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Introduction
Diabetes mellitus (DM) is a chronic disease characterized by high
blood glucose levels. It can be broadly categorized into type I
and type II. The former develops as a result of insulin deficiency
and the latter is due to defects in pancreatic secretion of insulin
and insulin action[1], and insulin resistance in target
tissues, mainly muscle and liver. The biological effect of insulin
is initiated with insulin binding to the ¦Á-subunit of insulin receptor
(IR) and activating the intrinsic tyrosine kinase activity of the
¦Â-subunit of the receptor[2]. Activated IR results in
the subsequent phosphorylation of intracellular substrates including
insulin receptor substrates (IRSs) such as IRS-1 and -2, phosphatidylinositol
(PI) 3-kinase, and protein kinase B (PKB)[3,4]. Normal
insulin action leads to increased glycogen synthesis, glucose transport,
and lipogenesis, and decreased gluconeogenesis, glycogenolysis,
and lipolysis[5-7]. The net effect on glucose metabolism
is that hepatic glucose production is reduced, whereas use of peripheral
glucose is increased[6,7].
An exquisite balance is required between kinases, which are involved
in transmitting signals to the downstream targets necessary for
the processes described above, and phosphatases, which are required
to shut down this signaling to prevent excessive or, in some cases,
insufficient activation[8]. Insulin resistance and diabetes
represent states in which the regulation of the signaling pathways
is altered so that the intracellular actions of insulin are absent
or reduced[9]. Overactivation of phosphatases is one
possible means of blocking the insulin signaling. One key phosphatase,
protein tyrosine phosphatase 1B (PTP1B), which dephosphorylates
the activated insulin receptor and IRS-1, has been shown to play
a major role in insulin resistance and type II diabetes[10].
Recently, it was shown that PTP1B-knock-out mice become more insulin
sensitive and fail to gain weight despite being fed with a fat-rich
diet[11]. Moreover, the role for certain PTPs, including
PTP1B, in the insulin resistance associated with diabetes and obesity,
has been suggested by some clinical studies in which correlations
between the levels of PTP1B expression in muscle and adipose tissue
and insulin resistant states were found[12,13]. Therefore,
PTP1B might be an attractive therapeutic target in the treatment
of type II diabetes and obesity.
Plants have always been usable sources of drugs, and many currently
available drugs are directly or indirectly derived from plants.
Currently available therapies for diabetes include insulin and various
oral anti-diabetic agents, such as sulfonylureas, metformin, ¦Á-glucosidase
inhibitors, and rosiglitazone. These drugs are used as monotherapies
or in combination to achieve better glycemic control. Each of these
oral agents suffers from implication in a number of serious adverse
effects[14]. Therefore, it is important to investigate
the hypoglycemic actions of plants that were originally used in
traditional medicine[15,16]. The biologically active
components of plants with hypoglycemic actions include flavonoids,
alkaloids glycosides, polysaccharides, and peptidoglycans[17,18].
Astragalus polysaccharides (APS) are one of the main efficacious
principles of Radix Astragali (Astragalus membranaceus),
which is reported to have anti-oxidant, anti-diabetic, anti-hypertensive,
and immunomo-dulatory activities [19].
To study natural products with anti-diabetes activity, we screened
our extract bank for inhibitors of PTP1B enzyme and found that a
fraction from an aqueous extract of the roots of A membranaceus
showed strong inhibitory bioactivity against PTP1B with IC50
equaling 7.50 µmol/L (positive control sodium orthovanadate
IC50=10 µmol/L). This study was to investigate the
effects of APS on the activity and expression of PTP1B in the livers
and skeletal muscles of non-diabetic rats and rats with high-fat
streptozotocin-induced diabetes. The effects of APS on insulin sensitivity
and insulin-induced tyrosine phosphorylation of IR ¦Â-subunit and
IRS-1 in non-diabetic and diabetic rats were also studied in
an effort to establish the mechanisms of its hypoglycemic actions.
Materials and methods
Plant materials and preparation of APS Astragalus
membranaceus (Fisch) Bunge var mongholicus (Bunge)
Hsiao was purchased from Shanghai Medicinal Materials Co (Shanghai,
China), and identified by the Department of Authentication of Chinese
Medicine, Hubei College of Chinese Traditional Medicine (Wuhan,
China). A representative specimen has been kept in our laboratory
for future reference.
APS were extracted with optimized techniques using direct water
decoction, as described previously[20]. The yield of
APS was 2.0%, and the total polysaccharide content was 38.5%. Three
subtypes of APS are defined by phytochemical screening: APSI, II,
and III (1.47:1.21:1). APSI consists of D-glucose, D-galactose,
and L-arabinose in molar ratios of 1.75:1.63:1 and has an
average molecular weight of 36 300. Both APSII and APSIII are dextrans,
the linkage mode of which is mainly ¦Á-(1¡ú4) linkages, and in
which ¦Á-(1¡ú6) linkages are exiguous.
APS is a hazel-colored and water-soluble powder. It was
diluted to 20 g/L in distilled water before use.
Materials and chemicals The reagents for sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting
were obtained from BioVision (Palo Alto, CA, USA) and the apparatus
from Bio-Rad (Richmond, CA, USA). Streptozotocin (STZ), Tris, Nonidet
P-40, porcine insulin, and nitrocellulose (NC) membranes were obtained
from Sigma Chemical Co (St Louis, MO, USA). Protein A-Sepharose
6 MB was from Pharmacia (Uppsala, Sweden). The monoclonal anti-phosphotyrosine
antibody (¦ÁPY, PY99) was purchased from Santa Cruz Biotechnology,
Inc (Santa Cruz, CA, USA). Rabbit poly-clonal anti-insulin receptor
¦Â-subunit (IR¦Â) antibody was purchased from Upstate Biotechnology
Inc (Lake Placid, NY, USA). The anti-rat carboxy-terminal IRS-1
antibody (clone 8-63) was from NeoMarkers (Fremont, CA, USA). ¦Á-PTP1B
polyclonal antibody was purchased from Upstate Biotechnology Inc.
Enhanced chemiluminescence (ECL) detection reagents were from Kirkegaard
& Perry Laboratories (Gaithersburg, MA, USA). All other chemicals
were of the highest analytical grade.
Type II diabetic (TIIDM) rat model and treatment protocol Male
Sprague-Dawley rats [from the experimental animal center of Wuhan
University, certificate No SCXK (Hubei) 2003-0003], 8 weeks
of age and weighing approximately 200 g, were used for all studies.
All procedures were in accordance with the Institute Ethical Committee
for the Experimental Use of Animals of Wuhan University. Rats were
housed five per cage in a room with a 12:12 hour light:dark cycle
and an ambient temperature of 22-25 °C. Animals were fed either
a normal chow diet consisting (as a percentage of total kcal) of
12% fat, 60% carbohydrate, and 28% protein, or a high-fat diet (HFD)
consisting of 41% fat, 41% carbohydrate, and 18% protein. After
2 weeks on either diet, animals (with the exception of noninjected
controls) were anesthetized with ketamine (65 mg/kg) and xylazine
(7 mg/kg) after an overnight fast and injected with STZ (30 mg/kg
in 0.1 mol/L citrate-buffered saline, pH 4.5) into the tail vein
via a temporary indwelling 24-gauge catheter. Animals had free access
to food and water after the STZ injection, and both STZ-injected
and noninjected animals continued on their original diets (chow
or fat) for the duration of the study. After a 12-h fast, animals
showing fasting glucose levels >6.7 mmol/L at 72 h after STZ
injection were considered diabetic. The control rats were fasted
in an identical manner and had a volume of citrate-buffered saline,
equal to that of the STZ solution, injected by the same route.
The control and diabetic groups were then further subdivided into
treated and untreated groups: control (n=10); control treated
with APS (Control+APS, n=10); TIIDM (n=12); and TIIDM
treated with oral APS (400 mg/kg) (TIIDM +APS, n=12). Treatment
was given daily for 5 weeks. The control group received an equal
volume of vehicle (saline). At the end of each week, individual
body weights were recorded, and glucose and insulin levels were
determined under fasting and nonfasting conditions. Glycemia was
assessed on blood collected from the tail vein using an OneTouch
Ultra blood glucose meter (LifeScan, Milpitas, CA, USA). Insulin
levels were determined by radio-immunoassay (RIA) with a kit from
Beifang Biotech Research Center (Beijing, China). Before necropsy,
saline- and APS-treated animals were intraperitoneally administered
insulin (5 U/kg) in saline with 0.1% bovine serum albumin (BSA)
or vehicle (saline with 0.1% BSA) after a 10-h fast. Tissue samples
from livers and soleus muscles were taken (10 min after treatment)
from both vehicle- and insulin-treated animals and snap frozen in
liquid nitrogen.
Insulin sensitivity Peripheral insulin resistance
was assessed with an insulin-tolerance test (ITT) [21]
that measured insulin sensitivity using KITT as
an index of insulin-mediated glucose metabolism. Rats were fasted
for 15 h before insulin challenge. Neutral insulin injections were
diluted with 0.9% saline to a final concentration of 2 kU/L, and
then administered at a dose of 2 U/kg body weight by slow intravenous
injection through the tail vein. Blood samples were collected at
0, 10, 20, 30, and 60 min after the administration of insulin. Serum
was separated and subjected to glucose estimation. Serum glucose
concentrations were determined using a commercial assay kit (Sigma
Diagnostics, St Louis, MO, USA) and 10 µL of serum was used
for each assay. KITT was determined from the
slope of a linear portion of the regression line of the natural
logarithm of glucose versus time[21], and calculated
using the formula[22]: KITT=(0.693/t1/2
)×100, where t1/2 represents the half-life
of plasma glucose decay, which was estimated by plotting plasma
glucose concentration versus time on semilogarithmic graph paper.
Lower insulin-sensitivity index (KITT) scores
mean higher degrees of insulin resistance.
Lysate preparation and protein assays Frozen
liver tissue (50 mg) was sonicated in 1 mL of lysis buffer (buffer
A) containing Tris-HCl 20 mmol/L (pH 7.4), 1% Triton X-100, 10%
glycerol, NaCl 150 mmol/L, edetic acid 2 mmol/L, b-glycerophosphate
25 mmol/L, sodium fluoride 20 mmol/L, sodium orthovanadate 1 mmol/L,
sodium pyrophosphate 2 mmol/L, leupeptin 10 mg/L, benzamidine 1
mmol/L, 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride
1 mmol/L, and microcystin 1 mmol/L, and rocked for 40 min at 4 °C.
Detergent-insoluble material was sedimented by centrifugation at
12 000×g at 4 °C for 10 min. Muscle samples (50
mg) were homogenized and centrifuged at 1000 000×g for
1 h in ice-cold Hepes buffer 50 mmol/L (pH 7.4) containing NaCl
150 mmol/L, sodium pyrophosphate 10 mmol/L, Na3VO4
2 mmol/L, NaF 10 mmol/L, edetic acid 2 mmol/L, phenylmethylsulfonyl
fluoride (PMSF) 2 mmol/L, leupeptin 5 mg/L, 1% Nonidet P-40, and
10% glycerol (buffer B).
Supernatants were collected, and protein concentrations were measured
with Bradford protein assay reagent (Bio-Rad), using BSA as the
standard.
Western blotting Aliquots (50 µg) of muscle
or liver homogenates were subjected to SDS-PAGE (7.5% gel) and transferred
electrophoretically onto nitrocellulose (NC) membranes for 5 h.
NC membranes were then blocked for 2 h at room temperature with
block solution provided in the ECL kits. This step was followed
by overnight incubation at 4 °C with anti-¦Á-PTP1B polyclonal
antibody or anti-phosphotyrosine (PY99) as the primary antibodies,
as described in the figure legends. The NC membranes were then washed
for 30 min with wash solution (ECL kits), followed by a 1-h incubation
with either anti-mouse or anti-rabbit IgG conjugated with horseradish-peroxidase
in block solution. The NC membranes were washed for 30 min in wash
solution, and the immunoreactive bands were detected with an enhanced
chemiluminescence method.
Tyrosine phosphorylation of IR-subunit and IRS-1 Muscle
or liver lysates (1 mg of protein) were immunoprecipitated overnight
at 4 °C with 2 µg of anti-IR¦Â or anti-IRS-1 coupled to
protein A-Sepharose. The immune complex was washed three times in
phosphate-buffered saline (PBS) (pH 7.4) containing 1% Nonidet P-40
and Na3VO4 2 mmol/L, resuspended in Laemmli
buffer, and boiled for 5 min. Proteins were resolved on SDS-PAGE
(7.5% gel), then electrotransferred from the gel to nitrocellulose
membranes. The nitrocellulose filters were incubated at 4 ºC
overnight with 1 mg/L PY99. Subsequent steps were performed as described
above.
PTP1B protein levels and activity The tissue
homogenate was assayed in a microtiter plate at 27 °C. The
PTP1B assay kit was obtained from Upstate Biotechnology Inc. The
protocol outlined by the manufacturer was followed rigidly. PTP1B
protein levels were assessed by immunoblotting using polyclonal
antibodies directed against PTP1B, as described above.
Statistical analysis All values are
expressed as mean± SEM. Statistical significance was determined
using analysis of variance (ANOVA) followed by Tukey's test. P<0.05
was considered statistically significant.
Results
Characteristics of experimental animals The glucose
levels of both fasting and fed rats were significantly higher (P<0.05)
in HFD/STZ rats (TIIDM) than in normal control rats (Control) (Table
1). The levels of glucose in fasting and fed TIIDM rats were significantly
reduced (P<0.05) after treatment with APS at a dose of
400 mg/kg po per day for 5 weeks (TIIDM+APS). It should be
noted that insulin concentrations in type II diabetic rats were
similar to values of control rats. Treatment with APS did not affect
insulin levels in control or type II diabetic rats. HFD/STZ diabetic
rats weighed 20 g more than the normal chow-fed controls (P<0.05).
APS treatment could significantly reduce body weights in diabetic
rats and had no effects on those of control rats. All data are mean±SEM
calculated from 10 rats.
Insulin sensitivity Insulin sensitivity in the
TIIDM rats was significantly lower than that in the control group.
The impaired insulin sensitivity in the TIIDM rats was improved
following APS treatment (Figure 1).
Effects of APS on PTP1B expression in muscles and livers of
control and TIIDM rats PTP1B protein levels in the
skeletal muscles of TIIDM rats were significantly increased (1.6-fold,
P<0.05) compared with those of controls. In TIIDM rats,
the level of PTP1B after treatment with APS was reduced by 30% (P<0.05),
compared with that of the TIIDM rats (Figure 2A and C). PTP1B protein
levels in the livers of TIIDM rats were also significantly increased
(1.7-fold, P<0.05) compared with those of control rats.
However, treatment with APS did not affect PTP1B protein levels
in the livers of TIIDM rats (Figure 2B and D). PTP1B levels in the
muscles and livers of normal control rats were not affected by APS
treatment.
Effects of APS on PTP1B activity in muscles and livers of control
and TIIDM rats There was a two-fold (P<0.01)
increase in the activity of PTP1B in the skeletal muscles of TIIDM
rats. APS reduced the activity of PTP1B by 25% (P<0.05)
in APS-treated TIIDM rats (Figure 3A). PTP1B activity in the livers
of TIIDM rats was significantly elevated (1.8-fold, P<0.01)
compared with that of the controls. Treatment with APS did not change
PTP1B activity in the livers of TIIDM rats (Figure 3B). Nor did
APS affect the activity of PTP1B in the skeletal muscles or livers
of APS-treated control rats.
Effects of APS on insulin-induced tyrosine phosphorylation of
IR ¦Â-subunit In each group of animals, the administration of
insulin resulted in an increase in the tyrosine phosphorylation
of the IR ¦Â-subunit (Figure 4). However, the levels of IR¦Â tyrosine
phosphorylation in the skeletal muscles and livers of TIIDM rats
after stimulation with insulin were reduced by 47% (P<0.05)
and 41% (P<0.05), respectively, compared with those of
the control rats (Figure 4). Treatment with APS significantly increased
the level of IR¦Â tyrosine phosphorylation stimulated by insulin
in the skeletal muscles of TIIDM rats, but did not affect that in
the livers of TIIDM rats. APS also did not affect the IR¦Â tyrosine
phosphorylation stimulated by insulin in the skeletal muscles or
livers of APS-treated control rats, as evaluated by scanning densitometry.
Effects of APS on insulin-induced tyrosine phosphorylation of
IRS-1 Insulin administration resulted in an increase
in IRS-1 tyrosine phosphorylation in each group of animals (Figure
5). TIIDM rats exhibited a reduced response to insulin, and the
levels of tyrosine phosphorylation of IRS-1 after insulin stimulation
in skeletal muscles and livers were reduced by 37% and 35% (P<0.05),
respectively, compared with the values measured in control rats
(Figure 5). APS treatment did not increase the level of tyrosine
phosphorylation of IRS-1 induced by insulin in the liver tissues
of TIIDM rats, but significantly improved that in skeletal muscles
(P<0.05) in TIIDM rats. APS had no effect on the tyrosine
phosphorylation of IRS-1 stimulated by insulin in the skeletal muscles
or livers of APS-treated control rats.
Discussion
In the present study, we developed an animal model for type II
diabetes, the HFD/STZ rat. These rats constitute a relatively inexpensive
and easily accessible rodent model that is not extremely obese and
simulates the natural history and metabolic characteristics of patients
with type II diabetes[23]. Recent reports[24]
have indicated that insulin resistance may be caused by an excess
nutrient supply. Both excess glucose and excess fat can cause insulin
resistance in muscle and fat tissues, whereas excess fat can cause
impaired suppression of endogenous glucose production. The present
animal model was based on the rationale described above. At first,
we used high-fat diets to induce insulin resistance in Sprague-Dawley
rats. Plasma glucose concentrations were similar in chow-fed and
HFD rats after two weeks of the high-fat diet, whereas insulin concentrations
in the HFD rats were significantly higher than those in the chow-fed
controls (HFD: 338±25 vs Control: 162±13 pmol/L,
P<0.05). The association of normoglycemia and hyperinsulinemia
suggested that the HFD rats were insulin-resistant. Conversion of
pre-diabetes to frank hyperglycemia in patients with type II diabetes
is associated with a decline in the secretory capacity of the pancreatic
¦Â-cells[25,26]. However, this fai-lure in ¦Â-cell compensation
is relative, not absolute[23]. We attempted to simulate
this evolution from a state of insulin resistance and absolute hyperinsulinemia
to a state of "relative" hypoinsulinemia by injecting
insulin-resistant HFD rats with a moderate amount of STZ, which
partly damaged ¦Â-cell and reduced the serum insulin concentration
to a relatively lower level (178±20 pmol/L), which is approximately
that of normal chow-fed rats. Comparison of the data in Table 1
indicates that we were successful in this attempt.
The signaling pathways affected by the inhibition of PTP1B, particularly
in a diabetic model, have been characterized. In this study, we
observed increased PTP1B activity in skeletal muscle and liver tissues
of HFD/STZ rats when para-nitrophenyl phosphate (pNPP)
was used as substrate. These changes were similar to those in PTP1B
protein levels in the tissues. Corresponding to these results, insulin-induced
tyrosine phosphorylation of the IR ¦Â-subunit and IRS-1 were significantly
decreased in muscle and liver tissues. This is the first study to
demonstrate the in vivo effects of changes in PTP1B activity
and expression on insulin signaling molecules in the skeletal muscles
and livers of HFD/STZ rats.
Using reduced carboxymethylated-maleylated (RCM)-lysozyme as a
substrate, Seiichi Tagami[27] and McGuire et al[28]
found increases in PTP1B activity in the skeletal muscles of insulin-resistant
Otsuka Long-Evans Tokushima fatty (OLETF) rats and insulin-resistant
human subjects, relative to the activity in the corresponding control
groups. Muscle PTP1B activity, measured with myelin basic protein
and the insulin-receptor cytoplasmic domain as substrates, was also
increased in an animal model of insulin-resistant obesity and DM[10].
Those findings accord with our results. It is reasonable to postulate
that increased PTP1B activity in the muscle and liver contributes
to insulin resistance and type II diabetes in HFD/STZ rats because
PTP1B inhibits tyrosine phosphorylation of the insulin receptor
and its substrates, including IRS-1.
Our own earlier ethnopharmacological studies of an aqueous extract
of Astragalus confirm the data reported for the plant A
membranaceus, which is traditionally used as an infusion of
(mainly) roots by the Chinese population to treat type II diabetes.
Most important of all, A membranaeus is an important component
of the majority of traditional herbal blend prescriptions used to
cure type II diabetes in traditional Chinese medicine. The unique
nature of the A membranaceus extract inspired us to work
towards the isolation of the active ingredient from the crude extract.
When subjected to sequential extraction, maximum activity was found
in APS. In the present study, we found that APS significantly reduced
both PTP1B protein levels and activity in the skeletal muscles,
but not in the livers of HFD/STZ rats. Consistent with this change,
the insulin-induced tyrosine phosphorylation of the IR ¦Â-subunit
and IRS-1 increased in the skeletal muscles, but not in the livers
of APS-treated TIIDM rats. Furthermore, blood glucose levels were
controlled and insulin sensitivity was apparently improved in APS-treated
TIIDM rats when KITT was used as an index of glucose
metabolism. This is a simple, reasonably accurate, and rapid method
for screening insulin resistance[29], and indicates the
net resistance to insulin at the target level, including receptor
and post-receptor defects. The data collected in this study indicate
that blood insulin levels are not altered by treatment with APS
in TIIDM rats, suggesting that the hypoglycemic effects of APS are
not mediated through insulin secretion. In addition, HFD/STZ rats
had a higher body weight than that of normal rats. The effect of
STZ injection plus high fat diet on body weight gain is in accordance
with the earlier findings[23,30]. APS could decrease
body weight in diabetic rats. It has been suggested that PTP1B can
act as a negative regulator of leptin signaling by dephosphorylating
the leptin receptor associated kinase Jak2[31,32]. So
one can postulate that the obesity resistance effect of APS be associated
with enhanced leptin sensitivity induced by its inhibitory action
on PTP1B.
PTP1B is a well-established drug target in the treatment of TIIDM
and obesity[33]. However, the PTP family of enzymes is
large, and all are highly specific for charged phosphotyrosine residues.
Finding a selective small-molecule inhibitor of PTP1B is therefore
proving difficult. Numerous PTP1B-inhibiting candidates are undergoing
trials, but significant success has yet to be achieved[34].
Our study showed that protein levels of PTP1B changed in parallel
with changes in its activities in skeletal muscle and liver. The
alterations in PTP1B activities may be partly due to changes in
its protein levels. Therefore, we postulate that the inhibitory
effect of APS on PTP1B activity is most likely mediated by the inhibition
of PTP1B protein expression. Detailed studies are in progress in
our laboratory to clarify the mechanism underlying the inhibitory
action of APS on PTP1B. In the present study, our data suggest that
APS induces normalization of PTP1B activity in the muscles, leading
to an improvement in insulin sensitivity. However, changes in PTP1B
activity alone may not account for the complete insulin-sensitizing
effect of APS. Other possible mechanisms cannot be excluded.
In conclusion, we have presented evidence to substantiate the anti-diabetic
and insulin-sensitizing effects of the A membranaceus
extract, APS. The insulin-enhancing effects of APS are at least
partially exerted through its effects on PTP1B in skeletal muscle.
The results of this study support this hypothesis, at least in this
animal model.
Acknowledgments
We thank R William CALDWELL, PhD, Ron DUNDORE, PhD, and Herman
RHEE, PhD for valuable discussion and critical review of the manuscript.
We also acknowledge the Young Investigator Award received by Yong
WU presented by the Division for Drug Discovery, Drug Development
and Regulatory Affairs of the American Society for Pharmacology
and Experimental Therapeutics.
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