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Department of Anatomy and Cell Biology, State University of New York, Health Science Center at Brooklyn, Brooklyn, New York 11203
Address all correspondence and requests for reprints to: Gladys Teitelman, Ph.D., State University of New York, Department of Anatomy and Cell Biology, Health Science Center at Brooklyn, 450 Clarkson Avenue, BSB2-94, Brooklyn, New York 11203. E-mail: gteitelman{at}hscbklyn.edu
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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cells),
insulin (ß cells), somatostatin (
cells), and pancreatic
polypeptide (PP cells), respectively. Insulin (IN) containing ß cells
form the core of the mature islets, which is surrounded by a rim of
, , and PP cells. Because the absence of ß cells leads to hyperglycemia and overt diabetes, a crucial question is whether the adult pancreas contains ß stem cells and, if so, the location of these cells in the tissue. One approach to answer this question is to characterize the phenotype of the presumptive precursor cells in embryos and then ascertain whether similar cells are present in adults. In the last few years, a molecular fingerprint of embryonic islet precursor cells has begun to emerge (Refs. 1, 2, 3, 4, 5, 6 and references herein). In embryos and young postnatal mice, precursors present in the pancreatic duct (7) migrate into the tissue parenchyma where they differentiate into mature islets. Previous studies indicated that the ductal precursors can be identified by the expression of the low affinity glucose transporter-2 (Glut-2) (8), which is also present in mature ß cells. This observation supported the view that ß cells are generated by Glut-2+ precursor cells (8). It is generally assumed that pancreatic duct cells retain the ability to generate endocrine cells and form new islets even late in life (reviewed in Ref. 9). Recently, cells expressing nestin, an intermediate filament protein expressed by neuronal stem cells, were located in pancreatic ducts of adult rats and were found to differentiate into IN-expressing cells in vitro (10). Whether the nestin+ cells display the ability to differentiated into mature ß cells in vivo remains to be determined.
There is also evidence suggesting the presence of presumptive ß precursor cells in mouse pancreatic islets that differentiate into insulin cells following injury. These cells differentiated in islets following depletion of the resident ß cell population by streptozotocin (SZ), a ß cell toxin. The first immature cell type to appear expressed somatostatin (SOM) and pancreatic and duodenal homeobox gene 1 (Pdx-1), a transcription factor expressed by islet progenitor cells (reviewed in Ref. 3) which it was followed in time by the appearance of cells coexpressing SOM and IN (10). It was proposed that the SOM/Pdx-1+ cells initiated IN expression, generating the SOM/IN+ cells that reappeared in islets following SZ treatment (11). However, the number of newly differentiated SOM/IN+ cells was low, they failed to differentiate into monospecific IN+ cells and the SZ-treated animals remained severely hyperglycemic at all stages examined.
High glucose levels have a negative effect on ß cell function (12) by decreasing the expression of Pdx-1 (13) and of ß cell genes, which are transactivated by Pdx-1, such as insulin, Glut2, glucokinase, a key enzyme in glucose metabolism, and islet amyeloid polypeptide (14, 15, 16, 17, 18, 19). These observations raised the possibility that hyperglycemia could also impair the differentiation of islet precursor cells into cells containing insulin. We reasoned that the reestablishment of normoglycemia would enhance ß cell neogenesis and maturation and tested this hypothesis in the present study. Our results indicate that prompt reestablishment of normoglycemia in SZ-treated mice by exogenously administered insulin allowed the differentiation of two sets of precursor cells in islets, the Glut-2 and Pdx-1/SOM cells, respectively, into IN-containing cells and the reappearance of morphologically normal islets.
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Materials and Methods |
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1) Hyperglycemic mice. These mice did not received IN treatment and were termed hyperglycemic x days post-SZ mice. They were examined at 1, 2, and 6 d post SZ (at least 6 animals per group).
2) Normoglycemic mice that received IN treatment from 1 d post SZ and were normoglycemic thereafter. These mice were injected with 2–3 U of NPH-ILETIN IN (Eli Lilly, Indianapolis, IN) ip in the afternoon of 1 d post SZ. The dosage of insulin was not adjusted to compensate for possible differences in body weight. Animals with blood glucose (bg) levels between 40 to 130 mg/dl at 2 d post SZ were considered normoglycemic. One group of normoglycemic mice was killed at 2 d post SZ (n = 40). Other normoglycemic 2 d post-SZ mice were anesthetized with Metofane (Schering-Plough Corp., Union, NJ), received two to three insulin implants (Linßit, Linshit Canada Inc., Ontario, Canada) following manufacturers’ instructions and were killed at 4 (n = 10) and 6 (n = 16) d post SZ.
Blood glucose levels were determined daily. From the morning of d 1 post SZ until the end of the experiment, all SZ mice were fed with Pregestimil, (Mead Johnson & Co., Evansville, IN), a hypoallergenic infant formula used to feed diabetic mice (20). Mice were perfused through the heart with 4% paraformaldehyde buffered to pH 7.4 with 0.1 M PBS. The fixed tissues were infiltrated overnight in 30% sucrose, mounted in embedding matrix (Lipshaw Co., Pittsburgh, PA) and 15–20 µm cryostat sections were collected onto gelatin-coated slides.
Source of antibodies and purified peptides
Primary antiserum. Guinea pig antibodies to bovine IN and
rat C-peptide were purchased from Linco Research, Inc.
(Eureka, MO). Rabbit antiserum to human glucagon was purchased from
Calbiochem (San Diego, CA). Rabbit antisera to human PP
and to somatostatin were supplied by Peninsula Laboratories, Inc. (Belmont, CA). Rabbit antisera to Ki 67p was purchased from
Novocastra. Mab antibody to rat PP was generously provided by
CURE/Gastroenteric Biology Center, Antibody/RIA Core (NIH Grant
DK-41301). Mab antibody to human glucagon was purchased from
Sigma (St. Louis, MO). Rabbit antisera to rat
L-amino acid decarboxylase (AADC) and to rat SOM were
purchased from Protos (New York, NY). Rabbit antisera to Glut2 was
purchased from Chemicon (Temecula, CA). Affinity purified antiserum to
the N-terminal domain of Pdx-1 was a generous gift from C. V. E.
Wright (21, 22). Antisera to peptide hormones were tested
by immunoadsorption and the dot blot technique according to criteria
previously described (23). Antibodies were used at the
following dilutions: guinea pig antibovine insulin and antirat
C-peptide, 1:400; rabbit antisera to Glut2, 1:1000; rabbit antihuman
glucagon, 1:12,000; rabbit antihuman somatostatin-1:8,000 for control
sections; and 1:20,000 for sections of SZ-treated pancreas; rabbit
antihuman pancreatic polypeptide, 1:100,000; rabbit antimouse PDX-1,
1:8000; rabbit antibovine AADC, 1:250; rat antisomatostatin, 1:2000;
Mab to glucagon, 1:6000; Mab to PP, 1:2000; rabbit anti Ki 67p,
1:1:1000.
Secondary antibodies. Biotinylated goat antirabbit IgG and avidin-labeled peroxidase were purchased from Vector Laboratories, Inc. (Burlingame, CA). Alexa Fluor 488 antimouse, antirat, and antirabbit IgG, Alexa Fluor 594 antiguinea pig, antirabbit and antimouse IgG and the nuclei acid dye Topro-3 were purchased from Molecular Probes, Inc. (Eugene, OR). Cy-5 donkey antirabbit and antimouse IgG were purchased from Jackson ImmunoResearch Laboratories, Inc. (West Grove, PA). For double-labeling using rabbit and guinea pig antisera, the secondary antibody used to visualize the guinea pig antibodies (purchased from Jackson ImmunoResearch Laboratories) did not cross-react with rabbit antibodies.
Immunolabeling of cryostat sections using peroxidase
techniques
These techniques have been previously described
(11). In brief, the sections were incubated sequentially
in an empirically derived optimal dilution of control serum or primary
antibody raised in species "X" containing 1% goat serum in
Tris-saline solution (TS; 0.9% NaCl in 0.1 M Tris, pH 7.4)
for 18 h; a 1:50 dilution of anti- (species x) biotinylated IgG
solution in 1% goat serum in TS for 30 min; and a 1:100 dilution of
peroxidase-avidin complex for 30 min (avidin-biotin complex: ABC
technique). Following these incubations, the bound peroxidase was
visualized by 3,3'-diaminobenzidine (DAB). After the DAB step, sections
were dehydrated and mounted with Permount (Fisher Scientific, Fairlawn, NJ).
Double label immunohistochemistry using two peroxidase
substrates
This technique allowed the simultaneous visualization of nuclear
and cytoplasmic antigens. Sections were incubated first with antisera
to PDX-1 and the bound antibody was visualized by DAB (brown
precipitate), followed by incubation with antisera to a hormone, which
was visualized with the blue reaction product of the Vector SG
substrate (Vector Laboratories, Inc.). Slides processed
for double immunohistochemical staining or for combined
immunohistochemistry and autoradiography (see below) were examined with
a Nikon Microphot SA microscope equipped with Nomarski
optics and using a 10x ocular and an oil immersion 100x
objective.
Determination of cell proliferation
For 5-bromo-2'deoxyuridine (BrdU) administration, mice were
injected ip with 200 mg/kg body weight of BrdU (Sigma, St
Louis, MO). Two hours after the injection, animals were perfused and
the pancreas collected and sectioned. Sections were first stained for a
hormone, rinsed and incubated with the corresponding IgG linked to an
Alexa fluorophore. Sections were rinsed overnight with 0.1
M PBS, pH 7.2, fixed with 4% paraformaldehyde for 30 min,
rinsed and treated with 2 N HCl at 37 C for 20 min and with
0.05 mg/ml pepsin in 0.1 N HCl at 37 C for 20 min. Sections
were then rinsed, blocked with 1:30 goat serum in PBS for 30 min, and
incubated overnight at 4 C with monoclonal antibody to BrdU (DAKO Corp., Carpinteria, CA; 1:100 diluted with a 1% solution of
goat serum in PBS). Sections were rinsed and incubated with goat
antimouse IgG labeled with Alexa fluorofore for 2 h. Sections were
mounted with coverslips using Prolong. In addition, in some
experiments, the presence of proliferating cells was examined in
sections processed for simultaneous visualization of a hormone and Ki
67p, a nucleolar protein expressed by cycling cells.
Confocal microscopy
For double and triple label immunofluorescence, SOM was
visualized with a rat antibody and Alexa Fluor goat 488 antirat IgG, IN
with a guinea pig antibody and Alexa Fluor 594 goat antiguinea pig IgG,
PDX-1 with a rabbit antibody and Cy5 donkey antirabbit IgG. PP,
glucagon (GLU), and SOM were visualized with a cocktail of specific rat
and mouse antisera and a mixture of antirat and antimouse Alexa Fluor
488. All secondary IgGs were used at 1:200 dilution and To-Pro 3 at 2
µM dilution for 30 min. After completion of the staining
procedure, sections were covered with two to three drops of Prolong
Antifade solution (Molecular Probes, Inc.) and were dried
at room temperature before examination.
A laser scanning confocal microscope, model LSM 510 (Carl Zeiss, Thornwood, NY), fitted with an Axiovert 100M microscope (Carl Zeiss) was used with a 63 x 1.4NA pan Apochromat objective (Carl Zeiss). Excitation on the laser scanning confocal microscope was with a 15 mW argon ion laser running at 75% power emitting at 488 nm, a 1.0-mW helium/neon laser emitting at 543 nm, and a 5.0 mW helium/neon laser emitting at 633 nm. Emissions were collected using a 505- to 530-nm band pass filter to collect Alexa green emissions, a 560- to 615-nm band pass filter to collect Alexa red emissions and a 650-nm long pass filter to collect Cy5 and To-Pro3 emissions. Typically, 0.7-µm vertical steps were used with a vertical optical resolution of <1.0 µm.
Processing of semithin sections
Severely hyperglycemic 1 d post-SZ mice (bg > 500
mg/dl) were anesthetized and the pancreas perfused through the common
pancreatic duct with fixative solution. This procedure produced a
distention of the pancreas that facilitated the dissection of tissue
surrounding the duct, which is rich in islets. The tissue was embedded
in epon and consecutive semithin sections (2 µm thick) were stained
with thionin. Although this technique gave clear cellular localization,
it precluded the survey of a large number of pancreatic islets. We
examined a total of 10 islets from two 1 d post-SZ mice.
Islet cultures
Mice were injected a solution of 200 mg/kg SZ prepared before
each individual injection. Studies in a test group of animals indicated
that this procedure assured the development of hyperglycemia (bg >350
mg/dl) in 100% of the injected mice (n = 30). Mice were
anesthetized with sodium pentobarbital 15 min after the injection, the
pancreas was perfused through bile duct with a 5 ml of a collagenase
solution (Worthington Biochemical Corp., Lakewood, NJ), 2
mg/ml in HBSS (Invitrogen, Carlsbad, CA), placed in a
Petri dish in a stationary 37 C water bath for 15 min, dissociated with
a 5 ml pipette, centrifuged, the pellet resuspended in 10 ml HBSS + 100
µl DNase [(Worthington Biochemical Corp.) 1 mg/ml in
HBSS: glycerol 1:1 vol/vol] and the islets handpicked under a
dissecting microscope. Islets were transferred to a 13-mm round
Thermanox coverslips (Nunc, Inc., Naperville, IL) placed in a 35-mm
plastic Petri dish and embedded in 20 µl of Matrigel (BD Biosciences,
Bedford, MA) diluted 1:1 with RPMI 1640 containing 5.6 mM
glucose. The dishes were placed in a CO2
incubator at 37 C for 30 min to allow the Matrigel to gel. Then, each
dish received 2 ml of culture media (RPMI 1640 containing 5.6
mM glucose, 10% heat inactivated FCS
(Invitrogen), 1% penicillin-streptomicin (10,000 U/ml)
and 15 mM HEPES buffer), and the dishes were incubated
overnight. The following day the cultures were fixed, processed for
immunostaining and examined by confocal microscopy.
Determination ß cell relative volume per islet and per tissue
To determine ß cell relative volume per islet and per tissue
each gland was sectioned throughout its length to avoid bias due to
regional variations in islet distribution. The relative volume of ß
cells per islet was determined in sections immunostained for insulin by
the point sampling method (24) using a 300 point ocular
grid according to the formula: F = h/n in which h was the number
of "hits" over ß cells and n was the number of points scored over
islets (24). The same formula was used to calculate the
relative ß cell volume per tissue. In this case, h was the number of
hits over ß cells and n the number of points over exocrine tissue
(24). Tissues were examined with a Nikon
Microphot SA microscope equipped with Nomarski optics and using a 10x
ocular and 40x objective.
Statistical analysis
All values are shown as mean ± SE. For
comparison between two groups, the unpaired t test (two
tail) was used. A P value < 0.05 was considered
significant.
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Results |
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). These
cells were located in the periphery of the islet, whereas the islet
core lacked stained cells but contained cell debris and scattered
nuclei (Fig. 1A). The islet rim is demarcated in Fig. 1B in a
consecutive section that was immunostained with a cocktail of antisera
against SOM, PP, and GLU. Examination of serial optical sections from
12 randomly selected islets from three 1 d post SZ/200 mice
confirmed the presence of 1-2 C-peptide+ cells
(not shown).
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Because it is known that chronic hyperglycemia induces ß cells degranulation and decreased levels of insulin, Glut-2 and Pdx-1 (14, 15, 16, 17), it could be argued that acute hyperglycemia, such as that in 1 SZ mice, had similar effects. If so, ß cells would become degranulated at 1SZ, thus evading detection by immunohistochemical staining. Following the reestablishment of normoglycemia by exogenous insulin injection, these cells would have restored the intracellular insulin concentration to normal levels and would stain again for insulin.
To address this issue, we sought to determine whether ß cells became
degranulated during acute hyperglycemia. First, we searched for cell
markers that continued to be expressed at high levels by islet cells of
hyperglycemic mice and, therefore, would stain degranulated ß cells.
One of the potentially useful markers was AADC, a neuronal enzyme
localized in the cytoplasm of the all the endocrine cells of the islet
(26, 27). We compared islets of obese mice with islets of
1d post-SZ/200 mice. In contrast to controls (Fig. 2, A and B), islets of 2-month-old obese
mice (n = 3; blood glucose levels between 250 and 270 mg/dl) were
lightly stained with insulin and PDX-1 (Figs. 2, C and D). This
staining pattern is characteristic of islets of chronically diabetic
animals (Ref. 28 and references therein). Islets of ob
mice, however, were darkly labeled with AADC (Fig. 2E). Similar levels
of AADC staining were observed in islets of controls (not shown),
indicating that AADC levels did not decrease with hyperglycemia. In
agreement with the immunofluorescent observations mentioned above,
examination of sections in which the bound insulin antibody was
visualized with DAB revealed that the core of islets of hyperglycemic
1 d post-SZ/200 mice (blood glucose levels > 350 mg/dl)
lacked ß cells but contained insulin-stained cell debris (Fig. 2F)
and was surrounded by PDX-1+ cells (Fig. 2G).
Importantly, in these mice, AADC+ cells were
found in the periphery but not in the center of the islets (Fig. 2H),
confirming that most, if not all ß cells, the endocrine cells present
this location, were deleted by the toxin.
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). Thus, a dose of 200 mg/kg eliminated almost all the
ß cells whether or not the islets were exposed, as in
vivo, to hyperglycemia. Taken together, these observations
demonstrate that ß cells would have been detected by immunostaining
if they were alive just before fixation of the tissue. Therefore, we
conclude that the almost total absence of ß cells in islets of
1SZ/200 was due to the fact that they were eliminated by the toxin.
Reappearance of morphological normal islets in normoglycemic SZ +
IN mice
To ascertain whether insulin-induced normoglycemia promoted ß
cell neogenesis, pancreas from 2 d post-SZ mice were examined. One
group consisted of mice that were injected with 200 mg/kg SZ at d 0,
with eluent at 1 d post SZ and killed the following day. These
mice were hyperglycemic at 1 and 2 d post SZ. A second group of
SZ-treated mice were injected with insulin at 1 d post SZ and
attained normoglycemia (bg levels between 40 and 120 mg/dl) at 2 d
post SZ. Pancreas of hyperglycemic 2 d post-SZ mice had islets
that were almost completely devoid of ß cells and still contained
peroxidase stained cellular debris (Fig. 3B). In contrast to the hyperglycemic
mice, more than half of the normoglycemic 2 d post SZ + IN mice
(25 out of 40) had islets that contained 5–60 insulin cells per islet
section (Fig. 3, C and D). Some of those islets contained
IN+ cells only in the periphery while, in other
islets, IN+ cells were also located in the core.
The core of these islets lacked cellular debris, which was replaced by
a bed of fibroblast-like unstained cells (Fig. 3D).
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). The ß cells stained with antibodies to either
insulin (Fig. 3) or C-peptide (not shown) indicating that the newly
formed ß cells were able to process proinsulin to completion. The
core of insulin-containing cells was partially surrounded by endocrine
nonß cells, as determined by SOM staining (Fig. 3F). At this time the
islets were irregularly shaped, suggesting that they had not fully
recovered from the SZ-induced injury. While in most instances islets of
6 d post-SZ mice contained a significant number of ß cells, in
only 5 out of 16 cases the islets were completely repopulated by ß
cells (Fig. 3E). This indicates that the success of recovery was not
uniform among normoglycemic mice, although each pancreas had similar
degree of islet reformation. Because SZ damages many tissues in
addition to the pancreas (29), it is probable that, in
most SZ-treated mice, its deleterious effect was only partially
reversed by the exogenous insulin treatment.
Morphometric analysis confirmed that, in addition to promoting rapid
islet recovery from injury, the restoration of euglycemia induced an
increase in the relative volume of ß cells per islet area. The effect
of insulin induced normoglycemia was already evident at 2 d post
SZ (24 h after injection of the hormone) because the relative volume of
ß cells per islet was almost 20-fold higher in normoglycemic mice
than in hyperglycemic 2 d post-SZ mice (Fig. 4, Ref. 11).
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). The relative ß cell volume per tissue also increased from 2% of
control values at 2 d post SZ to 40% of control values at 6
d post SZ (Fig. 5). These findings
indicates that insulin-induced normoglycemia promoted an early burst in
ß cell regeneration but that the number of newly formed ß cells per
islet did not reached control levels during the time period of this
study.
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), indicating that
the increased ß cell mass found during the first 4 d after SZ
was not due to proliferation of the insulin containing cells present at
those stages. Islets of 2 d post-SZ mice contained a significant
number of proliferating nonß cells. Some of these proliferating cells
were macrophages (Fig. 6B). Other proliferating cell types were the
(not shown) and (Fig. 6C) cells and cells located in the periphery
of the islet that expressed Glut-2 (Fig. 6D).
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, E–G) and
during regeneration (11) in hyperglycemic mice. The
present study confirmed the appearance of Pdx-1/SOM cells in SZ + IN
mice. The percentage of Pdx-1/SOM+ cells in
normoglycemic 2 d post SZ (60%, Fig. 7) was higher than in 2 d post-SZ
mice that did not receive the hormone and remained hyperglycemic (30%,
this study and Ref. 11). The percentage of
Pdx-1/SOM+ cells in normoglycemic SZ + IN mice
did not decrease with time, remaining at similar high levels at 6
d post SZ (Fig. 7). In contrast, no difference was found in the
percentage of Pdx-1/GLU+ and
Pdx-1/PP+ cells between experimental animals and
untreated controls (not shown).
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). While some islets contained several
SOM/IN+ cells (Figs. 8, A and B) in addition to the cells containing
only SOM, other islets in the same pancreas were populated mainly by
monospecific ß and cells (not shown).
SOM/IN+ cells were not detected in islets of
hyperglycemic 1 and 2 d post-SZ/200 mice or controls (data not
shown). No cells expressing GLU/IN (Fig. 8, C and D) or PP/IN (not
shown) were seen at this or other stages after SZ or in controls. These
observations indicate that cells coexpressing SOM and IN appeared
3 d earlier in normoglycemic 2 d post SZ + IN mice than in
animals that did not receive IN treatment (11). In
addition, islets of normoglycemic 2 and 6 SZ + IN mice also contained a
significant number of cells expressing only IN, a cell type that was
scarce in islets of hyperglycemic of 5 d post-SZ mice
(11).
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). The fact that in
controls Glut-2 expression is restricted to ß cells (Figs. 9A and 10, A–C), this observation confirmed
the toxin-mediated elimination of the original ß cell population.
Cells expressing Glut-2 in the cytoplasm reappeared in the periphery of
islets of normoglycemic 2 d post-SZ mice (Fig. 9C). At 4 d
post SZ, many islet cells distributed throughout the islet expressed
Glut-2 either in the cytoplasm or the cell membrane (Fig. 10H), whereas
at 6 d post SZ, most islet cells displayed membrane staining (Fig. 9D). These observations suggested a translocation of the transporter
from the cytoplasm to the cell membrane during cell maturation. In
contrast to euglycemic mice, islets of hyperglycemic 2 d post-SZ
mice (no insulin treatment) did not have Glut-2+
cells (Fig. 9E). In hyperglycemic mice, Glut-2+
cells were first seen at 6 d post SZ (Fig. 9F). However, the
number of Glut-2+ cells was significantly lower
in hyperglycemic than in normoglycemic animals at that stage and the
transporter had a diffuse, cytoplasmic localization. These observations
indicated that the reestablishment of normal blood glucose levels
accelerated the time of appearance of Glut-2+
cells in islets and promoted the maturation of these cells.
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, D–F) and the number of Glut-2/IN+
cells increased with time (Fig. 10, G–I). Islets of hyperglycemic
6 d post SZ + IN mice contained IN+
cells and some of these cells coexpressed Glut-2 (Fig. 10, J–L).
However, in these cells the transporter had a cytoplasmic localization,
indicating that they were not functionally mature.
At 2 d post SZ, islets also contained a significant number of
Glut-2/GLU+ cells and cells that expressed only
Glut-2 (Fig. 11). At this and later
stages, Glut-2 was localized to the cytoplasm of the cells. Glut-2
expression by GLU cells persisted in islets of pancreas with suboptimal
regeneration even at 6 d post SZ, but was transient in pancreas
with fully reformed islets (not shown). This observation indicated that
the inhibition of Glut-2 expression by GLU was correlated with the
success of the regeneration process. No Glut-2 cells expressed SOM or
the macrophage marker 4–80 (not shown). We did not test for
coexpression of Glut-2 and PP. As expected ß, but not cells of
controls expressed Glut-2 (Fig. 11, D–F).
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, A–C) in
addition to the Glut-2/IN cells reported by others
(8). We also confirmed the presence of cells coexpressing
SOM and IN at that stage of development (Ref. 32 and Fig. 12, E–G). These observations indicate that at least two sets of
embryonal cell types reappear in islets following SZ-induced injury,
i.e. the SOM/IN+ and the
Glut-2+ cells. Because both
SOM+ and Glut-2+ cells
proliferate, it is likely they generate the newly differentiated
IN+ cells. It has recently been reported that
adult rat and human islets contain nestin + precursor cells that
differentiate into insulin-containing cells in vitro
(33). It remains to be determined whether the Glut-2 and
SOM/IN+ cells of SZ-treated islets are generated
by precursor cells expressing nestin. |
Discussion |
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One of the possible ß cell precursors found in this model of neogenesis were the Glut-2+ cells that appeared in islets of SZ-treated normoglycemic mice. It has been previously reported that Glut-2 expression in ß cells was regulated by glucose levels and that it decreased during hyperglycemia (17, 28, 34, 35). Because islets of hyperglycemic 1 d post SZ lacked Glut-2+ cells, it could be argued that the absence of these cells could be due to the high circulating glucose levels. If this were true, ß cells could re-express the transporter following the restoration of euglycemia. However, the cells expressing Glut-2 in normoglycemic 2 d post-SZ mice are located in the periphery of the islets, the location of the nonß cells rather than in the core of the islet, supporting the view that this was a novel population of cells. Indeed, one unexpected finding in our study was that a significant number of Glut-2 cells present at 2 d post-SZ mice coexpressed GLU, a cell type unique to embryos and to islets of SZ mice. The fact that Glut-2 expression appeared sooner and was more abundant in islets of normoglycemic than hyperglycemic SZ-treated mice indicate that differences in glucose levels affected the initiation of Glut-2 expression. Expression of Glut-2 by GLU cells was transient and it disappeared at 6 d post SZ in mice with optimal regeneration but persisted in mice in which regeneration was less successful. Because mice with different degrees of regeneration were euglycemic, the fate of Glut-2 expression in nonß cells was related not only to variations in glucose levels but also to other unidentified factors affecting the outcome of the process of regeneration.
In addition to the Glut-2/GLU+ cells, islets of normoglycemic post-SZ mice contain Glut-2+/IN+ cells and the number of these cells increased with time. Our results suggest that cells expressing only Glut-2 appeared in islets of normoglycemic 2 d post SZ +IN mice and that, as in embryos, these cells were ß cell precursors and initiated IN synthesis. Because IN+ cells did not proliferate during the initial stages of regeneration, it is likely that the temporal increase in ß cell number was due to the augmentation in the number of Glut-2 cells that initiated IN expression. In contrast to IN cells, Glut-2 cells proliferated actively and could have generated ß cells without depleting the size of the precursor pool. In addition, GLU cells expressing the transporter could have converted into ß cells. However, this is improbable because islets of SZ-treated mice lacked cells coexpressing IN and GLU. Glut-2/IN+ cells appeared in islets of hyperglycemic 6 d post-SZ mice, but these cells were scarce and immature, suggesting that the presence of high circulating glucose levels prevented their maturation.
The restitution of euglycemia also promoted the differentiation of the PDX-1/SOM+ cells, a presumptive ß precursor cell type, into monospecific ß cells. Thus, islets of normoglycemic 2 d post SZ + IN mice contained a high percentage of PDX-1/SOM+ cells, and also had SOM/IN+ cells and monospecific IN+ cells. SOM/IN+ cells were also found during development but not in islets of adult, untreated mice. Conceivably, the process of transformation of SOM+ cells into insulin containing ß cells in normoglycemic SZ mice occurred in at least three separate stages. The first step would be the activation of PDX-1 expression in SOM+ cells and/or the stimulation of undifferentiated precursors present in or around islets to initiate PDX-1 and SOM expression. The second step would be the initiation of insulin expression by PDX-1/SOM+ cells and, in the third stage of the conversion, SOM/IN cells down regulated SOM expression and differentiated into fully mature, Glut-2+ ß cells. We previously reported that cells expressing SOM and IN eventually appeared in islets of hyperglycemic mice at 5 d post SZ, but these islets contained very few cells expressing only insulin (11). These observations and our present findings strongly suggest that hyperglycemia inhibited the maturation of the two types of precursor cells that appeared in islet of SZ-treated mice, namely Glut-2 and SOM/IN cells, into functional ß cells.
The previous discussion underscores the dramatic effect of the
elimination of ß cells by SZ treatment on the non-ß cells of the
islet. After exposure to SZ, both and cells reverted to an
immature phenotype characteristic of these cells during development.
Thus, the cells initiated expression of Pdx-1 and IN while the
started expressing Glut-2. In addition, following SZ-induced injury,
precursors present in islets presumably activated Glut-2 expression and
differentiated into ß cells. Although the signals responsible for
these phenotypic changes are presently unknown, the cellular
transformations did not occur until the islets were free of cellular
debris. Thus, in contrast to islets of normoglycemic 2 d post SZ,
which were mostly free of debris and showed regeneration, islets of
hyperglycemic mice at that stage still contained many dead ß cells
and lacked newly differentiated IN+ cells.
The removal of dead cells is likely to be accomplished by macrophages invading the islet following SZ treatment. Islets of both normoglycemic and hyperglycemic mice contained more macrophages than islets of controls (Refs. 35, 36, 37, 38 and this study). However, high glucose levels decrease macrophage phagocytic activity and affects the pattern of cytokine secretion (39, 40, 41). Our results suggests that, in normoglycemic mice, macrophages would rapidly remove cell debris and then secrete molecules that activate the proliferation and differentiation of precursor cells while hyperglycemia would delay or eliminate these activities. Macrophage-derived cytokines could be involved in the observed activation of proliferation of several islet cell types at 2 d post SZ and in the increase in the number of Pdx-1/SOM+ cells. The presence of a high percentage of Pdx-1/SOM cells in islets of nonobese diabetic mice (11), which are invaded by cells secreting proinflammatory cytokines (42), supports an involvement of hematopoietic cells in the regulation of Pdx-1 expression by SOM cells. Differences in macrophage function could also determine the appearance of Glut-2+ precursor cells in islets of normoglycemic but not of hyperglycemic mice. In addition to circulating blood glucose levels, macrophages respond to a multitude of signals that regulate their function (43), a response that could be skewed in the SZ-treated euglycemic mice. This would explain the fact that some, but not all of the normoglycemic mice showed islet regeneration.
Contrary to our expectation, the percentage of
SOM/Pdx-1+ cells did not decrease in regenerated
islets of 6 d post-SZ mice that appeared to have recovered from
the SZ-induced injury. This suggests that the mechanism/s involved in
the activation of Pdx-1 expression in SOM cells persist for a long time
in regenerated islets. Alternatively, it is possible that
Pdx-1/SOM+ cells were unable to down-regulate
Pdx-1. The fact that in embryos, a large percentage of cells
express Pdx-1 when they first differentiate and that this percentage
decreases during development (30, 31) indicates the
presence of mechanisms inhibiting Pdx-1 expression in
SOM+ cells. Conceivably, these regulatory
mechanisms were not active in the Pdx-1/SOM+
cells that appeared in adult islets following injury. Finally, because
the number of ß cells in regenerated islets at 6 d post SZ was
lower than in controls, it is also possible that the presumptive
precursor cells were still being generated even at that stage, and the
islets were primed to grow until they reached control values.
Our proposition that insulin therapy induced ß cell neogenesis is in agreement with previous reports indicating that insulin administration to SZ-treated neonatal rats promoted the recruitment of new ß cells from a precursor population (44). The question remains as to whether insulin directly promotes ß cell regeneration or if it acts indirectly through its role in the normalization of blood glucose levels. Recent studies on mildly diabetic SZ-treated rats indicated that short-term hyperinsulinemia had no effect on ß cell mass, whereas a 48 h glucose and insulin infusion sufficed to restore ß cell number to control values (45). The findings reported by Bernard et al. (45) and others (46) support the view that both mild hyperglycemia and hyperinsulinemia were required for maximal ß cell growth. In our model of acute hyperglycemic mice, the success of the regeneration process was dependant upon the restoration of normal glucose levels by exogenous insulin injection. However, due to the technique used to regulate blood sugar levels, we cannot rule out the possibility that the mice underwent periods of hyperglycemia and hyperinsulinemia, which could have had a positive effect on ß cell regeneration. Taken together, these observations stress the need for further examination of the roles of both insulin and glucose in the SZ and other models of ß cell neogenesis.
In conclusion, we found that islets of adults contain ß precursor cells and that the re-establishment of normoglycemia by exogenously administered insulin accelerates the differentiation of these cells into monospecific ß cells and the reappearance of morphologically normal islets less than 1 wk after SZ treatment. Two types of ß precursor cells were identified, the Glut-2+ cells and cells coexpressing PDX-1/SOM. It remains to be determined whether the neoformed ß cells have all the properties characteristic of mature insulin containing cells and are able to regulate circulating glucose levels within the physiologically normal range.
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Acknowledgments |
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Footnotes |
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Abbreviations: AADC, Aromatic L-amino acid decarboxylase; bg, blood glucose; BrdU, 5-bromo-2'deoxyuridine; DAB, 3,3'-diaminobenzidine; GLU, glucagon; Glut-2, glucose transporter-2; IN, insulin; Pdx-1, pancreatic and duodenal homeobox gene 1; PP, pancreatic polypeptide; SOM, somatostatin; SZ, streptozotocin.
Received April 11, 2001.
Accepted for publication July 26, 2001.
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