In vitro and in vivo evaluation of insulin-producing
TC6-F7
cells in microcapsules
D.
Zhou1,
A. M.
Sun1,
X.
Li1,2,
S. N.
Mamujee1,
I.
Vacek1,
J.
Georgiou1, and
M. B.
Wheeler1,2
Departments of 1 Physiology and
2 Medicine, Faculty of
Medicine, University of Toronto, Toronto, Ontario, Canada M5S 1A8
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ABSTRACT |
In the present study, the insulin secretory capacity of
TC6-F7 cells in microcapsules was evaluated. The cell mass within capsules was found to expand in a three-dimensional fashion, in contrast to cells seeded on plates that grew as a monolayer. In in
vitro studies, both free and encapsulated cells were found to secrete
insulin in the absence of glucose, at 13.6 ± 1.1 and 14.5 ± 0.9 ng · 106
cells
1 · 60 min
1, respectively, with
the response rising to a maximum of 26.0 ± 0.8 and 31 ± 2.3 ng · 106
cells
1 · 60 min
1 in the presence of
16.8 mM glucose. Encapsulated cells were able to produce
Ca2+ responses in the presence of
KCl (50 mM) and BAY K 8644 (100 µM). In in vivo studies,
intraperitoneal transplantation of 3.0 ×106 microencapsulated cells
into mice (n = 5) with
streptozotocin-induced diabetes resulted in the restoration of
normoglycemia up to 57 days. Insulin concentrations rose from 0.4 ± 0.1 ng/ml before the graft administration to 2.2 ± 0.8 ng/ml after
the transplantation in the normoglycemic recipients. An oral glucose
challenge in transplant recipients demonstrated a flat glucose
response, suggesting extremely high glucose clearance rates. These data
demonstrate the potential use of the immunoisolated
-cell lines for
the treatment of diabetes.
insulin-producing cells; microencapsulation; xenografts; transplantation
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INTRODUCTION |
THE RESULTS OF THE recent Diabetes Control and
Complications Trial (3) demonstrate that there is a major impact of
poor metabolic control on the rate of appearance of diabetic
complications and that hyperglycemia is the key factor in the
initiation of various pathological processes. Therefore, the tight
control and maintenance of euglycemia should be the fundamental measure
for preventing or delaying the development of diabetic complications. Consequently, it has become imperative to develop methods, applicable early in the course of the disease, for obtaining perfect metabolic control without increasing the risk of severe hypoglycemia. Currently, the transplantation of islet tissue, either as a whole pancreas or as
isolated islets, has been pursued because this technique can provide
normal blood glucose control and thus have the potential to prevent or
reduce diabetic complications (3, 19, 22).
To overcome the problem of immunorejection and the need for
immunosuppression and to circumvent the potential for disease recurrence, the concept of immunoisolation has been advanced. In our
approach to immunoisolation, we have developed
alginate-polylysine-alginate biocompatible capsules to enclose
individual pancreatic islets (5, 9, 12, 13, 16). In these earlier
studies, both allografts and xenografts of microencapsulated isolated
pancreatic islets were shown to reverse diabetes in long-term
experiments in spontaneously diabetic as well as chemically induced
diabetic rodents (5, 9, 12, 13, 16). These studies demonstrate the
potential for the use of allo- and xenotransplantation as therapeutic
alternatives to exogenous insulin therapy in insulin-dependent diabetics.
Although there is little question that transplanted islets offer a
potential alternative to exogenous insulin therapy, large-scale islet
isolation and cryopreservation, however, are both technically difficult
and expensive. In addition, consistent and reliable performance of
islet allografts and xenografts, from batch to batch, may be difficult
to achieve. Insulin-secreting cell lines derived from
-cells
represent a potential alternative approach to pancreatic islet
transplantation. These cells can be grown inexpensively and in
unlimited quantity. Several glucose-responsive
-cell lines have been
produced through the culturing of islet tumors; however, these cell
lines are, in general, phenotypically unstable as evidenced by a shift
in glucose responsivity and/or diminished insulin output
through passage (4, 7, 17, 20, 21). Recently, Knaack et al. (8)
reported the cloning of a phenotypically stable (>55 passages)
-cell line,
TC6-F7. This cell line has been shown to have
remarkably similar characteristics to
-cells of pancreatic islets,
including appropriate glucose-induced insulin responsivity (5-30
mmol range) and the expression of GLUT-2 but not GLUT-1 glucose
transporters, with high-glucokinase and low-hexokinase activities,
respectively. In the present study, we evaluated the
TC6-F7 cells both in vitro and in vivo and tested the hypothesis
that
TC6-F7 cells microencapsulated and transplanted into diabetic
mice would control hyperglycemia in long-term experiments. The results
of these studies are described.
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MATERIALS AND METHODS |
Cell culture.
TC6-F7 cells were grown and passaged as previously described (8)
with the following modifications: cells were grown in RPMI 1640 medium
(GIBCO) containing 5.5 mM glucose and supplemented with 10% FCS and 2 mM L-glutamine.
Cell encapsulation.
The encapsulation technique was a modification of the method of Lim and
Sun (11). This modification involved the use of an electrostatic
droplet generator (6), which produces smaller, stronger, and more
uniform capsules compared with the older air-jet technique. The cells
were suspended in 1.5% (wt/vol) purified sodium alginate (Kelco Gel
LV, Kelco, San Diego, CA) at a concentration of ~1.5 × 107 cells/ml. Spherical droplets
were formed by the electrostatic field interaction coupled with syringe
pump extrusion and were collected in a 100 mM calcium lactate solution.
The gelled droplets were suspended in 0.05% polylysine (Sigma,
molecular mass = 22-24 kDa) for 5 min. The droplets were washed
with 0.9% saline and suspended in 0.15% sodium alginate for 5 min.
After wash with 0.9% saline, the capsules were allowed to react with
55 mM sodium citrate for 5 min and finally washed with 0.9% saline and
with culture medium. On average, capsules were designed to contain ~300 cells and have an average diameter of 0.25-0.35 mm.
Cell viability assays.
To examine cell viability at different stages of culture (2 through 6 wk) cells were imaged by loading 1 µM calcein-AM (Molecular Probes,
Eugene, OR) with a final concentration of 0.1% pluronic acid. A
confocal laser scanning microscope (Bio-Rad 600) was used to analyze
viable cells using the 488-nm line of the argon laser, and emitted
fluorescence was detected through a low-pass filter with cutoff at 515 nm. To examine nonviable cells, encapsulated
TC6-F7 cells were
stained with 5 µM propidium iodide (Sigma) for 10 min. Propidium
iodide was excited using the 514-nm laser line, and the emitted
fluorescence was detected through a 550-nm long-pass filter.
In vitro insulin secretion studies.
For initial characterization of
TC6-F7 cells, insulin secretion
experiments were performed. Briefly, cells at passages
40-42 were plated in 24-well plates at a density
of 100,000 cells per well and cultured for an additional 48 h. Cells
were then preincubated in zero glucose RPMI 1640 (0.1% BSA, 0.05%
bacitracin) for 60 min, followed by a 60-min stimulation with various
concentrations of glucose (0-16.8 mM, 2 ml total volume). With
encapsulated cells, groups of 40 microcapsules, each initially
containing ~300 cells, were cultured in tissue culture multiwell
plates for 48 h following encapsulation. After a 60-min preincubation
period in RPMI 1640 (0 glucose), the encapsulated cells were exposed to
glucose (2 ml total volume) for 60 min. Samples of the medium were
collected following glucose exposure and stored at
2°C
before assay. In each case, the insulin content of the samples was
determined by radioimmunoassay (rat insulin kit, Linco Research, St.
Louis, MO); results (in ng/ml) were expressed as means ± SE of at
least three independent experiments.
Free cytosolic
Ca2+
measurements.
A confocal laser scanning microscope (Bio-Rad 600) was used to analyze
Ca2+ fluxes in encapsulated
TC6-F7 cells after ~2 wk under standard culture conditions, as
described. Cells were loaded with 10 µM fluo 3-AM (Molecular Probes)
for 1 h in DMEM, 0.5% DMSO, and 0.1% pluronic acid.
Subsequently, cells were washed twice in DMEM and changes in cytosolic
Ca2+ concentration
([Ca2+]i)
were analyzed in the presence of 50 mM KCl or BAY K 8644 (100 µM) for
250 ms. Fluo 3 was excited using the 488-nm line of the argon laser,
and emitted fluorescence was detected through a low-pass filter with
cutoff at 515 nm. The detected signals were digitized, forming an
eight-bit number (0-255) to represent each pixel. Images were thus
collected digitally, and a false color scale was generated for
quantitative purposes, where blue corresponds to lower and red to
higher Ca2+ levels. The changes in
fluorescence were measured using CFOCAL, a program for PC analysis and
the preparation of confocal images (written by T. A. Goldthorpe,
Department of Physiology, University of Toronto). Changes in
Ca2+ fluorescence (F) are
expressed as
F/F = (F
Frest)/Frest,
where Frest is the resting
fluorescence level.
Transplantation studies.
For the induction of diabetes, animals (C57/BL mice; Charles River, St.
Constant, PQ, Canada) were administered streptozotocin intravenously at
185 mg/kg; subsequently, these animals were considered diabetic and
therefore suitable for transplantation after registering three
consecutive, nonfasting blood glucose measurements above 20 mM. Before
the transplant, the microencapsulated cells were cultured overnight
under standard culture conditions. With the use of light anesthesia,
groups of mice with streptozotocin-induced diabetes received a single
transplant of either 3.0 × 106
(n = 5), 1.5 × 106
(n = 3), or 0.75 × 106
(n = 2) microencapsulated cells,
~300 cells per capsule, passages 40-42. A control group of diabetic mice
(n = 3) received corresponding numbers
of free, unencapsulated cells. Another diabetic control group
(n = 3) received equal numbers of
empty capsules. The grafts were administered by intraperitoneal
injection using an 18-gauge cannula. At regular intervals (2-3
days), blood samples were taken via tail vein from all animals for
blood glucose monitoring and analyzed with a glucometer (Miles,
Toronto, ON, Canada). To confirm that the normoglycemic condition of
these diabetic animals indeed resulted from the graft of the
microencapsulated cells, capsules were removed from the peritoneal
cavities of two normoglycemic, randomly selected recipients of
3.0 × 106
microencapsulated cells 35 days after transplantation. The procedure was performed under ether anesthesia. The microcapsules were retrieved by repeated washes with saline inside the peritoneal cavity.
Glucose tolerance test.
Oral glucose tolerance tests (OGTTs) were administered to transplant
recipients approximately 1 wk after normoglycemia had been established
as a result of the grafts. Nondiabetic animals were used as controls.
Before the OGTTs, all experimental animals were fasted overnight. In
the OGTT, 1.5 mg of glucose per gram of body weight were instilled
orally into each experimental mouse, and blood glucose concentrations
were established over a 120-min period.
Statistical analysis.
Results are given as means ± SE of at least three independent
observations unless otherwise stated. Data were analyzed with the
Student's t-test and
2 test for statistical
significance. P < 0.05 was
considered significant.
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RESULTS |
Imaging of encapsulated
TC6-F7 cells.
Encapsulated cells cultured for 2-6 wk were examined by confocal
microscopy to observe their growth activity in microcapsules (Fig.
1). Cells initially formed
several small aggregates within 1-2 wk (Fig.
1Ai) that would interfuse
primarily into one to a few large-core aggregates within 3-4 wk
(Fig. 1, Bi,
Ci, and
Ei). After 4 wk in culture, the
majority of capsules contained primarily one central aggregate, and, by
5-6 wk, the aggregate had expanded to fill the capsule (Fig. 1,
Di,
Fi, and
Gi). We used three approaches to
determine encapsulated
TC6-F7 viability within the capsule. First,
live cells were visualized by calcein fluorescence (Fig. 1,
Aii-Dii).
This method was employed because a laser confocal microscope can be
used to visualize the viability of cells within the central core of the
capsule, on the basis of the premise that enzymatic activity is
required to activate fluorescence of calcein-AM and cell membrane
integrity is required for its retention (14, 15) (Fig. 1,
Aii-Dii).
As shown, the majority of cells within the capsules were labeled
(viable cells emitting a green fluorescence); however, there was a
tendency for cells in the central core of the clusters to not label as
strongly. A trypan blue exclusion test was also performed on
encapsulated cells after 3 and 6 wk in culture. This test revealed that
at 3 wk of culture more than 90% of cells were viable and at 6 wk
~80% of cells appeared viable, in agreement with the calcein
results. To assess viability in an alternative way, expired or defunct
cells were visualized using propidium iodide. This red fluorescent DNA
binding agent is cell impermeant and thus will only gain access to
cells with compromised membranes (1). As shown in Fig. 1,
Eii-Gii,
a small proportion of cells have labeled nuclei, and there was no
correlation between labeling and cells within the central core of the
cell clusters. This pattern of fluorescence is in contrast to cells
cultured at room temperature in serum-free media for 2 days, in which most of the cells (Fig.
1Hii) have labeled nuclei. These
studies demonstrate that the majority of cells up to 6 wk
postencapsulation are viable, which is quite surprising given that the
cells do poorly in monolayer culture when confluence approaches values
>70% (unpublished results).

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Fig. 1.
Photomicrographs and fluorescent images of microencapsulated TC6-F7
cells. Cells were encapsulated and cultured for 2-6 wk to observe
their growth activity in microcapsules. Scale bar = 100 µm. Trans
images (i) refer to a light images
acquired using a transillumination adapter. Cells initially formed
several small aggregates within 2 wk
(Ai) that would form primarily into
a few large-core aggregates within 4 wk
(Bi,
Ci,
Ei). By 6 wk in culture, the
majority of capsules contained primarily 1 central core aggregate that
would ultimately fill the capsule (Di,
Fi-Hi).
Simultaneous to trans micrographs, images of calcein fluorescence from
the capsule series are shown
(Aii-Dii).
Live cells are detected by fluorescence using a confocal microscope.
Similarly, propidium iodide fluorescence images are shown
(far right) for a second series of
capsules
(Eii-Hii).
Cells with labeled nuclei are not viable.
Hi and
Hii are representative of a capsule
series cultured at room temperature in serum-free media for 2 days
before the experiment. Note the intense nuclear fluorescence
(Hii), indicating poor viability.
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In vitro glucose challenge experiments.
The results of the in vitro glucose challenge study comparing
microencapsulated and free cells are summarized in Fig.
2. In the absence of glucose, the
encapsulated cells and the free cells secreted 14.9 ± 0.9 and 13.6 ± 1.1 ng
insulin · 106
cells
1 · 60 min
1, respectively. The
exposure of the cells to 2.8 mM glucose resulted in insulin secretion
of 19.4 ± 2.6 and 17 ± 0.62 ng · 106
cells
1 · 60 min
1 for encapsulated and
free cells, respectively. On exposure to 5.6 mM glucose, the
encapsulated and free cells secreted insulin at a rate of 21.1 ± 2.5 and 22.7 ± 0.49 ng · 106
cells
1 · 60 min
1, respectively.
Finally, at the high-glucose concentration of 16.8 mM, the insulin
secretion for encapsulated and free cells amounted to 31.0 ± 2.3 and 26.0 ± 0.8 ng · 106
cells
1 · 60 min
1, respectively. These
results demonstrate that free and encapsulated cells respond similarly
to a glucose stimulus.

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Fig. 2.
In vitro glucose challenge study comparing free and encapsulated cells.
Free or encapsulated cells were preincubated in 0 glucose RPMI 1640 (0.1% BSA, 0.05% bacitracin) for 60 min, followed by a 60-min
stimulation with various concentrations of glucose (0-16.8 mM).
With free cells, * P < 0.05 for insulin secretion at 5.6 and 16.8 mM vs. insulin secretion at 0 mM
glucose. ** P < 0.05 for
encapsulated cell insulin secretion at 5.6 and 16.8 mM glucose vs.
secretion at 0 mM glucose (n = 5 independent observations per group).
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Measurement of
[Ca2+]i.
Cells in capsules were examined in vitro for their ability to
depolarize and regulate
[Ca2+]i
in response to test agents (Figs. 3 and
4 and Table 1). In cells
loaded with fluo 3-AM, the majority was observed to have similar
resting Ca2+ fluorescence
intensities; however, there was a trend toward cells in the interior of
larger aggregates to have significantly lower resting fluorescence
intensities (Fig. 3B, summarized in
Table 1). We attribute this lower resting intensity to the reduced efficiency with which fluo 3-AM penetrates the centers of the these
large aggregates and not to a reduced viability of the central portions
of the cores, based on the studies with calcein and propidium iodide.
In response to 50 mM KCl (Fig. 3C),
delivered by micropipette (Figs. 3 and
4A) to the interior of a capsule,
the majority of cells showed an increase in
Ca2+ fluorescence intensity, as
depicted using a false color scale. Fluorescence intensities were found
to return to prestimulatory levels within minutes of the KCl stimulus
(data not shown). The changes in fluorescence for four isolated cells
(2 from central and 2 from the periphery of large aggregates) were
normalized to respective control values (
F/F) and plotted against
time (Fig. 4A). The pattern of
fluorescence, typical of a depolarization event, was monophasic, with a
rapid transient peak of Ca2+. The
net increase above basal levels was similar for cells in the core or
periphery of large aggregates (Table 1). In Fig. 4B, KCl delivered to the medium
bathing capsules gave a similar response from representative
responders, but the response was slightly delayed. The L-type
Ca2+ channel agonist BAY K 8644 (100 µM) delivered to the bathing medium also elicited
Ca2+ responses (Fig.
4C). These studies suggest that most
of the cells express functional voltage-dependent
Ca2+ channels, as well as the
appropriate intracellular mechanisms for removing
Ca2+ from the cytoplasm.

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Fig. 3.
Ca2+ responses of encapsulated
TC6-F7 cells. A: nonconfocal image
acquired using the transmitted light attachment of the Bio-Rad 600 microscope, revealing a capsule containing TC6-F7 cells (~2 wk
postencapsulation). Note the microelectrode that has been inserted into
the capsule. Scale bar = 50 µm. B:
confocal image of the same encapsulated cells loaded with fluo 3-AM,
showing resting Ca2+ levels.
Relative Ca2+ fluorescence appears
in false color (see bar at right).
C: application of KCl (50 mM) induced
a Ca2+ response in most of the
cells. Data are representative of 5 capsule experiments.
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Fig. 4.
Time course of Ca2+ responses of
capsules containing TC6-F7 cells (measure in F/F, where F is the
detected fluorescence). A: time course
of Ca2+ signals evoked by KCl
application (50 mM) to the interior of the capsules. Each symbol in
A-C
represents a region of analysis with 2 cells from the central cores
(triangles) and 2 from the periphery of large aggregates (squares and
circles). Changes in Ca2+ from 4 regions were analyzed and plotted with respect to time, with KCl being
applied at time = 0 min. B: time
course experiment with KCl being applied to the bathing solution.
Independent areas within 4 capsules were analyzed.
C:
Ca2+ responses from 4 capsules on
addition of the L-type Ca2+
channel agonist BAY K 8644 (100 µM). Agonist was released at time = 0 min.
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Table 1.
Comparison of resting and peak [Ca2+]i
responses to KCl from TC6-F7 cells located on the
periphery or at the core of large cell aggregates
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In vivo studies with transplanted capsules.
The intraperitoneal transplantation of microencapsulated cells with the
highest dose of 3 × 106
cells resulted in the restoration of normoglycemia in all experimental animals (n = 5) within 5-7
posttransplantation days (Table
2). The blood glucose profiles
of these animals are shown in Fig. 5. One
out of three mice that received 1.5 × 106 microencapsulated cells became
normoglycemic (with blood glucose concentrations of <6.0 mM), whereas
only a partial lowering of blood glucose levels (6-11 mM) was
observed in the other two recipients. Finally, only a partial lowering
of the blood glucose concentrations was observed in the two recipients
with the lowest dose, 0.75 × 106 microencapsulated cells. In
the three recipients with 3.0 × 106 free unencapsulated
TC6-F7
cells (i.e., an equivalent of the highest dose of encapsulated cells),
normoglycemia was restored within 6 posttransplantation days; however,
during the next 7 days, the blood glucose concentrations returned to
the original hyperglycemic range, thus indicating the destruction of
the graft by the immune system of the recipient. No change in the
diabetic hyperglycemia was observed in the recipients of empty
capsules. The duration of normoglycemia in the five recipients of the
highest dose of the microencapsulated cells was at least 35 days (on
day 35, 2 animals of this group had
the capsules surgically removed), with the three remaining
normoglycemic from 55 to 57 days, at which time the grafts failed (Fig.
5). The removal of the microcapsules at 35 days posttransplantation
resulted in a return to hyperglycemia within 24 h. The recovered
capsules were free of cell overgrowth and physically intact, with
enclosed clusters of cells clearly visible. The body weights of the
transplant recipients in which normoglycemia had been established
increased during the period of normoglycemia, whereas hyperglycemic
animals lost weight. Although the serum insulin concentrations in the
diabetic mice before the transplants averaged 0.4 ± 0.1 ng/ml
following the graft administration, the insulin concentration increased
to 2.2 ± 0.8 ng/ml (n = 5, P < 0.05 compared with the
pretransplant concentration) in the recipients in which normoglycemia
was achieved. The average serum insulin concentration in healthy
control animals was determined at 2.0 ± 0.3 ng/ml. The OGTTs
indicated extremely high glucose clearance rates in the animals in
which normoglycemia had been restored as a result of the graft
administration (Fig. 6). These clearance
rates were reminiscent of glucose clearance rates of human patients
suffering from insulinoma.
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Table 2.
Restoration of normoglycemia within 5-7 days in
streptozotocin-induced diabetic mice transplanted with
microencapsulated TC6-F7 cells
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Fig. 5.
Blood glucose profiles of mice transplanted with microencapsulated
TC6-F7 cells. Streptozotocin-induced diabetic mice (initially,
n = 5) received a single transplant of
3.0 × 106 microencapsulated
cells, ~300 cells per capsule, administered by intraperitoneal
injection. Capsules were removed from 2 animals on day
35. With the remaining animals, implants failed to
normalize glucose on days 50,
55, and
57. Data are presented as means ± SE.
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Fig. 6.
Oral blood glucose tolerance test (OGTTs) of graft recipients. OGTTs
were administered to transplant recipients in which normoglycemia had
been established as a result of the grafts for a period of at least 1 wk. Nondiabetic animals and streptozotocin-induced diabetic mice
without capsules were used as control groups. Data are presented as
means ± SE of n = 5 animals per
experimental group.
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DISCUSSION |
Clinical transplantation of human islets from cadavers proved that this
can normalize hyperglycemia in diabetic patients (23). However, the
limited supply of human islets, especially when the procurement of
cadaver pancreata is not controlled, makes their use impractical.
Consequently, if the mass clinical transplantation of pancreatic islets
into human diabetics is to become a reality, xenografts of animal
islets will have to be used. We have shown in our past experiments that
xenotransplants do work in animal models (5, 9, 12, 13, 16). Thus
clinical transplantation of animal pancreatic endocrine tissue may be
used in the near future as a novel treatment for diabetes. Although
this approach could revolutionize the management of diabetes mellitus,
the use of animal tissue would also present a number of challenges. The possible existence of animal pathogens transmissible to the human graft
recipients must be carefully considered. The technique of islet
isolation from the pig, which now seems to represent the most likely
candidate for donor tissue, involves a number of technical difficulties. The process of isolation and purification of porcine islets is expensive and labor intensive. Although the use of animal islets may yet prove to be the right concept for the treatment of
diabetes, it also seems appropriate to search for possible alternatives. Because our concept of immunoisolation by
microencapsulation of cells and tissues has a vast scope of possible
applications, the use of bioengineered insulin-producing cells becomes
very attractive.
As a model, we have selected an insulin-producing clonal line
TC6-F7, originally derived from insulinomas arising in transgenic mice expressing the SV40 T antigen gene under control of the insulin promoter. Because
TC6-F7 cells are derived from the B6D2F1/J mouse
strain and C57/BL mice were used as transplant recipients, allotransplantation was performed in this study. After
microencapsulation, the
TC6-F7 cell mass expanded within the
capsules, displaying a pattern of distribution that was substantially
different from that of the free, unencapsulated cells (Fig. 1). In
monolayer culture, cells would grow to ~70% of confluence at which
point the cells would quite rapidly start necrotizing. The encapsulated cells grew in a three-dimensional fashion, initially in a few aggregates that would later (at 4-6 wk) expand and fill the entire capsule, visualized under light microscopy as a solid mass. Confocal microscopy in combination with the appropriate fluorescent dyes was
used to test for viability and to demonstrate that the cells, including
those at the most central point within the capsule, were viable. These
qualitative data are supported by trypan blue exclusion experiments
that show that the majority of the cells (>80%) were still living at
6 wk of culture.
In response to KCl and several secretagogues that stimulate
-cells,
the cell depolarizes, followed by an influx of
Ca2+ through voltage-dependent
Ca2+ channels (reviewed in Ref.
18). The increase in free
[Ca2+]i
is thought to be a trigger for insulin secretion (18). Because only
living cells can regulate Ca2+, we
used Ca2+ imaging to further
examine the viability of encapsulated cells. KCl and the L-type
Ca2+ channel agonist BAY K 8664 were able to stimulate
[Ca2+]i,
whether applied into a single capsule or to the medium adjacent to a
capsule (Figs. 3 and 4). Only a slight delay in the response was noted
on external application. Of interest was the observation that the
central cores of larger aggregates appeared to be equally responsive to
stimulation but less efficient at loading the
Ca2+ indicator, since the resting
Ca2+ fluorescence was lower in
central compared with peripheral cells. These data suggest that the
cells in the central cores may be less accessible and therefore less
responsive to larger molecules capable of crossing the capsule
membrane. It further suggests that this potential phenomenon would
become even more apparent as the cell mass expands and the aggregates
enlarge.
In an earlier publication by Knaack et al. (8),
TC6-F7 cells were
reported to have manifested a stable phenotype of insulin secretion in
response to physiological glucose concentrations. Our results are in
agreement with this previous study in that the cells were responsive to
glucose; however, our studies demonstrate that there is a progressive
increase in the insulin response to glucose through 16.8 mM (Fig. 2).
There are several differences between the two studies, including the
culture conditions, type of medium, glucose concentration, and the type
of serum conditions used. In addition, we did not use the
phosphodiesterase inhibitor IBMX in timed release assays. These
modifications may explain the differences. In the present study,
although the
TC6-F7 cells displayed a steady insulin secretion,
their response to glucose stimulation appeared rather limited compared
with pancreatic islets that we had previously characterized (10). In
our past studies on insulin secretion from microencapsulated rat and
porcine pancreatic islets, we showed that the kinetics of insulin
release from the free and microencapsulated islets were very
similar, thus indicating that the
alginate-polylysine-alginate membrane does not constitute an impediment to the free flow of insulin. These findings were confirmed in this study by demonstrating similar patterns of insulin secretion from the free and microencapsulated
TC6-F7 cells.
In vivo, the implantation of 3 × 106
TC6-F7 cells resulted in a
long-term restoration of normoglycemia in diabetic mice. In comparison,
in our past studies using islets of Langerhans, transplants of ~800
rat islets, comprising roughly 3 × 106 insulin-producing
-cells,
were sufficient to reverse diabetes in mice. Unencapsulated islets
implanted under the kidney capsule or into the portal vein may require
fewer islets compared with an intraperitoneal transplant of
immunoisolated islets (2). In our experience, at least 500 islets per
mouse, comprising perhaps 2 × 106
-cells, are necessary to
sustain euglycemia. Thus, in vivo, the insulin secretion of the grafted
TC6-F7 cells appears comparable to the transplants of natural islet
-cells. However, as the results of the glucose tolerance tests
indicate (Fig. 6), the very limited responsiveness of the
TC6-F7
cells to glucose causes the glucose clearance to be rather flat, very
similar to that displayed by many insulinoma patients. This observation
bears out the fact that the insulin release from the
TC6-F7 cells
most likely goes on continuously, with the cells not responding to the
dynamics of serum glucose concentrations. Thus, although not measured, we suspect that circulating insulin levels would be elevated in transplant recipients during the fasting state and before an OGTT.
One can only speculate about the factors responsible for the return of
diabetic hyperglycemia in the recipient animals and the failure of the
graft after ~55 days of normoglycemia, since in these initial studies
the viability of cells after failure was not examined. Failure may have
been caused by a decrease in insulin secretion or by an eventual death
of the grafted cells. Also, the capsule construction is of a critical
importance in this respect, as the capsules' strength determines the
duration of the graft function. Similarly, imperfectly constructed
capsules may limit the duration of the graft function because of
surficial cell overgrowth. After the graft failure, the absolute
majority of recovered capsules was found intact, with some displaying
varying degrees of cell overgrowth. In these initial experiments, the reported periods of normoglycemia were shorter than those we had earlier reported for transplants of pancreatic islets, possibly indicating a limited life span of the implanted cells. Future studies
could include those directed at prolonging graft survival and at
enhancing the glucose responsivity of the cells.
 |
ACKNOWLEDGEMENTS |
We thank Dr. M. P. Charlton for the use of the confocal facility in the
Department of Physiology at the University of Toronto.
 |
FOOTNOTES |
This work was funded by grants from the Medical Research Council of
Canada (MT-12843 and UI-13097 to A. M. Sun and MT-12898 to M. B. Wheeler), the Canadian Diabetes Association (A. M. Sun and M. B. Wheeler), and the Juvenile Diabetes Foundation International (A. M. Sun). J. Georgiou was supported by a Neuroscience Network Studentship.
Address for reprint requests: M. B. Wheeler and A. M. Sun, Dept. of
Physiology, Faculty of Medicine Univ. of Toronto, Medical Sciences
Bldg., 1 King's College Circle, Toronto, ON, Canada M5S 1A8.
Received 5 December 1997; accepted in final form 29 January 1998.
 |
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