose nor lipid infusion altered the transcription
of the genes measured in the present study, we have recently shown that
many of these genes are markedly altered in the 90% pancreatectomized
(Px) rat (20). Islet isolation, RNA preparation, and PCR
conditions were identical in the two studies (the same personnel
performed all procedures relating to the PCR). The important difference
between the two studies is that, here, the -cells were able to
maintain euglycemia, whereas in the 90% Px rat, graded levels of
hyperglycemia were observed that likely triggered a loss of
-cell
differentiation (20). Together, the two studies clearly
indicate that failure of
-cells to adapt to an increase in insulin
demands results in hyperglycemia and altered gene expression, whereas
successful
-cell adaptation leads to normal glucose levels and
normal gene expression.
The increase in -cell mass observed in the present study with
glucose infusion is comparable to the increase observed in our earlier
glucose infusion studies (7). However, in the present study, a second control group that was neither infused nor restrained was also studied. Compared with this group, saline infusion tended to
decrease the
-cell mass and significantly lowered the
-cell proliferation rate. Reasons for the decrease in proliferation and mass
are not entirely clear; however, during all of the substrate infusions,
the animals did not gain body weight, whereas noninfused nonrestrained
animals gain body weight at ~5-10 g per day. Thus it is possible
that the stress of the infusion led to a decrease in food intake (not
measured), leading to a lower insulin requirement and a subsequent
reduction in
-cell mass. Use of saline-infused animals as the
control group, as was done in the present study and in most studies of
this type, controls for this effect; together, the two controls
demonstrate that
-cell proliferation and mass are tightly coupled to
insulin demand. That is, during periods of reduced insulin demand
(saline infusion),
-cell mass and proliferation fall, whereas during
periods of increased insulin demand (glucose infusion), mass and
proliferation increase.
Consistent with the link between insulin demand and changes in -cell
mass, lipid infusion increased both the mass and proliferation rate
(Fig. 2). It is known that lipid infusion leads to insulin resistance
in the rat within ~5 h (8). The increase in
proliferation observed here is consistent with results obtained in
vitro (39). Interestingly, the increase in
-cell mass
observed with lipid infusion was smaller than that observed during
glucose infusion despite similar increases in
-cell proliferation.
This might suggest that the elevation in FFA also increased
apoptosis, as has been shown to occur in vitro
(43); however, this is speculative, given that changes in
islet neogenesis and hypertrophy were not measured. In any case, what
is clear is that, during the lipid infusion,
-cell mass was
maintained at a level sufficient to meet the demands of the animal.
Of all the changes in -cell mass observed in the present study, only
the change with glucose infusion resulted in a sustained increase in
insulin secretory capacity. In this group, during hyperglycemic clamps
that were performed 2 days after the chronic glucose infusion was
increased ~70% relative to all of the other groups (NON, SAL, and
LIH animals were all within 8% of each other; Fig. 3C).
This result, combined with our earlier finding that the increase in
-cell mass is stable for
1 wk (7), indicates that the
-cell mass formed during glucose infusion is functional. This
conclusion can be contrasted to numerous in vitro studies demonstrating
that the insulin response is reduced immediately after glucose infusion
(27, 28, 30-32). Although it is possible that a
glucotoxicity effect may had dissipated in the 2 days between the
infusion and the clamp, a study by Laury et al. (25)
demonstrated that the insulin response to a hyperglycemic clamp is
increased, even if the clamp is performed immediately after the glucose
infusion. The inconsistency between results obtained with the in vivo
hyperglycemic clamp and the in vitro isolated perfused pancreas has
been suggested to be due to the loss of neural stimulation during in
vitro perfusion (1). That the in vivo insulin response is
increased after glucose infusion is also supported by a recent study in
streptozotocin-diabetic rats indicating that
-cell mass and function
are both increased after glucose infusion in nondiabetic control rats
(3). Thus the present study, the Laury study
(25), and the control rats in Ref. 3 all
indicate that, during chronic glucose infusion, rats undergo
compensatory
-cell adaptations leading to an increased ability to
secrete insulin.
The change in plasma insulin response observed with glucose infusion cannot by itself be used to infer an increase in insulin secretion, because a decrease in insulin clearance will also appear as an increase in plasma insulin. Insulin clearance has been shown to be decreased during chronic glucose infusion (26). Although C-peptide has been used as a surrogate marker of clearance, no direct evidence exists to indicate that the clearance of this peptide is unaffected by chronic infusion. Furthermore, the rat has two insulin/C-peptide genes; this prevents the assessment of insulin secretion using traditional C-peptide deconvolution (13). In light of these issues, peripheral insulin clearance was evaluated by euglycemic-hyperinsulinemic clamp (Fig. 4) and determined to be normal. The normal clearance at this time point can be compared with the 50% reduction observed 1 h after glucose infusion (26). Together, the two studies suggest that insulin clearance may also be tightly linked to insulin demand.
The ability of the -cell to adapt to elevated FFA is controversial.
The two most recent studies using lipid-infused rats have concluded, in
one case, that the glucose-stimulated insulin response is increased
(34), and in the second case that it is decreased
(37). In the present study, lipid infusion did not result
in a postinfusion increase in insulin secretion, but compensatory increases in plasma insulin responses during feeding were observed during the infusion period (Fig. 1F). Although there are
methodological differences in the three studies (in Ref.
37 hyperglycemic clamps were performed while the lipid
infusions were in progress, in Ref. 34 intravenous glucose
tolerance tests were performed immediately after the infusions, and in
the present study hyperglycemic clamps were performed 2 days after the
infusions), several important observations can nonetheless be made.
First, the increased insulin response seen in Ref. 34
supports the contention that rat
-cells compensate during lipid
infusion. Interestingly, no change in mass was observed in Ref.
34 after 2 days of lipid infusion, whereas in the present
study a 54% increase in mass was observed after 4 days. This implies
that the adaptation in mass occurred on days 3 and
4. In contrast to the increased insulin response in Ref.
34, a decreased response was observed in Ref.
37 that the authors speculate may have been due to altered
gene expression or reduced
-cell mass. In this regard, we did not
find any alterations in gene expression at day 4, and the
mass was elevated on day 4. Thus the increased insulin
response observed in Ref. 34 and results of the present
study both support the contention that rat
-cells are able to adapt
to the increased need for insulin brought about by high FFA levels.
Although the elevation in FFA generated a clear adaptive response during the infusion period, no effect on the plasma insulin response was observed 2 days after the lipid infusion (Fig. 3C). Note that it is possible that secretion was elevated at this time and that the elevation was masked by an increase in insulin clearance [clearance was increased 22% at this time point (Fig. 4D), but this increase was not statistically significant; P = 0.17]. However, it is also possible that the normalized plasma insulin response was simply a result of insulin sensitivity having returned to a normal level. In this regard, one can argue that the elevated insulin response after glucose infusion was inappropriate given that the insulin sensitivity had also normalized in this group. Additional time points assessing insulin sensitivity, secretion, and clearance will need to be conducted to fully address this issue.
Finally, it is interesting to compare characteristics of the
substrate-infused rat with those of developing type 2 diabetes in
humans. In particular, it is noteworthy that the animals do not gain
body weight during the infusion period. Thus the model is not one of
caloric overconsumption or obesity-induced insulin resistance. However,
lipid infusion clearly results in peripheral insulin resistance
(8) and an inability of insulin to suppress hepatic
glucose output (4). Regarding the latter, overproduction of glucose by the liver is a well known characteristic of developing type 2 diabetes (10, 11, 35), and this overproduction is clearly mimicked by continuous glucose infusion. In the present study,
both the insulin resistance induced by lipid infusion and the elevated
rate of glucose appearance with glucose infusion were rapidly
compensated for by the -cell. This compensation included an increase
in
-cell mass, a functional increase in insulin secretion, and the
normal expression of key islet genes. Understanding the cellular and
molecular mechanisms of this adaptive response has important
implications for understanding why some individuals can adapt to
increases in insulin demand while others go on to develop diabetes.
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ACKNOWLEDGEMENTS |
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We thank Irene Reski and Marta Grinbergs of the Joslin Diabetes Center RIA Core for performing the insulin and C-peptide assays.
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FOOTNOTES |
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The RIA Core, Animal Facilities Core, and the Tissue Culture Core are part of the Diabetes Endocrinology Research Center of the Joslin Diabetes Center, supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-36836. N. Trivedi is the recipient of the American Diabetes Association (ADA) postdoctoral mentor-based training grant. This work was also supported by grants DK-35449 to G. C. Weir, DK-44523 to S. Bonner-Weir, and the ADA Career Development Award to A. Sharma. J.-C. Jonas is currently a Research Associate from the Fonds National de la Recherche Scientifique (Brussels, Belgium).
Address for reprint requests and other correspondence: G. M. Steil, MiniMed Inc., 18000 Devonshire St., Northridge, CA 91325-1219 (E-mail: GarryS{at}MiniMed.com).
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. Section 1734 solely to indicate this fact.
Received 26 July 2000; accepted in final form 23 January 2001.
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REFERENCES |
---|
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---|
1.
Balkan, B,
and
Dunning BE.
Muscarinic stimulation maintains in vivo insulin secretion in response to glucose after prolonged hyperglycemia.
Am J Physiol Regulatory Integrative Comp Physiol
268:
R475-R479,
1995
2.
Barnett, V,
and
Lewis T.
Outliers in Sstatistical Data. New York: Wiley, 1994.
3.
Bernard, C,
Thibault C,
Berthault MF,
Magnan C,
Saulnier C,
Portha B,
Pralong WF,
Penicaud L,
and
Ktorza A.
Pancreatic -cell regeneration after 48-h glucose infusion in mildly diabetic rats is not correlated with functional improvement.
Diabetes
47:
1058-1065,
1998[Abstract].
4.
Bevilacqua, S,
Bonadonna RC,
Buzzigoli G,
Boni C,
Ciociaro D,
Maccari F,
Giorico MA,
and
Ferrannini E.
Acute elevation of free fatty acid levels leads to hepatic insulin resistance in obese subjects.
Metabolism
36:
502-506,
1987[ISI][Medline].
5.
Bollheimer, LC,
Skelly RH,
Chester MW,
McGarry JD,
and
Rhodes CJ.
Chronic exposure to free fatty acid reduces pancreatic beta cell insulin content by increasing basal insulin secretion that is not compensated for by a corresponding increase in proinsulin biosynthesis translation.
J Clin Invest
101:
1094-1101,
1998
6.
Bonner-Weir, S.
Regulation of pancreatic -cell mass in vivo.
Recent Prog Horm Res
49:
91-104,
1994[ISI][Medline].
7.
Bonner-Weir, S,
Deery D,
Leahy JL,
and
Weir GC.
Compensatory growth of pancreatic -cells in adult rats after short-term glucose infusion.
Diabetes
38:
49-53,
1989[Abstract].
8.
Chalkley, SM,
Hettiarachchi M,
Chisholm DJ,
and
Kraegen EW.
Five-hour fatty acid elevation increases muscle lipids and impairs glycogen synthesis in the rat.
Metabolism
47:
1121-1126,
1998[ISI][Medline].
9.
Chen, C,
Bumbalo L,
and
Leahy JL.
Increased catalytic activity of glucokinase in isolated islets from hyperinsulinemic rats.
Diabetes
43:
684-689,
1994[Abstract].
10.
Consoli, A,
Nurjhan N,
Reilly JJ, Jr,
Bier DM,
and
Gerich JE.
Mechanism of increased gluconeogenesis in non-insulin-dependent diabetes mellitus. Role of alterations in systemic, hepatic, and muscle lactate and alanine metabolism.
J Clin Invest
86:
2038-2045,
1990[ISI][Medline].
11.
DeFronzo, RA,
Bonadonna RC,
and
Ferrannini E.
Pathogenesis of NIDDM: a balanced overview.
Diabetes Care
15:
318-368,
1992[Abstract].
12.
Dobbins, RL,
Chester MW,
Daniels MB,
McGarry JD,
and
Stein DT.
Circulating fatty acids are essential for efficient glucose-stimulated insulin secretion after prolonged fasting in humans.
Diabetes
47:
1613-1618,
1998[Abstract].
13.
Eaton, RP,
Allen RC,
Schade DS,
Erickson KM,
and
Standefer J.
Prehepatic insulin production in man: kinetic analysis using peripheral connecting peptide behavior.
J Clin Endocrinol Metab
51:
520-528,
1980[Abstract].
14.
Finegood, DT,
Scaglia L,
and
Bonner-Weir S.
Dynamics of -cell mass in the growing rat pancreas: estimation with a simple mathematical model.
Diabetes
44:
249-256,
1995[Abstract].
15.
Giddings, SJ,
Carnaghi LR,
and
Shalwitz RA.
Hypoglycemia but not hyperglycemia induces rapid changes in pancreatic -cell gene transcription.
Am J Physiol Endocrinol Metab
265:
E259-E266,
1993
16.
Gremlich, S,
Bonny C,
Waeber G,
and
Thorens B.
Fatty acids decrease IDX-1 expression in rat pancreatic islets and reduce GLUT2, glucokinase, insulin, and somatostatin levels.
J Biol Chem
272:
30261-30269,
1997
17.
Habener, JF,
and
Stoffers DA.
A newly discovered role of transcription factors involved in pancreas development and the pathogenesis of diabetes mellitus.
Proc Assoc Am Phys
110:
12-21,
1998[ISI][Medline].
18.
Hertz, R,
Magenheim J,
Berman I,
and
Bar-Tana J.
Fatty acyl-CoA thioesters are ligands of hepatic nuclear factor-4alpha.
Nature
392:
512-516,
1998[ISI][Medline].
19.
Hirose, H,
Lee YH,
Inman LR,
Nagasawa Y,
Johnson JH,
and
Unger RH.
Defective fatty acid-mediated -cell compensation in Zucker diabetic fatty rats: pathogenic implications for obesity-dependent diabetes.
J Biol Chem
271:
5633-5637,
1996
20.
Jonas, JC,
Sharma A,
Hasenkamp WM,
Ilkova H,
Laybutt DR,
Bonner-Weir S,
and
Weir GC.
Chronic hyperglycemia triggers loss of pancreatic cell differentiation in an animal model of diabetes.
J Biol Chem
274:
14112-14121,
1999
21.
Jonsson, J,
Carlsson L,
Edlund T,
and
Edlund H.
Insulin-promoter-factor I is required for pancreatic development in mice.
Nature
371:
606-609,
1994[ISI][Medline].
22.
Kahn, SE,
Beard JC,
Schwartz MW,
Ward WK,
Ding HL,
Bergman RN,
Taborsky GJJ,
and
Porte DJ.
Increased -cell secretory capacity as mechanism for islet adaptation to nicotinic acid-induced insulin resistance.
Diabetes
38:
562-568,
1989[Abstract].
23.
Kahn, SE,
Prigeon RL,
McCulloch DK,
Boyko EJ,
Bergman RN,
Schwartz MW,
Neifing JL,
Ward WK,
Beard JC,
and
Palmer JP.
Quantification of the relationship between insulin sensitivity and -cell function in human subjects. Evidence for a hyperbolic function.
Diabetes
42:
1663-1672,
1993[Abstract].
24.
Kirk, RE.
Experimental Design: Procedures for the Behavior Sciences. Pacific Grove, CA: Brooks/Cole, 1982.
25.
Laury, MC,
Takao F,
Bailbe D,
Penicaud L,
Portha B,
Picon L,
and
Kuikka J.
Differential effects of prolonged hyperglycemia on in vivo and in vitro insulin secretion in rats.
Endocrinology
128:
2526-2533,
1991[Abstract].
26.
Laybutt, DR,
Chisholm DJ,
and
Kraegen EW.
Specific adaptations in muscle and adipose tissue in response to chronic systemic glucose oversupply in rats.
Am J Physiol Endocrinol Metab
273:
E1-E9,
1997
27.
Leahy, JL,
Bonner-Weir S,
and
Weir GC.
Minimal chronic hyperglycemia is a critical determinant of impaired insulin secretion after an incomplete pancreatectomy.
J Clin Invest
81:
1407-1414,
1988[ISI][Medline].
28.
Leahy, JL,
Bonner-Weir S,
and
Weir GC.
Beta-cell dysfunction induced by chronic hyperglycemia. Current ideas on mechanism of impaired glucose-induced insulin secretion.
Diabetes Care
15:
442-455,
1992[Abstract].
29.
Leahy, JL,
Cooper HE,
Deal DA,
and
Weir GC.
Chronic hyperglycemia is associated with impaired glucose influence on insulin secretion. A study in normal rats using chronic in vivo glucose infusions.
J Clin Invest
77:
908-915,
1986[ISI][Medline].
30.
Leahy, JL,
Cooper HE,
and
Weir GC.
Impaired insulin secretion associated with near normoglycemia. Study in normal rats with 96-h in vivo glucose infusions.
Diabetes
36:
459-464,
1987[Abstract].
31.
Leahy, JL,
and
Weir GC.
Evolution of abnormal insulin secretory responses during 48-h in vivo hyperglycemia.
Diabetes
37:
217-222,
1988[Abstract].
32.
Leahy, JL,
and
Weir GC.
Beta-cell dysfunction in hyperglycaemic rat models: recovery of glucose-induced insulin secretion with lowering of the ambient glucose level.
Diabetologia
34:
640-647,
1991[ISI][Medline].
33.
Lee, Y,
Hirose H,
Ohneda M,
Johnson JH,
McGarry JD,
and
Unger RH.
-Cell lipotoxicity in the pathogenesis of non-insulin dependent diabetes of obese rats: impairment in adipocyte-
-cell relationships.
Proc Natl Acad Sci USA
91:
10878-10882,
1994
34.
Magnan, C,
Collins S,
Berthault MF,
Kassis N,
Vincent M,
Gilbert M,
Penicau L,
Ktorza A,
and
Assimacopoulos-Jeannet F.
Lipid infusion lowers sympathetic nervous activity and leads to increased -cell responsiveness to glucose.
J Clin Invest
103:
413-419,
1999
35.
Magnusson, I,
Rothman DL,
Katz LD,
Shulman RG,
and
Shulman GI.
Increased rate of gluconeogenesis in type II diabetes mellitus.
J Clin Invest
90:
1323-1327,
1992[ISI][Medline].
36.
Martin, BC,
Warram JH,
Krolewski AS,
Bergman RN,
Soeldner JS,
and
Kahn CR.
Role of glucose and insulin resistance in development of type 2 diabetes mellitus: results of a 25-year follow-up study.
Lancet
340:
925-929,
1992[ISI][Medline].
37.
Mason, TM,
Goh T,
Tchipahvili V,
Sandhu H,
Gupta N,
Lewis GF,
and
Giacca A.
Prolonged elevation of plasma free fatty acids desensitize the insulin secretory response to glucose in vivo in rats.
Diabetes
48:
524-530,
1999[Abstract].
38.
Matschinsky, FM.
Glucokinase as glucose sensor and metabolic generator in pancreatic -cell signal transduction.
Diabetes
39:
647-652,
1990[Abstract].
39.
Milburn, JL, Jr,
Hirose H,
Lee YH,
Nagasawa Y,
Ogawa A,
Ohneda M,
BeltrandelRio H,
Newgard CB,
Johnson JH,
and
Unger RH.
Pancreatic -cells in obesity: evidence for induction of functional, morphological, and metabolic abnormalities by increased long chain fatty acids.
J Biol Chem
270:
1295-1299,
1995
40.
Montana, E,
Bonner-Weir S,
and
Weir GC.
Beta cell mass and growth after syngeneic islet cell transplantation in normal and streptozotocin diabetic C57BL/6 mice.
J Clin Invest
91:
780-787,
1993[ISI][Medline].
41.
Naya, FJ,
Huang HP,
Qiu Y,
Mutoh H,
DeMayo FJ,
Leiter AB,
and
Tsai MJ.
Diabetes, defective pancreatic morphogenesis, and abnormal enteroendocrine differentiation in BETA2/neuroD-deficient mice.
Genes Dev
11:
2323-2334,
1997
42.
Shalwitz, RA,
Herbst T,
Carnaghi LR,
and
Giddings SJ.
Time course for effects of hypoglycemia on insulin gene transcription in vivo.
Diabetes
43:
929-934,
1994[Abstract].
43.
Shimabukuro, M,
Zhou YT,
Levi M,
and
Unger RH.
Fatty acid-induced cell apoptosis: a link between obesity and diabetes.
Proc Natl Acad Sci USA
95:
2498-2502,
1998
44.
Stein, DT,
Esser V,
Stevenson BE,
Lane KE,
Whiteside JH,
Daniels MB,
Chen S,
and
McGarry JD.
Essentiality of circulating fatty acids for glucose-stimulated insulin secretion in the fasted rat.
J Clin Invest
97:
2728-2735,
1996
45.
Stoffel, M,
and
Duncan SA.
The maturity-onset diabetes of the young (MODY1) transcription factor HNF4 regulates expression of genes required for glucose transport and metabolism.
Proc Natl Acad Sci USA
94:
13209-13214,
1997
46.
Warnotte, C,
Nenquin M,
and
Henquin JC.
Unbound rather than total concentration and saturation rather than unsaturation determine the potency of fatty acids on insulin secretion.
Mol Cell Endocrinol
153:
147-153,
1999[ISI][Medline].
47.
Weibel, ER.
Stereologic methods.
In: Practical Methods for Biological Morphometry. London: Academic, 1978, p. 101-161.
48.
Wisenthal, SR,
Sandhu H,
Mccall RH,
Tchipashvili V,
Yoshii H,
Polonsky K,
Shi ZQ,
Lewis GF,
Mari A,
and
Giacca A.
Free fatty acids impair hepatic insulin extraction in vivo.
Diabetes
48:
766-774,
1999[Abstract].
49.
Woerner, CA.
Studies on the islands of Langerhans after continuous injection of dextrose.
Anat Rec
71:
22-57,
1936.
50.
Zambon, A,
Hashimoto SI,
and
Brunzell JD.
Analysis of techniques to obtain plasma for measurement of levels of free fatty acids.
J Lipid Res
34:
1021-1028,
1993[Abstract].