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Section of Islet Transplantation and Cell Biology, Joslin Diabetes Center, Boston, Massachusetts 02215
Address all correspondence and requests for reprints to: Gordon C. Weir, MD, Section of Islet Transplantation and Cell Biology, Joslin Diabetes Center, One Joslin Place, Boston, Massachusetts 02215. E-mail: gordon.weir{at}joslin.harvard.edu
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Isolation of islets from adult pigs
Porcine pancreases from female Yorkshire pigs weighing 350–450
pounds were obtained at a local slaughterhouse (Blood Farms, West
Groton, MA). For islet isolation, the gland was removed from the
Eurocollins solution and trimmed of extraneous tissue. The pancreas was
then distention by infusion with collagenase (Liberase PI, 0.5 mg/ml,
Roche Molecular Biochemicals, Indianapolis, IN) suspended
in modified University of Wisconsin-D solution (18). The
gland was digested at 37 C with static incubation for 25 to 65 min.
Based on visual inspection of the gland, digestion was terminated by
the addition of cold M-199 media containing 10% newborn calf serum, 25
mM HEPES, 100 IU/ml of penicillin, and 100 µg/ml of
streptomycin. The digested tissue was passed over a bed of 4–6 mm
glass beads and through a 310 µm stainless steel mesh screen. The
tissue effluent was collected in 250 ml conical tubes and centrifuged
for 2 min at 700 rpm at 4 C. The islets were purified using a
discontinuous Ficoll 400 DL gradient (Sigma) with specific
gravities of 1.110, 1.096, and 1.060 respectively. Following
centrifugation, the islets were collected from interface between 1.060
and 1.096. Purified islets were washed, pooled, and aliquoted for
determination of islet yield with conversion to 150 µm
i.e. (17). The purity of islets ranged from
94–97%. This was determined by visual inspection with a light
microscope of dithizone stained islets isolated from seven adult pig
pancreases.
Transplantation, follow-up, and graft removal
Normoglycemic and diabetic male Swiss Webster nude mice
(Taconic Farms, Inc., Germantown, NY), aged 6–8 weeks,
were used as graft recipients. Diabetes was induced by ip injection of
STZ (250 mg/kg; Sigma), and only mice with blood glucose
>350 mg/dl at 1 wk were used. NPCCs (4000 IE) were transplanted
beneath the left kidney capsule under methoxyflurane anesthesia
(Metofane, Mallinckrodt Veterinary Inc., Mundelin, IL),
using PE-50 tubing (Becton Dickinson and Co.) connected to
a 1 ml Hamilton syringe (7). After transplantation, body
weight and fed blood glucose levels were determined weekly. In a third
group of nontransplanted normoglycemic Swiss nude mice, body weight and
blood glucose were measured weekly along with the transplanted groups
for 20 weeks. Blood glucose was measured with a portable glucose meter
(Precision QID, Medisense Inc., Bedford, MA) in blood obtained after
tail snipping. For measurement of porcine Cpeptide levels at 10
days, 6 weeks, and 20 weeks after transplantation, blood samples from
snipped tails were collected into heparinized capillary tubes on ice.
The capillary tubes were spun down in at 3000 rpm for 3 min and
supernatant plasma was collected and stored at -80 C until assayed for
C-peptide using RIA (Linco Research, Inc., St. Charles,
MO). Kidneys bearing the grafts were removed under methoxyflurane
anesthesia at 10 days, 6 weeks, and 20 weeks after transplantation.
Adult pig islets, 5000 or 10,000 IE were transplanted under the left
kidney capsule of diabetic nude mice and these grafts were removed 10
days, 6 weeks, or 20 weeks after transplantation. Diabetic mice
becoming normoglycemic following transplantation of either NPCCs or
adult islets underwent nephrectomy and were followed to verify a rise
in blood glucose level after graft removal. All other mice were killed
at the time of graft removal.
Graft insulin content
Insulin content was measured in grafts contained in kidneys
removed 10 days, 6 weeks, and 20 weeks after transplantation into
normoglycemic recipients. The kidneys were homogenized (Ultra-Turrax
T-25, IKA-Works, Cincinnati, OH) in 5 ml acid-ethanol, then stored
overnight at 4 C followed by centrifugation at 2500 rpm for 30 min. The
supernatant was removed and stored at -20 C pending measurement of
insulin content by RIA (Linco Research, Inc.).
Preparation of NPCCs, grafts, and pancreas for histology
Kidneys with NPCC grafts were fixed overnight in 10% buffered
formalin (Fisher Scientific, Fairlawn, NJ). Following
fixation, each kidney was sliced perpendicular to its long axis leaving
only the portion of kidney containing the grafts. These pieces of
tissues were stored in 0.1 M Sorenson’s buffer at 4 C
until embedded in paraffin. Pretransplantation NPCCs (approximately
2000 IE) were spun down (1200 rpm x 2 min) in
microcentrifuge tubes and the supernatants removed. The NPCCs were then
washed gently with PBS (x2) before being fixed in a 10% buffered
formalin solution (Fisher Scientific) for 30 min. NPCCs
were then suspended in 0.5 ml of a 2% agar solution in microcentrifuge
tubes and rapidly spun down to form pellets. The pellet was stored in
0.1 M Sorenson’s buffer until being embedded in
paraffin. For histological examination of neonatal pig pancreas small
pieces of pancreas obtained from head (n = 2) and tail (n =
2) were fixed for at least 6 h in 10% buffered formalin.
Immunofluorescent staining
Serial sections (5 µm) of the kidneys bearing grafts were
obtained and the first of these sections was stained with hematoxylin
for determination of graft size, location and infection. Double
immunofluorescent staining was performed in the rest of the graft
sections for morphometric analysis.
To quantify the percentage of ß-cells proliferating in the grafts,
sections were double-stained for Ki-67 and insulin (Fig. 5
). Ki-67, a
known marker of cell proliferation, is a protein that is expressed
during the cell cycle from mid-G1 until mitosis (19, 20, 21).
For immunostaining, sections were deparaffinized with xylene, and then
rehydrated in decreasing concentrations of ethyl alcohol; for antigen
retrieval, sections were microwaved in 0.01 M citrate (5
min x 3) and allowed to cool to room temperature. Sections were
incubated overnight at 4 C with anti Ki-67 polyclonal antibody (1:200
dilution, rabbit antihuman, Novocastra Laboratory, Newcastle
Upon Tyne, UK). After washing, indirect immunofluorescent staining was
performed by incubation with biotinylated secondary antibody
(Vector Inc., Burlingame, CA) for 1 h at room
temperature followed by FITC-Streptavidin (1:200 dilution, 1 h at
room temperature, Jackson ImmunoResearch Laboratories, Inc., West Grove, PA). Insulin staining was performed after
Ki-67 using direct immunofluorescence with 1 h incubation at room
temperature with anti-insulin antibody (1:200 dilution, Guinea pig
antihuman, Linco Research, Inc.), followed by incubation
with Texas Red-anti Guinea pig IgG for 1 h at room temperature
(1:200 dilution, Jackson ImmunoResearch Laboratories, Inc.). For evaluation of duct cell proliferation,
double-staining for cytokeratin 7 (CK7, duct cell marker) and Ki-67 was
performed (Fig. 6, panel B). For CK7 staining, anti CK7 antibody (1:100
dilution, mouse antihuman, DAKO Corp., Glostrup, Denmark)
was incubated overnight at 4 C. Amplification and indirect
immunostaining was performed using biotinylated secondary antibody for
1 h at room temperature (Vector Inc., Burlingame, CA) followed by
FITC-Streptavidin for 1 h at room temperature (1:200 dilution,
Jackson ImmunoResearch Laboratories, Inc.). Ki-67 staining
was performed on CK7 stained sections by overnight incubation with
polyclonal anti Ki-67 antibody (1:200 dilution, Novocastra Laboratory)
followed by Texas Red antirabbit IgG (1:200 dilution, Jackson ImmunoResearch Laboratories, Inc.) for 1 h at room
temperature. The relative proportions of ß- and duct cells were
assessed using sections double-stained for insulin and CK7.
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Determination of ß-cell cross-sectional area and ß-cell
number
From each immunostained graft section, double-stained
fluorescent images (range 7–13 images) of nonoverlapping fields (920x
final magnification) were captured systematically. Immunofluorescent
images were acquired on a Zeiss Laser Scanning Microscope 410
(nonconfocal mode) with an argon/krypton laser at excitations of 488 nm
and 568 nm using appropriate emission filters for simultaneous two
color immunofluoroscence with FITC and Texas Red. Brightness and
contrast parameters were optimized for each fluorochrome to cover the
entire range of intensity but were kept consistent for each
experiments.
Measurement of mean cross-sectional area was performed using IP Lab Spectrum software (Scanalytics Inc., Fairfax, VA). The mean cross-sectional area of individual ß-cells in a graft section, a measure of ß-cell size, was calculated after determining areas of randomly selected ß-cells having clear outlines (minimum of 30 ß-cells from 5–15 images). To determine the total number of ß-cells, the total area occupied by insulin staining was divided by the mean area of individual ß-cells.
Proliferation index of ß-cells and duct cells
The number of Ki-67 positive ß-cells in an image was
determined by manual counting. For calculating the proliferation index,
the minimum numbers of duct and ß-cells assessed at different time
points were 400, 600, and 1000 cells at 10 days, 6 weeks, and 20 weeks
after transplantation respectively, which gave us the desired relative
probable error of 15%.
The proliferation index of ß-cells in grafts was calculated after
determining the total number of ß-cells and total number of Ki-67
positive ß-cells from all images by:
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Maturation index of grafts
Estimation of relative area of ß-cells and duct cells of the
graft sections (maturation index) was performed by counting of CK7
positive duct cells, insulin positive ß-cells and cells staining for
both insulin and CK7 (costained cells) in sections double stained for
insulin and CK7:
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Apoptosis of duct cells
Apoptosis of the duct cells was determined in grafts 10 days
after transplantation by manually counting the total number of duct
cells and those with apoptotic nuclei under an immunofluorescent
microscope (630x magnification). Nonoverlapping fields were selected
systematically starting from one end of the graft and moving to other.
Between 453 and 607 duct cells were assessed with propidium iodide
staining in each of the 8 grafts that were studied.
Statistical analysis
All data are presented as mean ± SEM.
Statistical analysis was performed using Prism software GraphPad Software, Inc. (San Diego, CA). Comparison among three or more
groups was performed with one way ANOVA using Tukey’s or Dunnett’s
post hoc test. Paired and unpaired Student’s t tests were
also employed. A P value of <0.05 was considered
significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). The majority of mice became
normoglycemic (<200 mg/dl) after transplantation when followed for 20
weeks (5 of 6 mice); however, no mice (0 of 28 mice) became
normoglycemic at 10 days and only 5 of 20 mice became normoglycemic
between 10 days and 6 weeks of transplantation. The number of mice
decreased in the later time points after transplantation because
several mice were killed following graft explantation at earlier time
points to randomize the experimental protocol. In the subset of
diabetic mice that achieved normoglycemia between 10 days and 6 weeks
of transplantation, at 6 weeks their blood glucose levels (98 ±
13 mg/dl) were significantly lower than those of normoglycemic
recipients (134 ± 9 mg/dl) and normoglycemic nontransplanted mice
(136 ± 5 mg/dl, Fig. 1). Interestingly, blood glucose levels were
significantly lower in transplanted normoglycemic mice compared with
nontransplanted normoglycemic mice at all time points from 9 through 20
weeks after transplantation (Fig. 1). Furthermore, there was a
significant decline in blood glucose levels at all time points after 9
weeks of transplantation into normoglycemic mice compared with
pretransplantation levels (P < 0.05 by paired
t test, Fig. 1). Animals in all groups gained weight at a
similar rate after transplantation compared with normal nontransplanted
controls (data not shown).
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Plasma C-peptide and graft insulin content
Plasma porcine C-peptide levels were measured after
transplantation of 4000 IE NPCCs in both diabetic and normoglycemic
recipients (Fig. 2). At 6 and 20 weeks
after transplantation, the plasma C-peptide levels were higher in cured
diabetic mice than in those with persistent hyperglycemia (Fig. 2A). In
normoglycemic recipients, plasma C-peptide levels were higher 20 weeks
after transplantation than at 10 days and 6 weeks after transplantation
(Fig. 2B). Similarly, graft insulin content in normoglycemic recipients
also increased with time and was 20-fold higher at 20 weeks compared
with pretransplantation NPCCs (Fig. 3).
Graft insulin content in cured diabetic mice transplanted with 10,000
IE of adult pig islets was significantly lower (44.8 ± 6.6 µg,
n = 4) compared with the insulin content of 10,000 IE of
pretransplantation adult pig islets (140.4 ± 6.6 µg, n =
7, P < 0.01 by t test).
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). Following transplantation, the
ß-cell proliferation index progressively fell (Fig. 4). At 10 days
(Fig. 5A
), approximately 9% of ß-cells
are in cell cycle, which is not different than that of ß-cells in the
native neonatal pig pancreas (Fig. 6A),
but by 6 weeks only 5–6% of ß-cells are in cell cycle (Fig. 4, Fig. 5B). By 20 weeks, few Ki-67 positive ß-cells are seen even though at
this time the graft mainly consists of ß-cells (Fig. 5C). In
recipients with persistent hyperglycemia 6 weeks after transplantation,
the ß-cell proliferation index was higher than that in grafts from
normoglycemic and cured diabetic recipients (Fig. 4).
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) than proliferation
index of the duct cells in NPCCs pretransplantation (5.6 ± 0.5%,
n = 4), or in the grafts of mice 10 days after transplantation,
regardless of whether they were normoglycemic (7.3 ± 1.3, n
= 4) or diabetic recipients (5.6 ± 1.3%, n = 4;
P < 0.01 by ANOVA using Tukey’s post hoc test).
Proliferation of duct cells was not assessed in grafts at 6 weeks and
20 weeks after transplantation because so few duct cells were present
in the grafts at these time points. There was little evidence of apoptosis in duct cells evaluated in 10-day-old grafts of both diabetic and normoglycemic recipients. Four grafts were studied from each group. From the group of normoglycemic recipients, a total of 2018 duct cells were examined, and only 3 could be identified as apoptotic, as determined by nuclear fragmentation and condensation seen with propidium iodide staining. From the grafts of four diabetic recipients, a total of 2188 duct cells were evaluated with only 5 showing evidence of apoptosis. Assessment of apoptosis could not be performed in 6-week-old grafts because of the paucity of duct cells at that time point.
Adult pig islet grafts
In adult pig islets, the ß-cell proliferation index before
transplantation was much lower than that found in NPCCs (Fig. 7). Interestingly, the ß-cell
proliferation index was greatly increased in grafts from diabetic
recipients with persistent hyperglycemia, both at 10 days and 6 weeks
following transplantation, being 3- to 5-fold higher than in diabetic
recipients that became normoglycemic (Fig. 7). In cured diabetic
recipients, the ß-cell proliferation index was higher at 10 days
(0.9 ± 0.1%) after transplantation compared with
pretransplantation islets (0.3 ± 0.1%, P <
0.05, Fig. 7). However, at 6 weeks and 20 weeks after transplantation
the ß-cell proliferation index in the grafts was back to the low
level found in the pretransplantation islets (Fig. 7).
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, Fig. 8). The maturation index increased from
about 25% in the pretransplantation NPCCs to about 50% and 85% of
graft mass at 10 days and 6 weeks, respectively, in both diabetic and
nondiabetic recipients (Fig. 8). At 20 weeks after transplantation,
there were very few CK7positive cell, and most cells stained for
insulin. There was no significant difference in the graft maturation
between normoglycemic and diabetic recipients. With regards to
protodifferentiation, at 10 days after transplantation about 20%
of duct cells costained for CK7 and insulin in both diabetic and
normoglycemic recipients.
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). The cross sectional area of ß-cells
was higher at 20 weeks than in neonatal pancreas, 10-day-old grafts and
6-week-old grafts in noncured diabetic and normoglycemic recipients
(Table 1). Diabetic recipients cured at 6 weeks after transplantation
also had significantly higher mean ß-cell cross-sectional area
compared with that of neonatal pig pancreases and 10-day-old grafts in
normoglycemic recipients (Table 1).
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, P < 0.05, t test). The
cross-sectional area of individual ß-cells from adult pigs tended to
decrease at 10 days post transplantation followed by a significant
increase at 6 weeks post transplantation (Table 1). There was no
significant difference in the cross-sectional area of individual
ß-cells in grafts from cured and uncured diabetic recipients (Table 1). |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Graft insulin content increased over time in nondiabetic recipients, increasing by 20-fold in 20-week-old grafts, compared with pretransplantation NPCCs. The decrease in graft insulin content found 10 days after transplantation could be due to loss of some ß-cells that is known to occur in the transplantation process (23). At 20 weeks after transplantation of 4000 IE NPCCS in the present study, the graft insulin content (36 ± 6 µg) was similar to the insulin content of 4000 IE pretransplantation adult pig islets (38.4 ± 6.5 µg) reported by Davalli et al. (7). We have previously shown estimated the mass of successful grafts of NPCCs in nude mice to be 4 mg (14). Thus, the insulin content per IE would be about 17 ng, which is somewhat lower the values we usually obtain for fresh adult porcine islets (about 22 ng/IE, unpublished observation).
Proliferation was assessed with immunostaining for Ki-67. This endogenous protein has an advantage over BrdU in that a timed injection of exogenous marker before the study is not required. The disadvantage is that, unlike BrdU, it cannot be used to provide a precise proliferation rate. BrdU is incorporated into the DNA of cells during the S phase (DNA synthesis) of the cell cycle. Ki-67 is a protein that is expressed from mid-G1 until mitosis (19). Thus, because some cells in G1 do not necessarily divide, Ki-67 cannot be exactly equated with mitosis, even though there is certainly some correlation. Because of these concerns, we use the term proliferation index rather than proliferation rate. The most obvious finding is that the proliferation index of ß-cells in the pancreases of newborn pigs and in NPCCs that have been cultured for 8 days is far higher than that found in 20-week grafts. This fall in the ß-cell proliferation index is gradual, with the values in the newborn pancreases and in 10-day grafts being about 8%, those in grafts at 6 weeks being about 4%. At 20 weeks the index fell to a much lower value of only about 0.4%, which is virtually the same as that found in ß-cells of adult porcine pancreases. It is of interest that the ß-cells of cultured NPCCs have a higher proliferation index than those in the pancreas, a finding presumably due to the culture media that contains nicotinamide, IBMX and a relatively high glucose concentration of 10 mM. Another interesting point is that the ß-cell proliferation index is higher in the 6-week grafts of mice with persistent hyperglycemia than in those with normoglycemia. This increased ß-cell proliferation index is likely due to a stimulatory influence of high glucose levels (24, 25, 26).
Although adult porcine ß-cells have a very low proliferation index in the grafts of normoglycemic mice, they have a surprising capacity for increased proliferative activity when placed in a hyperglycemic environment. Thus, in the presence of high plasma glucose levels at 10 days and 6 weeks , the values were about 2.5% compared with about 0.3% in the normoglycemic situation. Thus, the given mass of transplanted adult ß-cells has some capacity to expand. Moreover, it is possible that ß-cell hypertrophy can lead to further increase in ß-cell mass. Hypertrophy received little study with human or porcine ß-cells, but has been convincingly shown in a variety of situations in rodents (22, 27, 28, 29).
Study of the proliferative capacity of duct cells also provided novel information. It is striking that the proliferation index of duct cells in the neonatal pancreas was more than twice as high as that found in NPCCs after 8 days of culture or in 10 days grafts (13% vs. about 6%). One must be cautious about comparing the proliferation index of duct cells with ß-cells because the length of the cell cycle of these two cell types is not known. The mechanisms responsible for the proliferation capacity of duct cells are unknown, but this fall in the index may be due to some maturation phenomenon or to a change in environment, that occurs with in vitro culture or in the engraftment site. We also suspect that the CK7 stained duct cells may represent a heterogeneous population, with some cells having greater capacity for proliferation and differentiation than others. A final point is that the proliferation index of the duct cells, in contrast to ß-cells was no higher in a hyperglycemic than normoglycemic environment. It seems likely that glucose levels have little influence on the proliferation and differentiation of duct cells, because the glucose-sensing machinery consisting of GLUT2, glucokinase, and other elements is underdeveloped.
It was also found that ß-cell size in NPCCs increased over time, presumably as part of the maturation process. In particular, the cross-sectional area, which correlates with cell size, of ß-cells in the neonatal pancreas is considerably smaller than that in an adult pancreas. Moreover, ß-cell size in grafts of NPCCs at 20 weeks is larger than that found in the neonatal pancreas or in 10 days grafts. Earlier studies in rodents have also found ß-cells in younger animals to be smaller (30). The question of ß-cell size is becoming more interesting now that a smaller size has been identified in less mature cells and hypertrophy has become a well established phenomenon in various rodent studies (22, 27, 28, 31) It is possible that heterogeneity in function could be related to differences in cell size (32, 33).
This study provides insight into the mechanisms responsible for the expansion of ß-cell mass in the grafts of NPCCs transplanted under the kidney capsule of nude mice. One mechanism is that duct cells actively proliferate and at least some of them become ß-cells, as evidenced by the finding of protodifferentiated cells staining for both CK7 and insulin. The importance of duct cells as a source is further supported by the low frequency of apoptosis in duct cells. A second mechanism is active proliferation of ß-cells, which must be the source of many of the ß-cells found at 20 weeks . A third mechanism is the increase in ß-cell size that was documented; this must contribute to the increase in ß-cell mass but less than the other two pathways. At the present time it is not possible to know the relative importance of these pathways. In a study of transplanted human fetal islet-like clusters, Beattie et al. (34) concluded that ß-cells mainly came from duct cells, but they did not assess ß-cell proliferation, so it is not clear their results are different from ours. Korsgren et al. has also reported low ß-cell proliferation in transplanted fetal pig islet like clusters (15) suggesting relatively less contribution from differentiated ß-cells to an increase in ß-cell mass.
A definitive study of the origin of ß-cells in grafts would require precise measurement of the birth rate of duct cells, non-ß-endocrine cells and ß-cells with BrdU, measurement of the mass and death rate of each of these cell types. At present, there is no accurate way to measure the rate of cell death, although it can be calculated if one knows the birth rate, cell mass, and cell size at different time points (30). Our present methods for measuring cell death rely on various markers for apoptosis and necrosis, but because these events are short-lived and variable, it is not possible to use them to determine an actual rate of death. In summary, differentiation of duct cells, proliferation of ß-cells and increased ß-cell size work together to turn transplanted NPCCs into grafts consisting almost completely of ß-cells.
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Acknowledgments |
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Footnotes |
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2 Has been a recipient of a Mentor-based Fellowship of the American
Diabetes Association and is currently a recipient of Joslin
Institutional Training Grant (T-32-DK-07260) from the NIH.
Received October 3, 2000.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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