U.S. patent application number 09/952522 was filed with the patent office on 2003-05-01 for adipose-derived stem cells and lattices.
Invention is credited to Benhaim, Prosper, Futrell, J. William, Hedrick, Marc H., Katz, Adam J., Llull, Ramon, Lorenz, Hermann Peter, Zhu, Min.
Application Number | 20030082152 09/952522 |
Document ID | / |
Family ID | 25492986 |
Filed Date | 2003-05-01 |
United States Patent
Application |
20030082152 |
Kind Code |
A1 |
Hedrick, Marc H. ; et
al. |
May 1, 2003 |
Adipose-derived stem cells and lattices
Abstract
The present invention provides adipose-derived stem cells
(ADSCs), adipose-derived stem cell-enriched fractions (ADSC-EF) and
adipose-derivedlattices, alone and combined with the ADSCs of the
invention. In one aspect, the present invention provides an ADSC
substantially free of adipocytes and red blood cells and clonal
populations of connective tissue stem cells. The ADSCs can be
employed, alone or within biologically-compatible compositions, to
generate differentiated tissues and structures, both in vivo and in
vitro. Additionally, the ADSCs can be expanded and cultured to
produce molecules such as hormones, and to provide conditioned
culture media for supporting the growth and expansion of other cell
populations. In another aspect, the present invention provides a
adipose-derived lattice substantially devoid of cells, which
includes extracellular matrix material from adipose tissue. The
lattice can be used as a substrate to facilitate the growth and
differentiation of cells, whether in vivo or in vitro, into anlagen
or even mature tissues or structures.
Inventors: |
Hedrick, Marc H.; (Encino,
CA) ; Katz, Adam J.; (Charlottesville, VA) ;
Llull, Ramon; (Mallorca, ES) ; Futrell, J.
William; (Pittsburgh, PA) ; Benhaim, Prosper;
(Encino, CA) ; Lorenz, Hermann Peter; (Belmont,
CA) ; Zhu, Min; (Los Angeles, CA) |
Correspondence
Address: |
MANDEL & ADRIANO
55 SOUTH LAKE AVENUE
SUITE 710
PASADENA
CA
91101
US
|
Family ID: |
25492986 |
Appl. No.: |
09/952522 |
Filed: |
September 10, 2001 |
Related U.S. Patent Documents
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Application
Number |
Filing Date |
Patent Number |
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09952522 |
Sep 10, 2001 |
|
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PCT/US00/06232 |
Mar 10, 2000 |
|
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60123711 |
Mar 10, 1999 |
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60162462 |
Oct 29, 1999 |
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Current U.S.
Class: |
424/93.21 ;
435/366 |
Current CPC
Class: |
C12N 2500/25 20130101;
C12N 2501/01 20130101; C12N 2500/42 20130101; C12N 2500/38
20130101; A61K 35/12 20130101; C12N 5/0068 20130101; A61K 48/00
20130101; C12N 5/0667 20130101; C12N 2510/00 20130101; C12N 2501/39
20130101; C12N 2501/33 20130101; C12N 2533/90 20130101; C12N
2501/15 20130101 |
Class at
Publication: |
424/93.21 ;
435/366 |
International
Class: |
A61K 048/00; C12N
005/08 |
Claims
What is claimed is:
1. An isolated adipose-derived stem cell (ADSC).
2. The stem cell of claim 1, which can be cultured for at least 15
passages without differentiating.
3. The stem cell of claim 1, that is multipotent.
4. The stem cell of claim 3, that differentiates into mesoderm,
ectoderm or endoderm.
5. An adipose-derived stem-cell enriched fraction (ADSC-EF) of an
adipose tissue sample from a subject, said fraction substantially
free of adipocytes.
6. The stem cell of claim 1 which is human.
7. The stem cell of claim 1, which is genetically modified.
8. A defined cell population comprising a plurality of the cell of
claim 1.
9. The defined cell population of claim 8 which is homogenous.
10. The defined cell population of claim 8 which is
heterogeneous.
11. The defined cell population of claim 8 which is clonal.
12. A progeny cell of the stem cell of claim 4, committed to
develop into a mesodermal cell.
13. A progeny cell of the stem cell of claim 4, committed to
develop into an ectodermal cell.
14. A progeny cell of the stem cell of claim 4, committed to
develop into an endodermal cell.
15. Tissue comprised of the stem cell of claim 4, and
differentiated mesodermal cells.
16. Tissue comprised of the stem cell of claim 4, and
differentiated ectodermal cells.
17. Tissue comprised of the stem cell of claim 4, and
differentiated into endodermal cells.
18. A method of inducing mesodermal tissue comprising culturing the
stem cell of claim 4 in a mesoderm-inducing medium.
19. A method of inducing ectodermal tissue comprising culturing the
stem cell of claim 4 in a ectoderm-inducing medium.
20. A method of inducing endodermal tissue comprising culturing the
stem cell of claim 4 in a endoderm-inducing medium.
21. A method of forming tissue in a subject comprising introducing
the progeny cell of claim 12, 13 or 14 into a subject in a
sufficient amount to form mesodermal, ectodermal or endodermal
tissue in said subject.
22. A method of regenerating or repairing tissue in a subject
comprising introducing a stem cell of claim 1, 12, 13 or 14 into a
subject in a sufficient amount to regenerate or repair tissue.
23. A method for obtaining an adipose-derived stem cell-enriched
fraction (ADSC-EF) comprising treating a sample of adipose tissue
from a subject to remove adipocytes forming an adipose-derived
stem-cell-enriched fraction (ADSC-EF).
24. The adipose-derived stem-cell enriched-fraction (ADSC-EF)
obtained by the method of claim 23.
25. The adipose-derived stem cells (ADSCs) obtained by separating
said cells from the ADSC-EF of claim 24.
26. The stem cells of claim 25, wherein said stem cells are
multipotent.
27. The stem cells of claim 26, wherein said stem cell
differentiate into mesoderm, ectoderm, or endoderm.
28. An adipose-derived lattice comprising adipose tissue
extracelluar matrix substantially devoid of cells.
29. The lattice of claim 28 which is substantially anhydrous.
30. The lattice of claim 28 which is hydrated.
31. A composition comprising the cell of claim 1 and a biologically
compatible lattice.
32. A composition comprising the cell of claim 1 and the lattice of
claim 29 or 30.
33. Progeny of the stem cell of claim 3.
34. A method of delivering a transgene to an animal comprising
introducing the stem cell of claim 1 containing a selected
transgene into a subject, such that the transgene is expressed in
the subject.
35. A method of inducing the differentiation of the cell of claim
1, comprising culturing the cell in a suitable medium effective to
induce differentiation under suitable differentiation
conditions.
36. The method of claim 35 wherein said medium is a conditioned
medium of a specific cell type.
37. A method of inducing the differentiation of the cell of claim
1, comprising co-culturing the cell with a cell of desired
lineage.
38. A method of conditioning culture medium comprising contacting
the medium with the cell of claim 1.
39. The cultured medium obtained by the method of claim 38.
40. A kit for obtaining adipose-derived stem cells (ADSCs) from
adipose tissues of a subject comprising means for separating the
ADSCs from the adipose tissue.
41. The kit of claim 40, further comprising a device for isolating
adipose tissue from a subject.
42. The kit of claim 40, further comprising a medium for inducing
differentiation of the adipose-derived stem cells.
43. The kit of claim 40, further comprising a medium for culturing
the ADSCs.
Description
[0001] This patent application is a continuation-in-part (CIP) of
U.S. Ser. No. not yet known, filed Sep. 10, 2001, which corresponds
to PCT application No. PCT/US00/06232, filed Mar. 10, 2000, which
claims the benfit of the filing dates of U.S. Ser. No. 60/123,711,
filed Mar. 10, 1999, and U.S. Ser. No. 60/162,462, filed Oct. 29,
1999. The contents of all of the foregoing application are
incorporated by refernce in their entireties into the present
patent application.
[0002] Throughout this application, various publications are
referenced. The disclosures of these publications are hereby
incorporated by reference herein in their entireties.
BACKGROUND OF THE INVENTION
[0003] In recent years, the identification of mesenchymal stem
cells, chiefly obtained from bone marrow, has led to advances in
tissue regrowth and differentiation. Such cells are pluripotent
cells found in bone marrow and periosteum, and they are capable of
differentiating into various mesenchymal or connective tissues. For
example, such bone-marrow derived stem cells can be induced to
develop into myocytes upon exposure to agents such as 5-azacytidine
(Wakitani et al., Muscle Nerve, 18(12), 1417-26 (1995)). It has
been suggested that such cells are useful for repair of tissues
such as cartilage, fat, and bone (see, e.g., U.S. Pat. Nos.
5,908,784, 5,906,934, 5,827,740, 5,827,735), and that they also
have applications through genetic modification (see, e.g., U.S.
Pat. No. 5,591,625). While the identification of such cells has led
to advances in tissue regrowth and differentiation, the use of such
cells is hampered by several technical hurdles. One drawback to the
use of such cells is that they are very rare (representing as few
as 1/2,000,000 cells), making any process for obtaining and
isolating them difficult and costly. Of course, bone marrow harvest
is universally painful to the donor. Moreover, such cells are
difficult to culture without inducing differentiation, unless
specifically screened sera lots are used, adding further cost and
labor to the use of such stem cells. U.S. Pat. No. 6,200,606 by
Peterson et al., describes the isolation of CD34+ bone or cartilage
precursor cells (of mesodermal origin) from tissues, including
adipose.
[0004] There remains a need for a more readily available source for
large numbers of stem cells, particularly cells that can
differentiate into multiple lineages of different germ layers, and
that can be cultured without the requirement for costly
prescreening of culture materials.
[0005] Other advances in tissue engineering have shown that cells
can be grown in specially-defined cultures to produce
three-dimensional structures. Spacial definition typically is
achieved by using various acellular lattices or matrices to support
and guide cell growth and differentiation. While this technique is
still in its infancy, experiments in animal models have
demonstrated that it is possible to employ various acellular
lattice materials to regenerate whole tissues (see, e.g., Probst et
al. BJU Int., 85(3), 362-7 (2000)). A suitable lattice material is
secreted extracellular matrix material isolated from tumor cell
lines (e.g., Engelbreth-Holm-Swarm tumor secreted
matrix--"matrigel"). This material contains type IV collagen and
growth factors, and provides an excellent substrate for cell growth
(see, e.g., Vukicevic et al., Exp. Cell Res, 202(1), 1-8 (1992)).
However, as this material also facilitates the malignant
transformation of some cells (see, e.g., Fridman, et al., Int. J.
Cancer, 51(5), 740-44 (1992)), it is not suitable for clinical
application. While other artificial lattices have been developed,
these can prove toxic either to cells or to patients when used in
vivo. Accordingly, there remains a need for a lattice material
suitable for use as a substrate in culturing and growing
populations of cells.
SUMMARY OF THE INVENTION
[0006] The present invention provides adipose-derived stem cells,
adipose-derived stem cell fractions, lattices, and method for
obtaining the cells, fractions, and lattices. In one aspect, the
present invention provides an adipose-derived stem cell fraction
substantially free of adipocytes and red blood cells and
populations of connective tissue cells. The present invention also
provides stem cells, isolated from the fraction, where the stem
cells are pluripotent. The pluripotent stem cells have the ability
to differentiate into mesoderm, ectoderm, or endoderm. The cells
can be employed, alone or within biologically-compatible
compositions, to generate differentiated tissues and structures,
both in vivo and in vitro. Additionally, the cells can be expanded
and cultured to produce growth factors and to provide conditioned
culture media for supporting the growth and expansion of other cell
populations. In another aspect, the present invention provides a
adipose-derived lattice substantially devoid of cells, which
includes extracellular matrix material from adipose tissue. The
lattice can be used as a substrate to facilitate the growth and
differentiation of cells, whether in vivo or in-vitro, into anlagen
or even mature tissues or structures.
[0007] Adipose tissue is plentiful and represent a ready source of
the stem cells, fractions, and lattices. Moreover, the stem cells
can be passaged in culture in an undifferentiated state under
culture conditions not requiring prescreened lots of serum; the
inventive cells can be maintained with considerably less expense
than other types of stem cells. These and other advantages of the
present invention, as well as additional inventive features, will
be apparent from the accompanying drawings and in the following
detailed description.
BRIEF DESCRIPTION OF THE FIGURES
[0008] FIG. 1. Morphology; growth kinetics and senescence of
adipose-derived stem cells over long-term culture. Panel A: The
morphology of adipose-derived stem cells (e.g., a processed
lipoaspirate or PLA) obtained from liposuctioned adipose tissue.
Panel B: adipose-derived stem cells (PLAs) obtained from 3 donors,
were cultured for an extended period and cumulative population
doubling was measured and expressed as a function of passage
number. Panel C: Senescence in adipose-derived stem cells (PLA)
cultures as detected by staining for .beta.-galactosidase
expression at pH 6.0. Representative senescent cells are shown
(arrows).
[0009] FIG. 2. Composition of the adipose-derived stem cells (PLA)
as determined by indirect immunofluorescence (IF). Adipose-derived
stem cells (PLA) and bone marrow stromal cells (BMS), were stained
with the following antibodies: 1) anti-Factor VIII (FVIII); 2)
anti-smooth muscle actin (SMA); and 3) ASO2 (ASO2). Factor VIII and
smooth muscle actin expressing cells are shown (arrows).
[0010] FIG. 3. Composition of the adipose-derived stem cells (PLA)
as determined by flow cytometry. Panel A: Flow cytometry of
adipose-derived stem cells (PLA) samples using forward and side
scatter (FS and SS, respectively). A representative adipose-derived
stem cells sample is shown. Panel B: The cell composition of a
representative adipose-derived stem cells (PLA) sample from one
donor was determined staining with the following monoclonal
antibodies: anti-Factor VIII (FVIII), anti-smooth muscle actin
(SMA), ASO2 and a monoclonal antibody to vimentin (VIM), an
additional marker for cells of mesenchymal origin. Panel C: Flow
cytometry data from 5 donors was collected and the mean number of
positive events for each cell-specific marker is expressed as a
percentage of total adipose-derived stem cells (PLA) cell
number.
[0011] FIG. 4. Adipose-derived stem cells (PLA) accumulate
lipid-filled droplets upon treatment with Adipogenic Medium (AM).
Adipose-derived stem cells (PLA), bone marrow-derived MSCs (MSC),
and 3T3-L1 pre-adipocyte cells (3T3-L1) were cultured for two weeks
in AM and stained with Oil Red O to identify lipid-filled
intracellular vacuoles. Undifferentiated PLA cells maintained in
Control Medium (-ve Control) were stained as a negative
control.
[0012] FIG. 5. Adipose-derived stem cells (PLA) induced with
Osteogenic Medium (OM) express Alkaline Phosphatase and are
associated with a calcified extracellular matrix (ECM).
Adipose-derived stem cells (PLA), bone marrow-derived MSCs (MSC)
and a human osteoblast cell line (NHOst) were cultured in OM to
induce osteogenesis. Cells were stained at 2 weeks for Alkaline
Phosphatase activity (AP; red). The presence of a calcified
extracellular matrix (black regions) was examined at 4 weeks (von
Kossa). Undifferentiated adipose-derived stem cells maintained in
Control Medium were examined for AP expression and matrix
calcification as a negative control (-ve Control).
[0013] FIG. 6. Adipose-derived stem cells (PLA) treated with
Chondrogenic Medium (CM) are associated with a proteoglycan-rich
matrix and express collagen type II. Adipose-derived stem cells
(PLA) and MSCs (MSC) were cultured for 2 weeks in CM using the
micromass technique to induce chondrogenesis. The cells were fixed
and processed for the presence of sulfated proteoglycans with
Alcian Blue under acidic conditions (Alcian Blue). Paraffin
sections of human cartilage were used as a positive control
(Cartilage) while undifferentiated PLAs maintained in Control
Medium were processed as a negative control (-ve Control). In
addition, the expression of cartilage-specific collagen type II
(Collagen II) was examined in PLA cells and human cartilage
sections. Adipose-derived stem cells cultured in Control Medium
(-ve Control) were stained with Alcian Blue and for collagen II
expression as a negative control.
[0014] FIG. 7. Adipose-derived stem cells (PLA) cultured in
Myogenic Medium (MM) express the myosin heavy chain and MyoD1.
Adipose-derived stem cells (PLA) were treated with MM and stained
with antibodies specific to skeletal muscle myosin heavy chain
(Myosin) or MyoD1 (MyoD1). A human skeletal muscle cell line (SKM)
was examined as a positive control. In addition, the presence of
multinucleated cells in adipose-derived stem cells cultures is
shown (PLA, inset box). Myosin and MyoD1 expression was also
assessed in undifferentiated adipose-derived stem cells (-ve
Control) as a negative control.
[0015] FIG. 8. Growth kinetics of adipose-derived stem cells (PLA).
Panel A: adipose-derived stem cells, isolated from each donor, were
seeded in triplicate at a density of 1.times.10.sup.4 cells per
well. Cell number was calculated after 24 hours (day 1) and every
48 hours subsequent to day 1 (days 3 through 11). Mean cell number
for each donor was expressed with respect to culture time. The
growth curves from 4 representative donors are shown (20
years--open squares, 39 years--open circles, 50 years--open
triangles and 58 years--crosses). Results are expressed as
mean.+-.SEM. Panel B: Population doubling was calculated in all
donors from the log phase of each growth curve (i.e. from day 3 to
day 9) and expressed according to age. The line of regression was
calculated (n=20; r=0.62)
[0016] FIG. 9. Histological confirmation of adipogenic and
osteogenic differentiation by adipose-derived stem cells (PLA). A:
To confirm adipogenesis, cells were stained at 2 weeks
post-induction with Oil Red O. Low and extensive adipogenic levels
are shown (Panel 1--low; Panel 2--high). Adipose-derived stem cells
cultured in non-inductive control medium were analyzed as negative
controls (Panel 3). B: To quantify adipogenic differentiation, the
number of Oil Red O-positive stained cells were counted within
three defined regions. Two samples were analyzed from each donor.
The mean number of Oil Red O-positive cells was determined and
expressed as a percentage of total adipose-derived stem cells
number as an indication of adipogenic differentiation.
Differentiation was expressed with respect to age and the line of
regression calculated (n=20; r=0.016).
[0017] FIG. 10. Osteogenic differentiation decreases with
increasing donor age. Panel A: To confirm osteogenesis,
adipose-derived stem cells (PLA) were stained at 2 weeks
post-induction for alkaline phosphotase (AP) activity (Panels 1 to
3) and at 4 weeks post-induction for matrix calcification using von
Kossa staining (Panels 4 to 6). Osteogenic differentiation levels
are shown (Panels 1/2--low; Panels 4/5--high). Adipose-derived stem
cells cultured in non-inductive control medium were analyzed as
negative controls (Panels 3 and 6). Panel B: To quantify osteogenic
differentiation, the number of AP-positive stained cells were
counted within three defined regions. Two samples were analyzed
from each donor. The mean number of AP-positive cells was
determined and expressed as a percentage of total adipose-derived
stem cells number as an indication of the osteogenic
differentiation. Differentiation was expressed with respect to age
and the line of regression calculated (n=18; r=-0.70). Panel C:
Based on the results of Panel B, the donor pool was divided into
two age groups [(20 to 36 years (n=7) and 37 to 58 years (n=11)].
The average level of osteogenic differentiation was calculated for
each group and expressed as a percentage of total adipose-derived
stem cells number. Statistical significance was determined using an
unpaired student t test assuming unequal variances (p<0.00 1).
Differentiation is expressed as mean.+-.SEM.
[0018] FIG. 11. Osteoprogenitor cell number within an
adipose-derived stem cell fraction (PLA fraction) does not
significantly change with age. Osteoprogenitor cell number within
the fraction was determined by identifying cells with osteogenic
potential. Two groups of donors were examined [Group A=20 to 39
years (n=5), Group B=40-58 years (n=6)]. Osteogenesis was confirmed
by staining for AP activity. Colonies containing more than 10
AP-positive cells (CFU/AP.sup.+) were counted and averaged as an
indicator of the number of osteogenic precursors within each age
group. Statistical significance was determined using an unpaired
student t test assuming unequal variances (p=0.11). Values are
expressed as mean CFU/AP.sup.+.+-.SEM.
[0019] FIG. 12. Human adipose-derived stem cells (PLA) placed in
micromass cultures and induced with chondrogenic media undergo
cellular condensation and nodule formation. Adipose-derived stem
cells induced under micromass conditions were stained with Alcian
blue staining at pH 1 to detect the presence of sulfated
proteoglycans. Panel A: cellular condensation; (Panel B) ridge
formation; (Panel C) formation of three-dimensional spheroids are
shown (magnification 100.times.); (Panel D) negative control
(control medium).
[0020] FIG. 13. Hematoxylin & Eosin, Goldner's trichrome, and
Alcian blue staining of nodule paraffin sections from
adipose-derived stem cells (PLA). Micromass cultures
adipose-derived stem cells were treated with chondrogenic medium to
form nodules, the nodules were embedded in paraffin and sectioned.
Nodule sections were stained using conventional hematoxylin and
eosin (Panels A and B) and a Goldner's trichrome stain to detect
collagens (green) (Panels C and D). Adipose-derived stem cells
induced for 2 days are shown at a magnification of 200.times.
(Panels A and C) and 14 days are shown at 100.times. (Panels B and
D). In addition, sections were stained with Alcian blue staining at
pH 1, to detect highly sulfated proteoglycans. Day two nodules
(Panel E) are shown at a magnification of 200.times. and day
fourteen nodules (Panel F) are shown at 100.times..
[0021] FIG. 14. Nodule differentiated from adipose-derived stem
cells (PLA) express chondroitin-4-sulfate and keratin sulfate as
well as cartilage-specific collagen type II. Nodules induced from
adipose-derived stem cells for 2 days (Panels A and C) and 14 days
(Panels B and D) were embedded in paraffin and sectioned. Sections
were stained with monoclonal antibodies to the sulfated
proteoglycans chondroitin-4-sulfate and keratin sulfate. Sections
were also stained with monoclonal antibodies to collagen type II
(Panels E and F) (magnification 200.times.).
[0022] FIG. 15. RT-PCR analysis of nodules induced from
adipose-derived stem cells confirms the expression of collagens
type II and type X as well as expression of cartilage-specific
proteoglycan and aggrecan. Adipose-derived stem cells induced for
2, 7, and 14 days in chondrogenic medium and non-inductive control
medium were analyzed by RT-PCR for the expression of collagen type
I (CN I), type II (CN II), and type X (CN X) as well as
cartilage-specific proteoglycan (PG), aggrecan (AG), and
osteocalcin (OC).
[0023] FIG. 16. Adipose-derived stem cells induced in Myogenic
Medium express MyoD1. Panels A to C: adipose-derived stem cells
(PLA) were stained with an antibody to MyoD1 following 1 week
(Panel A), 3 weeks (Panel B) and 6 weeks (Panel C) induction in MM.
Expression of MyoD1 in the nucleus of positive staining PLA cells
is shown (arrows, magnification 200.times.). Panels D to F: PLA
cells induced for 1 week (Panel D), 3 weeks (Panel E) and 6 weeks
(Panel F) in non-inductive control medium (CM) were processed as
above as a negative control (magnification 200.times.).
[0024] FIG. 17. Adipose-derived stem cells induced in Myogenic
Medium express skeletal muscle myosin heavy chain. Panels A to C:
adipose-derived stem cells (PLA) cells were stained with an
antibody to the myosin heavy chain (myosin) following 1 week (Panel
A), 3 weeks (Panel B) and 6 weeks (Panel C) induction in MM.
Myosin-positive staining PLA cells are shown (arrows, magnification
200.times.). Panels D to F: adipose-derived stem cells (PLA) cells
induced for 1 week (Panel D), 3 weeks (Panel E) and 6 weeks (Panel
F) in non-inductive CM were processed as above as a negative
control (magnification 200.times.).
[0025] FIG. 18. Adipose-derived stem cells cultured in Myogenic
Medium form multi-nucleated cells. Panel A: Phase contrast of
adipose-derived stem cells (PLA) at 3 weeks (1) and 6 weeks (2)
post-induction with MM (magnification 400.times.). Multi-nucleated
cells are shown (arrows). Panel B: Immunostaining of
adipose-derived stem cells (PLA) cells at 6 weeks post-induction
with an antibody to the myosin heavy chain. Myosin-expressing
multi-nucleated cells are shown (arrows).
[0026] FIG. 19: RT-PCR analysis of adipose-derived stem cells
induced in MM. RT-PCR was performed on adipose-derived stem cells
induced for 1, 3 and 6 weeks in MM (PLA-MM) or in CM (PLA-CM),
using primers to human MyoD1 and myosin. RT-PCR analysis of human
foreskin fibroblast (HFF) cells induced in MM (HFF-MM) was also
performed as a negative control. Duplicate reactions were performed
using a primer set to .beta.-actin as an internal control. PCR
products were resolved by agarose gel electrophoresis and equalized
using .beta.-actin levels.
[0027] FIG. 20. The proportion of MyoD1-positive adipose-derived
stem cells increases with induction time. Histogram showing the
mean number of MyoD1-positive, adipose-derived stem cells (PLA)
after a 1, 3 and 6 week induction in MM (% of total PLA
cells.+-.SEM--hatched bars). The mean number of MyoD1-positive
cells observed after induction of adipose-derived stem cells with
CM (black bars) and HFF cells in MM (open bars) was also measured.
The values for each experiment are shown in table format below. A
statistical comparison of MyoD1 values from 1 to 6 weeks using a
one-way ANOVA was performed (asterisks; P<0.001, F=18.9).
Furthermore, an ANOVA was performed comparing the experimental and
control values for each time point. The p-values are shown
(p<0.0001).
[0028] FIG. 21. A time-dependent increase in myosin expression is
observed in induced adipose-derived stem cells. Histogram showing
the mean number of myosin-positive adipose-derived stem cells (PLA)
after a 1, 3 and 6 week induction in myosin medium (MM) (% of total
PLA cells.+-.SEM--hatched bars). The mean number of myosin-positive
cells observed after induction of adipose-derived stem cells with
control medium (CM) (black bars), and human foreskin fibroblast
cells (HFF) in myosin medium (MM) (open bars) was also measured.
The values for each experiment are shown in table format below. A
statistical comparison of myosin values from 1 to 6 weeks using a
one-way ANOVA was performed (asterisks; P<0.0001, F=75.5).
Furthermore, an ANOVA was performed comparing the experimental and
control values for each time point. The p-values are shown
(p<0.0001).
[0029] FIG. 22. Long-term chrondrogenic potetial of adipose-derived
stem cells. Adipose-derived stem cells, at passage 1 (panel A), 3
(panel B), and 15 (panel C), were induced under micromass
conditions and stained with Alcian blue staining at pH 1 to detect
the presence of sulfated proteoglycans.
[0030] FIG. 23. The adipose-derived stem cells (PLA) express a
unique set of CD markers. PLA cell and MSCs from human bone marrow
were processed for IF for the indicated CD antigens. Cells were
co-stained with DAPI to visualize nuclei (blue) and the fluorescent
images combined.
[0031] FIG. 24. CD marker profile of adipose-derived stem cells
(PLA) and bone marrow MSCs using flow cytometry. Panel A:
Adipose-derived stem cells were analyzed by FC using forward and
side scatter to assess cell size and granularity (FSC-H and SSC-H,
respectively). MSCs were analyzed as a control. Panel B: PLA cells
were fixed and incubated for the indicated CD markers using
fluorochrome-conjugated primary antibodies. Stained PLA cells were
subsequently analyzed by FC. MSCs and PLA cells stained with
fluorochrome-conjugated non-specific IgG were examined as a
positive and negative control, respectively. All results were
corrected for senescence and represent a total of 10.sup.5
events.
[0032] FIG. 25. Osteogenic adipose-derived stem cells (PLA) can be
characterized by distinct proliferative, synthetic and
mineralization phases. Adipose-derived stem cells were harvested
and plated into 35 mm tissue culture dishes in two sets of four
plates per differentiation period. All dishes were maintained in
Control medium until approximately 50% confluence was reached. The
cells were induced with Osteogenic medium (OM) and cell number was
counted at the indicated days. Cell number was expressed as the
number of adipose-derived stem cells (# cells (10.sup.5 )) and
plotted versus differentiation time (Panel A). For each time
period, one dish was stained for alkaline phosphatase (AP) activity
and one dish was stained using a Von Kossa stain (VK) to detect
calcium phosphate (Panel B).
[0033] FIG. 26. Dexamethasone and 1,25-dihydroxyvitamin D.sub.3
differentially affect PLA osteogenesis: AP enzyme and calcium
phosphate quantitation. Triplicate samples of PLA cells, MSCs and
NHOsts were induced for up to 6 weeks in OM, containing either
10.sup.-7 M Dexamethasone (OM/Dex) or 10.sup.-8 M
1,25-dihydroxyvitamin D.sub.3 (OM/VD). Cells were assayed for AP
activity, total calcium content and total protein. AP levels were
expressed as nmol p-nitrophenol formed per minute per microgram
protein (nmol p-nitrophenol/min/ug). Calcium levels were expressed
as mM calcium per microgram protein (mM Ca.sup.2+/ug). Non-induced
PLA cells (Control) were analyzed as a negative control. Values
were expressed as the mean.+-.SD.
[0034] FIG. 27. Osteo-induced PLA cells express several genes
consistent with osteogenic differentiation: RT-PCR and Microarray
analyses. Panel A: PLA cells were cultured in either OM/Dex, OM/VD
or non-inductive Control medium (Control) for the indicated days.
Total RNA was isolated, cDNA synthesized and PCR amplification
performed for the indicated genes. MSCs were induced in OM/Dex or
OM/VD and NHOsts were induced for 2 and 3 weeks in OM/Dex as
controls. Duplicate reactions were amplified using primers to
.beta.-actin as an internal control. Panel B: PLA cells were
induced for 3 weeks in OM/Dex or maintained in non-inductive
control medium. Total RNA was isolated and subject to microarray
analysis using a customized array containing the genes, OC, OP, ON,
CBFA1, CNI and BSP.
[0035] FIG. 28. Osteo-induced PLA cells express several proteins
consistent with osteogenic differentiation: Immunofluorescent and
Western analyses. Panel A: PLA cells and MSCs were induced in
OM/Dex or maintained in non-inductive Control medium (Control) for
21 days. Cells were processed for IF for the expression of OC, OP
and ON. Cells were co-stained with DAPI to visualize nuclei (blue)
and the fluorescent images combined. Panel B: PLA cells were
cultured in OM/Dex or non-inductive Control medium (Control) for 7
and 21 days. Cell lysates were separated by electrophoresis and
analyzed by Western blotting using antibodies to OP (.alpha.OP), ON
(.alpha.ON), Decorin (.alpha.DEC), Biglycan (.alpha.BG) and CNI
(.alpha.CNI). The expression of the transferrin receptor
(.alpha.TfR) was used as an internal control.
[0036] FIG. 29. Adipogenic differentiation by adipose-derived stem
cells (PLA) is accompanied by growth arrest. Adipose-derived stem
cells were harvested and plated into 35 mm tissue culture dishes in
one set of four plates per differentiation period. All dishes were
maintained in Control medium until approximately 80% confluence was
reached. The cells were induced with Adipogenic medium (AM) and
cell number was counted at the indicated days. Cell number was
expressed as the number of PLA cells (# cells (10.sup.5)) and
plotted versus differentiation time (Panel A). For each time
period, one dish was stained with Oil Red O to detect lipid
accumulation (Panel B).
[0037] FIG. 30. Adipogenic PLA cells express GPDH activity.
Triplicate samples of PLA cells and 3T3-L1 cells were induced for
up to 5 weeks in AM (PLA--AM, 3T3--AM, respectively). The cells
were assayed for GPDH activity and total protein. GPDH levels were
expressed as units GPDH per microgram protein (GPDH/ug).
Non-induced PLA cells were analyzed as a negative control
(PLA--Control). Values were expressed as mean.+-.SD.
[0038] FIG. 31. Adipose-derived stem cells express several genes
consistent with adipogenic differentiation: RT-PCR: Adipose-derived
stem cells were induced in AM (AM) or maintained in non-inductive
Control medium (Control) for the indicated days. Cells were
analyzed by RT-PCR for the indicated genes. MSCs and 3T3-L1 cells
were induced in AM as controls. Duplicate reactions were amplified
using primers to .beta.-actin as an internal control.
[0039] FIG. 32. Adipose-derived stem cell induced toward the
chondrogenic lineage are associated with the proteoglycans keratan
and chondroitin sulfate: Immunohistochemistry and
Dimethyldimethylene blue assay. Panel A: Adipose-derived stem cells
(PLA), under micromass conditions, were induced in chondrogenic
medium (CM) or maintained in non-inductive Control medium (Control)
for 7 days. Nodules were fixed, embedded in paraffin, sectioned and
stained with Alcian Blue to identify sulfated proteoglycans.
Sections were also stained for the expression of CNII, keratan
sulfate (KS) and chondroitin-4-sulfate (CS), followed by
counter-staining using H&E. Panel B: Triplicate samples of PLA
cells and NHCK cells were induced for up to 3 weeks in CM (PLA--CM,
NHCK--CM, respectively). Proteoglycan levels (keratan sulfate and
chondroitin sulfate) were determined and expressed as microgram
proteoglycan per microgram total protein (ug PG/ug). Non-induced,
Adipose-derived stem cells (PLA--Control) were analyzed as a
negative control. Values were expressed as the mean.+-.SD.
[0040] FIG. 33. Chondrogenic PLA cells express several genes
consistent with cartilage differentiation: RT-PCR. PLA cells, under
micromass culture conditions, were induced in CM for 4, 7, 10 and
14 days or maintained in non-inductive Control medium for 10 days
(Control). Cells were analyzed by RT-PCR for the indicated genes.
NHCK cells were induced in a commercial pro-chondrogenic medium as
a positive control. Duplicate reactions were performed using
primers to .beta.-actin as an internal control.
[0041] FIG. 34. PLA cells induced toward the myogenic lineage
express several genes consistent with myogenic differentiation:
RT-PCR analysis. PLA cells were induced in MM (PLA--MM) for 1, 3
and 6 weeks. Cells were analyzed by RT-PCR for the expression of
MyoD1 (MD1), myosin (MYS), myogenin (MG) and myf5 (MYF5). Total RNA
prepared from human skeletal muscle (SKM) was analyzed as a
positive control. Duplicate reactions were amplified using primers
to .beta.-actin as an internal control.
[0042] FIG. 35. ADSCs express multiple markers consistent with
multi-lineage capacity. ADSC Isolation: PLA cells were plated at
extremely low confluency in order to result in isolated single
cells. Cultures were maintained in Control medium until
proliferation of single PLA cells resulted in the formation of
well-defined colonies. The single PLA-cell derived colonies were
termed Adipose Derived Stem Cells (ADSCs). ADSCs were harvested
using sterile cloning rings and 0.25% trypsin/EDTA. The harvested
ADSCs were amplified in Cloning Medium (15% FBS, 1%
antibiotic/antimycotic in F12/DMEM (1:1)). Tri-lineage ADSC clones
were differentiated in OM, AM and CM and multi-lineage capacity by
IH using the following histological and IH assays: Alkaline
Phosphatase (osteogenesis), Oil Red O (adipogenic) and Alcian Blue
(chondrogenic).
[0043] FIG. 36. Isolation of multi-lineage clones from PLA
populations does not alter the expression profile of CD markers.
Dual- and tri-lineage clones were isolated and expanded from single
PLA cells. The clone populations were processed for the expression
of the indicated CD markers using IF. The ADSCs were co-stained
with DAPI to visualize nuclei (blue) and the fluorescent images
combined.
[0044] FIG. 37. ADSCs express multiple genes consistent with
multi-lineage capacity. Tri-lineage ADSC clones were cultured in
OM/VD (ADSC--Bone), AM (ADSC--Fat) and CM (ADSC--Cartilage), in
addition to control medium (ADSC--Control), followed by RT-PCR
analysis for the indicated lineage-specific genes. .beta.-actin
levels were analyzed as an internal control.
[0045] FIG. 38. PLA cells appear to exhibit neurogenic capacity in
vitro. Panel A: Light micrographs of non-induced PLA cells (PLA--0
hrs) and PLA cells induced with NM for 2 and 8 hrs (PLA--2hrs,
PLA--8 hrs, respectively). Panel B: PLA cells were maintained in NM
or Control medium for 5 hours (PLA--NM, PLA--Control, respectively)
and analyzed by IH for expression of the following lineage-specific
markers: NSE, trk-A, NeuN and MAP-2 (neural), GFAP (astrocytic).
PC12 cells treated with NGF were also assessed as a positive
control. Panel C: PLA cells were induced in NM for 4.5 and 9 hrs
and analyzed by RT-PCR for the indicated genes. In addition, PLA
cells were induced in NM for 9 hrs and maintained in NPMM for 1
week (NPMM). Non-induced PLA cells (Control) were analyzed as a
negative control. PC12 cells were examined as a positive control,
together with total RNA prepared from human brain (Brain).
[0046] FIG. 39. Clones isolated from adipose-derived stem cell
fractions exhibit neurogenic potential. Clones were examined using
imunnohistochemistry for adipogenic (oil red O stain), osteogenic
(alkaline phosphotase), chondrogenic (Alcian blue stain), and
neurogenic (anti-trka expression) differntiation.
[0047] FIG. 40. Osteogenic differentiation of the adipose-derived
stem cells (PLA) does not significantly alter CD marker expression.
PLA cells (Panel A) and MSCs (Panel B) were induced in OM for 3
weeks (PLA--Bone, MSC--Bone respectively), or maintained in
non-inductive Control medium (PLA--Control, MSC--Control). Cells
were processed for IF for the expression of CD34, CD44, CD45 and
CD90, co-stained with DAPI to visualize nuclei (blue) and the
fluorescent images combined.
[0048] FIG. 41. Adipogenic differentiation results in subtle
changes to the adipose-derived stem cells (PLA) CD marker profile.
PLA cells (Panel A) and MSCs (Panel B) were induced in AM for 2
weeks (PLA--Fat, MSC--Fat, respectively) or maintained in
non-inductive Control medium (PLA--Control, MSC--Control). Cells
were processed for IF for the expression of CD34, CD44, CD45 and
CD90, co-stained with DAPI to visualize nuclei (blue) and the
fluorescent images combined. To visualize adipocytes and their
staining pattern, fluorescent images were combined with light
micrographs (inset). Lipid-filled cells (white arrows--fluorescent
image; black arrows--inset) and fibroblasts (filled white
arrows--fluorescent image; filled black arrows--inset) are
indicated.
[0049] FIG. 42. Differentiation alters the expression of specific
CD markers on adipose-derived stem cells (PLA): Flow cytometry.
Panel A: PLA cells were maintained for 2 weeks in Control medium
(Control), or in OM (Osteogenic) or AM (Adipogenic). Cells were
analyzed by FC using forward and side scatter to assess cell size
and granularity (FSC-H and SSC-H, respectively). Panels B and C:
PLA cells were maintained for 2 weeks in Control medium (PLA--CM),
or in OM (PLA--OM) or AM (PLA--AM). Cells were directly stained for
the indicated CD markers using fluorochrome-conjugated primary
antibodies and analyzed by FC. The adipose-derived stem cells,
stained with fluorochrome-conjugated non-specific IgG, were
examined as a negative control. All results were corrected for
senescence and represent a total of 10.sup.5 events.
[0050] FIG. 43. Differentiation of the adipose-derived stem cells
(PLA) results in a change in ECM composition. PLA cells were
induced for either 3 weeks in OM (PLA--Bone), 2 weeks in AM
(PLA--Fat) or maintained in Control medium (PLA--Control). Cells
were processed for IF using antibodies to collagen type 1 (CNI),
type 4 (CNIV) and type 5 (CNV). Cells were co-stained with DAPI to
visualize nuclei (blue) and the fluorescent images combined.
Fluorescent images were combined with light micrographs (inset).
Lipid-filled PLA cells (white arrows--fluorescent image; black
arrows--inset) are indicated. Osteo-induced MSCs (MSC--Bone),
adipo-induced MSCs (MSC--Fat) and non-induced MSCs (MSC--Control)
were also analyzed.
DETAILED DESCRIPTION OF THE INVENTION
Definitions
[0051] As used herein, "stem cell" defines an adult
undifferentiated cell that can produce itself and a further
differentiated progeny cell.
[0052] As used herein, the "lineage" of a cell defines the heredity
of the cell, i.e.; which cells it came from and what cells it can
give rise to. The lineage of a cell places the cell within a
hereditary scheme of development and differentiation.
[0053] As used herein, the term "differentiates or differentiated"
defines a cell that takes on a more committed ("differentiated")
position within the lineage of a cell. "Dedifferentiated" defines a
cell that reverts to a less committed position within the lineage
of a cell.
[0054] As used herein, "a cell that differentiates into a
mesodermal (or ectodermal or endodermal) lineage" defines a cell
that becomes committed to a specific mesodermal, ectodermal or
endodermal lineage, respectively. Examples of cells that
differentiate into a mesodermal lineage or give rise to specific
mesodermal cells include, but are not limited to, cells that are
adipogenic, chondrogenic, cardiogenic, dermatogenic, hematopoetic,
hemangiogenic, myogenic, nephrogenic, urogenitogenic, osteogenic,
pericardiogenic, or stromal.
[0055] Examples of cells that differentiate into ectodermal lineage
include, but are not limited to epidermal cells, neurogenic cells,
and neurogliagenic cells.
[0056] Examples of cells that differentiate into endodermal lineage
include, but are not limited to pleurigenic cells, and hepatogenic
cells, cell that give rise to the lining of the intestine, and
cells that give rise to pancreogenic and splanchogenic cells.
[0057] As used herein, a "pluripotent cell" defines a less
differentiated cell that can give rise to at least two distinct
(genotypically and/or phenotypically) further differentiated
progeny cells.
[0058] A "multi-lineage stem cell" or "multipotent stem cell"
refers to a stem cell that reproduces itself and at least two
further differentiated progeny cells from distinct developmental
lineages. The lineages can be from the same germ layer (i.e.
mesoderm, ectoderm or endoderm), or from different germ layers. An
example of two progeny cells with distinct developmental lineages
from differentiation of a multi-lineage stem cell is a myogenic
cell and an adipogenic cell (both are of mesodermal origin, yet
give rise to different tissues). Another example is a neurogenic
cell (of ectodermal origin) and adipogenic cell (of mesodermal
origin).
[0059] As used here, "adipose tissue" defines a diffuse organ of
primary metabolic importance made-up of white fat, yellow fat or
brown fat. The adipose tissue has adipocytes and stroma. Adipose
tissue is found throughout the body of an animal. For example, in
mammals, adipose tissue is present in the omentum, bone marrow,
subcutaneous space and surrounding most organs.
[0060] As used herein "conditioned media" defines a medium in which
a specific cell or population of cells have been cultured in, and
then removed. While the cells were cultured in said medium, they
secrete cellular factors that include, but are not limited to
hormones, cytokines, extracellular matrix (ECM), proteins,
vesicles, antibodies, and granules. The medium plus the cellular
factors is the conditioned medium.
[0061] As used herein "isolated" defines a substance, for example
an adipose-derived stem cell, that is separated from contaminants
(i.e. substances that differ from the stem cell).
[0062] The present invention provides adipose-derived stem cells
(ADSCs) and methods for obtaining them from a mesodermal origin
(e.g., adipose tissue) and using them. Surprisingly, the inventive
ADSCs can differentiate into cells that give rise to more than one
type of germ layer, e.g. mesoderm, endoderm or ectoderm, and
combinations thereof, and are thus "multilineage" or "multipotent"
cells.
[0063] In another embodiment, the ADSCs can differentiate into two
or more distinct lineages from different germ layers (such as
endodermal and mesodermal), for example hepatocytes and
adipocytes.
[0064] The ADSCs of the invention can differentiate into cells of
two or more lineages, for example adipogenic, chondrogenic,
cardiogenic, dermatogenic, hematopoetic, hemangiogenic, myogenic,
nephrogenic, neurogenic, neuralgiagenic, urogenitogenic,
osteogenic, pericardiogenic, peritoneogenic, pleurogenic,
splanchogenic, and stromal developmental phenotypes. While such
cells can retain two or more of these different linages (or
developmental phenotypes), preferably, such ADSCs can differentiate
into three or more different lineages. The most preferred ADSCs can
differentiate into four or more lineages.
[0065] The ADSCs of the invention have the capacity to
differentiate into mesodermal tissues, such as mature adipose
tissue, bone, various tissues of the heart (e.g., pericardium,
epicardium, epimyocardium, myocardium, pericardium, valve tissue,
etc.), dermal connective tissue, hemangial tissues (e.g.,
corpuscles, endocardium, vascular epithelium, etc.), hematopeotic
tissue, muscle tissues (including skeletal muscles, cardiac
muscles, smooth muscles, etc.), urogenital tissues (e.g., kidney,
pronephros, meta- and meso-nephric ducts, metanephric diverticulum,
ureters, renal pelvis, collecting tubules, epithelium of the female
reproductive structures (particularly the oviducts, uterus, and
vagina), mesodermal glandular tissues (e.g., adrenal cortex
tissues), and stromal tissues (e.g., bone marrow). Of course,
inasmuch as the ADSC can retain potential to develop into a mature
cell, it also can realize its developmental phenotypic potential by
differentiating into an appropriate precursor cell (e.g., a
preadipocyte, a premyocyte, a preosteocyte, etc.).
[0066] In another embodiment, the ADSCs have the capacity to
differentiate into ectodermal tissues, such as neurogenic tissue,
and neurogliagenic tissue.
[0067] In another embodiment, the ADSCs have the capacity to
differentiate into endodermal tissues, such as pleurogenic tissue,
and splanchnogenic tissue, and hepatogenic tissue, and pancreogenic
tissue.
[0068] In yet another embodiment, ADSCs have the capacity to
dedifferentiate into developmentally immature cell types. Examples
of ADSCs dedifferentiating into an immature cell type, include
embryonic cells and fetal cells.
[0069] In another embodiment, the inventive ADSCs can give rise to
one or more cell lineages from one or more germ layers such as
neurogenic cells (of ectodermal origin) and myogenic cells (of
mesodermal origin).
[0070] The inventive ADSCs are useful for tissue engineering, wound
repair, in vivo and ex vivo tissue regeneration, tissue
transplantation, and other methods that require cells that can
differentiate into a variety of phenotypes and genotypes, or can
support other cell types in vivo or in vitro.
[0071] One aspect of the invention pertains to an adipose-derived
stem cell-enriched fraction (ADSC-EF) that contains adipose-derived
stem cells (ADSCs) of the invention. Preferably, the ADSC-EF is
substantially free of other cell types (e.g., adipocytes, red blood
cells, and other stromal cells, etc.) and extracellular matrix
material. More preferably, the ADSC-EF is completely free of such
other cell types and matrix material. The ADSC-EF is obtained from
adipose tissue of a mammal. The preferred embodiment includes an
ADSC-EF obtained from adipose tissue of a higher primate (e.g., a
baboon or ape). The most preferred ADSC enriched fraction is
obtained from human adipose tissue, using the methods described
herein.
Methods of Obtaining ADSC-EF and ADSCs of the Invention
[0072] The ADSCs of the invention are isolated from adipose tissue.
The adipose tissue can be obtained from an animal by any suitable
method. A first step in any such method requires the isolation of
the adipose tissue from the source animal. The animal can be alive
or dead, so long as adipose stromal cells within the animal are
viable. Typically, human adipose tissue is obtained from a living
donor, using well-recognized protocols such as surgical or suction
lipectomy. The preferred method to obtain human adipose tissue is
by excision or liposuction procedures well known in the art.
Preferably, the inventive ADSCs are isolated from a liposuction
aspirate. The ADSCs of the invention are present in the initially
excised or extracted adipose tissue, regardless of the method by
which the adipose tissue is obtained.
[0073] Three deposits of lipoaspirates, each from a different
patient, identified as 1',2',3', have been deposited on Sep. 7,
2001, with the American Type Culture Collection (ATCC), 10801
University Blvd., Manassas, Va. 20110-2209, under the provisions of
the Budapest Treaty, and have been accorded ATCC deposit numbers
______, ______ and ______.
[0074] However obtained, the adipose tissue is processed to
separate the ADSCs of the invention from the remainder of the
adipose tissue. The ADSC-EF that contains the ADSCs is obtained by
washing the obtained adipose tissue with a
physiologically-compatible solution, such as phosphate buffer
saline (PBS). The washing step consists of rinsing the adipose
tissue with PBS, agitating the tissue, and allowing the tissue to
settle. In addition to washing, the adipose tissue is dissociated.
The dissociation can occur by enzyme degradation and
neutralization. Alternatively, or in conjunction with such
enzymatic treatment, other dissociation methods can be used such as
mechanical agitation, sonic energy, or thermal energy. Three layers
form after the washing, dissociation, and settling steps. The top
layer is a free lipid layer. The middle layer includes the lattice
and adipocyte aggregates. The middle layer is referred to as an
"adipose-derived lattice enriched fraction." The middle layer or
the lattice-enriched fraction is filtered to concentrate the
lattice of the invention. A method of filtration involves passing
the middle layer through a large pore filter. The material which
does not pass through the filter includes the inventive lattice and
aggregates of adipocytes. The adipose-derived lattice can be
manually separated from the other cells which did not pass through
the filter.
[0075] The bottom layer contains the ADSC-EF and the inventive
ADSCs. The bottom layer is further processed to isolate the ADSCs
of the invention. The cellular fraction of the bottom layer is
concentrated into a pellet. One method to concentrate the cells
includes centrifugation.
[0076] The bottom layer is centrifuged and the pellet is retained.
The pellet is designated the adipose-derived stem cell-enriched
fraction (ADSC-EF) which includes the adipose-derived stem
cell-enriched fraction (ADSC-EF). The ADSC-EF may contain
erythrocytes (RBCs). In a preferred method the RBCs are lysed and
removed. Methods for lysis and removed RBCs are well known in the
art (e.g., incubation in hypotonic medium). If the RBCs are
removed, then the RBC-free fraction contains the ADSC-EF fraction
and the ADSCs. However, the RBCs are not required to be removed
from the ADSC-EF.
[0077] The pellet is resuspended and can be washed (in PBS),
centrifuged, and resuspended one or more successive times to
achieve greater purity of the ADSCs. The ADSC-EF of the invention
maybe a heterogenous population of cells which include the ADSCs of
the invention and adipocytes. The cells of the washed and
resuspended pellet are ready for plating.
[0078] The ADSCs in the resuspended pellet can be separated from
other cells of the resuspended pellet by methods that include, but
are not limited to, cell sorting, size fractionation, granularity,
density, molecularly, morphologically, and
immunohistologically.
[0079] In one embodiment, the ADSCs are separated from the other
cells on the basis of cell size and granularity where ADSCs are
small and agranular. Alternatively, a molecular method for
separating the ADSCs from the other cells of the pellet is by
assaying the length of the telomere. Stem cells tend to have longer
telomeres than differentiated cells.
[0080] In another embodiment, a biochemical method for separating
the ADSCs from the other cells of the pellet is used by assaying
telomerase activity. Telomerase activity can serve as a stem
cell-specific marker.
[0081] In still another embodiment, the ADSCs are separated from
the other cells of the pellet immunohistochemically, for example,
by panning, using magnetic beads, or affinity chromatography.
[0082] Alternatively, the process of isolating the ADSC enriched
fraction with the ADSCs is with a suitable device, many of which
are known in the art (see, e.g., U.S. Pat. No. 5,786,207). Such
devices can mechanically achieve the washing and dissociation
steps.
Culturing ADSCs
[0083] The ADSCs in the ADSC-EF can be cultured and, if desired,
assayed for number and viability, to assess the yield.
[0084] In one embodiment, the stem cells are cultured without
differentiation using standard cell culture media (e.g., DMEM,
typically supplemented with 5-15% (e.g., 10%) serum (e.g., fetal
bovine serum, horse serum, etc.). Preferably, the stem cells are
passaged at least five times in such medium without
differentiating, while still retaining their developmental
phenotype, and more preferably, the stem cells are passaged at
least 10 times (e.g., at least 15 times or even at least 20 times)
while retaining multipotency. Thus, culturing the ADSCs, without
inducing differentiation, can be accomplished without specially
screened lots of serum. In contrast, mesenchymal stem cells (e.g.,
derived from bone marrow) would differentiate under the same
culturing conditions described above. Methods for measuring
viability and yield are known in the art and can be employed (e.g.,
trypan blue exclusion).
[0085] The ADSCs can be separated by phenotypic identification, to
identify those cells that have two or more of the aforementioned
developmental lineages. To phenotypically separate the ADSCs from
the ADSC-EF, the cells are plated at a desired density, such as
between about 100 cells/cm.sup.2 to about 100,000 cells/cm.sup.2
(such as about 500 cells/cm.sup.2 to about 50,000 cells/cm.sup.2,
or, more particularly, between about 1,000 cells/cm.sup.2 to about
20,000 cells/cm.sup.2).
[0086] In a preferred embodiment the ADSC-EF is plated at a lower
density (e.g., about 300 cells/cm.sup.2) to facilitate the clonal
isolation of the ADSCs. For example, after a few days, ADSCs plated
at such densities will proliferate (expand) into a clonal
population of ADSCs.
[0087] Such ADSCs can be used to clone and expand a multipotent
ADSC into clonal populations, using a suitable method for cloning
cell populations. The cloning and expanding methods include
cultures of cells, or small aggregates of cells, physically picking
and seeding into a separate plate (such as the well of a multi-well
plate). Alternatively, the stem cells can be subcloned onto a
multi-well plate at a statistical ratio for facilitating placing a
single cell into each well (e.g., from about 0.1 to about 1
cell/well or even about 0.25 to about 0.5 cells/well, such as 0.5
cells/well). The ADSCs can be cloned by plating them at low density
(e.g., in a petri-dish or other suitable substrate) and isolating
them from other cells using devices such as a cloning rings.
Alternatively, where an irradiation source is available, clones can
be obtained by permitting the cells to grow into a monolayer and
then shielding one and irradiating the rest of cells within the
monolayer. The surviving cell then will grow into a clonal
population. While production of a clonal population can be expanded
in any suitable culture medium, a preferred culture condition for
cloning stem cells (such as the inventive stem cells or other stem
cells) is about 2/3 F.sub.12 medium+20% serum (preferably fetal
bovine serum) and about 1/3 standard medium that haw been
conditioned with stromal cells (e.g., cells from the stromal
vascular fraction of liposuction aspirate), the relative
proportions being determined volumetrically).
[0088] In any event, whether clonal or not, the isolated ADSCs can
be cultured in a specific inducing medium to induce the ADSC to
differentiate and express its multipotency. The ADSCs give rise to
cells of mesodermal, ectodermal and endodermal lineage, and
combinations thereof. Thus, one or more ADSCs derived from a
multipotent ADSC can be treated to differentiate into a variety of
cell types.
[0089] In another embodiment, the ADSCs are cultured in a defmed
medium for inducing adipogenic differentiation. Examples of specifc
media that induce the ADSCs of the invention to take on a
adipogenic phenotype include, but are not limited to media
containing a glucocorticoid (e.g., dexamethasone, hydrocortisone,
cortisone, etc.), insulin, a compound which elevates intracellular
levels of cAMP (e.g., dibutyryl-cAMP, 8-CPT-cAMP
(8-(4)chlorophenylthio)-adenosine 3',5' cyclic monophosphate;
8-bromo-cAMP; dioctanoyl-cAMP, forskolin etc.), and/or a compound
which inhibits degradation of cAMP (e.g., a phosphodiesterase
inhibitor such as isobutyl methyl xanthine (IBMX), methyl
isobutylxanthine, theophylline, caffeine, indomethacin, and the
like), and serum. Thus, exposure of the ADSCs to between about 1
.mu.M and about 10 .mu.M insulin in combination with about
10.sup.-9 M to about 10.sup.-6 M to (e.g., about 1 .mu.M)
dexamethasone can induce adipogenic differentiation. Such a medium
also can include other agents, such as indomethacin (e.g., about
100 .mu.M to about 200 .mu.M), if desired, and preferably the
medium is serum-free.
[0090] In another embodiment, ADSCs cultured in DMEM, 10% FBS, 1 uM
dexamthasone, 10 uM insulin, 200 uM indomethacin, 1%
antibiotic/antimicotic,(ABAM), 0.5 mM IBMX, take on an adipogenic
phenotype.
[0091] Culturing media that can induce osteogenic differentiation
of the ADSCs include, but are not limited to, about 10.sup.-7 M and
about 10.sup.-9 M dexamethasone (e.g., about 1 .mu.M) in
combination with about 10 .mu.M to about 50 .mu.M
ascorbate-2-phosphate and between about 10 nM and about 50 nM
.beta.-glycerophosphate. The medium also can include serum (e.g.,
bovine serum, horse serum, etc.).
[0092] In another embodiment, ADSCs cultured in DMEM, 10%FBS, 5%
horse serum, 50 .mu.M hydrocortisone, 10.sup.-7M dexamethosone, 50
Mascorbate-2-phosphate, 1% ABAM, take on an osteogenic
phenotype.
[0093] Culturing medium that can induce myogenic differentiation of
the ADSCs of the invention include, but is not limited to, exposing
the cells to between about 10 .mu.M and about 100 .mu.M
hydrocortisone, preferably in a serum-rich medium (e.g., containing
between about 10% and about 20% serum (either bovine, horse, or a
mixture thereof)). Other glucocorticoids that can be used include,
but are not limited to, dexamethasone. Alternatively,
5'-azacytidine can be used instead of a glucocorticoid.
[0094] In another embodiment, ADSCs cultured in DMEM, 10% FBS,
10.sup.-7M dexamethosone, 50 uMascorbate-2-phosphate, 10
mMbeta-glycerophosphate, 1% ABAM, take on an myogenic
phenotype.
[0095] Culturing medium that can induce chondrogenic
differentiation of the ADSCs of the invention, include but is not
limited to, exposing the cells to between about 1 .mu.M to about 10
.mu.M insulin and between about 1 .mu.M to about 10 .mu.M
transferrin, between about 1 ng/ml and 10 ng/ml transforming growth
factor (TGF) .beta.1, and between about 10 nM and about 50 nM
ascorbate-2-phosphate (50 nM). For chondrogenic differentiation,
preferably the cells are cultured in high density (e.g., at about
several million cells/ml or using micromass culture techniques),
and also in the presence of low amounts of serum (e.g., from about
1% to about 5%).
[0096] In another embodiment, ADSCs cultured in DMEM, 50
uMascorbate-2-phosphate, 6.25 ug/ml transferrin, 100 ng/ml insulin
growth factor (IGF-1), 5 ng/ml TGF-beta-1, 5 ng/ml basic fibroblast
growth factor (bFGF; used only for one week), assume an
chondrogenic phenotype.
[0097] In yet another embodiment, ADSCs are cultured in a
neurogenic medium such as DMEM, no serum and 5-10 mM
.beta.-mercaptoethanol and assume an ectodernal lineage.
[0098] The ADSCs also can be induced to dedifferentiate into a
developmentally more immature phenotype (e.g., a fetal or embryonic
phenotype). Such an induction is achieved upon exposure of the ADSC
to conditions that mimic those within fetuses and embryos. For
example, the inventive ADSCs, or population of ADSCs, can be
co-cultured with cells isolated from fetuses or embryos, or in the
presence of fetal serum.
[0099] The ADSCs of the invention can be induced to differentiate
into a mesodermal, ectodermal, or an endodermal lineage by
co-culturing the ADSCs with mature cells from the respective germ
layer, or precursors thereof.
[0100] In an embodiment, induction of the ADSCs into specific cell
types by co-culturing with differentiated mature cells includes,
but is not limited to, myogenic differentiation induced by
co-culturing the ADSCs with myocytes or myocyte precursors.
Induction of the ADSCs into a neural lineage by co-culturing with
neurons or neuronal precursors, and induction of the ADSCs into an
endodermal lineage, may occur by co-culturing with mature or
precursor pancreatic cells or mature hepatocytes or their
respective precursors.
[0101] Alternatively, the ADSCs are cultured in a conditioned
medium and induced to differentiate into a specific phenotype.
Conditioned medium is medium which was cultured with a mature cell
that provides cellular factors to the medium such as cytokines,
growth factors, hormones, and extracellular matrix. For example, a
medium that has been exposed to mature myoctytes is used to culture
and induce ADSCs to differentiate into a myogenic lineage. Other
examples of conditioned media inducing specific differentiation
include, but are not limited to, culturing in a medium conditioned
by exposure to heart valve cells to induce differentiation into
heart valve tissue. In addition, ADSCs are cultured in a medium
conditioned by neurons to induce a neuronal lineage, or conditioned
by hepatoycytes to induce an endodermal lineage.
[0102] For co-culture, it may be desirable for the ADSCs and the
desired other cells to be co-cultured under conditions in which the
two cell types are in contact. This can be achieved, for example,
by seeding the cells as a heterogeneous population of cells onto a
suitable culture substrate. Alternatively, the ADSCs can first be
grown to confluence, which will serve as a substrate for the second
desired cells to be cultured within the conditioned medium.
[0103] Other methods of inducing differentiation are known in the
art and can be employed to induce the ADSCs to give rise to cells
having a mesodermal, ectodermal or endodermal lineage.
[0104] After culturing the stem cells in the
differentiating-inducing medium for a suitable time (e.g., several
days to a week or more), the ADSCs can be assayed to determine
whether, in fact, they have acquired the desired lineage.
[0105] Methods to characterize differentiated cells that develop
from the ADSCs of the invention, include, but are not limited to,
histological, morphological, biochemical and immunohistochemical
methods, or using cell surface markers, or genetically or
molecularly, or by identifying factors secreted by the
differentiated cell, and by the inductive qualities of the
differentiated ADSCs.
[0106] Molecular markers that characterize mesodermal cell that
differentiate from the ADSCs of the invention, include, but are not
limited to, MyoD, myosin, alpha-actin, brachyury, xFOG, Xtbx5
FoxF1, XNkx-2.5. Mammalian homologs of the above mentioned markers
are preferred.
[0107] Molecular markers that characterize ectodermal cell that
differentiate from the ADSCs of the invention, include but are not
limited to N-CAM, GABA and epidermis specific keratin. Mammalian
homologs of the above mentioned markers are preferred.
[0108] Molecular markers that characterize endodermal cell that
differentiate from the ADSCs include, but are not limited to,
Xhbox8, Endo1, Xhex, Xcad2, Edd, EF1-alpha, HNF3-beta, LFABP,
albumin, insulin. Mammalian homologs of the above mentioned markers
are preferred.
[0109] In an embodiment, molecular characterization of the
differentiated ADSCs is by measurement of telomere length.
Undifferentiated stem cells have longer telomeres than
differentiated cells; thus the cells can be assayed for the level
of telomerase activity. Alternatively, RNA or proteins can be
extracted from the ADSCs and assayed (via Northern hybridization,
RTPCR, Western blot analysis, etc.) for the presence of markers
indicative of a specific phenotype.
[0110] In an alternative embodiment, differentiation is assessed by
assaying the cells immunohistochemically or histologically , using
tissue-specific antibodies or stains, respectively. For example, to
assess adipogenic differentiation, the differentiated ADSCs are
stained with fat-specific stains (e.g., oil red O, safarin red,
sudan black, etc.) or with labeled antibodies or molecular markers
that identify adipose-related factors (e.g., PPAR-.gamma., adipsin,
lipoprotein lipase, etc.).
[0111] In another embodiment, ostogenesis can be assessed by
staining the differentiated ADSCs with bone-specific stains (e.g.,
alkaline phosphatase, von Kossa, etc.) or with labeled antibodies
or molecular markers that identify bone-specific markers (e.g.,
osteocalcin, osteonectin, osteopontin, type I collagen, bone
morphogenic proteins, cbfa, etc.).
[0112] Myogensis can be assessed by identifying classical
morphologic changes (e.g., polynucleated cells, syncitia formation,
etc.), or assessed biochemically for the presence of
muscle-specific factors (e.g., myo D, myosin heavy chain,
etc.).
[0113] Chondrogenesis can be determined by staining the cells using
cartilage-specific stains (e.g., Alcian blue) or with labeled
antibodies or molecular markers that identify cartilage-specific
molecules (e.g., sulfated glycosaminoglycans and proteoglycans,
keratin, chondroitin, Type II collagen, etc.) in the medium.
[0114] Alternative embodiments can employ methods of assessing
developmental phenotype, known in the art. For example, the cells
can be sorted by size and granularity. The cells can be used as an
immunogen to generate monoclonal antibodies (Kohler and Milstein),
which can then be used to bind to a given cell type. Correlation of
antigenicity can confirm that the ADSC has differentiated along a
given developmental pathway.
[0115] While an ADSC can be isolated, preferably it is within a
population of cells. The invention provides a defmed population of
ADSCs. In an embodiment, the population is heterogeneous. In
another embodiment, the population is homogeneous.
[0116] In another embodiment, a population of ADSCs can support
cells for culturing other cells. For example, cells that can be
supported by ADSC populations include other types of stem cells,
such as neural stem cells (NSC), hematopoetic stem cells (HPC,
particularly CD34.sup.+ stem cells), embryonic stem cells (ESC) and
mixtures thereof). In other embodiments, the population is
substantially homogeneous, consisting essentially of the inventive
adipose-derived stem cells.
Uses of the ADSC-EF, ADSCs and Methods of the Invention
[0117] The ADSC-EF can be used as a source of the ADSCs of the
invention. The ADSC-EF can be introduced into a subject for tissue
regeneration, wound repair or other applications requiring a source
of stem cells. In addition, the ADSC-EF can be treated to cause the
ADSCs therein to differentiate into a desired cell type for
introduction into a subject. The ADSC-EF can also be cultured in
vitro to maintain a source of ADSCs, or can be induced to produce
further differentiated ADSCs that can develop into a desired
tissue.
[0118] The ADSCs (and populations of ADSCs) can be employed for a
variety of purposes. The ADSCs can support the growth and expansion
of other cell types. The invention includes a method of
conditioning culture medium using the ADSCs in a suitable medium,
and the ADSC-conditioned medium produced by such a method.
Typically, the medium is used to support the in vitro growth of the
ADSCs, which secrete hormones, cell matrix material, and other
factors into the medium. After a suitable period (e.g., one or a
few days), the culture medium containing the secreted factors can
be separated from the cells and stored for future use. The ADSCs
can be re-used successively to condition medium, as desired. In
other applications (e.g., for co-culturing the ADSCs with other
cell types), the cells can remain within the conditioned medium.
Thus, the invention provides an ADSC-conditioned medium obtained
using this method, which either can contain the ADSCs, or be
substantially free of the ADSCs, as desired.
[0119] The ADSC-conditioned medium can be used to support the
growth and expansion of desired cell types, and the invention
provides a method of culturing cells (particularly stem cells)
using the conditioned medium. The method involves maintaining a
desired cell in the conditioned medium under conditions for the
cell to remain viable. The cell can be maintained under any
suitable condition for culturing them, such as are known in the
art. Desirably, the method permits successive rounds of mitotic
division of the cell to form an expanded population. The exact
conditions (e.g., temperature, CO.sub.2 levels, agitation, presence
of antibiotics, etc.) will depend on the other constituents of the
medium and on the cell type. However, optimizing these parameters
is within the ordinary skill in the art.
[0120] In another embodiment, the ADSCs can be genetically
modified, e.g., to express exogenous genes ("transgenes") or to
repress the expression of endogenous genes, and the invention
provides a method of genetically modifying such cells and
populations. In accordance with this method, the ADSC is exposed to
a gene transfer vector comprising a nucleic acid including a
transgene, such that the nucleic acid is introduced into the cell
under conditions appropriate for the transgene to be expressed
within the cell. The transgene generally is an expression cassette,
including a polynucleotide operably linked to a suitable promoter.
The polynucleotide can encode a protein, or it can encode
biologically active RNA (e.g., antisense RNA or a ribozyme). Thus,
for example, the polynucleotide can encode a gene conferring
resistance to a toxin, a hormone (such as peptide growth hormones,
hormone releasing factors, sex hormones, adrenocorticotrophic
hormones, cytokines (e.g., interfering, interleukins, lymphokines),
etc.), a cell-surface-bound intracellular signaling moiety (e.g.,
cell adhesion molecules, hormone receptors, etc.), a factor
promoting a given lineage of differentiation, (e.g., bone
morphogenic protein (BMP)) etc. Of course, where it is desired to
employ gene transfer technology to deliver a given transgene, its
sequence will be known.
[0121] Within the expression cassette, the coding polynucleotide is
operably linked to a suitable promoter. Examples of suitable
promoters include prokaryotic promoters and viral promoters (e.g.,
retroviral ITRs, LTRs, immediate early viral promoters (IEp), such
as herpesvirus IEp (e.g., ICP4-IEp and ICP0-IEp), cytomegalovirms
(CMV) lEp, and other viral promoters, such as Rous Sarcoma Virus
(RSV) promoters, and Murine Leukemia Virus (MLV) promoters). Other
suitable promoters are eukaryotic promoters, such as enhancers
(e.g., the rabbit .beta.-globin regulatory elements),
constitutively active promoters (e.g., the .beta.-actin promoter,
etc.), signal specific promoters (e.g., inducible promoters such as
a promoter responsive to RU486, etc.), and tissue-specific
promoters. It is well within the skill of the art to select a
promoter suitable for driving gene expression in a predefined
cellular context. The expression cassette can include more than one
coding polynucleotide, and it can include other elements (e.g.,
polyadenylation sequences, sequences encoding a membrane-insertion
signal or a secretion leader, ribosome entry sequences,
transcriptional regulatory elements (e.g., enhancers, silencers,
etc.), and the like), as desired.
[0122] The expression cassette containing the transgene should be
incorporated into a genetic vector suitable for delivering the
transgene to the cells. Depending on the desired end application,
any such vector can be so employed to genetically modify the cells
(e.g., plasmids, naked DNA, viruses such as adenovirus,
adeno-associated virus, herpesviruses, lentiviruses,
papillomaviruses, retroviruses, etc.). Any method of constructing
the desired expression cassette within such vectors can be
employed, many of which are well known in the art (e.g., direct
cloning, homologous recombination, etc.). Of course, the choice of
vector will largely determine the method used to introduce the
vector into the cells (e.g., by protoplast fusion,
calcium-phosphate precipitation, gene gun, electroporation,
infection with viral vectors, etc.), which are generally known in
the art.
[0123] The genetically altered ADSCs can be employed as bioreactors
for producing the product of the transgene. In other embodiments,
the genetically modified ADSCs are employed to deliver the
transgene and its product to an animal. For example, the ADSCs,
once genetically modified, can be introduced into the animal under
conditions sufficient for the transgene to be expressed in
vivo.
[0124] In addition to serving as useful targets for genetic
modification, many ADSCs and populations of ADSCs secrete hormones
(e.g., cytokines, peptide or other (e.g., monobutyrin) growth
factors, etc.). Some of the cells naturally secrete such hormones
upon initial isolation, and other cells can be genetically modified
to secrete hormones, as discussed herein. The ADSCs that secrete
hormones can be used in a variety of contexts in vivo and in vitro.
For example, such cells can be employed as bioreactors to provide a
ready source of a given hormone, and the invention pertains to a
method of obtaining hormones from such cells. In accordance with
the method, the ADSCs are cultured, under suitable conditions for
them to secrete the hormone into the culture medium. After a
suitable period of time, and preferably periodically, the medium is
harvested and processed to isolate the hormone from the medium. Any
standard method (e.g., gel or affinity chromatography, dialysis,
lyophilization, etc.) can be used to purify the hormone from the
medium, many of which are known in the art.
[0125] In other embodiments, ADSCs (and populations) secreting
hormones can be employed as therapeutic agents. Generally, such
methods involve transferring the cells to desired tissue, either in
vitro (e.g., as a graft prior to implantation or engrafting) or in
vivo, to animal tissue directly. The cells can be transferred to
the desired tissue by any method appropriate, which generally will
vary according to the tissue type. For example, ADSCs can be
transferred to a graft by bathing the graft (or infusing it) with
culture medium containing the cells. Alternatively, the ADSCs can
be seeded onto the desired site within the tissue to establish a
population. Cells can be transferred to sites in vivo using devices
such as catheters, trocars, cannulae, stents (which can be seeded
with the cells), etc. For these applications, preferably the ADSC
secretes a cytokine or growth hormone such as human growth factor,
fibroblast growth factor, nerve growth factor, insulin-like growth
factors, hemopoietic stem cell growth factors, members of the
fibroblast growth factor family, members of the platelet-derived
growth factor family, vascular and endothelial cell growth factors,
members of the TGFb family (including bone morphogenic factor), or
enzymes specific for congenital disorders (e.g., dystrophic).
[0126] In one application, the invention provides a method of
promoting the closure of a wound within a patient using ADSCs. In
accordance with the method, ADSCs secreting the hormone are
transferred to the vicinity of a wound under conditions sufficient
for the cells to produce the hormone. The presence of the hormone
in the vicinity of the wound promotes closure of the wound. The
method promotes closure of both external (e.g., surface) and
internal wounds. Wounds to which the present inventive method is
useful in promoting closure include, but are not limited to,
abrasions, avulsions, blowing wounds, bum wounds, contusions,
gunshot wounds, incised wounds, open wounds, penetrating wounds,
perforating wounds, puncture wounds, seton wounds, stab wounds,
surgical wounds, subcutaneous wounds, or tangential wounds. The
method need not achieve complete healing or closure of the wound;
it is sufficient for the method to promote any degree of wound
closure. In this respect, the method can be employed alone or as an
adjunct to other methods for healing wounded tissue.
[0127] Where the ADSCs secrete an angiogenic hormone (e.g.,
vascular growth factor, vascular and endothelial cell growth
factor, etc.), they (as well as populations containing them) can be
employed to induce angiogenesis within tissues. Thus, the invention
provides a method of promoting or inhibiting neovascularization
within tissue using such ADSCs. The presence of the hormone within
the tissue promotes or inhibits neovascularization. In accordance
with this method, the ADSC is introduced the desired tissue under
conditions sufficient for the cell to produce the angiogenic
hormone. The presence of the hormone within the tissue promotes
neovascularization within the tissue.
[0128] Because the ADSCs have a developmental phenotype, they can
be employed in tissue engineering. In this regard, the invention
provides a method of producing animal matter comprising maintaining
the ADSCs under conditions sufficient for them to expand and
differentiate to form the desired matter. The matter can include
mature tissues, or even whole organs, including tissue types into
which the inventive cells can differentiate (as set forth herein).
Typically, such matter will comprise adipose, cartilage, heart,
dermal connective tissue, blood tissue, muscle, kidney, bone,
pleural, splanchnic tissues, vascular tissues, and the like. More
typically, the matter will comprise combinations of these tissue
types (i.e., more than one tissue type). For example, the matter
can comprise all or a portion of an animal organ (e.g., a heart, a
kidney) or a limb (e.g., a leg, a wing, an arm, a hand, a foot,
etc.). Of course, in as much as the cells can divide and
differentiate to produce such structures, they can also form
anlagen of such structures. At early stages, such anlagen can be
cryopreserved for future generation of the desired mature structure
or organ.
[0129] To produce such structures, the ADSCs and populations are
maintained under conditions suitable for them to expand and divide
to form the desired structures. In some applications, this is
accomplished by transferring them to an animal (i.e., in vivo)
typically at a sight at which the new matter is desired. Thus, for
example, the invention can facilitate the regeneration of tissues
(e.g., bone, muscle, cartilage, tendons, adipose, etc.) within an
animal where the ADSCs are implanted into such tissues. In other
embodiments, and particularly to create anlagen, the ADSCs can be
induced to differentiate and expand into tissues in vitro. In such
applications, the ADSCs are cultured on substrates that facilitate
formation into three-dimensional structures conducive for tissue
development. Thus, for example, the ADSCs can be cultured or seeded
onto a bio-compatible lattice, such as one that includes
extracellular matrix material, synthetic polymers, cytokines,
growth factors, etc. Such a lattice can be molded into desired
shapes for facilitating the development of tissue types. Also, at
least at an early stage during such culturing, the medium and/or
substrate is supplemented with factors (e.g., growth factors,
cytokines, extracellular matrix material, etc.) that facilitate the
development of appropriate tissue types and structures. Indeed, in
some embodiments, it is desired to co-culture the ADSCs with mature
cells of the respective tissue type, or precursors thereof, or to
expose the cells to the respective conditioned medium, as discussed
herein.
[0130] To facilitate the use of the ADSCs and populations for
producing such animal matter and tissues, the invention provides a
composition including the ADSCs (and populations) and biologically
compatible lattice. Typically, the lattice is formed from polymeric
material, having fibers as a mesh or sponge, typically with spaces
on the order of between about 100 .mu.m and about 300 .mu.m. Such a
structure provides sufficient area on which the cells can grow and
proliferate. Desirably, the lattice is biodegradable over time, so
that it will be absorbed into the animal matter as it develops.
Suitable polymeric lattices, thus, can be formed from monomers such
as glycolic acid, lactic acid, propyl fumarate, caprolactone,
hyaluronan, hyaluronic acid, and the like. Other lattices can
include proteins, polysaccharides, polyhydroxy acids,
polyorthoesters, polyanhydrides, polyphosphazenes, or synthetic
polymers (particularly biodegradable polymers). Of course, a
suitable polymer for forming such lattice can include more than one
monomers (e.g., combinations of the indicated monomers). Also, the
lattice can also include hormones, such as growth factors,
cytokines, and morphogens (e.g., retinoic acid, aracadonic acid,
etc.), desired extracellular matrix molecules (e.g., fibronectin,
laminin, collagen, etc.), or other materials (e.g., DNA, viruses,
other cell types, etc.) as desired.
[0131] To form the composition, the ADSCs are introduced into the
lattice such that they permeate into the interstitial spaces
therein. For example, the matrix can be soaked in a solution or
suspension containing the cells, or they can be infused or injected
into the matrix. A particularly preferred composition is a hydrogel
formed by crosslinking of a suspension including the polymer and
also having the inventive cells dispersed therein. This method of
formation permits the cells to be dispersed throughout the lattice,
facilitating more even permeation of the lattice with the cells. Of
course, the composition also can include mature cells of a desired
phenotype or precursors thereof, particularly to potentate the
induction of the ADSCs to differentiate appropriately within the
lattice (e.g., as an effect of co-culturing such cells within the
lattice).
[0132] The composition can be employed in any suitable manner to
facilitate the growth and generation of the desired tissue types,
structures, or anlagen. For example, the composition can be
constructed using three-dimensional or sterotactic modeling
techniques. Thus, for example, a layer or domain within the
composition can be populated by cells primed for osteogenic
differentiation, and another layer or domain within the composition
can be populated with cells primed for myogenic and/or chondrogenic
development. Bringing such domains into juxtaposition with each
other facilitates the molding and differentiation of complex
structures including multiple tissue types (e.g., bone surrounded
by muscle, such as found in a limb). To direct the growth and
differentiation of the desired structure, the composition can be
cultured ex vivo in a bioreactor or incubator, as appropriate. In
other embodiments, the structure is implanted within the host
animal directly at the site in which it is desired to grow the
tissue or structure. In still another embodiment, the composition
can be engrafted on a host (typically an animal such as a pig,
baboon, etc.), where it will grow and mature until ready for use.
Thereafter, the mature structure (or anlagen) is excised from the
host and implanted into the host, as appropriate.
[0133] Lattices suitable for inclusion into the composition can be
derived from any suitable source (e.g., matrigel), and some
commercial sources for suitable lattices exist (e.g., suitable of
polyglycolic acid can be obtained from sources such as Ethicon,
N.J.). Another suitable lattice can be derived from the acellular
portion of adipose tissue--i.e., adipose tissue extracellular
matrix matter substantially devoid of cells, and the invention
provides such a adipose-derived lattice. Typically, such
adipose-derived lattice includes proteins such as proteoglycans,
glycoproteins, hyaluronins, fibronectins, collagens (type I, type
II, type III, type IV, type V, type VI, etc.), and the like, which
serve as excellent substrates for cell growth. Additionally, such
adipose-derived lattices can include hormones, preferably cytokines
and growth factors, for facilitating the growth of cells seeded
into the matrix.
[0134] The adipose-derived matrix can be isolated form adipose
tissue similarly as described above, except that it will be present
in the acellular fraction. For example, adipose tissue or
derivatives thereof (e.g; the lattice enriched supernant fraction
of the method described above) can be subjected to sonic or thermal
energy and/or enzymatic processed to recover the matrix material.
Also, desirably the cellular fraction of the adipose tissue is
disrupted, for example by treating it with lipases, detergents,
proteases, and/or by mechanical or sonic disruption (e.g., using a
homogenizer or sonicator). However isolated, the material is
initially identified as a viscous material, but it can be
subsequently treated, as desired, depending on the desired end use.
For example, the raw matrix material can be treated (e.g., dialyzed
or treated with proteases or acids, etc.) to produce a desirable
lattice material. Thus the lattice can be prepared in a hyrated
form or it can be dried or lyophilized into a substantially
anhydrous form or a powder. Thereafter, the powder can be
rehydrated for use as a cell culture substrate, for example by
suspending it in a suitable cell culture medium. In this regard,
the adipose-derived lattice can be mixed with other suitable
lattice materials, such as described above. Of course, the
invention pertains to compositions including the adipose-derived
lattice and cells or populations of cells, such as the inventive
ADSCs and other cells as well (particularly other types of stem
cells).
[0135] As discussed above, the ADSCs, populations, lattices, and
compositions of the invention can be used in tissue engineering and
regeneration. Thus, the invention pertains to an implantable
structure (i.e., an implant) incorporating any of these inventive
features. The exact nature of the implant will vary according to
the use to which it is to be put. The implant can be or comprise,
as described, mature tissue, or it can include immature tissue or
the lattice. Thus, for example, one type of implant can be a bone
implant, comprising a population of the inventive cells that are
undergoing (or are primed for) osteogenic differentiation,
optionally seeded within a lattice of a suitable size and
dimension, as described above. Such an implant can be injected or
engrafted within a host to encourage the generation or regeneration
of mature bone tissue within the patient. Similar implants can be
used to encourage the growth or regeneration of muscle, fat,
cartilage, tendons, etc., within patients. Other types of implants
are anlagen (such as described herein), e.g., limb buds, digit
buds, developing kidneys, etc, that, once engrafted onto a patient,
will mature into the appropriate structures.
[0136] The adipose-derived lattice can conveniently be employed as
part of a cell culture kit. Accordingly, the invention provides a
kit including the inventive adipose-derived lattice and one or more
other components, such as hydrating agents (e.g., water,
physiologically-compatible saline solutions, prepared cell culture
media, serum or derivatives thereof, etc.), cell culture substrates
(e.g., culture dishes, plates, vials, etc.), cell culture media
(whether in liquid or powdered form), antibiotic compounds,
hormones, and the like. While the kit can include any such
ingredients, preferably it includes all ingredients necessary to
support the culture and growth of desired cell types upon proper
combination. Of course, if desired, the kit also can include cells
(typically frozen), which can be seeded into the lattice as
described herein.
[0137] While many aspects of the invention pertain to tissue growth
and differentiation, the invention has other applications as well.
For example, the adipose-derived lattice can be used as an
experimental reagent, such as in developing improved lattices and
substrates for tissue growth and differentiation. The
adipose-derived lattice also can be employed cosmetically, for
example, to hide wrinkles, scars, cutaneous depressions, etc., or
for tissue augmentation. For such applications, preferably the
lattice is stylized and packaged in unit dosage form. If desired,
it can be admixed with carriers (e.g., solvents such as glycerine
or alcohols), perfumes, antibiotics, colorants, and other
ingredients commonly employed in cosmetic products. The substrate
also can be employed autologously or as an allograft, and it can
used as, or included within, ointments or dressings for
facilitating wound healing. The ADSCs can also be used as
experimental reagents. For example, they can be employed to help
discover agents responsible for early events in differentiation.
For example, the inventive cells can be exposed to a medium for
inducing a particular line of differentiation and then assayed for
differential expression of genes (e.g., by random-primed PCR or
electrophoresis or protein or RNA, etc.).
[0138] As any of the steps for isolating the inventive ADSCs or the
adipose-derived lattice, the, the invention provides a kit for
isolating such reagents from adipose tissues. The kit can include a
means for isolating adipose tissue from a patient (e.g., a cannula,
a needle, an aspirator, etc.), as well as a means for separating
stromal cells (e.g., through methods described herein). The kit can
be employed, for example, as an immediate source of ADSCs that can
then be re-introduced from the same individual as appropriate.
Thus, the kit can facilitate the isolation of adipose-derived stem
cells for implantation in a patient needing regrowth of a desired
tissue type, even in the same procedure. In this respect, the kit
can also include a medium for differentiating the cells, such as
those set forth herein. As appropriate, the cells can be exposed to
the medium to prime them for differentiation within the patient as
needed. In addition, the kit can be used as a convenient source of
ADSCs for in vitro manipulation (e.g., cloning or differentiating
as described herein). In another embodiment, the kit can be
employed for isolating a adipose-derived lattice as described
herein.
[0139] While one of skill in the art is fully able to practice the
instant invention upon reading the foregoing detailed description,
the following examples will help elucidate some of its features. In
particular, they demonstrate the isolation of a human
adipose-derived stem cell substantially free of mature adipocytes,
the isolation of a clonal population of such cells, the ability of
such cells to differentiate in vivo and in vitro into all cells
with a mesodermal phenotype, endodermal phenotype, and extodermal
phenotype, and the capacity of such cells to support the growth of
other types of stem cells. The examples also demonstrate the
isolation of a adipose-derived lattice substantially free of cells
that is capable of serving as a suitable substrate for cell
culture. Of course, as these examples are presented for purely
illustrative purposes, they should not be used to construe the
scope of the invention in a limited manner, but rather should be
seen as expanding upon the foregoing description of the invention
as a whole.
[0140] The procedures employed in these examples, such as surgery,
cell culture, enzymatic digestion, histology, and molecular
analysis of proteins and polynucleotides, are familiar to those of
ordinary skill in this art. As such, and in the interest of
brevity, experimental details are not recited in detail.
EXAMPLE 1
[0141] This example demonstrates the isolation of a human
adipose-derived stem cell substantially free of mature
adipocytes.
[0142] Raw liposuction aspirate was obtained from patients
undergoing elective surgery. Prior to the liposuction procedures,
the patients were given epinephrine to minimize contamination of
the aspirate with blood. The aspirate was strained to separate
associated adipose tissue pieces from associated liquid waste.
Isolated tissue was rinsed thoroughly with neutral phosphate
buffered saline and then enzymatically dissociated with 0.075% w/v
collagenase at 37.degree. C. for about 20 minutes under
intermittent agitation. Following the digestion, the collagenase
was neutralized, and the slurry was centrifuged at about 260 g for
about 10 minutes, which produced a multi-layered supernatant and a
cellular pellet. The supernatant was removed and retained for
further use, and the pellet was resuspended in an
erythrocyte-lysing solution and incubated without agitation at
about 25.degree. C. for about 10 minutes. Following incubation, the
medium was neutralized, and the cells were again centrifuged at
about 250 g for about 10 minutes. Following the second
centrifugation, the cells were suspended, and assessed for
viability (using trypan blue exclusion) and cell number.
Thereafter, they were plated at a density of about at about
1.times.10.sup.6 cells/100 mm dish. They were cultured at
37.degree. C. in DMEM+fetal bovine serum (about 10%) in about 5%
CO.sub.2.
[0143] The majority of the cells were adherent, small, mononucleic,
relatively agranular fibroblast-like cells containing no visible
lipid droplets and were CD34-negative. The majority of the cells
stained negatively with oil-red O and von Kossa. The cells were
also assayed for expression of telomerase (using a commercially
available TRAP assay kit), using HeLa cells and HN-12 cells as
positive controls. Human foreskin fibroblasts and HN-12 heated cell
extracts were used as negative controls. Telomeric products were
resolved onto a 12.5% polyacrylamide cells and the signals
determined by phosphorimaging. Telemeric ladders representing
telomerase activity were observed in the adipose-derived stem cells
as well as the positive controls. No ladders were observed in the
negative controls.
[0144] Thus, these cells were not identifiable as myocytes,
adipocytes, chondrocytes, osteocytes, or blood cells. These results
demonstrate that the adipose-derived cells express telomerase
activity similar to that previously reported for human stem
cells.
[0145] Subpopulations of these cells were then exposed to the
following media to assess their developmental phenotype:
1 Adipogenesis Osteogenesis Myogenesis Chondrogenesis DMEM DMEM
DMEM DMEM 10% FETAL 10% FETAL 10% FETAL 1% FETAL BOVINE BOVINE
BOVINE BOVINE SERUM SERUM SERUM SERUM 0.5 mM ISO- 5% horse serum 5%
horse serum 6.25 .mu.g/ml insulin BUTYL- 0.1 .mu.M 50 .mu.M 6.25
.mu.g/ml METHYL- dexamethasone hydrocortisone transferrin XANTHINE
50 .mu.M ascorbate- 1% ABAM 10 ng/ml 1 .mu.M 2-phosphate TGF.beta.1
dexamethasone 10 mM .beta.- 50 nM 10 .mu.M insulin glycerophosphate
ascorbate-2- 200 .mu.M 1% ABAM phosphate indomethacin 1% ABAM 1%
ABAM
[0146] A population was cultured at high density in the
chondrogenic medium for several weeks. Histological analysis of the
tissue culture and paraffin sections was performed with H&E,
alcian blue, toludene blue, and Goldner's trichrome staining at 2,
7, and 14 days. Immunohistochemistry was performed using antibodies
against chondroitin-4-sulfate and keratin sulfate and type II
collagen. Qualitative estimate of matrix staining was also
performed. The results indicated that cartilaginous spheroid
nodules with a distinct border of perichondral cells formed as
early as 48 hours after initial treatment. Untreated control cells
exhibited no evidence of chondrogenic differentiation. These
results confirm that the stem cells have chondrogenic developmental
phenotype.
[0147] A population was cultured until near confluence and then
exposed to the adipogenic medium for several weeks. The population
was examined at two and four weeks after plating by calorimetric
assessment of relative opacity following oil red-O staining.
Adipogenesis was determined to be underway at two weeks and quite
advanced at four weeks (relative opacity of 1 and 5.3,
respectively). Bone marrow-derived stem cells were employed as a
positive control, and these cells exhibited slightly less
adipogenic potential (relative density of 0.7 and 2.8,
respectively).
[0148] A population was cultured until near confluence and then
exposed to the osteogeneic medium for several weeks. The population
was examined at two and four weeks after plating by colorimetric
assessment of relative opacity following von Kossa staining.
Osteogenesis was determined to be underway at two weeks and quite
advanced at four weeks (relative opacity of 1.1 and 7.3,
respectively. Bone marrow-derived stem cells were employed as a
positive control, and these cells exhibited slightly less
osteogenic potential (relative density of 0.2 and 6.6,
respectively).
[0149] A population was cultured until near confluence and then
exposed to the myogenic medium for several weeks. The population
was examined at one, three, and six weeks after plating by
assessment of multinucleated cells and expression of
muscle-specific proteins (MyoD and myosin heavy chain). Human
foreskin fibroblasts and skeletal myoblasts were used as controls.
Cells expressing MyoD and myosin were found at all time points
following exposure to the myogenic medium in the stem cell
population, and the proportion of such cells increased at 3 and 6
weeks. Multinucleated cells were observed at 6 weeks. In contrast,
the fibroblasts exhibited none of these characteristics at any time
points.
[0150] These results demonstrate the isolation of a human
adipose-derived pluripotent stem cell substantially free of mature
adipocytes.
EXAMPLE 2
[0151] This example demonstrates that the adipose-derived stem
cells do not differentiate in response to 5-azacytidine.
[0152] Adipose-derived stem cells obtained in accordance with
Example 1 were cultured in the presence of 5-azacytidine. In
contrast with bone marrow-derived stem cells, exposure to this
agent did not induce myogenic differentiation (see Wakitani et al.,
supra).
EXAMPLE 3
[0153] This example demonstrates the generation of a clonal
population of human adipose-derived stem cells from an
adipose-derived stem cell enriched fraction.
[0154] Cells isolated in accordance with the procedure set forth in
Example 1 were plated at about 5,000 cells/100 mm dish and cultured
for a few days as indicated in Example 1. After some rounds of cell
division, some clones were picked with a cloning ring and
transferred to wells in a 48 well plate. These cells were cultured
for several weeks, changing the medium twice weekly, until they
were about 80% to about 90% confluent (at 37.degree. C. in about 5%
CO.sub.2 in 2/3 F.sub.12 medium+20% fetal bovine serum and 1/3
standard medium that was first conditioned by the cells isolated in
Example 1, "cloning medium"). Thereafter, each culture was
transferred to a 35 mm dish and grown, and then retransferred to a
100 mm dish and grown until close to confluent. Following this, one
cell population was frozen, and the remaining populations were
plated on 12 well plates, at 1000 cells/well.
[0155] The cells were cultured for more than 15 passages in cloning
medium and monitored for differentiation as indicated in Example 1.
The undifferentiated state of each clone remained true after
successive rounds of culturing.
[0156] Populations of the clones then were established and exposed
to adipogenic, chondrogenic, myogenic, and osteogenic medium as
discussed in Example 1. It was observed that at least one of the
clones was able to differentiate into bone, fat, cartilage, and
muscle when exposed to the respective media, and most of the clones
were able to differentiate into at least three types of tissues.
The capacity of the cells to develop into muscle and cartilage
further demonstrates the pluripotentiality of these adipose-derived
stem cells.
[0157] These results demonstrate that the adipose-derived stem
cells can be maintained in an undifferentiated state for many
passages without the requirement for specially pre-screened lots of
serum. The results also demonstrate that the cells retain
pluripotentiality following such extensive passaging, proving that
the cells are indeed stem cells and not merely committed progenitor
cells.
EXAMPLE 4
[0158] This example demonstrates the adipose-derived stem cells
from an adipose-devired stem cell enriched fraction can support the
culture of other types of stem cells.
[0159] Human adipose-derived stem cells were passaged onto 96 well
plates at a density of about 30,000/well, cultured for one week and
then irradiated. Human CD34.sup.+ hematopoetic stem cells isolated
from umbilical cord blood were then seeded into the wells.
Co-cultures were maintained in MyeloCult H5100 media, and cell
viability and proliferation were monitored subjectively by
microscopic observation. After two weeks of co-culture, the
hematopoetic stem cells were evaluated for CD34 expression by flow
cytometry.
[0160] Over a two-week period of co-culture with stromal cells, the
hematopoetic stem cells formed large colonies of rounded cells.
Flow analysis revealed that 62% of the cells remained CD34.sup.+.
Based on microscopic observations, human adipo-derived stromal
cells maintained the survival and supported the growth of human
hematopoetic stem cells derived from umbilical cord blood.
[0161] These results demonstrate that stromal cells from human
subcutaneous adipose tissue are able to support the ex vivo
maintenance, growth and differentiation of other stem cells.
EXAMPLE 5
[0162] This example demonstrates that the adipose-derived stem
cells from an adipose-devired stem cell enriched fraction can
differentiate in vivo.
[0163] Four groups (A-D) of 12 athymic mice each were implanted
subcutaneously with hydroxyapatite/tricalcium phosphate cubes
containing the following: Group A contained adipose-derived stem
cells that had been pretreated with osteogenic medium as set forth
in Example 1. Group B contained untreated adipose-derived stem
cells. Group C contained osteogenic medium but no cells. Group D
contained non-osteogenic medium and no cells. Within each group,
six mice were sacrificed at three weeks, and the remaining mice
sacrificed at eight weeks following implantation. The cubes were
extracted, fixed, decalcified, and sectioned. Each section was
analyzed by staining with hematoxylin and eosin (e.g., H&E),
Mallory bone stain, and immunostaining for osteocalcin.
[0164] Distinct regions of osteoid-like tissue staining for
osteocalcin and Mallory bone staining was observed in sections from
groups A and B. Substantially more osteoid tissue was observed in
groups A and B than in the other groups (p<0.05 ANOVA), but no
significant difference in osteogenesis was observed between groups
A and B. Moreover, a qualitative increase in bone growth was noted
in both groups A and B between 3 and 8 weeks. These results
demonstrate that the adipose-derived stem cells can differentiate
in vivo.
EXAMPLE 6
[0165] This example demonstrates the isolation of an
adipose-derived lattice substantially devoid of cells.
[0166] In one protocol, the lattice-enriched fraction from Example
1 was subjected to enzymatic digestion for three days in 0.05%
trypsin EDTA/100 U/ml deoxyribonuclease to destroy the cells. Every
day the debris was rinsed in saline and fresh enzyme was added.
Thereafter the material was rinsed in saline and resuspended in
0.05% collagease and about 0.1% lipase to partially digest the
proteins and fat present. This incubation continued for two
days.
[0167] In another protocol, the withheld supernatant from Example 1
was incubated in EDTA to eliminate any epithelial cells. The
remaining cells were lysed using a buffer containing 1% NP40, 0.5%
sodium deoxycholate, 0.1% SDS, 5 mM EDTA, 0.4M NaCl, 50 mMTris-HCL
(pH 8) and protease inhibitors, and 10 .mu.g/ml each of leupeptin,
chymostatin, antipain, and pepstatin A. Finally, the tissue was
extensively washed in PBS without divalent cations.
[0168] After both preparatory protocols, remaining substance was
washed and identified as a gelatinous mass. Microscopic analysis of
this material revealed that it contained no cells, and it was
composed of high amounts of collagen (likely type IV) and a wide
variety of growth factors. Preparations of this material have
supported the growth of cells, demonstrating it to be an excellent
substrate for tissue culture.
EXAMPLE 7
[0169] The following description provides adipose-derived stem
cells enriched fractions which exhibit mesodermal multi-tissue
potential, and methods for isolating said stem cells.
Materials and Methods
[0170] All materials were purchased from Sigma (St. Louis, Mo.)
unless otherwise stated. All tissue culture reagents were purchased
from Life Technologies (New York, N.Y.). Fetal Bovine Serum (FBS)
and Horse Serum (HS) were purchased from Hyclone (Logan, Utah) and
Life Technologies, respectively.
Cell Lines
[0171] Normal Human Osteoblasts (NHOsts), human Skeletal Muscle
(SkM) cells and a population of Mesenchymal Stem Cells derived from
bone marrow (MSCs) were purchased from Clonetics (Walkersville,
Md.). The murine 3T3-L1 pre-adipocyte cell line (Green H., and
Meuth, M., 1974, Cell 3: 127-133) was obtained from ATCC
(Rockville, Md.). Human Foreskin Fibroblasts (HFFs) were obtained
from Cascade Biologics (Portland, Oreg.).
Isolation and Culture of Stem Cells
[0172] Human adipose tissue was obtained from elective liposuction
procedures under local anesthesia according to patient consent
protocol, HSPC #98-08 011-02 (Univerisity of California Los
Angeles). In this procedure, a hollow blunt-tipped cannula was
introduced into the subcutaneous space through small (.about.1 cm)
incisions. The cannula was attached to a gentle suction and moved
through the adipose compartment, mechanically disrupting the fat
tissue. A solution of saline and the vasoconstrictor, epinephrine,
was infused into the adipose compartment to minimize blood loss and
contamination of the tissue by peripheral blood cells. The raw
lipoaspirate (approximately 300 cc) was processed according to
established methodologies in order to obtain a stromal vascular
fraction (SVF) (Hauner H., et al., 1987, J. Clin. Endocrinol.
Metabol. 64: 832-835; Katz, A. J., et al., 1999 Clin. Plast. Surg.
26: 587-603, viii). To isolate the SVF, lipoaspirates were washed
extensively with equal volumes of Phospho-Buffered Saline (PBS) and
the extracellular matrix (ECM) was digested at 37.degree. C. for 30
minutes with 0.075% collagenase. Enzyme activity was neutralized
with Dulbecco's Modified Eagle's Medium (DMEM), containing 10%
Fetal Bovine Serum (FBS) and centrifuged at 1200.times.g for 10
minutes to obtain a high-density SVF pellet. The pellet was
resuspended in 160 mM NH.sub.4Cl and incubated at room temperature
for 10 minutes to lyse contaminating red blood cells. The SVF was
collected by centrifugation, as detailed above, filtered through a
100 .mu.m nylon mesh to remove cellular debris and incubated
overnight at 37.degree. C./5% CO.sub.2 in Control Medium (DMEM, 10%
FBS, 1% antibiotic/antimycotic solution). Following incubation, the
plates were washed extensively with PBS to remove residual
non-adherent red blood cells. The resulting cell population was
termed an adipose-derived stem cell enriched fraction (ADSC
enriched fraction), in order to distinguish it from the SVF
obtained from excised adipose tissue. The adipose-derived stem
cells were maintained at 37.degree. C./5% CO.sub.2 in non-inductive
Control Medium. Cells did not require specific FBS sera lots for
expansion and differentiation . For immunofluorescent studies, a
population of MSCs was obtained from human bone marrow aspirates
according to the protocol of Rickard et al. (Rickard D. J., et al.,
1996, J. Bone Min. Res. 11: 312-324) and maintained in Control
medium. To prevent spontaneous differentiation, cells were
maintained at subconfluent levels.
Indirect Immunofluorescence of Stem Cells
[0173] Stem cells and MSCs obtained from human bone marrow
aspirates were plated onto glass chamber slides and fixed for 15
minutes in 4% paraformaldehyde in 100 mM sodium phosphate buffer
(pH 7.0). The cells were washed for 10 minutes in 100 mM glycine in
PBS (PBS/glycine) and blocked for 1 hour in Immunofluorescent
Blocking Buffer (IBB; 5% BSA, 10% FBS, 1.times.PBS, 0.1% Triton
X-100). The cells were subsequently incubated for 1 hour in IBB
containing the following cell-specific monoclonal antibodies: 1)
anti-smooth muscle actin (anti-SMA; Cedarlane Inc, Homby Ont), to
identify smooth muscle cells and pericytes (Skalli, O., et al.,
1986, J. Cell Biol. 103:2787-2796; Schurch, W., et al., 1987, Am J.
Pathol 128:91-103; Nehls, A. and D. Drenckhahn 1991, J. CellBiol.
113:147-154; Barghom, A. et al., 1998, Pediatr. Pathol. Lab. Med.
18:5-22)); 2) anti-Factor VIII (anti-FVIII; Calbiochem, San Diego,
Calif.), to identify endothelial cells (Jaffe, E A, et al., 1973,
J. Clin. Envest. 52:2757-2764; Nagle, R B, et al., 1987 Lymphology
20:20-24); and 3) ASO2 (dianova, Hamburg, Germany), to identify
fibroblasts and cells of mesenchymal origin (Saalbach, A., et al.,
1996 J. Invest.Dermatol. 106:1314-1319; Saalbach, A., et al., 1997
Cell and Tiss. Res. 290:595-599). The cells were washed extensively
with PBS/glycine and incubated for 1 hour in IBB containing an
FITC-conjugated secondary antibody. The cells were washed with
PBS/glycine and mounted with a solution containing DAPI to
visualize nuclei (VectaShield, Vector Labs, Burlingame,
Calif.).
Flow Cytometry
[0174] Adipose-derived stem cells samples from 5 donors were
cultured in Control Medium for 72 hours prior to analysis. Flow
cytometry was performed on a FACScan argon laser cytometer (Becton
Dickson, San Jose, Calif.). Cells were harvested in 0.25%
trypsin/EDTA and fixed for 30 minutes in ice-cold 2% formaldehyde.
Following fixation, cells were washed in Flow Cytometry Buffer
(FCB; 1.times.PBS, 2% FBS, 0.2% Tween-20). Cell aliquots
(1.times.10.sup.6 cells) were incubated in FCB containing
monoclonal antibodies to Factor VIII, smooth muscle actin or ASO2.
In addition, cells were also incubated with FCB containing a
monoclonal antibody to vimentin (anti-VIM; Biogenesis, Brentwood,
N.H.), to identify mesenchymal cells (Lazarides, E. 1982 Annu. Rev.
Biochem. 51:219-250; Suza, S., et al., 1996 Eur. J. Cell Biol.
70:84-91). To assess viability, duplicate samples were harvested,
fixed for 30 minutes with ice-cold 1% paraformaldehyde,
permeabilized with 0.05% Nonidet-40 and incubated with propidium
iodide (PI) at a concentration of 25 .mu.g/ml. Debris and dead
cells were excluded by eliminating PI-positive events. All
subsequent adipose-derived stem cell samples were corrected
accordingly.
Cumulative Population Doubling
[0175] Adipose-derived stem cells cells were maintained in Control
Medium until 80% confluent. Cells were harvested at confluence and
population doubling calculated using the formula log
N.sub.1/logN.sub.2, where N.sub.1 is the number of cells at
confluence prior to passaging and N.sub.2 is the number of cells
seeded after passaging. Cumulative population doubling was
determined in cultures maintained until passage 13 (approximately
165 days). The mean cumulative population doubling obtained from 3
donors was expressed as a function of passage number.
Cell Senescence Assay
[0176] Senescence was assessed using a .beta.-gal staining assay,
in which .beta.-galactosidase activity is detected in senescent
cells at pH 6.0 but is absent in proliferating cells (Dimri, G P,
et al., 1995 Proc. Natl. Acad. Sci. USA 92:9363-9367). Cells from
each culture passage (passage 1 to passage 15) were fixed for 5
minutes in 2% formaldehyde/glutaraldehyde and incubated in a
.beta.-Gal Reaction Buffer (1 mg/ml X-Gal, 40 mM citric acid/sodium
phosphate buffer (pH 6.0), 5 mM each of potassium ferrocyanide and
potassium ferricyanide, 150 mM NaCl and 2 mM MgCl.sub.2). Senescent
cells (blue) were identified by light microscopy.
Confirmation of Multi-lineage Differentiation of Adipose-Derived
Stem Cells
[0177] Adipose-derived stem cells at passage 1 were analyzed for
their capacity to differentiate toward the adipogenic, osteogenic,
chondrogenic and myogenic lineages. To induce differentiation, the
stem cells were cultured with specific induction media as detailed
in Table 1. Each media has been previously described and shown to
induce multi-lineage differentiation of MSCs (Pittenger, M F., et
al., 1999 Science 284:143-147; Grigoradis, A., et al., 1988 J. Cell
Biol. 106:2139-2151; Cheng, S-L., et al., 1994 Endo 134:277-286;
Loffler, G., et al., 1987 Klin. Wochenschr. 65:812-817; Hauner, H.,
et al., 1987 J. Clin. Endocrinol. Metabol. 64:832-835).
Differentiation was confirmed using the histological and
immunohistological assays outlined in Table 2. A commercial source
of bone marrow-derived MSCs and lineage-specific precursors were
examined as positive controls. The adipose-derived stem cells were
maintained in Control Medium and HFFs were analyzed as negative
controls.
[0178] 1. Adipogenesis: Adipogenic differentiation was induced by
culturing the stem cells for 2 weeks in Adipogenic Medium (AM) and
assessed using an Oil Red O stain as an indicator of intracellular
lipid accumulation (Preece, A. 1972 A Manual for Histologic
Technicians, Boston, Mass.: Little, Brown, and Co.). Prior to
staining, the cells were fixed for 60 minutes at room temperature
in 4% formaldehyde/1% calcium and washed with 70% ethanol. The
cells were incubated in 2% (w/v) Oil Red O reagent for 5 minutes at
room temperature. Excess stain was removed by washing with 70%
ethanol, followed by several changes of distilled water. The cells
were counter-stained for 2 minutes with hematoxylin.
[0179] 2. Osteogenesis: Osteogenic differentiation was induced by
culturing the stem cells for a minimum of 2 weeks in Osteogenic
Medium (OM) and examined for Alkaline Phosphatase (AP) activity and
ECM calcification by von Kossa staining. To detect AP activity,
cells were incubated in OM for 2 weeks, rinsed with PBS and stained
with a 1% AP solution (1% napthol ABSI phosphate, 1 mg/ml Fast Red
TR) at 37.degree. C. for 30 minutes. For von Kossa staining, the
cells were incubated in OM for 4 weeks and fixed with 4%
paraformaldehyde for 60 minutes at room temperature. The cells were
rinsed with distilled water and then overlaid with a 1% (w/v)
silver nitrate solution in the absence of light for 30 minutes. The
cells were washed several times with distilled water and developed
under UV light for 60 minutes. Finally, the cells were
counter-stained with 0.1% eosin in ethanol.
[0180] 3. Chondrogenesis: Chondrogenic differentiation was induced
using the micromass culture technique (Ahrens, P B, et al., 1977
Develop. Biol. 60:69-82; Reddi, A H 1982 Prog. Clin. Biol. Res. 110
(part B):261-268; Denker, A E., et al., 1995 Differentiation
59:25-34). Briefly, 10 .mu.l of a concentrated adipose-derived stem
cell suspension (8.times.10.sup.6 cells/ml) was plated into the
center of each well and allowed to attach at 37.degree. C. for two
hours. Chondrogenic medium (CM) was gently overlaid so as not to
detach the cell nodules and cultures were maintained in CM for 2
weeks prior to analysis. Chondrogenesis was confirmed using the
histologic stain Alcian Blue at acidic pH. The stem cell nodules
were fixed with 4% paraformaldehyde for 15 minutes at room
temperature and washed with several changes of PBS. Studies have
shown specific staining of sulfated proteoglycans, present in
cartilagenous matrices, at pH levels of 1 and below (Lev, R. and S.
Spicer 1964 J. Histochem. Cytochem. 12:309-312). In light of this,
the cells were incubated for 30 minutes with 1% (w/v) Alcian Blue
(Sigma A-3157) in 0.1N HCl (pH 1.0) and washed with 0.1N HCl for 5
minutes to remove excess stain. In addition to Alcian Blue
staining, expression of the cartilage-specific collagen type II
isoform was also determined. The stem cells were fixed in 4%
paraformaldehyde for 15 minutes at room temperature. Cells were
incubated in 0.2 U/ml chondroitinase ABC for 40 min at 37.degree.
C. to facilitate antibody access to collagen II. The cells were
rinsed in PBS and endogenous peroxidase activity quenched by
incubating for 10 minutes in 3% hydrogen peroxide in methanol.
Following a wash in PBS, non-specific sites were blocked by
incubating cells for 1 hour in Blocking Buffer (PBS, containing 10%
Horse Serum). The cells were subsequently incubated for 1 hour in
Blocking Buffer containing a monoclonal antibody specific to human
collagen type II (ICN Biomedical, Costa Mesa, Calif.). The cells
were washed extensively in Blocking Buffer and collagen type II
visualized using a commercially available kit for the detection of
monoclonal antibodies according to the manufacturer (VectaStain ABC
kit, Vector Labs Inc., Burlingame, Calif.).
[0181] 4. Myogenesis: Myogenic differentiation was induced by
culturing the adipose-derived stem cells in Myogenic Medium (MM)
for 6 weeks and confirmed by immunohistochemical staining for the
muscle-specific transcription factor, MyoD1 and the myosin heavy
chain. Cells were rinsed twice with PBS, fixed for 20 minutes with
4% paraformaldehyde and washed several times with PBS. The cells
were incubated with 3% hydrogen peroxide in PBS for 10 minutes to
quench endogenous peroxidase enzyme activity and non-specific sites
were blocked by incubation in Blocking Buffer (PBS, 10% HS, 0.1%
Triton X-100) for an additional 60 minutes. The cells were washed 3
times for 5 minutes each in Blocking Buffer and incubated for 1
hour in Blocking Buffer containing a either a monoclonal antibody
specific to skeletal muscle myosin heavy chain (Biomeda, Foster
City, Calif.) or to MyoD1 (Dako Corp, Carpenteria, Calif.). The
cells were washed extensively in Blocking Buffer and the monoclonal
antibodies visualized using the VectaStain ABC kit according to
manufacturer's specifications. The cells were counter-stained with
hematoxylin for 3 minutes.
Results
[0182] Human adipose tissue was obtained by suction-assisted
lipectomy (i.e. liposuction) and the lipoaspirates were processed
based on adapted methodologies (Katz, A J, et al., 1999 Clin.
Plast. Surg. 26:587-603, viii), in order to obtain a Processed
Lipoaspirate or PLA cell (adipose-derived stem cells) population,
containing the putative stem cell fraction. Processing of 300 cc of
liposuctioned tissue routinely yielded stem cell samples of
2-6.times.10.sup.8 cells. The cultures were maintained in DMEM
supplemented with 10% Fetal Bovine Serum (FBS). Supplementation
with FBS has been shown to be important for human and animal MSC
attachment and proliferation in vitro (Haynesworth, S E, et al.,
1992 Bone 13:81-88; Lennon, D P, et al., 1995 Exp. Cell Res.
219:211-222; Lennon, D P, et al., 1996 In Vitro Cell Dev. Biol.
32:602-611). However, studies suggest that proliferation and
differentiation of human MSCs may be dependent upon FBS source and
quality, making sera screening critical (Lennon, D P, et al., 1995
Exp. Cell Res. 219:211-222; Lennon, D P, et al., 1996 In Vitro Cell
Dev. Biol. 32:602-611). The stem cells expanded easily in vitro and
exhibited a fibroblast-like morphology, consistent with that of
MSCs obtained from bone marrow and a commercial source (FIG. 1A).
The stem cells did not appear to require specific sera lots for
expansion and multi-lineage differentiation. Ten FBS lots from
three manufacturers were tested and did not appear to alter the
stem cell morphology, proliferation rate or their differentiative
capacity in vitro.
Growth Kinetics and Composition of the PLA
[0183] The adipose-derived stem cells, obtained from 20 donors and
cultured under standard conditions (i.e. 10% FBS), exhibited an
average population doubling time of 60 hours using several sera
sources and lots. Following isolation, an initial lag time of 5 to
7 days was observed in stem cell cultures. Cells then entered a
proliferative phase reaching confluence within 48 hours. To examine
long-term growth kinetics of the stem cell cultures, we measured
cumulative population doublings with respect to passage number in
multiple donors. Consistent with the observed lag time upon initial
culture, the stem cells underwent an average of three population
doublings prior to the first passage (FIG. 1B). An average of 1.5
population doublings was observed upon subsequent passages. A
linear relationship between cumulative population doubling and
passage number was observed, indicating a relatively constant
population doubling rate over the range studied. Furthermore, no
appreciable decrease in cumulative population doublings was
observed at later passages (P13=165 days in culture), suggesting
that the stem cell cultures maintain their proliferative potential
during extended culture periods.
[0184] In addition to cumulative population doubling, we also
examined cell senescence in long-term stem cell cultures using a
.beta.-gal staining protocol, in which .beta.-galactosidase
expression is absent in proliferating cells but can be detected in
senescent cells at a pH of 6.0 (Dimri, G P, et al., 1995 Proc.
Natl. Acad. Sci. USA 92:9363-9367). Using this assay, the stem cell
cultures were examined for senescence at each passage. The stem
cell cultures at passage 1 exhibited no appreciable .beta.-gal
staining (FIG. 1C, P1). An increase in .beta.-gal staining was
observed at later passages, however the percentage of senescent
cells remained below 5% through 10 passages and increased to 15% at
passage 15. Taken together, the data indicates that adipose-derived
stem cell samples are relatively stable over long-term culture,
maintaining a consistent population doubling rate and exhibiting
low levels of senescence.
[0185] The SVF processed from excised adipose tissue is a
heterogenous population including mast cells, endothelial cells,
pericytes, fibroblasts and lineage-committed progenitor cells, or
pre-adipocytes (Pettersson, P. et al., 1984 Acta Med. Scand.
215:447-453; Hauner, H., et al., 1987 J. Clin. Endocrinol. Metabol.
64:832-835). These components may also be present, together with
the putative stem cell fraction obtained from liposuctioned adipose
tissue. However, no literature regarding this has been published.
To phenotypically characterize the stem cells, samples from several
donors were examined by indirect immunofluorescence using
antibodies specific to established cell-surface markers. A bone
marrow stromal fraction obtained from human marrow aspirates was
also examined as a control. To identify endothelial cells, the stem
cells were incubated with a monoclonal antibody to Factor VIII
(Jaffe, E A, et al., 1973 J. Clin. Invest. 52:2757-2764; Nagle, R
B, et al., 1987 Lymphology 20:20-24). Smooth muscle cells were
identified using a monoclonal antibody to smooth muscle actin
(Lazarides, E. 1982 Annu. Rev. Biochem. 51:219-250; Suza, S., et
al., 1996 Eur. J. Cell Biol. 70:84-91). This antibody has also been
shown to react with transitional pericytes (i.e. pericytes of pre-
and post-capillaries) and the contractile apparatus of pericytes
committed to the smooth muscle lineage (Nehls, A. and D. Drenckhahn
1991 J. Cell Biol. 113:147-154; Herman, I M and P A D'Amore 1985 J.
Cell Biol. 101:43-52). Low levels of endothelial cells, smooth
muscle cells and pericytes were observed in the stem cell fraction
(FIG. 2). In comparison, no staining for these markers was observed
in processed bone marrow stromal samples. In addition to Factor
VIII and smooth muscle actin, cells were also incubated with a
monoclonal antibody (ASO2) specific to fibroblasts and mesenchymal
cells (Saalbach, A., et al., 1996 J. Invest. Dermatol.
106:1314-1319; Saalbach, A., et al., 1997 Cell and Tiss. Res.
290:595-599). The majority of the stem cells and bone marrow
stromal cells stained positively with ASO2, suggesting a
mesenchymal origin (FIG. 2, ASO2 panels).
[0186] To quantitatively determine the stem cell composition,
samples were analyzed by flow cytometry using the cell surface
markers described above. The samples were obtained and cultured for
72 hours in Control Medium. Cell size and granularity were measured
using forward and side scatter settings (FIG. 3A). The majority of
the stem cell sample was comprised of small, agranular cells. In
addition, the stem cells were incubated with monoclonal antibodies
to Factor VIII, smooth muscle actin and ASO2 and a monoclonal
antibody to vimentin, an intermediate filament protein found
predominantly in cells of mesenchymal origin (Lazarides, E. 1982
Annu. Rev. Biochem. 51:219-250; Suza, S., et al., 1996 Eur. J. Cell
Biol. 70:84-91). Viability was assessed using propidium iodide and
samples were corrected for viability, non-specific fluorescence and
autofluorescence. Data from a representative patient is shown (FIG.
3B). Cytometry data was collected from 5 donors and the number of
positive events for each cell-specific marker was expressed as a
percentage of the total stem cell number. Consistent with the
immunofluorescent data, a fraction of the stem cells expressed
Factor VIII (FVIII-positive cells=24.9%.+-.8.2 of total stem cell
number) and smooth muscle actin (SMA-positive cells=29.2%.+-.2.1 of
total PLA cell number) (FIG. 3C), indicating that the stem cell
fraction contains endothelial cells, smooth muscle cells and,
possibly, pericytes. Furthermore, the majority of the stem cells
stained positively for ASO2 (ASO2-positive cells=85.0%.+-.12.8 of
total PLA cell number) and vimentin (VIM-positive
cells=63.2%.+-.5.6 of total cell number), indicative of cells of
mesenchymal origin. Taken together, the results suggest that the
stem cell fraction is a relatively homogenous population of
mesodermal or mesenchymal cells with low contamination by
endothelial cells, pericytes and smooth muscle cells.
Adipose-Derived Stem Cells Exhibit Multi-Lineage Potential
[0187] To study the multi-lineage capacity of the adipose-derived
stem cells cells, cells were differentiated toward the adipogenic,
osteogenic, chondrogenic and myogenic lineages using
lineage-specific induction factors (Table 1). Human and animal bone
marrow-derived MSCs have been shown to differentiate toward the
adipogenic, osteogenic and chondrogenic lineages with appropriate
medium supplementation (Pittenger, M F., et al., 1999 Science
284:143-147; Grigoradis, A., et al., 1988 J. Cell Biol.
106:2139-2151; Cheng, S-L., et al., 1994 Endo 134:277-286; Loffler,
G., et al., 1987 Klin. Wochenschr. 65:812-817; Hauner, H., et al.,
1987 J. Clin. Endocrinol. Metabol. 64:832-835). Following
induction, differentiation was assessed using histology and
immunohistochemistry (Table 2). Commercially available MSCs and
lineage-committed progenitor cells served as positive controls
while the stem cells maintained in Control Medium and HFF cells
were examined as negative controls.
[0188] Pre-adipocytes and MSCs treated with adipogenic induction
medium, containing cAMP agonists and induction agents such as
isobutyl-methylxanthine (IBMX), indomethacin, insulin and
dexamethasone, develop lipid-containing droplets that accumulate
the lipid dye Oil Red-O (Pittenger, M F., et al., 1999 Science
284:143-147; Rubin, C S, et al., 1978 J. Biol. Chem. 253:7570-7578;
Deslex, S, et al., 1987 Int. J. Obesity 11:19-27). To determine if
PLA cells undergo adipogenesis, cells were cultured in medium
containing these agents (Adipogenic Medium, AM) and stained with
Oil Red-O. The stem cells cultured in AM were reproducibly induced
toward the adipogenic lineage as early as two weeks post-induction
(FIG. 4). A significant fraction of the cells contained multiple,
intracellular lipid-filled droplets that accumulated Oil Red-O. The
Oil Red O-containing stem cells exhibited an expanded morphology
with the majority of the intracellular volume (90-98%) occupied by
lipid droplets, consistent with the phenotype of mature adipocytes.
The mean level of adipogenic differentiation measured in 6 donors
under 35 years of age was 42.4%.+-.10.6% (% Oil Red 0-positive
cells /total PLA cell number). Prolonged culture times (i.e. 4
weeks) resulted in the detachment of differentiating cells from the
culture plate arid flotation to the surface. The observed
morphology and lipid accumulation of differentiated stem cells were
similar to that observed upon treatment of bone marrow-derived MSCs
and the pre-adipocyte cell line, 3T3-L1, in AM. No lipid droplets
were observed in undifferentiated stem cells or in HFF negative
controls. In contrast to MSCs, in which adipogenic differentiation
dramatically decreases beyond the third culture passage (Conget, P
A and J J Minguell 1999 J. Cell. Physiol. 181:67-73), the
adipogenic potential of the stem cells was maintained over
long-term culture (i.e. passage 15=175 days culture). Taken
together, the results indicate that the stem cells undergo
adipogenic differentiation.
[0189] Differentiation of osteoprogenitor cells and marrow-derived
MSCs into osteoblasts is induced in vitro by treating cells with
low concentrations of ascorbic acid, .beta.-glycerophosphate and
dexamethasone (Pittenger, M F, et al., 1999 Science 284:143-147;
Cheng, S-L, et al., 1994 Endo 134:277-286; Conget, P A and J J
Minguell 1999 J. Cell. Physiol. 181:67-73). Early differentiation
of these cells into immature osteoblasts is characterized by
Alkaline Phosphatase (AP) enzyme activity with human MSCs
expressing AP as early as 4 days and maximum levels observed at 12
days post-induction (Jaiswal, N, et al., 1997 J. Cell Biochem.
64:295-312). To confirm their osteogenic capacity, the stem cells
were treated with osteogenic medium (OM) for 14 days and the
expression of AP was examined. The stem cells cultured in OM formed
an extensive network of dense, multi-layered nodules that stained
positively for AP (FIG. 5). The mean number of AP-positive staining
cells measured in 6 donors was 50.2%.+-.10.8% of total stem cell
number. Expression of AP was also observed in both MSCs and NHOst
positive controls maintained in OM. In contrast, undifferentiated
stem cells and HFF negative controls did not show evidence of AP
expression. While AP expression is dramatically upregulated in
osteogenic tissues, its expression has been observed in several
non-osteogenic cell types and tissues such as cartilage, liver and
kidney (Henthom, P S, et al., 1988 J. Biol. Chem. 263:12011; Weiss,
M J, et al., 1988 J. Biol. Chem. 263:12002; Leboy, P S, et al.,
1989 J. Biol. Chem. 264:17281). Therefore, AP expression is
frequently used, in conjunction with other osteogenic specific
markers, as an indicator of osteogenesis. One such indicator is the
formation of a calcified extracellular matrix (ECM). Mature
osteoblasts secrete a collagen I-rich ECM that becomes calcified
during the later stages of differentiation (Scott, D M 1980 Arch.
Biochem. Biophys. 201:384-391). Therefore, in order to confirm
osteogenic differentiation, calcification of the ECM matrix was
assessed in the stem cells using a von Kossa stain. Calcification
appears as black regions within the cell monolayer. Consistent with
osteogenesis, several black regions, indicative of a calcified ECM,
were observed in the stem cells treated for 4 weeks in OM.
Calcification was also identified in MSC and NHOst positive
controls, while no calcification was observed in undifferentiated
stem cells or HFF cells. The osteogenic potential of the stem cells
was maintained over long-term culture, with cells expressing AP as
late as 175 days of culture. Taken together, the expression of AP
by the adipose-derived stem cells and the production of a calcified
ECM strongly suggests that these adipose-derived cells can be
induced toward the osteogenic lineage.
[0190] Chondrogenic differentiation can be induced in vitro using a
micromass culture technique, in which cellular condensation (a
critical first event of chondrogenesis) is duplicated (Ahrens P.
B., et al., 1977 Develop. Biol. 60: 69-82; Reddi A. H. 1982 Prog.
Clin. Biol. Res. 110 Pt B: 261-268; Denker, A. E., et al., 1995
Differentiation 59, 25-34; Tacchetti, C, et al., 1992 Exp Cell Res.
200:26-33). Enhanced differentiation can be obtained by treating
cells with dexamethasone and TGF.beta.1 (Iwasaki, M. et al., 1993
Endocrinology 132:1603-1608). Marrow-derived MSCs, cultured with
these agents under micromass conditions, form cell nodules
associated with a well-organized ECM rich in collagen II and
sulfated proteoglycans (Pittenger, M F, et al., 1999 Science
284:143-147; Mackay, A M, et al., 1998 Tissue Eng. 4:415-428).
These sulfated proteoglycans can be specifically detected using the
stain Alcian Blue under acidic conditions (Lev, R and S. Spicer
1964 J. Histochem. Cytochem. 12:309-312). To assess the
chondrogenic capacity of the stem cells, the cells were cultured
via micromass in Chondrogenic Medium (CM), containing dexamethasone
and TGF.beta.1. Micromass culture of the stem cells resulted in the
formation of dense nodules consistent with chondrogenic
differentiation. The stem cell nodules were associated with an
Alcian Blue-positive ECM, indicative of the presence of sulfated
proteoglycans within the matrix (FIG. 6). Cartilaginous nodules
were also observed upon micromass culture of MSC controls. To
confirm the specificity of Alcian Blue for cartilaginous matrices,
human cartilage and bone sections were stained with Alcian Blue
under acidic conditions. As expected, human cartilage sections
stained positively with Alcian Blue, while no staining was observed
in bone sections. In addition to the presence of sulfated
proteoglycans within the ECM, both stem cells and human cartilage
sections expressed the cartilage-specific collagen type II isoform,
while no staining was observed in undifferentiated stem cells.
Consistent with adipogenic and osteogenic differentiation, the stem
cells retained their chondrogenic differentiation potential after
extended culture periods (i.e. up to 175 days). The above results
suggest that the adipose-derived stem cells possess the capacity to
differentiate toward the chondrogenic lineage.
[0191] Myogenesis is characterized by a period of myoblast
proliferation, followed by the expression of muscle-specific
proteins and fusion to form multinucleated myotubules. Early
myogenic differentiation is characterized by the expression of
several myogenic regulatory factors including Myogenic
Determination factor1 (MyoD1; (Davis, R. L., et al., 1987 Cell
51:987-1000; Weintraub, H., et al., 1991 Science 251:761-763; Dias,
P., et al., 1994 Semin. Diagn. Pathol. 11:3-14). Terminally
differentiated myoblasts can be characterized by the expression of
myosin and the presence of multiple nuclei (Silberstein, L., et
al., 1986 Cell 46:1075-1081). Proliferation and myogenic
differentiation of muscle precursors and bone marrow-derived stem
cells can be induced by dexamethasone and results in the expression
of muscle-specific proteins (Grigoradis, A, et al., 1988 J. Cell
Biol. 106:2139-2151; Ball, E H and B D Sanwal 1980 J. Cell. Physiol
102:27-36; Guerriero, V and J R Florini 1980 Endocrinology
106:1198-1202). Furthermore, addition of hydrocortisone is known to
stimulate human myoblast proliferation, prior to their transition
into differentiated myotubules (Zalin, R J 1987 Exp. Cell Res.
172:265-281). To examine if the stem cells undergo myogenesis, the
cells were cultured for 6 weeks in the presence of dexamethasone
and hydrocortisone, and incubated with antibodies specific to MyoD1
and myosin (heavy chain). Consistent with early myogenic
differentiation, treatment of the stem cells with MM for 1 week
induced the expression of MyoD1 (FIG. 7). The stem cells treated
for longer time periods (6 weeks) stained positively for myosin. In
addition to myosin expression, the presence of discrete `patches`
of large, elongated cells with multiple nuclei were also observed,
suggesting that the stem cells underwent myoblast fusion (PLA
panel, inset). MyoD1 and myosin heavy chain expression were also
detected in human skeletal muscle positive control cells. Using
Myogenic Medium, myogenic differentiation was not observed in MSC
controls even at 6 weeks of induction. These cells may be adversely
affected by hydrocortisone and may require alternate conditions to
induce differentiation. Myogenic differentiation levels in the stem
cells averaged 12%. Multi-nucleation, myosin heavy chain and MyoD1
expression were not observed in undifferentiated stem cells nor in
HFF negative controls. The presence of multi-nucleated cells and
the expression of both MyoD1 and myosin heavy chain suggests that
the adipose-derived stem cells have the capacity to undergo
myogenic differentiation.
2TABLE 1 Lineage-specific differentiation induced by media
supplementation Medium Media Serum Supplementation Control DMEM 10%
FBS none Adipogenic DMEM 10% FBS 0.5 mM isobutyl-methylxanthine
(AM) (IBMX), 1 .mu.M dexamethasone, 10 .mu.M insulin, 200 .mu.M
indomethacin, 1% antibiotic/ antimycotic Osteogenic DMEM 10% FBS
0.1 .mu.M dexamethasone, 50 .mu.M (OM) ascorbate-2-phosphate, 10 mM
.beta.-glycerophosphate, 1% anti- biotic/antimycotic Chondrogenic
DMEM 1% FBS 6.25 .mu.g/ml insulin, 10 ng/ml (CM) TGF.beta.1, 50 nM
ascorbate-2- phosphate, 1% antibiotic/anti- mycotic Myogenic DMEM
10% FBS, 0.1 .mu.M dexamethasone, 50 .mu.M (MM) 5% HS
hydrocortisone, 1% antibiotic/anti- mycotic
[0192]
3TABLE 2 Differentiation markers and assays of lineage-specific
differentiation Lineage-specific Histologic/immunohisto- Lineage
determinant chemical assay Adipogenic Lipid accumulation Oil Red O
stain Osteogenic 1. Alkaline phosphatase 1. Alkaline Phosphatase
activity stain 2. Calcified matrix 2. Von Kossa stain production
Chondrogenic 1. Sulfated proteoglycan- 1. Alcian Blue (pH 1.0) rich
matrix stain 2. Collagen II synthesis 2. Collagen II-specific
monoclonal antibody Myogenic 1. Multi-nucleation 1. Phase contrast
2. Skeletal muscle microscopy myosin heavy 2. Myosin & MyoD1
chain & MyoD1 specific monoclonal expression antibodies
Discussion
[0193] Conceptually, there are two general types of stem cells:
Embryonic Stem Cells (ESCs) and autologous stem cells. Although
theoretically appealing because of their pluripotentiality, the
practical use of ESCs is limited due to potential problems of cell
regulation and ethical considerations. In contrast, autologous stem
cells, by their nature, are immunocompatible and have no ethical
issues related to their use. For the engineering of mesodermally
derived tissues, autologous stem cells obtained from bone marrow
have proven experimentally promising. Human bone marrow is derived
from the embryonic mesoderm and is comprised of a population of
Hematopoeitic Stem Cells (HSCs), supported by a mesenchymal stroma
(Friedenstein A. J., et al., 1968 Transplantation 6: 230-47;
Friedenstein A. J., et al., 1974 Transplantation 17: 331-40; Werts
E. D., et al., 1980 Exp. Hematol. 8: 423; Dexter T. M. 1982 J. Cell
Physiol. 1: 87-94; Paul S. R., et al., 1991 Blood 77: 1723-33).
While the proliferation and differentiation of HSCs has been well
documented, less is known about the stromal component. The bone
marrow stroma, in both animals and humans, is heterogenous in
composition, containing several cell populations, including a stem
cell population termed Mesenchymal Stem Cells or MSCs (Caplan A I
1991 J. Orthop. Res. 9:641-650). Studies on MSCs have demonstrated
their differentiation into adipocytes (Beresford J. N., et al.,
1992 J. Cell Sci. 102; 341-351; Pittenger M. F., et al., 1999
Science 284: 143-147), chondrocytes (Caplan A. I. 1991 J. Orthop.
Res. 9: 641-50; Pittenger M. F., et al., 1999 Science 284: 143-147;
Berry L., et al., 1992 J. Cell Sci. 101: 333-342; Johnstone B., et
al., 1998 Exp. Cell Res. 238: 265-272; Yoo J. U., et al., 1998 J.
Bone Joint Surg. Am. 80: 1745-1757), myoblasts (Wakitani S., et
al., 1995 Muscle Nerve 18: 1417-1426; Ferrari G., et al., 1998
Science 279: 1528-1530) and osteoblasts (Caplan A. I. 1991 J.
Orthop. Res. 9: 641-50; Pittenger M. F., et al., 1999 Science 284:
143-147; Grigoradis A., et al., 1988 J. Cell Biol. 106: 2139-2151;
Cheng S-L., et al., 1994 Endo 134: 277-286, 1994; Haynesworth S.
E., et al., 1992 Bone 13: 81-8; Rickard D. J., et al., 1996 J. Bone
Min. Res. 11: 312-324; Prockop D. J. 1997 Science 276: 71-74;
Dennis J. E., et al., 1999 J. Bone Miner. Res. 14: 700-709). These
cells represent a promising option for future tissue engineering
strategies. However, traditional bone marrow procurement procedures
may be painful, frequently requiring general or spinal anesthesia
and may yield low numbers of MSCs upon processing (approximately 1
MSC per 10.sup.5 adherent stromal cells (Pittenger, M F et al.,
1999 Science 284:143-147; Rickard, D J, et al., J. Bone Min. Res.
11:312-324: Bruder, S P, et al., 1997 J. Cell. Biochem.
64:278-294)). From a practical standpoint, low stem cell numbers
necessitate an ex vivo expansion step in order to obtain clinically
significant cell numbers. Such a step is time consuming, expensive
and risks cell contamination and loss. An ideal source of
autologous stem cells would, therefore, be both easy to obtain,
result in minimal patient discomfort yet be capable of yielding
cell numbers substantial enough to obviate extensive expansion in
culture.
[0194] We report that a cellular fraction with multiple mesodermal
lineage capabilities can be processed from human lipoaspirates.
This cellular fraction is the adipose-derived stem cells which is
designated a Processed Lipoaspirate (PLA), comprising
fibroblast-like cells that can be expanded easily in vitro without
the need for specific sera lots or media supplementation. The stem
cell samples maintained a linear growth rate with no appreciable
senescence over extended culture periods. The stem cell population
was heterogenous in nature, with the majority of the cells being
mesenchymal in origin. However, contaminating endothelial, smooth
muscle and pericyte cell populations were identified. The stem
cells also exhibited multi-lineage potential in vitro,
differentiating toward the adipogenic, osteogenic, chondrogenic and
myogenic lineages when cultured in the presence of established
lineage-specific differentiation factors. The differentiation
results were consistent with those observed upon lineage-specific
differentiation of bone marrow-derived MSCs and lineage-committed
precursors.
[0195] While the apparent multi-differentiative capacity of the
stem cells suggests the presence of a stem cell population within
human liposuctioned adipose tissue, it is not conclusive.
Multi-lineage differentiation may also be due to the presence of:
(1) multiple lineage-committed progenitor cells; (2) multi-potent
cells from other sources (e.g. pericytes, marrow-derived MSCs from
peripheral blood); or (3) a combination of the above.
[0196] The observed differentiation may be due to the presence of
lineage-committed progenitor cells, such as pre-osteoblasts,
pre-myoblasts or pre-adipocytes within the stem cell fraction.
Cellular fractions (i.e. SVFS) obtained from excised adipose tissue
are known to contain pre-adipocytes that differentiate into mature
adipocytes (Pettersson, P, et al., 1984 Acta Med. Scand.
215:447-453; Pettersson, P, et al., 1985 Metabolism 34:808-812). It
is possible that the observed adipogenic differentiation by the
stem cells is simply the commitment of existing pre-adipocytes and
not the differentiation of a multi-potent cell. However, we do not
believe this to be the case. As little as 0.02% of the SVF obtained
from excised adipose tissue have been identified as pre-adipocytes
capable of adipogenic differentiation (Pettersson, P, et al., 1984
Acta Med. Scand. 215:447-453). If pre-adipocyte numbers within the
stem cell fraction are comparable to those levels measured in the
SVF from excised tissue, one would expect a relatively low level of
adipogenesis. However, the degree of adipogenesis observed in the
stem cells is significant (approximately 40% of the total PLA cell
number) and may result from the differentiation of additional cell
types.
[0197] Damage to the underlying muscle during liposuction may
introduce myogenic precursor cells or satellite cells into the stem
cell fraction, resulting in the observed myogenic differentiation
by these cells. Located between the sarcolemma and the external
lamina of the muscle fiber, myogenic precursor cells in their
undifferentiated state are quiescent and exhibit no distinguishing
features, making their identification difficult. Several groups
have attempted to identify unique markers for these precursors with
limited success. Currently, the expression of the myogenic
regulatory factors, MyoD1 and myogenin have been used to identify
satellite cells during embryogenesis and in regenerating adult
muscle in rodents (Cusella-DeAngelis, M. C., et al., 1992 Cell
Biol. 116:1243-1255; Grounds, M. D., et al., 1992 Cell Tiss. Res.
267:99-104; Sassoon, D. A. 1993 Develop. Biol. 156:11; Maley, M. A.
L., et al., 1994 Exp. Cell Res. 211:99-107; Lawson-Smith, M. J. and
McGeachie, J. K. 1998 J. Anat. 192:161-171). In addition, MyoD1
expression has been identified in proliferating myoblasts prior to
the onset of differentiation (Weintraub, H, et al., Science
251:761-763). While these markers have not been used to identify
myogenic precursors in human subjects, MyoD1 is expressed during
early myogenic differentiation in normal skeletal muscle and has
been used to identify the skeletal muscle origin of rhabdosarcomas
in humans 77-79 (Dias, P., et al., 1990 Am. J. Pathol.
137:1283-1291; Rosai, J., et al., 1991 Am. J. Surg. Pathol. 15:974;
Nakano, H., et al., 2000 Oncology 58:319-323). The absence of MyoD1
expression in the stem cells maintained in non-inductive Control
Medium (see FIG. 28), suggests that our observed myogenic
differentiation is not due to the presence of myogenic precursors
or proliferating myoblasts within the stem cell fraction.
Consistent with this, the blunt contour of the liposuction cannula
would make it extremely difficult to penetrate the fibrous fascial
cavity surrounding the muscle and introduce these precursors into
the adipose compartment.
[0198] Human adipose tissue is vascularized and, as such, contains
potential systemic vascular `conduits` for contamination by
multi-potent cells, such as pericytes and marrow-derived MSCs.
Disruption of the blood supply during liposuction may result in the
release of pericytes, known to possess multi-lineage potential both
in vivo and in vitro (Schor, A M, et al., 1990 J. Cell Sci.
97:449-461; Doherty, M J 1998 J. Bone Miner. Res. 13:828-838;
Diefenderfer, D L and C T Brighton 2000 Biochem. Biophys. Res.
Commun. 269:172-178). Consistent with this, our immunofluorescent
and flow cytometry data show that a small fraction of the stem
cells is comprised of cells that express smooth muscle actin, a
component of transitional pericytes and pericytes committed to the
smooth muscle lineage (Nehls, A. and D Drenckhahn 1991 J. Cell
Biol. 113:147-154). The multi-lineage differentiation observed in
the stem cells may be, in part, due to the presence of pericytes.
Disruption of the blood supply may also introduce MSCs into the
stem cell fraction. However, the literature is conflicted as to the
presence of these stem cells in the peripheral blood Huss, R 2000
Stem Cells 18:1-9; Lazaras, H M, et al., 1997 J. Hematother.
6:447-455). If the peripheral blood does indeed represent a source
of MSCs, our observed multi-lineage differentiation may be due to
the contamination of adipose tissue by these stem cells (MSCs).
However, MSCs are a small constituent of the bone marrow stroma in
humans (approximately 1 MSC per 10.sup.5 adherent stromal cells
(Pittenger, M F, et al., 1999 Science 284:143-147; Rickard, D J
1996 J. Bone Min. Res. 11:312-324; Bruder, S P, et al., 1997 J.
Cell. Biochem. 64:278-294). If these cells do exist in peripheral
blood, they are likely to be in even smaller quantities than in the
bone marrow and contamination levels of the adipose-derived stem
cells fraction by these cells may be negligible.
[0199] While these arguments may provide support for the presence
of a multi-potent stem cell population within liposuctioned adipose
tissue, definitive confirmation requires the isolation and
characterization of multiple clones derived from a single cell.
Preliminary data confirms that clonal stem cell populations possess
multi-lineage potential, capable of adipogenic, osteogenic, and
chondrogenic differentiation.
[0200] Current research has demonstrated positive results using
bone marrow-derived MSCs. MSCs can differentiate into osteogenic
and chondrogenic tissues in vivo (Benayahu, D. et al., 1989 J. Cell
Physiol 140:1-7; Ohgushi, H M 1990 Acta Orthop. Scand. 61:431-434;
Krebsbach, P H, et al., 1997 Transplantation 63:1059-1069; Bruder,
S P, et al., 1998 J. Orthop. Res. 16:155-162) and preliminary data
suggests that these cells can be used to repair bony and
cartilagenous defects (Wakitani, S., et al., 1995 Muscle Nerve
18:1417-1426; Krebsbach, P H, et al., 1997 Transplantation
63:1059-1069; Bruder, S P, et al., 1998 J. Orthop. Res. 16:155-162;
Bruder, et al., 1998 Clin. Orthop. S247-56; Krebsbach, P H 1998
Transplantation 66:1272-1278; Johnstone and Yoo 1999 Clin. Orthop.
S156-162). The stem cells obtained from liposuctioned adipose
tissue may represent another source of multi-lineage mesodermal
stem cells. Like the bone marrow stroma, these data suggests that
adipose tissue may contain a significant fraction of cells with
multi-lineage capacity. These adipose-derived stem cells may be
readily available in large quantities with minimal morbidity and
discomfort associated with their harvest.
EXAMPLE 8
[0201] The following description provides adipose-derived stem
cells which differentiate into osteogenic tissue, and methods for
isolating said stem cells. The osteogenic potential of the stem
cells decreases with the age of the donor. However, adipogenesis is
not affected by age of the donor.
Materials and Methods
Lipoaspirate Collection and Processing
[0202] Human adipose tissue was obtained from elective liposuction
procedures under local anesthesia according to patient consent
protocol HSPC #98-08 011-02 (University of California Los Angeles).
The raw lipoaspirate was processed to obtain the adipose-derived
stem cells population (Zuk, P, et al., 2001 Tissue Engineering
7:209-226). Briefly, raw lipoaspirates were washed extensively with
equal volumes of Phospho-Buffered Saline (PBS) and the
extracellular matrix (ECM) was digested at 37.degree. C. for 30
minutes with 0.075% collagenase. Enzyme activity was neutralized
with Dulbecco's Modified Eagle's Medium (DMEM; Life Technologies),
containing 10% Fetal Bovine Serum (FBS; HyClone) and centrifuged at
1200.times.g for 10 minutes. The stem cell pellet was resuspended
in DMEM/10% FBS and filtered through a cell strainer to remove any
remaining tissue. The cells were incubated overnight at 37.degree.
C., 5% CO.sub.2 in non-inductive control medium (DMEM, 10% FBS, 1%
antibiotic/antimycotic solution). Following incubation, the plates
were washed extensively with PBS to remove residual non-adherent
red blood cells. The stem cells were maintained at 37.degree. C.,
5% CO.sub.2 in control medium (Table 3). To prevent spontaneous
differentiation, cultures were maintained at sub-confluent
levels.
4TABLE 3 Lineage-Specific Differentiation Induced By Media
Supplementation Medium Media Serum Supplementation Control DMEM 10%
FBS none Adipogenic DMEM 10% FBS 0.5 mM isobutyl-methylxanthine
(IBMX), 1 .mu.M dexamethasone, 10 .mu.M insulin, 200 .mu.M
indomethacin, 1% antibiotic/antimycotic Osteogenic DMEM 10% FBS 0.1
.mu.M dexamethasone, 50 .mu.M ascorbate-2-phosphate, 10 mM
.beta.-glycerophosphate, 1% antibiotic/ antimycotic
Induction and Analysis of Differentiation
[0203] 1. Adipogenic Differentiation: PLA cells (passage 1) were
seeded into six well plates (Costar, Cambridge, Mass.) at a density
of 4.times.10.sup.4 cells per well and cultured in control medium
for 72 hours. To induce adipogenic differentiation, PLA cells were
cultured for 2 weeks in Adipogenic Medium (Table 3). PLA cells, at
the same density, were maintained in control medium as a negative
control.
[0204] Oil Red O Staining: Adipogenesis was confirmed at two weeks
post-induction by staining with Oil Red O to identify intracellular
lipid vacuoles. Cells were fixed for 60 minutes at room temperature
in 4% formaldehyde/1% calcium and washed with 70% ethanol. The
cells were incubated in 2% (w/v) Oil Red O reagent (Sigma, St
Louis, Mo.) for 5 minutes at room temperature. Excess stain was
removed by washing with 70% ethanol, followed by several changes of
distilled water. The cells were counter-stained for 1 minute with
hematoxylin.
[0205] 2. Osteogenic Differentiation: PLA cells (passage 1) were
seeded into six well plates at a density of 1.times.10.sup.4 cells
per well and cultured for 72 hours in control medium. Based on
previous studies on bone marrow-derived Mesenchymal stem cells
(Pittenger, M F 1999 Science 284:143-147), PLA cells were
maintained for a minimum of two weeks in Osteogenic Medium (Table
3) to induce osteogenesis. PLA cells were maintained in control
medium as a negative control.
[0206] Alkaline Phosphatase Staining: Alkaline phosphatase (AP)
activity was examined at 14 days post-induction. PLA cells were
rinsed with PBS and incubated for 30 minutes at 37.degree. C. in
0.05M Tris-HCl (pH 9) containing 1% (v/v) of a 50 mg/ml solution of
naphthol AS-Biphosphate (Sigma) dissolved in dimethyl sulfoxide
(DMSO) and 1 mg/ml Fast Red TR salt (Sigma). Following incubation,
cells were fixed for 10 minutes with an equal volume of 8%
paraformaldehyde, followed by a rinse with distilled water.
[0207] Von Kossa Staining: Extracellular matrix calcification was
detected at four weeks post-induction by von Kossa staining. PLA
cells were fixed with 4% paraformaldehyde at room temperature for 1
hour, followed by a 30 minute incubation with a 5% (w/v) silver
nitrate solution (Sigma) in the absence of light. The cells were
washed several times with distilled water, developed under UV light
for 60 minutes and counter-stained with 0.1% cosin in ethanol.
Matrix calcification was identified by the presence of black
extracellular deposits.
Quantitation of Differentiation
[0208] Adipogenic and osteogenic differentiation levels in each
donor were quantified using a Zeiss Axioscope 2 microscope fitted
with a Spot 2 digital camera and a 2.times. objective
(magnification 200.times.). The total number of Oil Red O- and
AP-positive cells (adipogenesis and osteogenesis, respectively) in
duplicate samples from each donor were counted within three
consistent regions from each well (e.g., at positions 3, 6 and 9
o'clock). The total number of positive-staining cells was expressed
as a percentage of total PLA cell number counted within each
region. Values from the three regions were averaged to give the
mean differentiation level for each donor. The mean level of
differentiation was expressed with respect to patient age. Von
Kossa identifies regions of matrix production rather than
individual differentiated cells, therefore this staining procedure
was used to confirm osteogenic differentiation.
Quantitation of Osteogenic Precursors within PLA Samples
[0209] In order to estimate the number of osteogenic precursors
within the PLA population, cells with osteogenic activity were
counted and related to patient age. Specifically, two age groups
were examined: Group A=20 to 39 years and Group B=40 to 60 years.
First-passage PLA cells (P1) were seeded onto 100 mm dishes,
induced toward the osteogenic lineage as described above and
stained for AP activity to confirm differentiation. Precursor
number within each PLA sample was determined by counting the number
of AP-positive colony forming units (CFU/AP.sup.+). Based on a
previous study, a minimum of ten AP-positive cells was used to
identify a CFU/AP.sup.+ (Long, M, et al., 1999 J. Gerontol. A.
Biol. Sci. Med. Soc. 54:B54-62). The average number of CFU/AP.sup.+
was determined and expressed with respect to age group. The optimal
number of PLA cells required for osteogenic differentiation was
determined empirically (1.times.10.sup.4, 5.times.10.sup.4,
1.times.10.sup.5 and 5.times.10.sup.5 cells plated per dish). While
osteogenic differentiation levels were greatest at 5.times.10.sup.5
cells per dish, confluency levels prevented accurate colony
counting. Data was therefore obtained using a density of
1.times.10.sup.5 cells per dish.
Growth Kinetics
[0210] To measure PLA cell growth kinetics (population doubling)
with respect to donor age, PLA cells from each donor (P1) were
seeded at a density of 1.times.10.sup.4 cells into multiple dishes.
Cell number was determined from triplicate samples 24 hours after
plating and every 48 hours until day 11. A growth curve (cell
number vs. culture time) was derived and population doubling was
calculated from the log phase.
Statistical Analysis
[0211] Significant differences in PLA cell osteogenesis and
adipogenesis according to donor age were determined by linear
regression analysis (r value). In addition, the mean levels of
differentiation across donor age were compared using an unpaired
student t-test (assuming unequal variances) and a one way analysis
of variance (ANOVA).
Results
PLA Cell Growth Kinetics Vary with Respect to Donor Age
[0212] Initial PLA cultures were relatively homogenous in
appearance, with the majority of cells (85 to 90%) exhibiting a
fibroblast-like morphology. A small fraction of endothelial cells,
macrophages and pre-adipocytes could be identified (less than 10%
of the total population). PLA cells reached 80-90% confluency
within 14 days of culture in control medium. Growth curves derived
from first-passage PLA cell cultures (P1) were characterized in
each donor by an initial lag phase (typically 48 hrs), followed by
a log phase (average=7 days) and a plateau phase. Representative
growth kinetic curves are shown in FIG. 8A. No significant
difference in the duration of the lag and log phases was observed
in any donor. Similarly, no significant differences in PLA growth
kinetics were observed in younger patients (20 years vs. 39 years).
However, a decrease in the log phase of PLA cells was observed in
older patients (eg. day 13; 58 years--12.6.times.10.sup.4 cells, 20
years--26.9.times.10.sup.4 cells). Based on the growth kinetics
data, the average PLA cell population doubling time calculated from
15 donors was 52.67.+-.8.67 hours. PLA cell population doubling
time ranged from 38 to 77 hours (FIG. 8B; 20 years vs. 53 years).
Regression analysis of population doubling and donor age yielded a
positive correlation of r=0.62 (n=15), indicating a trend toward
increasing population doubling (i.e. decreasing proliferative
potential) with age. However, statistical analysis of donors
grouped according to decade (i.e. 20-30 years, 30-40 years, 40-50
years, 50-60 years), using an unpaired t-test, did not show a
significant difference in population doubling (p>0.05),
suggesting that PLA proliferation does not significantly decrease
with increasing donor age.
Adipogenic Differentiation Potential Does Not Change with PLA
Age
[0213] Adipogenesis and lipid vacuole formation in PLA cells were
confirmed by staining cells with the lipid dye Oil Red O. In all
donors, low levels of adipogenic differentiation in PLA cells were
apparent as early as 5 days induction in Adipogenic Medium.
Differentiating cells assumed an expanded morphology consistent
with adipocytes and accumulated lipid-rich intracellular vacuoles
that stained with Oil Red O (FIG. 9A, Panels 1 and 2). By 14 days
post-induction, differentiating cells contained large, Oil Red
O-positive lipid droplets within the cytoplasm. Differentiation
levels varied from donor to donor with several donors exhibiting
low levels of adipogenesis, in which individual Oil Red O-positive
cells containing a moderate amount of stain were easily identified
(FIG. 9A, Panel 1). In addition, several donors exhibited enhanced
levels of adipogenesis, in which both the number of Oil Red
O-positive cells and the accumulation of the stain increased
dramatically (FIG. 9A, Panel 2). Cells cultured in non-inductive
control media exhibited no change in morphology and did not
accumulate Oil Red O, confirming the specificity of the inductive
medium conditions (FIGS. 9A, Panel 3). To measure changes in
adipogenic differentiation potential with respect to donor age, the
number of Oil Red O-positive cells was directly counted within a
defined region and expressed as a percentage of the total number of
PLA cells counted. Values from each region were averaged to give
the mean level of adipogenic differentiation and expressed with
respect to donor age (FIG. 9B). Adipogenic differentiation levels
ranged from 4.51% to 57.78% of the total PLA cells (n=20). An
average differentiation potential of 26.55.+-.18.14% was
calculated. However, a negligible regression value was obtained
upon analysis (r=0.016), suggesting that no significant changes in
adipogenic differentiation occur with increasing donor age.
Osteogenic Differentiation Decreases with Donor Age
[0214] To confirm osteogenesis, cells were stained for Alkaline
Phosphatase (AP) activity and extracellular matrix calcification
using a silver nitrate/Von Kossa stain. PLA cells, cultured in
Osteogenic Medium, underwent a dramatic change in cellular
morphology as early as 4 days post-induction, changing from
spindle-shaped to cuboidal, characteristic of osteoblasts. Low
levels of osteogenesis were characterized in some donors by the
formation of a monolayer of AP-positive cells (FIG. 10A, Panel 1).
Higher levels of osteogenesis were characterized in some patients
by the presence of multi-layered AP-positive nodular structures
with well-defined inter-nodular regions containing no cells (FIG.
10A, Panel 2). In addition to AP activity, regions of
mineralization, as detected by von Kossa staining, were evident
after 3 weeks of culture, further substantiating osteogenic
differentiation (FIG. 10A, Panels 4 and 5). Control PLA cells did
not exhibit AP activity or matrix mineralization (FIG. 10A, Panels
3 and 6). To measure potential changes in osteogenic
differentiation with donor age, the mean level of osteogenesis
(i.e. AP-positive cells) was determined using the same method to
calculate adipogenic levels.
[0215] In contrast to adipogenesis, a significant decrease in
osteogenesis was observed in older donors. Osteogenic
differentiation ranged from 11.64% to 64.69% of the total PLA cells
(FIG. 10B). Regression analysis of donor age and osteogenesis
yielded a significant negative correlation (r=-0.70, n=19),
suggesting that osteogenic differentiation decreases with respect
to age. A similar trend was observed using von Kossa staining.
Interestingly, a distinct decrease in osteogenic differentiation
was observed in donors older than 36 years of age (FIG. 10B, dashed
line). Consistent with this, a significant difference in
osteogenesis (p<0.001) was observed when the subjects were
divided into two age groups. Donors from the younger age group (20
to 36 years; n=7) exhibited a mean osteogenic potential of
50.7.+-.10% (total PLA cells) while a significantly lower level of
osteogenesis (20.7.+-.7.9% total PLA cells) was measured in the
older age group (37 to 58 years; n=11) (FIG. 10C). Based on this
data, cells from the younger group exhibited a 2.4-fold increase in
osteogenic potential, forming 59% more AP-positive cells.
Relative Proportion of Osteogenic Precursors Within PLA
[0216] In order to determine if the decrease in PLA osteogenesis is
due to a decrease in the number of PLA cells with osteogenic
potential, the relative proportion of osteogenic precursor cells
within the PLA was calculated with respect to donor age. PLA cells
were induced for 2 weeks in Osteogenic Medium and the number of
precursors within the PLA determined by calculating the number of
AP-positive Colony Forming Units (CFU/AP.sup.+) (Grigoradis, A, et
al., 1988 J. Cell Biol. 106:2139-2151; Pittenger, M F, et al., 1999
Science 284:143-147; Jaiswal, N., et al., 1997 J. Cell Biochem.
64:295-312). The number of precursors was calculated in two age
groups (Group A=20-39 years, n=5 and Group B=40-58 years, n=6).
Consistent with the diminished osteogenic potential observed in
older PLA samples, a slight decrease in CFU/AP.sup.+ number was
observed with increasing age. The average number of CFU/AP.sup.+ in
Group A was 194.+-.61 per 10.sup.5 PLA cells, while the number of
CFU/AP.sup.+ in Group B decreased to 136.+-.32 per 10.sup.5 PLA
cells (FIG. 11). While a decreasing trend in osteoprogenitor cells
was observed, this decrease was not statistically significant
(p=0.11), suggesting that the decrease in osteogenic potential by
PLA cells may not be directly due to a decrease in the number of
osteogenic precursors.
Discussion
[0217] Mesenchymal stem cells can be isolated from bone marrow.
Mesenchymal stem cells are a component of the bone marrow stroma
and possess the capacity to differentiate into various mesodermal
tissues including fat, bone and cartilage (Grigoradis, A., et al.,
1988 J. Cell Biol. 106:2139-2151; Caplan, A. 1. 1991 J. Orthop.
Res. 9:641-650; Beresford, J. N., et al., 1992 J. Cell Sci.
102:341-351; Berry, L., et al., 1992 J. Cell Sci. 101:333-342;
Ferrari, G., et al., 1998 Science 279:1528-1530; Johnstone, B., et
al., 1998 Exp. Cell Res. 238:265-272; Pittenger, M. F., et al.,
1999 Science 284:143-147). This multi-lineage potential may be
clinically useful for the repair of complex post-traumatic and
congenital defects. Indeed, several in vitro and in vivo studies
have suggested the clinical potential for these stem cells
(Benayahu, D., et al., 1989 J. Cell Physiol. 140:1-7; Wakitani, S.,
et al., 1995 Muscle Nerve 18:1417-1426; Krebsbach, P. H., et al.,
1997 Transplantation 63:1059-1069; Bruder, S. P., et al., 1998
Clin. Orthop. (355 Suppl):S247-56; Johnstone, B., and Yoo, J. U.
1999 Clin. Orthop. (367 Suppl):S156-62). However, bone marrow
procurement is painful, requires general anesthesia and yields low
numbers of mesenchymal stem cells upon processing (Pittenger, M.
F., et al., 1999 Science 284:143-147; Rickard, D J, et al., 1996 J.
Bone Miner. Res. 11:312-324; Bruder, S P, et al., 1997 J. Cell.
Biochem. 64:278-294), thus requiring an ex vivo expansion step
prior to clinical use. In light of these factors, an additional
source of multi-lineage stem cells may be desirable. We have
identified a population of stem cells in the stromal-vascular
fraction of liposuctioned human adipose tissue (Example 7, supra).
This cell population is designated a Processed Lipoaspirate (PLA),
and appears to be similar to bone marrow-derived mesenchymal stem
cells in many aspects. Like mesenchymal stem cells, PLA cells are
stable over long-term culture, expand easily in vitro and possess
multi-lineage potential, differentiating into adipogenic,
osteogenic, myogenic and chondrogenic cells.
[0218] PLA cells possess a fibroblast-like morphology, expand
stably in vitro, and proliferate with an average population
doubling time of 53 hours. Previous studies have shown that the
size and number of adipocytes within adipose tissue increases with
age (Hauner, H. et al., 1987 J. Clin. Endocrinol. Metabol.
64:832-835) suggesting an overall increase in adipogenesis in the
adipose stores with advancing age. In contrast to these studies, we
do not observe a significant age-related change in adipogenesis by
PLA cells, suggesting that the adipogenic potential of older PLA
cells is unaffected by advancing age. The development of adipose
tissue requires the activity of several growth factors and steroid
hormones (Hatmer, H. et al., 1987 J. Clin. Endocrinol. Metabol.
64:832-835). Therefore, the adipogenic potential of PLA cells may
be influenced by the genetic background and/or hormonal levels
within each donor. Proenza et al. has reported that adipogenesis
can be affected by alterations in the expression of several genes,
including lipoprotein lipase, adrenoreceptor and uncoupling protein
(Rickard, D J, et al., 1996 J. Bone Miner. Res. 11:312-324;
Glowacki J. 1995 Calcif. Tiss. Int. 56 (Suppl 1):S50-51). In
addition, Chen et al. has shown that the expression of specific
obesity-related genes in pre-adipocytes is related to the
differentiation of these cells into mature adipocytes (Chen, X., et
al., 1997 Biochim. Biophys. 1359:136-142). Therefore, gene
expression levels, together with hormonal activity, may differ from
donor to donor, influencing the adipogenic potential of PLA cells
and resulting in varying levels of adipogenesis, irrespective of
donor age.
[0219] In contrast to adipogenesis, a decrease in PLA osteogenic
potential (as measured by AP activity) is observed with increasing
donor age. A significant negative correlation between osteogenesis
and donor age is found by regression analysis (r=-0.70).
Furthermore, a significant difference in osteogenesis is observed
when donors are segregated into two age groups (20 to 36 years and
37 to 58 years), with cells from the younger age group possessing
over a two-fold greater osteogenic potential.
[0220] Osteogenesis is defined by three phases: the proliferation
of osteogenic precursors, maturation of these precursors into
osteoblasts (accompanied by matrix deposition) and a mineralization
phase. Each phase is essential and can dramatically affect the
development of mature bone. The decrease in osteogenesis observed
in older donors may be due to three possibilities: 1) a decrease in
PLA cell proliferation, 2) a decrease in the number of PLA-derived
osteogenic precursors themselves or 3) a decrease in osteogenic
differentiation capacity. As shown in FIG. 8, PLA population
doubling time increases slightly in older donors suggesting that
the proliferative capacity of older PLA cells diminishes with age.
However, this increase in population doubling time is not
statistically significant and is not likely to contribute to the
age-dependent decrease in osteogenic potential.
[0221] In order to determine if a decrease in the number of
osteogenic precursors within the PLA contributed to our results,
the average number of CFU/AP.sup.+ colonies was determined.
Colonies with AP activity are considered to be osteoprogenitors and
have been previously used to determine the number of osteogenic
precursors and/or stem cells in bone marrow (Owen, T A, et al.,
1990 J. Cell Physiol. 143:420-430). While animal studies indicate a
decrease in the number of osteoprogenitors in bone marrow with
advancing age (Bergman R J, et al., 1996 J. Bone Miner. Res.
11:568-577; Huibregtse, B A, et al., 2000 J. Orthop. Res.
18:18-24)), conflicting results have been reported for human
samples. Work by Glowacki and Rickard et al. indicate no
age-related changes in bone marrow osteoprogenitor cells (Rickard,
D J, et al., 1996 J. Bone Miner. Res. 11:312-324; Glowacki, J. 1995
Calcif. Tiss. Int. 56(Suppl 1):S50-51)). In support of these
studies, we find a small, but statistically insignificant, change
in the number of CFU/AP.sup.+ colonies with age. This suggests that
the observed age-dependent decrease in osteogenic potential may not
be due to a drop in the number of osteogenic precursors and/or stem
cells within the PLA.
[0222] The decrease in PLA osteogenesis may be due to the loss of
osteogenic capacity. Several factors may influence the osteogenic
capacity of stem cells, including: 1.) cell-cell and cell-matrix
interactions; and 2.) growth factors and hormones. A recent study
by Becerra et al. demonstrates a significant decrease in the
osteogenic response of older Mesenchymal stem cells to
demineralized bone matrix in rats (Becerra, J., et al., 1996 J.
Bone Miner. Res. 11:1703-1714), suggesting age-related alterations
in MSC-matrix interactions. Similarly, decreases in osteogenic
potential in older donors have been correlated to the degradation
of the extracellular matrix (Bailey, A J, et al., 1999 Calcif.
Tiss. Int. 65:203-210). The microenvironment surrounding PLA cells
may change with increasing age, altering cell-cell and
cell-extracellular matrix interactions that could inhibit
osteogenic differentiation of PLA cells or favor their
differentiation to another lineage (e.g adipogenic). Furthermore,
the diminishment of osteogenic potential in PLA cells may be due to
gender. All donors in this study were female. It is well documented
that aging in the female is accompanied by the loss of estrogen,
coupled to a decrease in skeletal bone mass (Parfitt, A M 1990 in
Bone, ed B K Hall, Vol. 1, 351-431, New Jersey: Caldwell; Hahn, T J
1993 in Textbook of Rheumatology, ed W N Kelly, 1593-1627, New
York: Saunders). Since osteocytes do not replicate, bone remodeling
and repair requires a continuous supply of osteoblasts, the
principal source of which is the bone marrow stroma. Estrogen is
known to regulate the differentiation of bone marrow-derived stem
cells and decreases in circulating estrogen levels can be linked to
a loss of stem cell osteogenic potential (Robinson, J A, et al.,
1997 Endocrinology 138:2919-2927; Ankrom, M A, et al., 1998
Biochem. J. 333:787-794). Like bone marrow stem cells, the loss of
osteogenic capacity by PLA cells in older female donors may simply
reflect the changes that are associated with estrogen loss. A
possibility is that the decrease in PLA osteogenic potential may be
due to relatively small changes in all three factors discussed
above, reflecting a general phenomenon observed in aging women.
[0223] A reduction in osteoblast number and bone-forming activity,
coupled to an increase in marrow cavity adipogenesis, contributes
to type II or age-related, osteoporosis (Parfitt, A M 1990 in Bone,
ed B K Hall, Vol. 1, 351-431, New Jersey: Caldwell; Hahn, T J 1993
in Textbook of Rheumatology, ed W N Kelly, 1593-1627, New York:
Saunders). While current research is focusing on the role of bone
marrow-derived Mesenchymal stem cells in osteoporosis, the
age-related loss of osteogenic capacity by adipose-derived PLA
cells may provide researchers with an alternate model system for
the study of this disease. Furthermore, PLA cells may represent
another viable cell-based therapeutic paradigm for the treatment of
osteoporosis and other metabolic bone disorders.
[0224] The use of stem cells for tissue engineering applications
may be dramatically influenced by stem cell number, growth kinetics
and differentiation potential. Each of these factors, in turn, may
be affected by the age of the donor. Several studies on bone
marrow-derived mesenchymal stem cells have reported alterations in
MSC number, population doubling and differentiation potential with
respect to donor age in both animal and human models (Lansdorp, P.
M., et al., 1994 Blood Cells 20:376-380; Becerra, J., et al., 1996
J. Bone Miner. Res. 11:1703-1714; Bergman, R. J., et al., 1996 J.
Bone Miner. Res. 11:568-77; Gazit, D., et al., 1998 J. Cell
Biochem. 70:478-88; Oreffo, R. O., et al., 1998 Clin. Sci (Colch.)
94:549-555; D-Ippoliot, G., et al., 1999 J. Bone Miner. Res.
14:1115-1122; Long, M. W., et al., 1999 J. Gerontol. A. Biol. Sci.
Med. Soc. 54:B54-62; Huibregtse, B. A., et al., 2000 J. Orthop.
Res. 18:18-24). We have characterized several PLA populations by
determining population doubling, differentiation potential and
average colony forming unit number with respect to donor age.
EXAMPLE 9
[0225] The following description provides adipose-derived stem
cells which differentiated into chondrogenic tissue, and method for
isolating said stem cells.
Materials and Methods
Reagents and Antibodies
[0226] Sodium acetate, bovine serum albumin (BSA), N-ethylmaleimide
(NEM), 6-aminocaproic acid, phenylmethyl-sulfonyl fluoride (PMSF),
and benzamidine hydrochloride were all purchased from Sigma (St.
Louis, Mo.). Monoclonal antibodies to type II collagen (clone
II-4C11), chondroitin-4-sulfate, and keratan sulfate (clone 5-D-4)
were purchased from ICN Biomedical (Aurora, Ohio).
Lipoaspirate Processing
[0227] Human liposuction aspirates were obtained from ten healthy
elective cosmetic surgery patients ranging in age from 20-55 years,
and processed to obtain the processed lipoaspirate (PLA) cell
populations. All procedures were approved by the Human Subject
Protection Committee (HSPC) under protocol number HSPC
#98-08-011-02. Raw lipoaspirates were processed based on the method
described in Example 7, supra. Briefly, the lipoaspirates were
washed extensively in phosphate-buffered saline (PBS) and then
incubated with 0.075% collagenase (Sigma, St. Louis, Mo.) at
37.degree. C. for thirty minutes with gentle agitation. The
collagenase was neutralized by adding an equal volume of Dulbecco's
Modified Eagle Medium (DMEM, Cellgro, Herndon, Va.), and FBS, and
the cellular suspension was centrifuged at 260 g for five minutes.
The resultant cell pellet was resuspended in 1% erythrocyte lysis
buffer (0.16 M NH.sub.4Cl) to lyse the contaminating reb blood
cells. The cell suspension was centrifuged at 260 g for five
minutes to isolate the PLA fraction. The PLA pellet was resuspended
in control medium (DMEM, 10% FBS, and 1% antibiotics-antimycotics)
and maintained at subconfluent concentrations at 37.degree. C. with
5% CO.sub.2. Human foreskin fibroblasts (HFFs) were similarly
harvested through enzymatic digestion with collagenase and
maintained at subconfluent levels in control medium.
Chondrogenic Differentiation
[0228] After culture expansion to three passages (P3), the PLA
cells were trypsinized and resuspended in control medium at a
concentration of 10.sup.7 cells/ml. Chondrogenic differentiation
was induced using a micromass culture protocol as previously
described with some modifications (Ahrens, P B, et al., 1977 Dev.
Biol. 60:69-82; Denker, A E 1995 Differentiation 59:25-34). Ten
microliter drops of the PLA cellular suspension were placed in the
center of each well of a 24-well tissue culture plate and on
chamber slides. The cells were placed in an incubator at 37.degree.
C. at 5% CO.sub.2 for two hours to allow cell adherence. The
pellets were gently overlaid with control medium and incubated
overnight. The medium was replaced by chondrogenic medium [DMEM
with 1% FBS supplemented with 10 ng/ml TGF-.beta.1 l (R&D
Systems, Minneapolis, Minn.), 6.25 .mu.g/ml insulin (Sigma), and
6.25 .mu.g/ml transferrin (Sigma)]. The pellets were induced for
six days in chondrogenic medium. At day six and thereafter, 50
.mu.g/ml ascorbic acid-2-phosphate (Sigma) was added to the
chondrogenic medium mixture. PLA pellets were harvested at days
two, seven, and fourteen after initial induction for analysis. In
order to identify optimal culture conditions for the induction of
chondrogenic differentiation, PLA cells were also induced with
dexamethasone (Sigma) alone at a concentration of 0.1 .mu.M and in
combination with TGF-.beta.1. HFF cells were cultured as above
under micromass and monolayer conditions as a negative control. PLA
cells, incubated as monolayer cultures, did not form
three-dimensional nodules and were unavailable for paraffin
embedding and histologic and immunohistological analysis.
Differential Cell Density Plating
[0229] In order to assess the relationship of chondrogenic
induction to PLA cell, micromass cultures were plated in
chondrogenic medium at cell concentrations of 1.times.10.sup.5,
1.times.10.sup.6, 2.5.times.10.sup.6, 5.times.10.sup.6,
1.times.10.sup.7, 2.times.10.sup.7, and 5.times.10.sup.7 cells per
milliliter (cells/ml). The micromass cultures were then subjected
to chondrogenic culture conditions and the onset of nodule
formation noted.
Alcian Blue Staining
[0230] In order to detect the presence of highly sulfated
proteoglycans, characteristic of cartilaginous matrices, induced
PLA pellets were stained using Alcian blue at acidic pH (Lev, R and
S Spicer 1964 J. Histochem Cytochem. 12:309). Micromass cultures
were fixed with 4% paraformaldehyde in PBS for fifteen minutes,
followed by a five minute incubation in 0.1 N HCl to decrease the
pH to 1. The cultures were stained overnight with 1% Alcian blue
8GX (Sigma) in 0.1 N HCl (pH 1). The cells were washed twice with
0.1 N HCl to remove nonspecific staining and then air-dried. For
paraffin sections, cellular nodules were harvested, washed twice in
PBS and fixed in 4% paraformaldehyde for one hour. The nodules were
embedded in paraffin and cut into five-micrometer sections.
Paraffin sections of PLA nodules were prepared as described and
stained with standard Alcian blue staining at pH 1 in order to
determine the spatial distribution of sulfated proteoglycans within
the three-dimensional structure of the nodules. Digital images were
acquired with a Zeiss Axioskop II microscope (Carl Zeiss, Munich,
Germany) and Spot software.
Histology And Immunohistochemistry
[0231] Histologic evaluation of PLA paraffin sections was performed
using standard hematoxylin & eosin (H&E) to determine
cellular morphology and Goldner's trichrome stain to detect the
presence of collagen in the extracellular matrix. For
immunohistochemistry, paraffin sections were first deparaffinized
in xylene and then hydrated in decreasing ethanol solutions (100%
to 70%). To facilitate antibody access to epitopes, sections were
predigested for one hour at 37.degree. C. in 0.5 ml chondroitinase
ABC (Sigma) in 50 mM Tris (Gibco BRL), pH 8.0, 30 mM sodium acetate
containing 0.5 mg/ml BSA, 10 mM NEM. The sections were incubated in
3% H.sub.2O.sub.2 for fifteen minutes to quench endogenous
peroxidase activity, followed by incubation in 10% horse serum to
block nonspecific binding. The sections were subsequently incubated
for one hour at 37.degree. C. with primary antibodies to the
following: human type II collagen, chondroitin-4-sulfate, and
keratan sulfate at dilutions of 1:10, 1:50, and 1:250,
respectively. Incubation in normal horse serum in lieu of
monoclonal antibodies was performed to serve as a negative control.
Reactivity was detected with the Vectastain ABC kit (Vector
Laboratories, Burlingame, Calif.) according to the
manufacturer.
cDNA Synthesis and RT-PCR
[0232] Total RNA was isolated from untreated PLA cells, PLA
nodules, and HFFs. Briefly, RNA was isolated using the following
method (RNA-Easy, Qiagen). The RNA was used for oligo dT-primed
cDNA synthesis using MMLV-RT enzyme (Promega). Equivalent amounts
of cDNA were subjected to PCR amplification using primer pairs
designed to: human type I collagen .alpha.1 chain (CN I), human
type II collagen .alpha.1 chain (CN II), human type X collagen
.alpha.1 chain (CN X), human large aggregating proteoglycan or
aggrecan (AG) and human osteocalcin (OC). The primer pairs used
were obtained from published GeneBank sequences (Table 4) and are
as follows:
5TABLE 4: Expected Product Gene accession # Primer #1 Primer #2
Size CN I NM.sub.--000088 5'-CAT CTC CCC 5'-CTG TGG AGG AGG 598 bp
TTC GTT TTT GA- GTT TCA GA-3' 3' CN II Published 5'-CTG CTC GTC
5'-AAG GGT CCC AGG IIA*: 432 bp (148) GCC GCT GTC TTC TCC ATC-3'
IIB*: 225 bp CTT-3' N X NM_000493 5'-TGG AGT GGG 5'-GTC CTC CAA CTC
601 bp AAA AAG AGG CAG GAT CA-3' TG-3' AG X17406 5'-GCA GAG ACG
5'-GGT AAT TGC AGG 504 bp CAT CTA GAA GAA CAT CAT T-3' ATT G-3' OC
X04143 5'-GCT CTA GAA 5'-GCG ATA TCC TAG 310 bp TGG CCC TCA ACC GGG
CCG TAG-3' CAC TC-3' *Collagen type IIA splice--prechondrocytes and
mesenchymal chondrocytic precursors; type IIB--mature chondrocytes
(148).
[0233] Primer pairs for type II collagen, type X collagen and
aggrecan were confirmed against articular cartilage samples as a
positive control. Calculated optimal annealing temperatures (OLIGO
Primer Analysis Software, National Biosciences Inc., Plymouth,
Minn.) were used for each primer pair. Templates were amplified for
35 cycles and the PCR products were analyzed using conventional
agarose gel electrophoresis.
Effect Of Passage On The Chondrogenic Potential Of PLA Cells
[0234] To examine the effect of multiple cell passaging on the
chondrogenic potential of human PLA cells, monolayer cultures were
passaged fifteen times, with cell fractions taken at the first,
third and fifteenth passages. The cell fractions were placed in
micromass cultures, grown in chondrogenic medium and chondrogenic
differentiation was assessed by Alcian blue staining.
PLA Clonal Isolation
[0235] Freshly isolated PLA cells were plated out at a density of
100 cells per 100 mm.sup.2 tissue culture dish, to promote the
formation of colonies from single cells. Cultures were expanded in
control medium until the appearance of distinct colonies. Colonies
derived from single PLA cells were isolated using sterile cloning
rings, then harvested with 0.25% trypsin digestion. The dissociated
cells were seeded into 24-well plates and expanded. PLA clones were
induced toward the chondrogenic lineage as described above and
chondrogenic differentiation was confirmed by Alcian blue staining
and type II collagen immunohistochemistry.
Results
[0236] Human lipoaspirates were processed to obtain the PLA cell
population. The PLA was placed into high-density micromass cultures
supplemented with TGF-.beta.1, insulin, transferrin, and ascorbic
acid to induce chondrogenic differentiation. Chondrogenesis was
assessed histologically at two, seven, and fourteen days using
standard histologic assays. In addition, immunohistochemistry was
performed with antibodies to type II collagen,
chondroitin-4-sulfate, and keratan sulfate. Finally, RT-PCR
analysis was performed to confirm the expression of type I, type
II, and type X collagen as well as cartilage-specific proteoglycan
and aggrecan.
[0237] All TGF-.beta.1-treated micromass cultures formed
three-dimensional spheroids within 48 hours of induction that
stained positively with Alcian blue, suggestive of cartilaginous
nodule formation. Immunohistochemistry confirmed the presence of
type II collagen, chondroitin-4-sulfate, and keratan sulfate
throughout the extracellular matrix of the nodules. Finally, RT-PCR
analysis confirmed the expression of cartilage-specific type II
collagen, aggrecan, and cartilage-specific proteoglycan.
PLA Cells Form Chondrogenic Nodules
[0238] Pre-cartilage mesenchymal cells and multi-lineage stem cells
can be induced toward the chondrogenic lineage using a high-density
micromass culture technique, followed by induction with
pro-chondrogenic factors (Ahrens, P B, et al., 1977 Dev. Biol.
60:69-82; Denker, A E, et al., 1995 Differentiation 59:25-34;
Johnstone, B, et al., 1998 Exp. Cell Res. 238:265-272). Consistent
with these studies, human Processed Lipoaspirate (PLA) cells,
cultured under high-density micromass conditions and induced with
chondrogenic medium, containing transforming growth factor-beta 1
(TGF-.beta.1), insulin, and transferrin, condensed into
three-dimensional spheroids as early as twenty-four hours
post-induction. At this time period, the PLA nodules were visible
to the naked eye as white, round structures measuring approximately
1-2 mm in diameter. Nodules formed in 100% of over 500 treated
micromass cultures. Small spheroids formed in untreated micromass
cultures occasionally (10%) and may be an effect of the culture
conditions themselves. No PLA nodules were observed in TGF-.beta.1
-treated or untreated PLA monolayer cultures. PLA nodules became
larger in size with culture time and smaller adjacent nodules could
be visualized under a microscope after seven days in culture. In
some cases, adjacent PLA nodules coalesced into a larger, cellular
aggregates with increased culture time and is consistent with the
proposed cellular interactions and recruitment that are essential
to chondrogenesis (Ahrens, P B, et al., 1977 Dev. Biol.
60:69-82).
[0239] In order to assess the effect of cell number on PLA nodule
formation, differential plating studies were performed. No evidence
of spheroid formation was seen in cultures plated at a density of
less than 5.times.10.sup.6 cells/ml. PLA cells plated at increasing
densities (i.e. above 1.times.10.sup.7 cells/ml) underwent nodule
formation more rapidly and, in some cases, were more likely to
undergo spheroid formation in the absence of TGF-.beta.1. The
addition of dexamethasone to chondrogenic medium, containing
TGF-.beta.1, has been shown to lead to the formation of larger
cartilaginous aggregates (Johnstone, B., et al., 1998 Exp. Cell
Res. 238:265-272). Consistent with this, the addition of
dexamethasone resulted in larger spheroids when compared to nodules
formed with TGF-.beta.1 stimulation. Cultures treated with
dexamethasone alone did not form nodules, suggesting that
TGF-.beta.1 is crucial to nodule formation by PLA cells. Finally,
no evidence of nodule formation was observed in micromass and
monolayer HFF cultures treated with chondrogenic medium, confirming
the specificity of our chondrogenic conditions.
PLA Nodules Contain an Extracellular Matrix Rich in Sulfated
Proteoglycans
[0240] Cartilagenous matrices contain very high quantities of
polyanionic sulfated glycoasminoglycans (GAGs), such as chondroitin
4- and 6-sulfate, and are characterized by the ability to stain
positively with Alcian blue at low pH (R Lev and S Spicer 1964 J.
Histochem. Cytochem. 12:309). In order to confirm the cartilaginous
nature of the PLA nodules, histologic analysis was performed on
whole-mount PLA nodules, plated on chamber slides, and paraffin
sections. Initial treatment of PLA cultures with chondrogenic
medium resulted in cellular condensation within 24 hours (FIG. 12,
Panel A). Condensing PLA cells exhibited a low level of Alcian Blue
staining, suggesting the initial formation of a sulfated
extracellular matrix. PLA condensation was followed by ridge
formation and increased staining by Alcian Blue, indicating an
increase in matrix secretion (Panel B). Intense Alcian Blue
staining and spheroid formation was observed after 48 hours
post-induction (Panel C). In contrast, untreated PLA cells in
micromass cultures did not show any regions of positive Alcian blue
staining (Panel D).
[0241] In addition to whole-mount PLA samples, paraffin sections of
PLA nodules were prepared in order to assess the three-dimensional
architecture of the nodule. The morphology of the paraffin-embedded
sections, as analyzed by hematoxylin and eosin staining, showed a
flat, peripheral layer of fibroblast-like cells that resembled
perichondral cells, surrounding an inner core of rounder cells at
two days post-induction (FIG. 13, Panel A). After fourteen days of
treatment, nodules became more hypocellular with increasing
deposits of extracellular matrix into the core (Panel B). Goldner's
trichrome staining, which indicates the presence of collagenous
matrix (green color), confirmed the H&E pattern (Panels C and
D). Faint background levels of collagenous matrix were observed in
the nodule sections at two days (Panel C), compared with higher
levels of collagen seen in the nodule core at fourteen days
post-induction (Panel D). Alcian blue staining of the paraffin
sections was similar to the whole-mount preparations, confirming
the formation of cartilaginous matrix rich in sulfated
proteoglycans after two days induction (Panel E). Increased
staining intensity in the central core region was observed at
fourteen days post-induction (Panel F), suggesting an increased
secretion of sulfated proteoglycans as the cells mature down the
chondrocytic pathway. In summary, our histological staining results
confirm the formation of cartilage-like PLA nodules, associated
with an extracellular matrix rich in collagens and sulfated
proteoglycans.
PLA Nodules Express Cartilage-Specific Proteins
[0242] Immunohistochemical analysis was used to detect the presence
of type II collagen, an extracellular matrix component highly
specific for cartilaginous tissue, and chondroitin-4-sulfate and
keratan sulfate, two of the main monomeric components of cartilage
proteoglycans. After two days induction, areas of strong
immunoreactivity to chondroitin-4-sulfate and keratan sulfate were
seen along the outer periphery of the spheroids and throughout the
core and is supportive of our Alcian Blue staining results (FIG.
14, Panels A and C). A significant increase in
chondroitin-4-sulfate and keratan sulfate expression within the
nodule core was noted over the course of two weeks (Panels B and
D). In contrast, positive type II collagen immunoreactivity was not
evident in the PLA nodules at day two (Panel E). Rather, collagen
type II expression appeared at day seven post-induction, with
strong expression appearing at day fourteen (Panel F). Whole-mount
cultures of TGF-.beta.1-treated PLA micromass cultures also showed
intense type II collagen reactivity while untreated micromass PLA
cultures showed no staining . In addition, no staining was observed
in paraffin sections incubated in normal horse serum instead of
primary monoclonal antibodies, supporting the specificity of the
type II collagen, chondroitin-4-sulfate, and keratan sulfate
antibodies. Taken together, the immunohistochemical results support
the histological staining data and suggest the presence of a
cartilaginous matrix in PLA nodules.
Chondrogenic Differentiation of Single-Cell Derived Clonal
Populations
[0243] The apparent chondrogenic differentiation by PLA cells may
result from contamination of the lipoaspirate by pre-chondrogenic
cells rather from the presence of a multipotential cell. Therefore
to determine if our results are due to differentiation of
multipotential PLA cells, we isolated and confirmed the
multilineage potential of single-cell derived PLA clones. PLA
clonal populations (i.e. adipo-derived mesodermal stem cells or
ADSCs) demonstrated the ability to undergo chondrogenic
differentiation in addition to osteogenic and adipogenic
differentiation. PLA clonal populations induced toward the
osteogenic and adipogenic lineages exhibited classic
lineage-specific histological markers (alkaline phosphatase
activity-osteogenesis; Oil-Red-O accumulation-adipogenesis)
(unpublished data). Like the heterogeneous PLA cultures, PLA clonal
populations also underwent spheroid formation within forty-eight
hours of induction in chondrogenic medium. In addition, the PLA
nodules secreted an extracellular matrix rich in type II collagen
and highly sulfated proteoglycans.
PLA Cells Retain Chondrogenic Potential After Extended Culture
[0244] Culture time and passage number can affect the
differentiative capacity of many cell types. To assess the effect
of passaging on the chondrogenic potential of PLA cells, PLA cells
were passaged in monolayer cultures as many as fifteen times (175
culture days) and cultured under high-density conditions to induced
chondrogenesis. PLA cells retained their chondrogenic
differentiation potential throughout this extended culture period,
as evidenced by their ability to form three-dimensional spheroids
after induction with chondrogenic medium. Finally, both early and
late passage PLA nodules secreted an extracellular matrix rich in
highly-sulfated proteoglycans as evidenced by the positive staining
with Alcian blue (FIG. 22). Cellular nodules from all culture
passages (i.e. P1 to P15) had a very similar appearance: a flat,
peripheral layer of fibroblast-like cells resembling the
perichondrium surrounding an inner core of rounder cells.
RT-PCR Analysis Confirms the Expression Of Cartilage-Specific
Collagens
[0245] RT-PCR analysis of PLA nodules was performed using primers
specific to the genes for human type I collagen, type II collagen,
and type X collagen, as well as aggrecan and osteocalcin. Untreated
HFF and human PLA cells cultured under micromass conditions were
analyzed as negative controls. RT-PCR analysis of PLA nodules
confirmed the expression of type II collagen.alpha.1 (CN II) at day
7 and day 14 only (FIG. 15). Moreover, decrease in CN II expression
was observed between 7 and 14 days induction. Both splice variants
of CN II (IIA and IIB--type IIB variant shown) were observed at
both time points.
[0246] In contrast to day seven and fourteen nodules, CN II
expression was not observed in 2-day nodules, confirming our
immunohistochemical data. As expected, CN II was not observed in
HFF micromass cultures . However, small amounts of CN II mRNA were
present in the untreated PLA cells. Chondrogenic differentiation
was further confirmed by examining nodules for the expression of
the large aggregating proteoglycan, or aggrecan. Aggrecan has been
shown to be specific to cartilage and accumulates at the onset of
over chondrogenesis (Kosher, R A, et al., 1986 J. Cell Biol.
102:1151-1156). Aggrecan expression was observed at both 2 and 7
days induction and was absent in 14 day PLA nodules. Aggrecan
expression was specific to treated PLA nodules, as no expression
was noted in control PLA cells or in HFF cultures.
[0247] Further characterization of PLA nodules was performed by
assessing the expression of the .alpha.1 chains of type I and type
X collagen. Collagen type I expression is known to be up-regulated
in osseous tissues and is down-regulated during chondrogenic
differentiation (Kosher, R A, et al., 1986 J. Cell Biol.
102:1151-1156; Shukunami, C., et al., 1998 Exp. Cell Res.
241:1-11). Consistent with this, CN I expression was observed in
2-day treated PLA nodules only. Similar to CN II, low levels of CN
I were observed in untreated PLA cells, suggesting that
undifferentiated PLA cells are associated with a collagenous matrix
that is dramatically remodeled as differentiation proceeds. CN X
expression was not observed in PLA nodules at two and seven days
post-induction but appeared at the 14-day time point. No CN X was
observed in untreated PLA cells or in HFF controls. Collagen type X
is specific to hypertrophic chondrocytes and may signal the
progression to endochondral ossification and bone formation
(Linsenmayer, T F, et al., 1988 Pathol. Immunopathol. Res.
7:14).
[0248] To confirm the absence of ossification and bone formation
within the PLA nodules, RT-PCR analysis was performed using primers
to osteocalcin, a bone-specific gene (Price P A 1989 Connect.
Tissue Res. 21:51-57). As expected, osteocalcin expression was
absent in all treated and untreated PLA samples. Taken together,
the expression of cartilage-specific aggrecan, both type II and X
collagen, together with the decreased expression of type I collagen
supports the chondrogenic differentiation by PLA cells.
Discussion
[0249] The repair of cartilaginous defects remains a significant
clinical challenge. Damaged articular cartilage has a limited
potential for repair and large defects do not heal spontaneously.
When the damage extends into the subchondral bone, the repair
process is sporadic and the original articular cartilage is
replaced by fibrocartilage and scar tissue, which are structurally
inferior to the hyaline architecture of normal articular
cartilage.
[0250] Conventional treatment modalities for cartilage defects
include marrow stimulation techniques (e.g. subchondral drilling)
and joint arthroplasty (I H Beiser and O I Knat 1990 J. Foot Surg.
29:595-601; Gilbert, J E 1998 Am. J. Knee Surg. 11:42-46; T Minas
and S Nehrer 1997 Orthopedics 20:525-538; O'Driscoll, S W 1998 J.
Bone Joint Surg. Am. 80:1795-1812). More recently, newer strategies
have been developed, such as the use of osteochondral,
perichondral, and periosteal allografts (Bouwmeester, S J, et al.,
1997 Int. Orthop. 21:313-317; Carranza-Bencano, A, et al., 1999
Calcif. Tissue Int. 65:402-407; Ghazavi, M T, et al., 1997 J. Bone
Joint Surg. Br. 79:1008-1013; Homminga, G N, et al., 1990 J. Bone
Joint Surg. Br. 72:1003-1007). Unfortunately, these options do not
result in complete regeneration of the original hyaline
architecture. More importantly, the joint is not capable of normal
weight-bearing and physical activity over prolonged periods of
time.
[0251] Cell-based tissue engineering strategies represent a
promising alternative to conventional techniques. First-generation
tissue engineering strategies are currently employed clinically
using autologous chondrocyte implantation (Brittberg, M., et al.,
1994 New Engl. J. Med. 331:889-95; Chen, F S, et al., 1997 Am. J.
Orthop. 26:396-406; Gilbert J E 1998 Am. J. Knee Surg. 11:42-46;
Richardson J B, et al., 1999 J. Bone Joint Surg. Br. 81:1064-1068).
However, limited availability of donor sites for chondrocyte
harvest, the requirement for lengthy in vitro culture expansion,
and donor site morbidity limit the practicality of this technique.
It is important to identify other sources of chondrocytic
precursors.
[0252] Several cell types have been shown to undergo in vitro and
in vivo chondrogenesis, including rat calvarial clonal cell lines
and primary cells, the murine embryonic C3H10T1/2 cells, and
periosteum-derived and bone marrow-derived precursors from several
animals including rabbits, rats, horses, and goats (Denker, A E, et
al., 1995 Differentiation 59:25-34; Fortier, L A, et al., 1998 Am
J. Vet. Res. 59:1182-1187; Grigoriadis, et al., 1996
Differentiation 60:299-307; Grigoriadis, et al., 1988 J. Cell.
Biol. 30 106:2139-2151; Iwasaki, et al., 1995 J. Bone Joint Surg.
Am. 77:543-554; Johnstone, et al., 1998 Exp. Cell Res. 238:265-272;
Nakahara, et al., 1990 Bone 11:181-188; Shukunami, et al., 1996 J.
Cell. Biol. 133:457-468). However, there remains a large potential
reservoir of osteochondrogenic precursors from other tissue types
that have yet to be studied. The interconversion ability of various
mesodermal cell types has been reported in many studies.
Specifically, both mature human adipocytes and adipocytes isolated
from bone marrow exhibit the potential to differentiate into bone
(Bennett, J H, et al., 1991 J. Cell Sci. 99(Ptl):131-139; Park, et
al., 1999 Bone 24:549-554). In addition, osteoblasts
transdifferentiate into chondrocytes and muscle cells are capable
of commitment to the cartilage lineage (Manduca, et al., 1992 Eur.
J. Cell Biol. 57:193-201; Nathanson, M A 1985 Clin. Orthop.
200:142-158; Sampath, et al., 1984 Proc. Natl. Acad. Sci. USA
81:3419-3423).
[0253] The presence of mesenchymal stem cells capable of
osteochondrogenic differentiation in human bone marrow has been
well-documented (Mackay, et al., 1998 Tissue Eng. 4:414-428;
Pittinger, et al., 1999 Science 284:143-147; Yoo, et al., 1998 J.
Bone Joint Surg. Am. 80:1745-1757). Some of the advantages of using
mesenchymal stem cells include their ability to proliferate rapidly
in culture, their ability to differentiate into chondrogenic cells
even after multiple passages, their regenerative capacity, and a
broad range of resultant chondrogenic cell types (i.e.
prechondroctyes, mature chondrocytes, and hypertrophic
chondrocytes). Researchers have anticipated that the differentiated
chondrogenic tissue derived from stem cells will more closely
resemble that seen in developing embryonic limb buds. Moreover,
chondrocytes proliferate poorly in culture, are difficult to
maintain, and dedifferentiate when expanded in monolayer cultures
(von der Mark, et al., 1977 Nature 267:531-532). The use of
autologous stem cells in place of harvested chondrocytes in tissue
engineering may be a more efficacious alternative in the future for
treatment of cartilage defects. Unfortunately, the limited
availability of donor sites and the discomfort and pain associated
with bone marrow procurement remain a concern.
[0254] The presence of a multipotential cell population within
adipose tissue, capable of differentiation into several mesenchymal
tissues may be an important finding. Adipose tissue is available in
large quantities and relatively easy to obtain. Moreover,
liposuction procedures have minimal donor site morbidity and
patient discomfort. Because of these practical advantages as a cell
source, we sought to determine if PLA cells, like bone marrow- and
periosteum-derived mesenchymal stem cells, represent a cell
population with the ability to undergo chondrogenic
differentiation.
[0255] We have confirmed the chondrogenic potential of multilineage
human processed lipoaspirate (PLA) cells. Human PLA cells in
high-density micromass cultures treated with TGF-.beta.1 resulted
in the formation of three-dimensional cellular nodules with
cartilaginous characteristics. The chondrogenic nature of the
differentiated cells was supported by several findings: 1)
whole-mount PLA nodules and histologic sections stained positively
with Alcian blue, 2) H&E morphology revealing a perichondral
border of cells surrounding a hypocellular chondrogenic core, 3) a
collagen-rich extracellular matrix as shown by Goldner's trichrome
staining, 4) expression of type II collagen, chondroitin-4-sulfate,
and keratan sulfate as confirmed by immunohistochemistry, and 5)
expression of collagen type II as well as cartilage-specific
aggrecan as shown by RT-PCR.
[0256] One of the earliest features of cartilage development in
vivo is the formation of cellular condensations that represent
skeletal primordia. Cartilage initially differentiates in the
center of these condensations and is followed by a period in which
the cells secrete and are surrounded by a characteristic
extracellular matrix. Similar to this situation, chondrogenic
differentiation in vitro is characterized by the formation of
multi-layered cellular aggregates, called spheroids or nodules.
Primary nodule formation is followed by ridge formation, the
accumulation of matrix and the recruitment of adjacent cells,
resulting in the expansion of the original nodule (Ahrens, et al.,
1977 Dev. Biol. 60:69-82; Denker, et al., 1995 Differentiation
59:25-34; Stott, et al., 1999 J. Cell Physiol. 180:314-324;
Tacchetti, et al., 1992 Exp. Cell Res. 200:26-33; Tavella, et al.,
1994 Exp. Cell Res. 215:354-362). Consistent with these studies,
PLA cells began condensing within twenty-four hours induction with
TGF-.beta.1-containing chondrogenic medium and formed well-defined
three-dimensional spheroids by forty-eight hours post-induction.
The appearance of smaller adjacent nodules in addition to the
original cartilage nodule was noted after cultures were treated for
extended periods in chondrogenic medium, suggesting the presence of
further chondrogenic induction through possible paracrine growth
factor signaling by the maturing cartilaginous nodule. PLA nodule
formation was evident only in micromass cultures plated at a cell
density higher than 5.times.10.sup.6 cells/ml, consistent with
previous studies describing the high cell density requirement for
chondrogenesis (Rodgers, et al., 1989 Cell Differ. Dev. 28:179-187;
Tsonis and Goetinck 1990 Exp. Cell Res. 190:247-253 ).
[0257] Cartilage is comprised of a mixture of collagen fibrils and
proteoglycans that give the tissue high tensile strength and
internal swelling pressure. The predominant collagen of cartilage
is collagen type II. Although this collagen is not specific to
cartilage it is highly characteristic of this tissue, as collagen
type II is produced by a limited number of non-chondrogenic cell
types. Positive staining using a Goldner Trichrome stain, specific
for collagens in general, confirmed these proteins within the PLA
nodule after both 2 and 14 days induction with chondrogenic medium.
Specifically, PLA nodules treated with TGF-.beta.1 for 48 hours
were associated with an extracellular matrix containing low levels
of collagen type II, suggesting that PLA cells have undergone
preliminary chondrogenic differentiation. Collagen type II levels
appeared to increase with induction time. In addition to collagen
type II, cartilagenous matrices also contain high levels of
sulfated GAGs, such as chondroitin-4- and -6-sulfate that are
typically associated with proteoglycans such as aggrecan.
Consistent with this, histological staining with Alcian Blue
confirmed the presence of sulfated proteoglycans as early as 24
hours induction, increasing as the PLA nodule became more defined
(i.e. 2 days). Increased Alcian Blue staining was also observed as
far as 14 days induction, localizing to the nodule core and
surrounding individual cells. Similar results were also observed
when nodules were stained with antibodies specific to keratan- and
chondroitin-sulfate confirmed and immunohistochemical studies
confirmed the presence of these components and further supports the
presence of chondrogenic cells within the PLA nodule.
[0258] In support of our immunohistochemical results, RT-PCR
analysis confirmed the expression of CN II in PLA nodules induced
for 7 and 14 days, with a lower level of this gene being observed
at day 14. No CN II expression was observed after 48 hours
induction with chondrogenic medium. The expression of CN II in PLA
nodules is supportive of the chondrogenic phenotype. Our
immunohistochemical results showed a significant level of both
chondroitin- and keratan-sulfate specifically in induced PLA
nodules. It is known that chondroitin-4- and 6- sulfate are the
main monomeric components of the cartilage-specific protein,
aggrecan (Hall, B K 1981 Histochem. J. 13:599-614). Aggrecan has
been shown to be cartilage-specific and accumulates at the onset of
overt chondrogenesis, coincident with cellular condensation
(Kosher, et al., 1986 J. Cell Biol. 102:1151-1156).
[0259] We confirmed the chondrogenic nature of the PLA nodule by
assessing the expression of aggrecan. As shown in FIG. 22, the
expression pattern of aggrecan overlapped with that of CN II at day
7. In addition, the expression of aggrecan preceded that of CN II.
However, in contrast to CN II, aggrecan was not observed in PLA
nodules induced for 14 days. Aggrecan is a cartilage-specific
protein that consists of a multidomain protein core containing
binding sites for sulfated proteoglycans (Hardinghamn, et la., 1984
Prog. Clin. Biol. Res. 151:17-29). The primers used to detect
aggrecan in this study were designed to the C-terminus, which
contains the G3 globular domain, a site that undergoes alternative
splicing and is proteolytically cleaved in mature cartilage (Fulop,
et al., 1993 J. Biol Chem. 268:17377-17383). The absence of
aggrecan in day 14 PLA nodules may therefore represent an
alternatively spliced form of aggrecan that lacks the C-terminus.
However, no aggrecan at day 14 was observed when RT-PCR was
performed using primers designed to the N-terminus, suggesting that
aggrecan is no longer expressed after two weeks induction with
chondrogenic medium.
[0260] In addition to aggrecan and CN II, PLA nodules expressed
both type I and type X collagen at distinct time points. Day 2 PLA
nodules were characterized by the expression of both CN I and CN
II. However, in contrast to aggrecan and CN II, the expression
pattern of CN I was highly restricted and did not appear beyond the
two day time point. Interestingly, low levels of both CN I and CN
II were observed in untreated PLA control cells. Consistent with
this, both type I and type II collagen mRNA have been found in many
developing embryonic tissues and basal levels of these collagen
subtypes can be detected in pre-cartilage mesenchymal precursors
prior to chondrogenic differentiation (Lisenmeyer, et al., 1973
Dev. Biol. 35:232-239; Dessau, et al., 1980 J. Embryol. Exp.
Morphol. 57:51-60; Cheah, et al., 1991 Development 111:945-953;
Kosher and Solursh 1989 Dev. Biol 131:558-566; Poliard, et al.,
1995 J. Cell Biol. 130:1461-1472). Finally, the decrease in CN II
expression in day fourteen nodules coincided with the appearance of
CN X, a collagen indicative of hypertrophic chondrocytes (Kirsch,
et al., 1992 Bone Miner. 18:107-117; Linsenmayer, et al., 1988
Pathol. Immunopathol. Res. 7:14-19).
[0261] The appearance of collagen type X and the hypertrophic
phenotype may precede possible nodule ossification and bone
formation. However, PLA nodules did not express osteocalcin, a
bone-specific gene expressed in cells differentiating toward the
osteogenic lineage (Price, et al., 1983 Biochem. Biophys. Res.
Commun. 117:765-771). Despite the expression of collagen type X,
mature hypertrophic chondrocytes with their characteristic lacunae
were not seen in PLA nodules. However, a similar result has been
described by Denker et al. when C3H10T1/2 murine pluripotent cells
were placed in micromass cultures and treated with TGF-.beta.1
(Denker, et al., 1995 Differentiation 59:25-34). Hypertrophic
chondrocytes were only observed in place of nodules when cultures
were treated with BMP-2 (139). It therefore may be necessary to
induce PLA micromass cultures with BMP-2 to fully induce
hypertrophy and induce the formation of mature chondrocytes.
[0262] Taken together, our histologic, immunohistochemical, and
RT-PCR data support the differentiation of PLA cells toward the
chondrogenic lineage. However, the processed lipoaspirate is a
heterogeneous population of cells and may contain several cell
types of various mesodermal lineages. Specifically, there may exist
chondrogenic precursors in the lipoaspirate that are capable of
spontaneous differentiation, as well as a subpopulation of
multipotential cells (i.e. PLA stem cells). The isolation of PLA
clones derived from single PLA cells and their multilineage
differentiation (chondrogenesis, osteogeneis, and adipogenesis)
supports the presence of multipotential stem cells (adipo-derived
mesodermal stem cells) within this heterogeneous cell
population.
[0263] In order to apply cell-based tissue engineering techniques
to the clinical setting, a number of criteria must be met. The cell
population used as the cellular vehicle should be abundant and easy
to obtain, expandable in tissue culture, able to maintain its
differentiative ability through multiple passages, and exhibit
properties equivalent to the native target tissue. The healing of
articular cartilage defects using stem cells harvested from bone
marrow has been successfully reported in various animal models
(Angele, et al., 1999 Tissue Eng. 5:545-554; Butnariu-Ephrat, M.,
et al.,1996 Clin. Orthop. 330:234-43; Wakitani, et al., 1994 J.
Bone Joint Surg. Am. 76:579-592). However, the bone marrow harvest
is painful and yields low number of stem cells for clinical use,
usually requiring in vitro expansion. Adipose tissue is plentiful
and easy to obtain with relatively minimal discomfort. PLA cells
can be harvested from a relatively small amount of adipose tissue
in large numbers (Zuk, P., et al., 2001 Tissue Engineering
7:209-226), thereby, obviating the need for lengthy culture
expansions. While elective cosmetic surgery is the most common
source of lipoaspirates, sufficient adipose tissue could also be
obtained through a small-bore cannula for non-cosmetic surgery
patients requiring reconstruction, making this technique available
to a wide variety of patients. In this example, the chondrogenic
capacity of multipotential adipose-derived stem cells is
demonstrated and shows that the stem cells retain their ability to
differentiate even after long-term culture. Finally,
adipose-derived stem cell nodules exhibit many properties
consistent with native cartilage tissue.
[0264] The fulfillment of these properties, together with the
potential abundance of PLA cells, make these multipotential cells
an ideal system for tissue engineering strategies. In addition,
these cells may be appropriate for the study of chondrogenesis in
both in vitro culture studies and in vivo animal models. The
identification of chondrogenic precursors has important
implications for the repair of articular cartilage defects. The
abundance and easy accessibility of adipose tissue makes it a
feasible alternative for cartilage reconstruction (Asahina, et al.,
1996 Exp. Cell Res. 222:38-47; Atkinson, et al., 1997 J. Cell
Biochem. 65:325-339; Chimal-Monroy and Diaz de Leon 1999 Int. J.
Dev. Biol. 43:59-67; Denker, et al., 1999 Differentiation 64:67-76;
Klein-Nulend, et al., 1998 Tissue Eng. 4:305-313; Martin, et al.,
1999 Exp. Cell Res. 253:681-688; Quarto, et al., 1997 Endocrinology
138:4966-4976; Shukunami, et al., 1998 Exp. Cell Res.
241:1-11).
EXAMPLE 10
[0265] The following description provides methods for isolating
stem cells from adipose tissues, where the stem cells differentiate
into myogenic tissue.
Methods
Differentiation and Tissue Culture Reagents
[0266] Hydrocortisone, collagenase and paraformaldehyde were
purchased from Sigma (St. Louis, Mo.). Horse Serum (HS) was
purchased from Life Technologies (Grand Island, N.Y.).
Phospho-Buffered Saline (PBS), 0.25% trypsin/l mM EDTA
(trypsin/EDTA), Dulbecco's Modified Eagle's Medium (DMEM) and
antibiotic/antimycotic solution were purchased from CellGro
(Herndon, Va.). Fetal Bovine Serum (FBS) was purchased from Hyclone
(Logan, Utah).
PLA Preparation and Culture
[0267] Human adipose tissue, obtained from eight patients (mean
age=39.3 years, range 25-58 years) undergoing elective
Suction-Assisted Lipectomy (SAL), according to patient consent
protocol HSPC #98-08 011-02 (University of California Los Angeles)
was processed as described, according to the method described in
Example 7, supra, to obtain the Processed Lipoaspirate (PLA) cell
population. Briefly, the raw liposuctioned aspirates were washed
extensively with sterile PBS in order to remove blood cells, saline
and local anesthetics. The extracellular matrix was digested with
0.075% collagenase 37.degree. C. for 30 minutes to release the
cellular fraction. Collagenase was inactivated with an equal volume
of DMEM containing 10% FBS. The infranatant was centrifuged at
250.times.g for 10 minutes to obtain a high-density PLA cell
pellet. The pellet was resuspended in DMEM/10% FBS and an
Erythrocyte Lysis Buffer (0.16M NH.sub.4Cl) was added for 10
minutes to lyse contaminating erythrocytes. Following an additional
centrifugation step, the PLA cell pellet was resuspended in
DMEM/10% FBS and plated in 100 mm tissue culture dishes at a
density of 1.times.10.sup.6 cells per plate. PLA cells were
maintained in Control Medium (CM--DMEM, 10% FBS, 1%
antibiotic/antimycotic) at 37.degree. C. and 5% CO.sub.2. The
culture medium was changed twice weekly. Confluent PLA cultures
(approximately 80% confluence) were passaged at a ratio of 1:3 in
trypsin/EDTA. For control studies, a human foreskin fibroblast cell
line, HFF (American Type Culture Collection, Manassas, Va.) and a
human skeletal muscle cell line, SKM (Clonetics, Walkersville, Md.)
were maintained at 37.degree. C./5% CO.sub.2 in CM and a myogenic
maintenance medium (SKM--Clonetics), respectively.
Myogenic Differentiation
[0268] To induce optimal myogenesis, PLA cells were plated at a
density of 1.times.10.sup.4 cells onto 35 mm tissue culture dishes
and incubated overnight in CM to allow adherence. Optimal
myogenesis was obtained by incubating PLA cells in Myogenic Medium
(MM=CM supplemented with 5% Horse Serum and 50 .mu.m hydrocortisone
to promote proliferation, a key event in myogenic differentiation)
(196). PLA cells were induced in MM for 1, 3 and 6 weeks. Medium
was changed twice weekly until the experiment was terminated. SKM
and HFF cells were induced for 1, 3 and 6 weeks in MM as positive
and negative controls, respectively.
Immunohistochemistry
[0269] To assess myogenic differentiation, PLA cells were seeded
onto 8-well chamber slides at a density of 5.times.10.sup.3 cells
per well and allowed to adhere in CM overnight. Cells were induced
in MM for 1, 3 and 6 weeks. Following induction, the cells were
rinsed twice with PBS and fixed with 4% paraformaldehyde for 20
minutes at 4.degree. C. The cells were incubated with 3% hydrogen
peroxide for 5 minutes to quench endogenous peroxidase activity.
Non-specific epitopes were blocked by a 30 minute incubation in
Blocking Buffer (BB; PBS, 1% HS, 0.1% Triton X-100). The cells were
incubated at 4.degree. C. overnight with either a monoclonal
antibody to human MyoD 1 (Dako; Carpenteria, Calif.) or monoclonal
antibody to human fast twitch skeletal muscle myosin heavy chain
(Biomeda Corp.; Foster City, Calif.). Following incubation, the
cells were washed with BB and incubated at room temperature for 2
hours in BB containing a horse anti-mouse IgG secondary antibody
conjugated to biotin. The secondary antibody was visualized using
the VectaStain ABC kit (Vector Labs; Burlingame, Calif.) according
to manufacturer's specifications. The cells were counterstained
with hematoxylin for 3 minutes. SKM cells induced in MM were
processed as above as a positive control. PLA cells cultured in CM
and HFF cells induced in MM were analyzed as negative controls.
RT-PCR Analysis
[0270] Total RNA was isolated from PLA cells treated with MM for 1,
3 and 6 weeks. RNA was isolated according to the method described
in Example 9 above. Five micrograms (5 ug) of total RNA was used
for oligo dT-primed cDNA synthesis using Murine Maloney Leukemia
Virus Reverse Transcriptase (MMLV-RT; Promega; Madison, Wis.). The
resulting cDNA was used as a template for PCR analysis using primer
pairs designed to human MyoD1 (Accession; NM.sub.--002478) and
human skeletal muscle myosin heavy chain (Accession; X03740). The
primer pairs used and the expected PCR product sizes were as
follows: MyoD1: 5'-AAGCGCCATCTCTTGAGGTA-3' (forward primer) and
5'-GCGCCTTTATTTTGATCACC-3' (reverse primer); 500 bp; myosin heavy
chain: 5'-TGTGAATGCCAAATGTGCTT-3' (forward primer) and
5'-GTGGAGCTGGGTATCCTTGA-3' (reverse primer); 750 bp. MyoD1 and
myosin were amplified using Taq polymerase (Promega) for 35 cycles
in a total reaction volume of 100 ul. Duplicate reactions were
performed using primers designed to the housekeeping gene,
.beta.-actin, as an internal control. PCR products were resolved by
agarose gel electrophoresis. PCR amplification of cDNA obtained
from PLA cells cultured in CM and HFF cells induced in MM was
performed as negative controls.
Immunohistochemical Quantification And Data Analysis
[0271] To quantitate myogenesis, a total of five hundred PLA cells
from each induction time point were manually counted at
200.times.magnification using an "Axioskop 2" inverted microscope
(Carl Zeiss Inc; Thornwood, N.Y.) and the number of MyoD1 and
myosin positive cells determined. The number of MyoD1 and
myosin-positive cells was expressed as a percentage of the total
500 cells (% total PLA cells) and was used as an indication of the
degree of myogenic differentiation. All studies were performed on
eight patients and the mean number of MyoD1 and myosin-positive
cells calculated, together with the standard error of the mean
(.+-.SEM). Myogenic differentiation in both the experimental and
control groups described above was analyzed for statistical
significance using a one-way analysis of variance (ANOVA). A p
value of less than 0.05 was considered significant.
Results
Induced Stem Cells Express Myodl and Myosin Heavy Chain
[0272] Consistent with the previous examples, no qualitative
changes in PLA growth kinetics and morphology between the 8
patients used in this study, suggesting that the isolated PLA
populations are relatively consistent between all patients. PLA
cells were isolated from raw lipoaspirates and induced using MM,
containing hydrocortisone. Myogenesis by PLA cells was specific to
the myogenic conditions used in this study, as no differentiation
was observed in non-inductive control medium, or in media inductive
for alternate mesodermal lineages (i.e. osteogenic and adipogenic).
Furthermore, no osteogenic or adipogenic differentiation was noted
in PLA cells induced for up to 6 weeks in MM.
[0273] To confirm PLA myogenic potential, the expression of
established muscle-specific markers was determined by
immunohistochemistry. Differentiation of myogenic precursors and
stem cells into myogenic precursor cells can be confirmed by the
expression of several transcription factors, that include MyoD1,
Myf-5, myogenin and structural proteins such as myosin heavy chain
(Butler-Browne, et al., 1990 Anat. Embryol. (Berl) 181:513-522;
Thornell, et al., 1984 J. Neurol. Sci. 66:107-115; Megeney, et al.,
1996 Genes Dev. 10:1173-1183; Seale and Rudnicki 2000 Dev. Biol.
218:115-124; Tapscott, et al., 1988 Science 242:405-41 1;
Weintraub, et al., 1991 Science 251:761-766; Molkentin and Olson
1996 Curr. Opin. Genet. Dev. 6:445-453). Commitment to the myogenic
lineage was identified by staining cells with a monoclonal antibody
specific to MyoD1. Nuclear expression of MyoD1 in PLA cells was
observed at 1, 3 and 6 weeks induction with MM, suggesting
initiation of the myogenic differentiation pathway in these cells
(FIG. 16, Panels A to C). Similar to the PLA results, nuclear
expression of MyoD1 was observed in positive control SKM cells as
early as 1 week post-induction with MM and increased MyoD1
expression was observed in SKM cells by 6 weeks induction. In
contrast to induced PLA cells, MyoD1 expression was not observed in
PLA cells treated for 1, 3 and 6 weeks with CM (FIG. 16, Panels D
to F). Similarly, no MyoD1 expression was observed in HFF cells
treated with MM . The expression of MyoD1 in induced PLA cells
suggests that these cells have initiated a program of myogenic
differentiation.
[0274] To further confirm myogenesis, cells were stained with a
monoclonal antibody specific to skeletal muscle myosin heavy chain
(myosin), in order to identify terminally differentiated myoblasts
(Butler-Browne, et al., 1990 Anat. Embryol. (Berl) 181:513-522;
Thornell, et al., 1984 J. Neurol. Sci. 66:107-115). Consistent with
the nuclear expression of MyoD1, PLA cells induced with MM also
expressed myosin (FIG. 17, Panels A to C). However, in contrast to
MyoD1, myosin expression was restricted to later induction time
points (3 and 6 weeks only), consistent with the expression of this
marker in mature, fully differentiated myoblasts (Butler-Browne, et
al., 1990 Anat. Embryol. (Berl) 181:513-522; Thomell, et al., 1984
J. Neurol. Sci. 66:107-115). Similar to our MyoD1 results, no
myosin expression was observed in PLA cells cultured in CM (FIG.
17, Panels D to F) or in HFF cells induced with MM. Extensive
myosin expression was also observed in SKM positive controls
induced for 3 and 6 weeks with MM. Taken together, the expression
of both MyoD1 and myosin in induced PLA cells suggests that these
cells possess myogenic potential.
[0275] Terminal differentiation of myogenic precursors is
accompanied by the fusion of the differentiated myoblast into long,
multi-nucleated myotubes. Therefore, we examined induced PLA
cultures for the formation of putative myotubes. Treatment of PLA
cells with MM for a minimum of three weeks resulted in the
formation of long multi-nucleated cells (FIG. 18A). The number and
size of these multi-nucleated cells gradually increased with
induction time with multi-nucleated cells observed in all PLA
cultures at 6 weeks induction. No fusion was observed at 1-week
post-induction with MM Furthermore, multi-nucleation was not
observed in PLA cells cultured for similar time periods in CM or in
HFF cells treated with MM . To confirm the myogenic origin of these
putative myotubes, the expression of myosin was examined in PLA
cultures at 6 weeks post-induction. As shown in FIG. 18B,
multi-nucleated PLA cells at 6 weeks also expressed the myosin
heavy chain. The formation of multi-nucleated cells expressing
myosin upon induction with MM suggests that PLA cells underwent
fusion to form myotubes and further confirms their myogenic
potential in vitro.
RT-PCR Analysis
[0276] Finally, myogenic differentiation was confirmed using RT-PCR
(FIG. 19). Consistent with our immunohistochemistry data, RT-PCR
analysis confirmed the expression of MyoD1 in PLA cells induced for
1, 3 and 6 weeks in MM. In contrast, MyoD 1 expression was not
observed in PLA cells cultured in CM nor in HFF cells induced with
MM. Low levels of myosin expression were observed in induced PLA
cells at 3 weeks, while increased expression of this marker was
seen at 6 weeks post-induction with MM. Myosin was not detected in
these cells after 1 week of induction and was supportive of the
immunohistochemistry results. The expression of myosin was specific
to induced PLA cells as no expression was detected in control PLA
cells or in myo-induced HFF cells. The RT-PCR results confirm our
immunohistochemistry data and further support the myogenic
potential of PLA cells.
Quantitation of Myogenic Differentiation by PLA Cells
[0277] In order to determine the degree of myogenic differentiation
by induced PLA cells, the immunohistochemistry data was
quantitated. To do so, the number of MyoD1- or myosin-positive
cells was counted as an indicator of myogenic marker expression
level and expressed as a percentage of total PLA cells
counted.+-.the standard error of the mean (% total PLA.+-.SEM). The
number of MyoD1-positive PLA cells upon MM induction is shown in
FIG. 20. Low levels of MyoD1-positive PLA cells were observed after
1 week induction in MM (4.11.+-.0.51% total PLA cells). By 3 weeks
post-induction, 10.11.+-.3.85% of the total PLA cells were MyoD1
positive while 15.37.+-.4.33% of the total PLA cells were MyoD1
positive at 6 week post-induction. Based on the cell count, a
2.4-fold increase in the number of MyoD1 -positive cells was
observed within the first 3 weeks of induction. In contrast to the
first 3 weeks, MyoD1 expression levels only increased 1.5-fold in
the last 3 weeks of myogenic induction. The greater number of
MyoD1-positive cells in the first 3 weeks of induction relative to
the last 3 weeks may reflect the early role this regulatory factor
plays in myogenic differentiation (Megeney, et al., 1996 Genes Dev.
10:1173-1183; Seale and Rudnicki 2000 Dev. Biol. 218:115-124;
Tapscott, et al., 1988 Science 242:405-411; Weintraub, et al., 1991
Science 251:761-766).
[0278] In contrast to myo-induced PLA cells, no appreciable
myogenic differentiation was observed at any time point upon
treatment of PLA cells with CM or in HFF cells with MM, confirming
the specificity of the induction conditions. To confirm if the
increase in MyoD1 expression in PLA cells over time was
significant, statistical analysis was performed using a one-way
ANOVA. Comparison of the MyoD1 experimental values only, from 1 to
6 weeks, yielded statistical significance (P<0.001; F=18.9). In
addition, analysis of 1 and 3 week MyoD1 levels using an unpaired
t-test confirmed a significant difference (p=0.0021). A reduced
level of significance was determined between 3 and 6 weeks
(p=0.0335) and is likely a reflection of the reduced role MyoD1
plays in maturing myoblasts. Finally, the increased expression of
MyoD1 in the experimental group versus controls within each
differentiation time period was found to be statistically
significant using a one-way ANOVA (p<0.0001).
[0279] A time-dependent increase in the number of myosin-positive
PLA cells was also observed upon induction with MM (FIG. 21).
Negligible levels of myosin expression were observed at 1 week
post-induction, consistent with expression of this protein in
maturing myoblasts. Following 3 weeks induction, 3.88.+-.0.46% of
the total PLA cells counted were positive for myosin expression,
while 8.42.+-.0.71% were myosin positive at 6 weeks, a 2.2-fold
increase in the number of myosin-positive cells in the last 3 weeks
of induction. The increased number of myosin expressing cells from
3 to 6 weeks post-induction was greater than that measured for
MyoD1 and is consistent with a shift from differentiating to
maturing myogenic cells (Butler-Browne, et al., 1990 Anat. Embryol.
(Berl) 181:513-522; Thornell, et al., 1984 J. Neurol. Sci.
66:107-115). No myosin expression was observed in PLA cells
cultured with CM or in HFF cells induced with MM, confirming the
specificity of the induction conditions. Analysis of the increase
in myosin expression levels from 1 to 6 weeks confirmed statistical
significance (one-way ANOVA--P<0.0001; F=75.5). Statistical
significance was also observed using an unpaired t-test to compare
3 and 6 week myosin expression levels only (p<0.0001). As in the
MyoD1 studies, statistical analysis of both PLA and control
cultures confirmed statistical significance (one-way
ANOVA--P<0.0001). Finally, myogenic differentiation levels, as
measured using both MyoD1 and myosin expression levels, were found
to be consistent from patient to patient. Furthermore, regression
analysis did not demonstrate a significant correlation of myogenic
differentiation with patient age (MyoD1, correlation=0.27; myosin,
correlation=0.30)
Discussion
[0280] Muscle loss due to trauma, vascular insult, tumor resection
or degenerative muscle disease such as muscular dystrophy
represents a significant clinical problem with few solutions. For
focal muscle loss, vascularized muscle transplantation has been
performed, but incumbent donor site morbidity is both cosmetically
and functionally limiting. System muscle disorders, such as
degenerative muscle loss, are generally considered to be fatal
disorders resulting in progressive muscle loss, diaphragmatic
paralysis or dysfunction and eventual suffocation. Current
therapeutic approaches, such as gene therapy, have proven
unsuccessful thus far. However, recent developments in the field of
tissue engineering may allow eventual replacement or repair of both
focal and generalized muscle tissue loss.
[0281] Two cell types are generally considered candidate cells for
muscle tissue engineering: embryonic stem cells and post-natally
derived progenitor cells or stem cells. Unfortunately, ethical
issues and potential problems with cell regulation have limited the
use of embryonic stem cells (Baker, et al., 1996 Curr. Top. Dev.
Biol. 33:263-279; Dinsmore, et al., 1996 Cell Transplant 5:131-143;
Lenoir, N. 2000 Science 287:1425-1427; Rohwedel, et al., 1994 Dev.
Biol. 164:87-101; Young, FE 2000 Science 287:1424). Post-natal
skeletal muscle progenitors or satellite cells have been introduced
for the treatment of Duchenne's muscular dystrophy by Myoblast
Transfer Therapy (MTT) (Karpati, et al., 1989 Am J. Pathol.
135:27-32; Law, et al., 1988 Muscle Nerve 11:525-533; Rando, et
al., 1995 Exp. Cell Res. 220:383-389; Partridge, et al., Nature
273:306-308). Although potentially beneficial, the practical use of
satellite cells is limited primarily due to cell availability (such
cells must be harvested from viable donor muscle tissue), as well
as decreased self-renewal potential with increasing age (Rando, et
al., 1994 J. Cell Biol. 125:1275-1287; Satoh, et al., 1993 J.
Histochem. Cytochem. 41:1579-1582; Schultz and Lipton 1982 Mech.
Ageing Dev. 20:377-383). In addition to satellite cells,
mesenchymal stem cells derived from bone marrow (MSCs) have also
been reported to have myogenic capability under special culture
conditions (Ferrari, et al., 1998 Science 279:1528-1530; Wakitani,
et al., 1995 Muscle Nerve 18:1417-1426).
[0282] In this study, we show that human Processed Lipoaspirate
(PLA) cells obtained from Suctioned-Assisted Lipectomy (SAL) have
myogenic potential in vitro. Immunohistochemical and RT-PCR
analyses reveal that PLA cells induced with MM express both MyoD1
and myosin heavy chain, markers that are expressed in skeletal
muscle precursors undergoing differentiation and maturation. MyoD1
expression in PLA cells is highest during the first 3 weeks of
induction, consistent with its early role in myogenic
differentiation. A time-dependent increase in myosin is also
observed, with the highest number of myosin-positive cells observed
during the latter stages of differentiation (i.e. 3 to 6 weeks
post-induction). Such an increase may reflect the maturation of PLA
cells into myoblasts. Consistent with terminal differentiation and
myoblast fusion, long, multi-nucleated myotubes, expressing myosin,
are first observed at three weeks post-induction, with the number
and the size of these multi-nucleated cells increasing over time.
Finally, immunohistochemical quantification showed that
approximately 15% of PLA cells undergo myogenesis.
[0283] In post-natal life, mature skeletal muscle fibers cannot
regenerate if damaged. In response to muscle injury or in
individuals with chronic degenerative myopathies, satellite cells,
located between the sarcolemma and the basal lamina of the muscle
fiber, activate to become myogenic precursor cells. These
precursors divide and fuse to repair the damaged muscle (Campion,
DR 1984 Int. Rev. Cytol. 87:225-251). However, the number of
satellite cells within mature muscle is only 1-5% of the total cell
number and their self-renewal potential decreases with age (Schultz
and Lipton 1982 Mech. Ageing Dev. 20:377-383; Alameddine, et al.,
1989 Muscle Nerve 12:544-555). For focal muscle loss, vascularized
muscle transplantation has been performed, but incumbent donor site
morbidity is both cosmetically and functionally limiting.
Furthermore, for systemic muscle diseases, autologous skeletal
tissue transplantation cannot be used because of the generalized
nature of the disease process. Therefore, other cell-based
therapeutic approaches are required.
[0284] One such emerging treatment strategy is Myoblast Transfer
Therapy or MTT. Myoblast Transfer Therapy involves implanting large
numbers of healthy myoblasts. This method was first performed in
1978 and has been shown to be a promising treatment for Duchenne's
muscular dystrophy patients (Karpati, et al., 1989 Am J. Pathol.
135:27-32; Law, et al., 1988 Muscle Nerve 11:525-533; Rando, et
al., 1995 Exp. Cell Res. 220:383-389; Partridge, et al., Nature
273:306-308). Although theoretically beneficial for muscle tissue
replacement or augmentation, its success has been limited (Rando,
et al., 1994 J. Cell Biol. 125:1275-1287; Satoh, et al., 1993 J.
Histochem. Cytochem. 41:1579-1582). As an alternative,
multipotential stem cells have become promising candidates for
future cell-based therapeutic strategies since they can rapidly
proliferate in culture and retain the ability to differentiate into
several mesodermal cell types (Caplan 1991 J. Orthop. Res.
9:641-650; Pittenger, et al., 1999 Science 284:143-147).
[0285] Previous reports have demonstrated that mesodermal stem
cells can be isolated from both prenatal and post-natal organisms
(Ferrari, et al., 1998 Science 279:1528-1530; Caplan 1991 J.
Orthop. Res. 9:641-650; Elmer, et al., 1981 Teratology 24: 215-223;
Swalla, et al., 1986 Dev. Biol. 116: 31-38; Hauschka, et al., 1974
Dev. Biol. 37: 345-68; Solursh, et al., 1981 Dev. Biol. 83: 9-19;
Nakahara, et al., 1991 Exp. Cell Res. 195: 492-503; Goshima, et
al., 1991 Clin. Orthop. 274-283; Goshima, et al., 1991 Clin.
Orthop. 298-311; Benayahu, et al., 1989 J. Cell Physiol. 140: 1-7;
Bennett, et al., 1991 J. Cell Sci. 99: 131-139; Calcutt, et al.,
1993 Clin. Res. 41: 536A; Lucas, et al., 1992 In Vitro Cell Dev.
Biol. 28: 154A; Lucas, et al., 1993 J. Cell Biochem. 17E: 122).
Williams et al. has shown that post-natal cells isolated from
skeletal muscle tissue possess adipogenic, osteogenic, chondrogenic
and myogenic potential (Williams, et al., 1999 Am Surg. 65:22-26).
Moreover, several groups have demonstrated the differentiation of
Mesenchymal Stem Cells (MSCs) obtained from both human and animal
bone marrow into adipogenic, osteogenic and chondrogenic lineage
cells (Pittenger, et al., 1999 Science 284:143-147; Grigoriadis, et
al., 1988 J. Cell Biol. 106:2139-2151; Beresford, et al., 1992 J.
Cell Sci. 102:341-351; Cheng, et al., 1994 Endocrinology
134:277-286; Johnstone, et al., 1998 Exp. Cell Res. 238:265-272;
Yoo, et al., 1998 J. Bone Joint Surg. Am. 80:1745-1757). These
findings suggest that bone marrow and skeletal muscle may be a
promising source of stem cells. However, there are drawbacks to the
use of bone marrow and skeletal muscle as sources of myogenic
cells. Bone marrow procurement is painful and yields a low number
of MSCs, often requiring ex vivo expansion prior to clinical use.
Moreover, only a few stem cells can be obtained from skeletal
muscle without a functional loss to patients.
[0286] In this example we demonstrate the expression of established
myogenic markers by adipose-derived stem cells (MyoD1, myosin,
multi-nucleation), confirming and quantitating their myogenic
potential. Since adipose tissue is plentiful and liposuction
procedures are relatively safe with minimal patient discomfort,
human adipose-derived stem cells can provide an an additional
source of multi-lineage cells, together with those obtained from
bone marrow and skeletal muscle, for treating muscular
disorders.
[0287] While the expression of myogenic markers in stem cells was
shown, the exact origin of these cells cannot be confirmed. It is
possible, though unlikely, that our results are due to the
contamination of the adipose compartment with satellite cells or
myogenic precursors from a non-adipose tissue source. One
possibility is the contamination of the adipose compartment with
myogenic precursor cells from skeletal muscle. However, it is very
unlikely from a technical standpoint that the investing fascia of
the skeletal muscle could be entered with the blunt-tip liposuction
cannula. Another possibility is the contamination of the adipose
compartment by MSCs from the peripheral blood. Conflicting reports
have been presented as to the presence of MSCs in peripheral blood
(Lazarus, et al., 1997 J. Hematother. 6:447-455; Huss 2000 Stem
Cell 18:1-9), although we believe that the level of myogenesis
observed in our study is inconsistent with the low percentage of
MSCs that might be contributed by peripheral blood. Finally, while
no clear marker exists for the identification of satellite cells
and myogenic precursors, MyoD1 is one of the earliest markers
expressed during differentiation and has been used to identify
myogenic precursors (Weintraub, et al., 1991 Science 251:761-766;
Grounds, et al., 1992 Cell Tiss. Res. 267:99-104; Sassoon, D A 1993
Develop. Biol. 156:11). As shown in FIG. 16, MyoD1 expression was
not observed in non-induced PLA cultures, suggesting that our
results are not due to the presence of myogenic precursor cells in
the PLA, but are due to the myogenic differentiation of a
multi-lineage stem cell.
[0288] The goal of skeletal muscle tissue engineering is the
treatment of intrinsic skeletal muscle diseases and the loss of
skeletal muscle following trauma or ischemia. Present medical and
surgical therapies for these disorders are either ineffective or
impractical. The use of human PLA cells in these areas is
promising. Human PLA cells are plentiful, easily obtainable with
minimal morbidity and discomfort and exhibit myogenic potential. As
such, these cells may have important applications for myogenic
tissue engineering and repair.
[0289] While the degree of myogenic differentiation of PLA cells is
relatively low compared to observed levels of adipogenic and
osteogenic differentiation (Zuk, P., et al., 2001 Tissue
Engineering 7:209-226), application of exogenous factors such as
passive and active mechanical forces (Periasamy, et al., 1989
Biochem. J. 257:691-698; Vandenburgh and Kaufman 1981 J. Cell
Physiol. 109:205-214; Vandenburgh 1983 J. Cell Physiol.
116:363-371; Vandenburgh, et al., 1988 In Vitro Cell Dev. Biol.
24:166-174; Vandenburgh 1989 In Vitro Cell Dev. Biol. 25: 607-616)
and refinement of culture conditions may augment myogenic
differentiation, making these cells clinically useful.
EXAMPLE 11
[0290] The following description provides adipose-derived stem
cells which differentiate into osteogenic, chondrogenic,
adipogenic, myogenic, and neurogenic tissues. The description also
provides methods for isolating and inducing differentiation of said
stem cells.
Materials and Methods
[0291] All materials were purchased from Sigma (St. Louis, Mo.),
VWR (San Dimas, Calif.) and Fisher Scientific (Pittsburgh, Pa.)
unless otherwise stated. All tissue culture reagents were purchased
from Life Technologies (New York, N.Y.). Fetal Bovine Serum (FBS)
and Horse Serum (HS) were purchased from Hyclone (Logan, Utah) and
Life Technologies, respectively. 1,25-dihydroxyvitamin D.sub.3 was
purchased from BioMol (Plymouth Meeting, Pa.).
Cell Lines
[0292] Normal human osteoblasts (NHOsts), normal human chondrocytes
from the knee (NHCK) and a population of MSCs from human bone
marrow were purchased from Clonetics (Walkersville, Md.). The
murine 3T3-L1 preadipocyte cell line (Green and Meuth 1974 Cell
3:127-133) was obtained from ATCC (Rockville, Md.). The human
neuroendocrine cell line, PC 12, was the generous gift of Dr.
Harvey Herschman (UCLA, Los Angeles, Calif.).
Antibodies
[0293] A monoclonal antibody to human osteocalcin was purchased
from TaKaRa Shizo Co. (Japan). The polyclonal antibodies to human
osteopontin (.alpha.OP-LF123), osteonectin (.alpha.ON--LF37),
biglycan, (.alpha.BG--LF51), decorin (.alpha.DEC--LF136) and
alkaline phosphatase (.alpha.AP) were obtained from Dr. Larry
Fisher (NIH). Monoclonal antibodies to MAP2 (.alpha.MAP),
neurofilament 70 (.alpha.NF70) and .tau.-tau (.alpha.tau) were
purchased from Leinco Technologies (St. Louis, Mo.). Monoclonal
antibodies to trk-a (.alpha.TRK) and NeuN (.alpha.Neu) were
purchased from Santa Cruz Biotech (Santa Cruz, Calif.) and Chemicon
(Temecula, Calif.), respectively. Polyclonal antibodies to glial
fibrillary acidic protein (.alpha.GFAP) and were purchased from
Dako and Stressgen (Victoria, BC), respectively. Secondary
antibodies conjugated to alkaline phosphatase were obtained from
Zymed, while secondary antibodies conjugated to FITC were purchased
from BioSource (Camarillo Calif.).
Cell Harvest, Culture and Differentiation Conditions
[0294] Adipose-derived stem cells (PLA) cells were obtained from
raw lipoaspirates and cultured as described in a previous study
(Zuk, 2001 Tissue Engineering 7(2):209-226). Adipose-derived stem
cells and 3T3-L1 cells were maintained in non-inductive Control
medium (Table 5). NHOst, MSC and NHCK cells were maintained in
specialized commercial Control media (Clonetics). Adipose-derived
stem cells cells were induced toward the desired mesenchymal
lineages as outlined in Table 5. MSCs were induced using commercial
control medium supplemented with the growth factors outlined in
Table 5. 3T3-L1 cells were induced toward using Adipogenic Medium
(AM). NHOst and NHCK cells were induced using commercially
available induction media (Clonetics).
Histology, Immunohistochemistry and Indirect Immunofluorescence
[0295] Indirect Immunofluorescence (IF): PLA cells and MSCs were
processed for IF as described in Zuk, P. et al., 2001 Tissue
Engineering 7:209-226 using monoclonal antibodies to specific CD
markers (Table 6).
[0296] Histology and Immunohistochemistry (IH): To confirm
lineage-specific differentiation, differentiated cells were
processed as described in Zuk, P. et al., 2001 Tissue Engineering
7:209-226, using the following histological assays: Alkaline
Phosphatase (osteogenesis), Oil Red O (adipogenic) and Alcian Blue
(chondrogenic). In addition, PLA nodules induced toward the
chondrogenic lineage were processed by IH for the expression of
collagen type 2, keratan sulfate (KS) and chondroitin-4-sulfate
(CS), as previously described in Zuk, P. et al., 2001 Tissue
Engineering 7:209-226.
Spectrophotometric Assays
[0297] Alkaline Phosphatase (AP): Triplicate samples of PLA cells
were differentiated in Osteogenic Medium (OM) for up to 6 weeks.
Cells were washed with PBS and harvested into PBS/0.1% Triton X-100
(PBS/TX10O). AP enzyme activity was assayed using a commercial AP
enzyme kit (Sigma) and measured at an absorbance of 405nm. Total
protein in each sample was measured based on the Bradford method
(Bradford 1976 Anal. Biochem. 72:248-254) using a BCA Protein Assay
Kit (Pierce, Rockford, Ill.). AP activity was expressed a nmol
p-nitrophenol produced/minute/ug protein. The assay was calibrated
using standard p-nitrophenol solutions. Differentiated MSC and
NHOst cells were assayed as positive controls while non-induced PLA
cells were assayed as a negative control. Values are expressed as
the mean.+-.SD.
[0298] Total Calcium (Ca.sup.2+): Triplicate samples of PLA cells
were differentiated in OM as described above. Cells were washed
with PBS (no Ca2+, no Mg.sup.2+) and harvested in 0.1N HCl. Cells
were extracted for a minimum of 4 hours at 4.degree. C. and
centrifuged at 1000.times.g for 5 minutes. Total calcium in the
supernatant was determined using Sigma kit #587 and measured at
A575 nm. The assay was calibrated using calcium standard solutions.
Total protein was determined and the samples were expressed as mM
Ca/ug protein.
[0299] Differentiated MSC and NHOst cells were assayed as positive
controls, while non-induced PLA cells were assayed as a negative
control. Values are expressed as the mean.+-.SD.
[0300] Glycerol-3-Phosphate Dehydrogenase (GPDH): Triplicate
samples of PLA cells were differentiated in AM for up to 5 weeks.
The cells were harvested in 25 mM Tris-Cl, 1 mM EDTA (pH 7.5), 0.1
mM .beta.-mercaptoethanol and sonicated for 5 sec and 40 W to lyze.
The suspension was centrifuged at 10000.times.g and GPDH in the
supernatant assayed by measuring the oxidation of NADH at A340 nm,
according to the method of Wise and Green (Wise, 1979). One unit of
GPDH was defined as the oxidation of 1 nmol of NADH per minute.
Samples were normalized with respect to protein and expressed as
units GPDH/ug. Differentiated MSC and 3T3-L1 cells were assayed as
positive controls while non-induced PLA cells were assayed as a
negative control. Values are expressed as the mean.+-.SD.
[0301] Dimethyldimethylene Blue (DMMB): Triplicate samples of PLA
cells were differentiated in Chondrogenic Medium (CM) for up to 3
weeks using established micromass protocols (Ahrens, et al., 1977
Develop. Biol. 60:69-82; Denker, et al., 1995 Differentiation
59:25-34; Reddi, et al., 1982 Prog. Clin. Biol. Res. 110(Part
B):261-268). PLA nodules were harvested and assayed for the
sulfated glycosaminoglycans keratan sulfate (KS) and chondroitin
sulfate (CS) according to the method of Farndale et al.(Farndate,
et al., 1986 Biochimica et Biophysica Acta 883:173-177). The assay
was calibrated by the use of standard KS and CS solutions. Samples
were normalized with respect to protein and expressed as .mu.g KS
or CS per .mu.g protein. Non-induced PLA cells were assayed as a
negative control. Values are expressed as the mean.+-.SD.
RT-PCR Analysis
[0302] PLA cells were induced toward the five lineages outlined in
Table 5 for defmed time periods. Total cellular RNA was isolated
from the differentiated cells using a commercially available kit
(QiaEasy, Qiagen). RNA was reverse transcribed using an oligo-dT
primer and MMLV-Reverse Transcriptase (Promega, Madison, Wis.) for
60 minutes at 42.degree. C. PCR amplification was performed by the
addition of Taq buffer (Promega), 2.5 mM MgCl.sub.2, 1 mM dNTPs and
50 pmol of the appropriate primer set (Table 7). The mix was
incubated for 1 minute at 94.degree. C. and 2.5 units of Taq
polymerase (Promega) was added. PCR was performed for 40 cycles (1
minute--94.degree. C., 1 minute--57.degree. C., 1
minute--72.degree. C.: final extension--5 minutes at 72.degree.
C.). All primer sequences were determined using established GenBank
sequences and the Primer3 program. PCR reactions with primers
designed to the housekeeping gene .beta.-actin were amplified for
35 cycles as an internal control. The sequence of each PCR product
was confirmed using automated sequencing. Non-induced PLA cells
were examined as a negative control. Lineage-specific cell lines
were analyzed as a positive controls for the osteogenic, adipogenic
and chondrogenic lineages. Total human skeletal muscle and brain
RNA were reverse-transcribed and amplified by PCR as a positive
control for the myogenic and neurogenic lineages, respectively.
Western Blotting
[0303] PLA cells were differentiated and harvested in 1% SDS.
Lysates were homogenized and total protein assayed. Equivalent
amounts of protein from each lineage were denatured for 5 minutes
at 100.degree. C. in SDS Load Buffer (0.5M Tris-Cl (pH 6.8), 1%SDS,
1 mM DTT, 50% Glycerol, 1% Bromophenol Blue). Lysates were resolved
by polyacrylamide gel electrophoresis (10% separating gel, 5%
stacking gel), according to standard protocols. Proteins were
transferred overnight to nitrocellulose membranes and the membranes
blocked for a minimum of 60 minutes in Western Blocking Buffer
(WBB: 5% non-fat milk, 1.times.PBS, 0.1% Tween-20). Membranes were
incubated for a minimum of 60 minutes in WBB, supplemented with the
following antibodies: osteogenesis: .alpha.OP, .alpha.ON,
.alpha.CNI, .alpha.DEC and .alpha.BG and adipogenesis: .alpha.G4
and .alpha.LEP. Membranes were also incubated with antibodies to
the transferrin receptor and the soluble heat shock protein HSC70
as internal controls. Membranes were washed a minimum of 3 times
with PBS/0.1% Tween-20 and then incubated for 60 minutes with WBB
supplemented with the appropriate secondary antibody conjugated to
alkaline phosphatase. The membranes were washed, as described
above, and the secondary antibodies visualized using a commercial
kit (CSPD Ready-To-Use, Tropix, Bedford, Mass.) according to the
manufacturer. Non-induced PLA cells were also analyzed as a
negative control.
Neurogenic Differentiation
[0304] Immunohistochemistry: Subconfluent PLA cells were cultured
in Preinduction Medium (DMEM, 20% FBS, 1 mM .beta.-mercaptoethanol)
for 24 hours. Following preinduction, cells were induced for up to
8 hours in Neurogenic Medium (NM) and assessed by IH in order to
determine specific neurogenic lineages (Table 8).
[0305] RT-PCR: PLA cells were pre-induced for 24 hours in
Preinduction Medium, followed by induction in NM for up to 9 hours.
PLA samples were harvested, RNA isolated (QiaEasy, Qiagen) and
analyzed by RT-PCR for the expression of specific neurogenic genes
(Table 7) as detailed above.
Isolation and Analysis of PLA Clones
[0306] Clone Isolation: PLA cells were plated at extremely low
confluency in order to result in isolated single cells. Cultures
were maintained in Control medium until proliferation of single PLA
cells resulted in the formation of well-defined colonies. The
single PLA-cell derived colonies were harvested using sterile
cloning rings and 0.25% trypsin/EDTA. The harvested clones were
amplified in Cloning Medium (15% FBS, 1% antibiotic/antimycotic in
F12/DMEM (1:1)).
[0307] Confirmation of Multi-lineage capacity: Expanded clones were
analyzed for multi-lineage potential as described earlier (see
Histology and Immunofluorescence).
[0308] Molecular Characterization: Clones were analyzed by RT-PCR
for the expression of several lineage-specific genes as described
above.
Results
Stem Cells Share Many Similarities With MSCs
[0309] In order to characterize the PLA population further, cells
were examined using indirect IF and compared to a commercial
population of human MSCs. MSCs have been shown to express a unique
set of cell surface markers that can be used to help identify this
stem cell population (Table 6) (Bruder, et al., 1998 J. Orthop.
Res. 16:155-162; Cheng, et al., 1994 Endo 134:277-286; Jaiswal, et
al., 1997 J. Cell Biochem. 64:295-312; Pittenger, et al., 1999
Science 284:143-147). Like MSCs, PLA cells expressed several of
these proteins (FIG. 23), supporting the characterization of these
cells as stem cells. Approximately 100% of the PLA and MSC cultures
were positive for the expression of CD29, CD44, CD90 and CD105/SH2
with high expression levels for each of these markers being
observed in both cell populations. Both cell populations also
expressed the SH3 antigen, which, together with SH2, is considered
a specific marker for MSCs (Haynesworth, et al., 1992 Bone
13:69-80). In addition, the majority of PLA cells and MSCs were
also positive for the transferrin receptor, CD71, indicating that a
fraction of these cell populations were replicating. PLA and MSCs
did not express the haematopoietic lineage markers, CD31 and CD34.
A small number of PLA samples did show negligible staining for
CD45, although the number of CD45-positive cells did not exceed 5%
of the total PLA cell number. Unlike MSCs, no staining for the
adhesion molecule CD58 was observed in PLA cells. Flow cytometric
analysis for CD marker expression confirmed the IF results (FIG.
24). Taken together, the immunofluorescent and flow results
demonstrate several similarities in CD expression profiles between
PLA populations and bone marrow-derived MSCs.
PLA Cells Undergo Distinct Changes Upon Osteogenic Induction
[0310] In this Example, we demonstrate that PLA cells undergo
distinct proliferative, synthetic and mineralization phases upon
osteogenic induction. In order to characterize the osteogenic
capacity of PLA cells further, the proliferation of osteo-induced
PLA cells was measured and correlated to AP activity and calcium
phosphate formation (FIG. 25, Panel A). PLA cell number increased
upon initiation of osteogenic differentiation (day 1 to day 3),
however, negligible AP and Von Kossa staining was observed (FIG.
25, Panel B). A linear increase in PLA cell number was observed
from day 3 to day 9 and minimal AP staining was observed up until
day 13. PLA proliferation rates leveled off briefly between day 13
and day 15, a phenomenon that was observed in several PLA
populations. A dramatic increase in AP activity was seen between
day 15 and day 19 and the first appearance of calcium phosphate
deposits were observed by 3 weeks induction. An enhanced rate of
PLA proliferation was measured from day 15 to day 25 and coincided
with a time-dependent increase in both AP and VK staining. In
addition, the formation of multilayered nodular structures and
increased matrix synthesis were also observed during this time
period . PLA cell number decreased from day 25 onward and was
accompanied by the development of intemodular regions lacking
adherent cells, together with increased matrix mineralization.
Together, these results suggest that PLA cells may undergo distinct
proliferative and metabolic phases as osteogenic differentiation
proceeds.
Alkaline Phosphatase Activity and Time-Dependent Increase in Matrix
Mineralization
[0311] Bone formation in vivo is a complex process involving
morphogens, hormones and growth factors. Recent work has questioned
the efficacy of synthetic glucocorticoids, like dexamethasone
(Dex), in mediating osteogenesis. Glucocorticoids appear to inhibit
the action of several osteogenic genes including osteocalcin,
CBFA-1 and CNI (as reviewed in Cooper, et al., 1999 J. Endocrinol.
163:159-164). It is well established that bone tissue and
osteoprogenitor cells are targets of vitamin D action (Chen, et
al., 1983 J. Biol. Chem. 258:4350; Chen, et al., 1979 J. Biol.
Chem. 254:7491; Narbaitz, et al., 1983 Calcif. Tiss. Int. 35:177)
and this metabolite stimulates both AP activity and CNI synthesis
by human bone cell populations (Beresford, et al., 1986 Endo
119:1776-1785). Therefore, PLA cells were induced using two
osteogenic media compositions: containing either dexamethasone (Dex
at 10.sup.-7 M) or 1,25-dihydroxyvitamin D.sub.3 (VD at 10.sup.-8
M). AP activity and Ca.sup.2+ accumulation were measured over time
using commercial kits and normalized with respect to protein and/or
time.
[0312] Induced PLA, MSC and NHOst cells were measured for AP
activity and stage-specific induction levels presented in Table 9.
AP activity in PLA cells, resulting from either Dex or
VD-induction, first appeared at 3 weeks and undifferentiated PLA
cells exhibited negligible AP levels at all time points (FIG. 26,
Panel A). AP activity from 3 to 6 weeks was bi-phasic upon both Dex
and VD stimulation of PLA cells, with peak activities at days 21
and 42 and decreasing levels at day 35. VD induction of PLA cells
resulted in higher enzyme activities at 3, 4 and 5 weeks, while no
significant difference could be measured between the two induction
conditions at 6 weeks. Maximum AP levels were detected at 3 weeks
in VD-induced PLA cells, whereas no significant maximum was
detected upon Dex treatment. Moreover, PLA cells appeared to be
more responsive to VD stimulation at 3 weeks with an enhanced level
of enzyme induction being measured in these cells compared to Dex
treatment (17.2-fold induction/Dex vs. 71.3-fold induction/VD).
Like PLA cells, negligible AP activity was measured in Dex-treated
MSCs until day 21. In contrast to PLA cells, Dex stimulation of
MSCs resulted in higher enzyme activities at 3, 4 and 5 weeks. The
overall pattern of MSC AP activity observed under Dex and VD
induction was similar to VD-induced PLA cells (i.e. bi-phasic).
However, decreasing levels were measured at day 28 in MSCs rather
than day 35. Moreover, maximum enzyme activities in MSCs were
detected at a later differentiation stage (i.e. 5/6 weeks).
Finally, as observed in PLA cells, induction of MSCs from 2 to 3
weeks resulted in the greatest induction of AP activity. However,
dexamethasone, rather then VD treatment, affected enzyme levels
more in these cells (54.2-fold induction/Dex vs. 1.1-fold
induction/VD). The pattern of AP enzyme activity was dramatically
different in NHOst osteoblasts. Maximum AP levels were observed at
7 days in these cells and enzyme levels decreased after this time
point to reach minimum levels at 6 weeks. Negligible enzyme
activity could be detected in VD-treated osteoblasts at day 35 and
42. Furthermore, no significant difference in AP activity could be
measured from day 7 to day 28 under either induction condition.
[0313] Induction of PLA cells and MSCs with either Dex or VD
resulted in a time-dependent increase in matrix mineralization
(FIG. 26 and Table 10). Consistent with AP activity, PLA cells were
more responsive to VD-induction, producing a greater overall
increase in matrix mineralization (122-fold/VD vs. 56-fold/Dex),
with maximum levels detected at 6 weeks. A similar effect, was also
observed in VD-treated MSCs, although maximum levels were reached
one week earlier. As with AP activity, negligible mineralization in
both PLA cells and MSCs was observed until 3 weeks. A true effect
of induction condition was only observed in PLA cells at 6 weeks,
with VD-treated cells associated with significantly more calcium
phosphate. In contrast to PLA cells, induction condition
significantly affected mineralization in MSC samples with Dex
treatment resulting in greater calcium levels early in
differentiation (3 and 4 weeks), consistent with the AP levels
under this induction condition. This trend was reversed at 5 and 6
weeks, with VD resulting in enhanced mineral levels. Interestingly,
a decrease in Ca.sup.2+ was observed in both Dex- and VD-treated
MSCs at 28 days and appeared to correlate with the decrease in AP
activity at this time point. In contrast to PLA cells and MSCs,
maximum Ca.sup.2+ accumulation occurred at 2 weeks in induced NHOst
cells and decreased beyond this time point, consistent with
observed NHOst AP activity. Like MSCs, Dex induction resulted in
greater Ca.sup.2+ levels at all induction points with the exception
of 5 weeks. Control osteoblasts were associated with minimal levels
of Ca.sup.2+, indicating that these cells do not spontaneously
mineralize without appropriate induction. Taken together, the AP
and Ca.sup.2+ spectrophotometric data further supports the
osteogenic phenotype of PLA cells.
Osteo-Induced PLA Cells Express Osteocalcin and CBFA-1
[0314] To confirm the osteogenic phenotype of PLA cells at the
molecular level, osteo-induced PLA cells were analyzed using RT-PCR
and Western blotting. For RT-PCR analysis, PLA cells were induced
for increasing time periods in OM containing either Dex or VD. Dex
and VD-induced MSCs were also analyzed, in addition to NHOst cells.
Osteogenic differentiation did not appear to affect PLA cells, as
.beta.-actin levels did not differ significantly from control cells
(FIG. 27). Induction with Dex or VD did not significantly affect
the expression of the majority of the genes examined. However, a
dramatic effect was observed in the expression of the bone-specific
gene, OC. OC expression was not detected in Dex-treated and control
PLA cells, nor in control and Dex-induced NHOst osteoblasts.
Treatment of PLA cells with VD produced a bi-phasic OC expression
pattern. Negligible levels of OC were detected at 4 and 14 days of
induction, whereas a significant increase was observed at day 7.
Finally, a relatively consistent level of OC expression was
detected from day 21 to day 42. Elimination of Dex and replacement
with VD for the last 48 hours of PLA induction was sufficient to
overcome the effects of Dex and induce significant levels of OC
expression. In contrast to the RT-PCR results, analysis using a
gene microarray detected a slight increase in OC expression in
Dex-treated PLA cells versus non-induced controls (FIG. 27, Panel
B). OC was not specific to osteo-induced MSCs, as detectable levels
were observed in control MSCs. Dex treatment of MSCs appeared to
increase OC levels at 4 and 7 days and, like control cells, was
followed by a decrease at 2 and 4 weeks. Unlike PLA cells, VD
induction of MSCs. did not result in a bi-phasic OC expression
pattern. Rather, expression levels of this gene appeared to remain
consist across the 4 week induction period and were elevated when
compared to Dex treatment.
[0315] In addition to OC, expression of the bone-specific
transcription factor CBFA1 was observed in osteo-induced PLA cells
using RT-PCR. CBFA1 was expressed at all induction points and no
discernible effect on expression was observed upon Dex or VD
induction. In addition, CBFA1 expression was not specific to
osteo-induced PLA cells as a lower level of this gene was seen in
controls. However, an increased level of CBFA1 expression (approx.
two-fold) was measured in osteo-induced PLA cells using gene arrays
(FIG. 27, Panel B). Like PLA cells, CBFA1 was expressed in Dex- and
VD-induced MSCs at each induction point, in addition to being
expressed in undifferentiated MSC controls at a decreased level.
CBFA1 expression was more restricted in osteoblasts, detected in 4
week osteo-induced NHOst cells only. AP expression was observed at
all time points in differentiated and control PLA cells, MSCs and
NHOst cells. In addition to CBFA1 and AP, high levels of CNI were
observed in these cells. While, no appreciable difference in CNI
expression level was seen upon Dex or VD induction of PLA cells by
RT-PCR, gene array analysis confirmed a decrease in CNI level upon
osteogenic induction (FIG. 27, Panel B). In addition to OC, CBFA1,
AP and CNI, differentiated PLA cells, MSCs and NHOsts expressed
other markers consistent with bone differentiation, including OP
and ON . As seen with CNI, decreased levels of ON and OP were also
measured in Dex-treated PLA cells using arrays (FIG. 27, Panel B).
An increased expression of the transcription factor, PPAR.gamma.1
was also observed in Dex-treated PLA and MSCs when compared to
non-induced controls. In addition, a lower level of PPAR.gamma.1
was seen in the early stages of VD induction of PLA cells (day 4 to
day 14) and was followed by increased expression beyond four weeks
induction. Osteogenic induction did not result in the expression of
genes consistent with fat and cartilage differentiation
(PPAR.gamma.2 and CNII, respectively). Together, the expression of
bone-specific OC and other genes characteristic of osteogenic
differentiation in osteo-induced PLA cells further supports the
osteogenic capacity of these cells.
[0316] Finally, osteogenic differentiation by PLA cells was
confirmed at the protein level by immunofluorescent analysis and
Western blotting. Osteo-induced PLA cells (OM/Dex) were analyzed by
IF for the expression of OP, ON and OC. MSCs, induced under
identical conditions were also examined as a control. As shown in
FIG. 28, non-induced and osteo-induced PLA cells specifically
expressed both OP and ON (FIG. 28, Panel A). OP was distributed
evenly throughout control and induced cells, in addition to a
distinct perinuclear concentration. No extracellular OP staining
could be observed. A punctate intracellular pattern was also
observed for ON in both cell types. In addition, increased nuclear
staining for this protein was also observed in controls. Upon
osteo-induction, ON staining appeared to increase in areas of
concentrated cells and was expressed both intracellularly and
extracellularly. No expression of OC could be observed in
non-induced cells and was consistent with the RT-PCR results. A
small percentage of the osteogenic PLA cells appeared to express
low levels of OC intracellularly. Similar expression patterns for
these proteins were observed in MSCs. Control and induced MSCs
expressed high levels of both OP and ON. Like PLA cells, a punctate
intracellular and nuclear staining pattern were observed for ON
with the nuclear staining decreasing upon induction (FIG. 28, Panel
B). Control and induced MSCs expressed OP intracellularly. However,
unlike PLA cells, no perinuclear concentration for this protein
could be seen. Finally, consistent with the RT-PCR data, both
control and induced MSCs expressed OC.
[0317] To confirm the expression of osteogenic proteins by Western
analysis, PLA cells were maintained in OM for 7, 14 and 21 days and
lyzed. Cell lysates were analyzed for CNI, OP, ON, Decorin and
Biglycan expression. In addition, lysates were also analyzed for
the transferrin receptor (TfR) as internal controls. OC was not
assessed due to the small size of the protein (6 kDa). Osteogenic
differentiation did not appear to alter the expression of the TfR
as equivalent levels were seen in osteo-induced cells and controls
maintained for 3 weeks in Control medium. Comparable levels of CNI,
Decorin and Biglycan were seen in osteo-induced PLA cells at all
three induction periods. In addition, these proteins were also seen
in controls, suggesting that both differentiated and
undifferentiated PLA cells are associated with a proteinaceous ECM.
Like these matrix proteins, both ON and OP were seen in
differentiated cells and undifferentiated controls. However, ON
levels appeared to decrease upon initial induction of PLA cells and
returned to control levels by 3 weeks. In addition, osteogenic
induction was accompanied by a slight increase in OP at day 21.
Taken together, the immunofluorescent and Western data confirms the
expression of proteins consistent with osteogenic differentiation
by PLA cells.
PLA Cells Undergo Adipogenic Differentiation
[0318] Adipogenic differentiation is associated with the growth
arrest of preadipocytes before commitment to the differentiation
program (reviewed in Ailhaud, et al., 1992 Annu. Rev. Nutr.
12:207-233; MacDougald and Lane 1995 Annu. Rev. Biochem.
64:345-373; Smyth, et al., 1993 J. Cell Sci. 106:1-9). To determine
the correlation between PLA proliferation and adipogenic
differentiation, PLA cells were induced toward the adipogenic
lineage in AM for up to 3 weeks and cell numbers determined,
together with the degree of differentiation using Oil Red O
staining. The differentiation (i.e. appearance of intracellular
lipid vacuoles) first appeared as early as 4 days induction.
Consistent with the commitment of preadipocytes, no appreciable
increase in PLA cell number was detected over the course of
adipogenic induction (FIG. 29, Panel A) despite a time-dependent
increase in Oil Red O staining/lipid accumulation levels (FIG. 29,
Panel B). Differentiation levels were greatest in culture regions
in which the PLA cells were confluent and in contact with one
another. These results suggest that the commitment of PLA cells to
the adipogenic lineage is influenced by cell-cell contact and
coincides with growth arrest.
[0319] Adipose conversion of preadipocyte cells lines, such as
3T3-L1, is also characterized by an increase in the activity of
lipogenic enzymes, including glycerol-3-phosphate dehydrogenase
(GPDH) (Wise, et al., 1979 J. Biol. Chem. 254:273-275). PLA cells
were therefore induced with AM and the level of GPDH activity
determined. 3T3-L1 cells were similarly induced as a positive
control. Initial induction of PLA and 3T3-L1 cells (day 4 to day 7)
resulted in comparable GPDH activities and were similar to control
PLA levels (FIG. 30). Induction of PLA cells for two weeks resulted
in a decrease in enzyme activity and no significant difference
between control, induced PLA and 3T3-L1 cells was observed. This
decrease was also observed in adipo-induced MSCs. Adipogenic
differentiation beyond 2 weeks resulted in an increase in GPDH
activity specifically in induced PLA and 3T3-L1 cells. Enzyme
activity leveled off between 4 and 5 weeks and was significantly
higher than control PLA cells. The time-dependent increase in GPDH
activity correlated with the increased percentage of lipid-filled
PLA cells within adipo-induced cultures (FIG. 29) and was
consistent with adipogenic differentiation by these cells.
[0320] To confirm PLA adipogenesis, adipo-induced cells were
analyzed by RT-PCR. As shown in (FIG. 31), PLA induction resulted
in the expression of the adipose-specific transcription factor
PPAR.gamma.2. Expression of PPAR.gamma.2 was observed at day 7 and
the levels appeared to remain consistent throughout the remainder
of the 5 week induction period. No expression of this gene was
detected in non-induced PLA cells. In addition to PPAR.gamma.2, low
levels of the adipogenic genes LPL and aP2 were expressed in
induced PLA cells. Low levels of these genes were observed upon
early adipose induction (4 days) and were followed by significant
increase at 1 week. Increased levels were maintained in these cells
as far as 5 weeks induction. Basal expression of LPL and aP2 was
also observed in control PLA cells, although at a significantly
lower level than induced samples. Like osteo-induced PLA cells,
PPAR.gamma.1 was expressed in adipo-induced cells. However, the
expression pattern of this gene appeared to be distinct from
osteo-induced cells, with low expression levels observed at early
time periods (day 4 to 14) followed by increased expression from 3
to 6 weeks. Adipogenic induction of MSCs resulted in similar gene
expression patterns. Like PLA cells, PPAR.gamma.2 expression was
specific to adipo-induced MSCs and did not appear at the earliest
stages of induction. Extremely low levels of aP2 and LPL were also
observed in control MSCs and adipogenic induction resulted in a
significant increase in these genes beyond 7 days. However, in
contrast to PLA cells, PPAR.gamma.1 was not observed in control
MSCs. Rather, expression of this transcription factor was
restricted to adipo-MSCs. Furthermore, the expression pattern of
this gene paralleled that of PPAR.gamma.2, with no expression being
observed until 1 week of induction. Finally, expression of these
genes was examined in 3T3-L1 cells induced toward the adipogenic
lineage or induced via growth to confluence. Expression of aP2 and
LPL were observed in adipo-induced 3T3-L1 cells, while an apparent
inhibition of PPAR.gamma.2 expression was seen. Adipogenic
differentiation of PLA cells, MSCs and 3T3-L1 cells did not result
in the expression of the bone-specific gene, OC and the
cartilagenous marker, CNII, confirming the specificity of the
adipogenic induction conditions. In summary, the restricted
expression of PPAR.gamma.2 by adipo-induced PLA cells, together
with the increased expression of aP2 and LPL upon induction
supports the in vitro adipogenic capacity of these cells.
Chondrogenic Differentiation
[0321] Induction of PLA cells cultured under micro-mass conditions
with CM resulted in the formation of well-defined, compact nodules
consistent with those seen upon chondrogenic induction of MSCs
(Johnstone, et al., 1998 Exp. Cell Res. 238:265-272; Yoo, et al.,
1998 J. Bone Joint Surg. Am. 80:1745-1757) Chondrogenic
differentiation of PLA cells was dependent upon cell density and
induction conditions. Specifically, PLA nodules formed in induction
medium containing TGF.beta.1 alone, while the addition of
dexamethasone increased the size of TGFP1-induced PLA nodules.
Nodule formation was not observed in the presence of dexamethasone
alone. Attempts to initiate PLA chondrogenesis in monolayer culture
was unsuccessful. To assess the ECM produced by chondrogenic PLA
cells, nodules were examined by IH for the expression of CNII and
sulfated proteoglycans. PLA nodules, induced for 14 days in CM,
stained positively using Alcian Blue, which specifically identifies
sulfated proteoglycans (FIG. 32, Panel A, AB). In support of this,
14 day PLA nodules also stained positively using monoclonal
antibodies specific for keratan and chondroitin-4-sulfate (Panel A,
KS and CS, respectively). Expression of CNII was also observed in
these nodules. Alcian Blue and CNII staining were also detected in
sections of human cartilage and were not seen in high density PLA
cultures maintained in Control medium, confirming the specificity
of our histologic and IH protocols.
[0322] In addition to IH staining for KS and CS, the level of these
sulfated proteoglycans was measured using a quantitative
dimethyldimethylene blue assay (FIG. 32, Panel B). PLA nodules and
NHCK controls were predigested with papain to eliminate possible
interference by proteins and glycoproteins prior to assay. A
time-dependent increase in KS and CS was observed in PLA nodules up
to 2 weeks of chondrogenic induction. A slight decrease was
observed at 3 weeks for both PGs. Non-induced PLA cells, maintained
under high-density conditions, were also associated with an ECM
containing these proteoglycans. Furthermore, control PLA cells at 4
and 7 days induction contained more KS and CS in comparison to
induced samples. However, significantly more proteoglycan
accumulation was observed in induced PLA cells at days 14 and
21.
[0323] Treatment of PLA cells for 2 weeks in CM resulted in the
expression of several genes consistent with chondrogenesis as shown
by RT-PCR (FIG. 33). CNII expression was observed specifically in
induced PLA cells and was restricted to day 7 and 10. RT-PCR
analysis confirmed the presence of both the IIA and IIB splice
variants of CNII, although the IIB variant only is shown in FIG.
33. A low level of CNII expression was also observed upon
chondrogenic induction of NHCK controls. A similar expression
pattern to CNII was observed in induced PLA nodules using primers
designed to the amino terminus of the large proteoglycan, aggrecan
(AG). Expression of this proteoglycan was also observed using
primers to the carboxy terminus (PG). However, aggrecan expression
by the PLA nodule using the carboxy primers was observed from day 7
to day 14. In support of the PLA results, chondrogenic induction of
NHCK controls also resulted in the expression of aggrecan using
both primer sets. Finally, like CNII, aggrecan expression was
specific to induced PLA nodules and NHCK cells. In addition to
CNII, chondrogenic induction of PLA nodules resulted in the
specific expression of CNX, a marker of hypertrophic chondrocytes,
at day 14 only. In contrast to this, no expression of CNX could be
observed in NHCK controls and may be due to their derivation from
articular cartilage. PLA cells were also associated with additional
collagen types. Both induced and control PLA cells expressed CNI
and CNIII. While the majority of PLA samples examined exhibited a
restricted collagen expression pattern (day 4 only), a few PLA
samples showed expression of CNI and CNIII up to day 14. Induced
PLA cells also expressed the proteoglycans, decorin and biglycan
and the gene Cbfa-1. Expression of these genes was observed
throughout the entire induction period and was also seen in control
PLA cells. While decorin and biglycan levels remained consistent, a
slight decrease in CBFA-1 levels appeared at later stages of
induction (i.e. days 10 and 14). No expression of OC was seen at
any time point, confirming the absence of osteogenic
differentiation. Taken together, the specific expression of CNII,
aggrecan and CNX in induced PLA nodules, in addition to the
presence of keratan- and chondroitin-sulfate within the ECM
supports the chondrogenic phenotype of these cells.
PLA Cells Express Myod1, Myf5, Myogenin And Myosin Transcipts
[0324] MSCs from rat have been shown to possess myogenic potential
(Saito, 1995; Wakitani, 1995). To examine if PLA cells possess this
capacity, cells were examined for the expression of the early
myogenic regulatory factors, myoD1 and myf5, in addition to
myogenin and the myosin heavy chain, a later marker of myogenic
differentiation. Expression of myod1, myogenin and myosin was
observed at all induction points, while expression of myf5 appeared
to be restricted to 1 and 3 weeks only (FIG. 34). Consistent with
the role of myodl myogenic determination, increased levels of this
gene were observed at 1 week. Furthermore, while myogenin levels
appeared to remain consistent, a time dependent increase in
expression was detected for myosin, consistent with the expression
of this protein in mature myoblasts. In support of the PLA results,
expression of these four myogenic genes was also observed in
samples of total RNA prepared from human skeletal muscle. Therefore
the expression of these myogeic regulatory proteins indicates
possible myogenic differentiation by PLA cells.
Clones Derived from Single PLA Cells Possess Multi-Lineage
Capacity: Adipose Derived Stem Cells (ADSCs)
[0325] The presence of multiple mesodermal potential in PLA cells
is strong support for the characterization of these cells as stem
cells. However, this phenomenon may simply be due to the
contamination by lineage-specific precursors. To determine if this
is the case, PLA cells were cultured at a low enough confluence to
promote the formation of colonies derived from single PLA cells.
Several multi-lineage clones were isolated and those possessing
tri-lineage potential were termed termed Adipose Derived Stem Cells
or ADSCs. Like PLA cells, ADSCs were fibroblastic in morphology.
Following expansion, no evidence of other cell morphologies could
be observed, confirming the homogeneity of ADSC cultures. Analysis
of 500 PLA clonal isolates confirmed differentiation potential in
approximately 6% of the total number of clones examined. Seven ADSC
isolates exhibited tri-lineage potential, differentiating into
cells of the osteogenic, adipogenic and chondrogenic lineages
(Table 11). In addition to tri-lineage ADSCs, several dual-lineage
clones (O/A, O/C and A/O) and single adipogenic lineage clones were
also isolated (FIG. 35). A qualitative increase in differentiation
level, as measured by histologic staining, was observed in all PLA
clonal populations . Finally, isolation and expansion of
tri-lineage ADSCs did not alter the CD expression profile as shown
by IF, nor could differences be detected in the dual lineage clones
(FIG. 36). RT-PCR analysis of tri-lineage ADSCs confirmed their
multi-lineage potential (FIG. 36).
[0326] Induction of ADSCs in OM/VD for 2 to 4 weeks resulted in the
expression of OC and 3 and 4 weeks only, consistent with
osteo-induced PLA cells, in which no OC expression could be
detected at 2 weeks. In addition to OC, expression of ON, OP, CNI
and AP were seen at all induction points. Like PLA cells,
expression of OC was specific to induced ADSCs, nor could the fat
marker PPAR.gamma.2 be detected in both induced and control clones.
Fat induction of ADSCs for 2 and 4 weeks resulted in the specific
expression of aP2 and LPL. Interestingly, a dramatic decrease in
PPAR.gamma.2 was observed in fat ADSCs, expressed weakly at 4 weeks
only. As seen in the heterogenous PLA population, no osteogenic
differentiation was detected in adipogenic ADSCs. Finally,
expression of aggrecan, CNX, decorin and biglycan was detected upon
2 weeks of chondrogenic induction. No expression of CNII could be
observed in these cells at this induction point. Like PLA cells,
expression of aggrecan and CNX was restricted to chondrogenic
ADSCs, nor could OC expression be detected. Together with the IH
data, the RT-PCR results confirms the multi-lineage capacity of
ADSC isolates and suggests that the multi-lineage capacity of the
PLA population may be due to the presence of a putative stem cell
population.
PLA Cells May Possess Neurogenic Potential
[0327] The mesodermal embryonic layer gives rise to several
connective tissues while the overlying ectoderm is the progenitor
of multiple neural tissues and cell types. Recent evidence suggests
that MSCs can be induced toward non-mesodermal lineages,
differentiating to cells with putative neurogenic potential (Deng,
et al., 2001 Biochem. Biophys. Res. Commun. 282:148-152;
Sanches-Ramos, et al., 2000 Exp. Neurol. 164:247-256; Woodbury, et
al., 2000 J. Neurosci. Res. 61:364-370). It is possible that the
similarities between PLA and MSCs may extend beyond mesodermal
potential. Therefore, PLA cells were induced toward the
neuroectodermal lineage based on the protocol of Woodbury et al.
(Woodbury, et al., 2000 J. Neurosci. Res. 61:364-370) and examined
for the expression of neural markers, including NSE, trk-a and
MAP-2, or the expression of GFAP and GaIC, markers of astrocytes
and oligodendrocytes, respectively. To induce the PLA cells,
subconfluent cultures were pre-treated with 1 mM
.beta.-mercaptoethanol (.beta.ME) and 20% FBS for a maximum of 24
hours (pre-induction), followed by induction in serum-free medium
with 5-10 mM .beta.ME (Neurogenic Medium/NM) for up to 8 hours.
Pre-induction did not change the fibroblastic morphology of the PLA
cells (FIG. 38, Panel A--PLA/0 hrs). A morphologic change was noted
as early as 30 minutes induction in NM, with 10% of the cultures
assuming a neuronal-like phenotype. No morphological changes were
observed if FBS was added to the NM. Sixty minutes of induction
increased the proportion of neuronal-like cells to 20% of the
culture. Induction for three hours increased this phenotype to a
maximum of 70% and no significant increase was observed beyond this
induction time. NM-induced PLA cells underwent retraction, forming
compact cells bodies with multiple extensions. Cell bodies became
more spherical and cell processes exhibited secondary branches with
increasing induction time (Panel A--PLA/2 hrs vs. PLA/8 hrs).
Induction in NM resulted in significant expression of NSE, trk-a
and NeuN, consistent with the neuronal lineage (Panel B). Virtually
100% of the PLA culture stained positively for both NSE and trk-a.
In contrast to the NSE results, not all PLA cells appeared to be
NeuN positive, and may represent a more defined subset of
neuronal-like cells within the PLA culture. No expression of the
mature neuronal markers MAP-2 and NF-70 were observed, suggesting
that induced PLA cells represent an early developmental stage. In
addition, induced PLA cells did not express GalC and GFAP,
indicating that PLA cells did not differentiate into
oligodendrocytes and astrocytes, respectively. Finally, control PLA
cells did not express any neuronal, oligodendrictyic or astrocytic
markers, confirming the specificity of our induction conditions and
staining protocol.
[0328] To further assess the resulting lineage upon NM induction,
PLA cells were analyzed by RT-PCR (FIG. 38, Panel C). PLA cells
induced for 4.5 hours in NM expressed significant amounts of
nestin, an intermediate filament protein expressed in significant
quantities in neural stem cells and precursors (Lendahl, 1990).
Nestin expression was also detected in non-induced PLA cells and in
total RNA prepared from human brain. No expression of ChaT, a
marker of peripheral nerves, was observed in NM-induced cells or in
brain. In addition, NM-induced PLA cells did not express GAD65, a
marker of mature neurons and was consistent with the lack of IH
staining using antibodies to this neuronal stage (eg. MAP-2,
NF-70). As seen in the IH results, PLA cells also did not express
GFAP. Similar expression patterns were observed in PLA cells
induced for 9 hours. The expression of nestin, NSE, NeuN and trk-a,
together with the lack of ChaT, or GFAP expression suggests that
PLA cells may be capable of differentiating into an early neuronal
phenotype, characteristic of the CNS. Thus the PLA cultured in NM
can differentiate into an ectodermal lineage. Furthermore, recent
data by Lumelsky et al. (Lumelsky, N., et al. 2001 Science
292:1389) show that an embryonic stem cell can be induced to
differentiate into a cell that expresses nestin. The nestin-postive
cell was further characterized to be a pancreatic precursor cell.
Therefore, Lumelsky's data suggests that nestin-positive cells can
differentiate into both an endodermal lineage and an ectodermal
lineage. An endodermal phenotype can be further confirmed by the
additional expression of one or more of the following: insulin,
glucose transporter 2, islet amyloid polypeptide, GATA4, GATA6,
albumin, tyrosin aminotransferase.
6TABLE 5 Lineage-specific differentiation induced by media
supplementation Medium Media Serum Supplementation Control DMEM 10%
FBS none Adipogenic DMEM 10% FBS 0.5 mM isobutyl-methylxanthine
(AM) (IBMX), 1 .mu.M dexamethasone, 10 .mu.M insulin, 200 .mu.M
indomethacin, 1% antibiotic/ antimycotic Osteogenic DMEM 10% FBS
0.1 .mu.M dexamethasone, 50 .mu.M (OM) ascorbate-2-phosphate, 10 mM
.beta.-glycerophosphate, 1% antibiotic/ antimycotic Chondrogenic
DMEM 1% FBS 6.25 .mu.g/ml insulin, 10 ng/ml (CM) TGF.beta.1, 50 nM
ascorbate-2- phosphate, 1% antibiotic/ antimycotic Myogenic DMEM
10% FBS, 0.1 .mu.M dexamethasone, 50 .mu.M (MM) 5% HS
hydrocortisone, 1% antibiotic/ antimycotic Neurogenic DMEM none
5-10 mM .beta.-mercaptoethanol (NM)
[0329]
7TABLE 6 Monoclonal antibodies to CD antigens: Reported cell
specificity and distribution CD Antigen Clone Cell Specificity 29
Integrin .beta.1 MAR4 broad distribution--lymphocytes, monocytes,
granulocytes NOT on erythrocytes 31 PECAM-1 9G11 endothelial cells,
platelets, monocytes, granulocytes, haematopoietic precursors 34 --
581 endothelial cells, some tissue fibroblasts, haematopoietic
precursors 44 Pgp-1 G44-26 leucocytes, erythrocytes, epithelial
cells, platelets 45 LCA HI30 leucocytes, haematopoietic cells 58
LFA-3 L306.4 wide distribution--haematopoietic cells, endothelial
cells, fibroblasts 71 TfR H68.4 most dividing cells 90 Thy-1 5E10
immature CD34+ cells, cells capable of long term culture, primitive
progenitor cells 105 Endoglin -- endothelial cells, B cell
precursors, MSCs SH3 -- -- mesenchymal stem cells
[0330]
8TABLE 7 Oliogonucleotide primer sequences and expected PCR product
sizes Product Lineage Gene Oligonucleotide primers size BONE
Osteonectin (ON) 5' TGTGGGAGCTAATCCTGTCC 400 bp 3'
TCAGGACGTTCTTGAGCCAGT Osteopontin (OP) 5' GCTCTAGAATGAGAATTGCACTG
270 bp 3' GTCAATGGAGTCCTGGCTGT Osteocalcin (OC) 5'
GCTCTAGAATGGCCCTCACACTC 300 bp 3' GCGATATCCTAGACCGGGCCGTAG Bone
sialoprotein (BSP) 5' GCTCTAGAATGAAGACTGCTTTAATT 185 bp 3'
ACTGCCCTGAACTGGAAATC Core binding factor 5' CTCACTACCACACCTACCTG
320 bp .alpha.-1 (CBFA-1) 3' TCAATATGGTCGCCAAACAGATTC Collagen I
(CNI) 5' GAGAGAGAGGCTTCCCTGGT 300 bp (.alpha.1 chain) 3'
CACCACGATCACCACTCTTG Alkaline phosphatase 5' TGAAATATGCCCTGGAGC 475
bp (AP) 3' TCACGTTGTTCCTGTTTAG FAT aP2 5' TGGTTGATTTTCCATCCCAT 150
bp 3' TACTGGGCCAGGAATTTGAT LPL 5' GAGATTTCTCTGTATGGCACC 275 bp 3'
CTGCAAATGAGACACTTTCTC PPAR gamma1 5' GCTCTAGAATGACCATGGTTGAC 3'
ATAAGGTGGAGATGCAGGCTC PPAR gamma2 5' GCTGATATGGGTGMACTCTG 3'
ATAAGGTGGAGATGCAGGTTC PPAR delta 5' GCCAACGGCAGTGGCAATGTC 3'
TTAGTACATGTCCTTGTAGATCTC CARTILAGE Collagen II (.alpha.1' chain) 5'
ATGATTCGCCTCGGGGCTCC 260 bp 3' TCCCAGGTTCTCCATCTCTG Aggrecan 5'
GCAGAGACGCATCTAGAAATT 505 bp 3' GGTAATTGCAGGGAACATCAT Decorin 5'
CCTTTGGTGAAGTTGGAACG 300 bp 3' AAGATGTAATTCCGTAAGGG Biglycan 5'
TGCAGAACAACGACATCTCC 475 bp 3' AGCTTGGAGTAGCGAAGCAG Collagen X 5'
TGGAGTGGGAAAAAGAGGTG 600 bp 3' GTCCTCCAACTCCAGGATCA MUSCLE MyoD1 5'
AAGCGCCATCTCTTGAGGTA 500 bp 3' GCGCCTTTATTTTGATCACC Myf5 5'
CCACCTCCAACTGCTCTGAT 250 bp 3' GGAGTTCGAGGCTGTGAATC Myogenin 5'
TGGGCGTGTAAGGTGTGTAA 130 bp 3' TTGAGCAGGGTGCTTCTCTT Myosin 5'
TGTGAATGCCAAATGTGCTT 750 bp 3' GTGGAGCTGGGTATCCTTTGA NERVE CHaT 5'
TACAGGCTCCACCGAAGACT 375 bp 3' AGCAGAACATCTCCGTGGTT Synaptophysin
(SYN) 5' TTCAGGCTGCACCAAGTGTA 350 bp 3' CAGGGTCTCTCAGCTCCTTG Glial
Fibrillary Acidic Protein 5' AATGCTGGCTTCAAGGAGAC 405 bp (GFAP) 3'
CCAGCGACTCAATCTTCCTC GAD65 5' TGGCGATGGGATATTTTCTC 300 bp 3'
GCACTCACGAGGAAAGGAAC Nestin 5' GGAGTCGTTTCAGATGTGGG 240 bp 3'
AGCTCTTCAGCCAGGTTGTC
[0331]
9TABLE 8 Assessment of neurogenic differentiation by PLA cells:
antibodies and established neurogenic lineages Antibody Name
Protein Lineage NeuN Neuron-specific nuclear Neurons & Neural
protein progenitors NF-70 Neurofilament 70 kDa Neurons trk-A trk-A
(NGF receptor) Neurons MAP2 microtubule associated Neuron (mature)
protein-2 GalC galactocerebroside Oligodendrocytes GFAP glial
acidic fibrillary Astrocytes protein .tau.-tau tau Neurons,
Oligodendrocytes, Astrocytes
[0332]
10TABLE 9 Alkaline phosphatase induction levels AP Induction
("x"-fold) Cell line day 14-21 day 21-28 day 28-35 day 35-42
PLA-Dex +17.2 +1.6 -2.9 +3.2 PLA-VD +71.3 -1.3 -1.9 +1.9 MSC-Dex
+54.2 -1.2 +2.7 NS MSC-VD NS -1.5 +3.5 +1.5 NHOst-Dex -1.4 NS -2.8
-1.2 NHOst-VD +1.4 -2.2 -25.5 ND "+" upregulated enzyme induction
"-" downregulated enzyme induction NS no significant difference
detected ND Not Determined
[0333]
11TABLE 10 Quantitation of calcium phosphate levels Change in
Overall Calcium Content Cell line ("x"-fold increase/decrease)
PLA-Dex 56 PLA-VD 122 MSC-Dex 12 MSC-VD 67 NHOst-Dex ND NHOst-VD ND
ND: Not Determined
[0334]
12TABLE 11 Summary of Lineage-Specific ADSC Differentiation Lineage
Specific Differentiation O, A, C O, C A, O A, C O only A only C
only # ADSC 7 10 3 3 0 6 0 Clones
[0335]
13TABLE 12 Flow cytometric analysis of CD marker expression on
control PLA cells CD Antigen Geometric Mean CD4 2.44 CD8 2.31 CD11c
2.49 CD13 148.88 CD14 2.43 CD16 2.38 CD19 2.92 CD31 2.22 CD33 2.61
CD34 3.55 CD44 16.92 CD45 2.52 CD49d 5.33 CD56 2.66 CD61 3.68 CD62E
2.30 CD71 3.76 CD90 25.96 CD104 2.31 CD105 8.39 CD106 2.45 SH3 8.95
STRO-1 31.26 -ve 2.59
Discussion
[0336] To further confirm if PLA cells represent a mesenchymal stem
cell population, we conducted an extensive molecular and
biochemical characterization of this cell population and several
PLA clones termed Adipose-Derived Stem Cells, or ADSCs. PLA
populations were induced toward multiple mesodermal lineages,
including bone and fat, and the expression of lineage-specific
genes and proteins confirmed by RT-PCR, indirect immunofluorescence
(IF) and Western blotting. In addition, established biochemical
assays were used to measure the activities of alkaline phosphatase,
a marker for bone metabolism, the lipogenic enzyme
glycerol-3-phosphate dehydrogenase (GPDH), together with the
accumulation of sulfated proteoglycans upon chondrogenic induction.
Histological analysis and RT-PCR were also used to confirm the
multi-lineage differentiation of ADSCs. Finally, the potential of
PLA cells to differentiate into cells of the neurogenic lineage was
also examined.
[0337] We have demonstrated the multi-lineage capacity of the
heterogenous PLA cell population and its clonal derivatives, ADSCs,
obtained from human lipoaspirates. In agreement with this work, we
confirm that PLA cells and ADSC clones are capable of osteogenic,
adipogenic, chondrogenic and myogenic differentiation as shown by
the expression of several lineage-specific genes and proteins. In
addition to mesodermal lineages, PLA cells also appeared to undergo
differentiation to a lineage consistent with the neurogenic
phenotype. Taken together, the molecular and biochemical data
suggest that PLA cells may represent a putative stem cell
population that can be isolated from human adipose tissue.
PLA Cells Express a Similar Complement of CD Markers as Observed in
MSCs
[0338] Characterization of a cell population can be accomplished
through identification of unique proteins expressed on the cell
surface. Several groups have subsequently characterized MSCs based
on their expression of cell-specific proteins (e.g. STRO-1, SH2,
SH3, SH4) and "cluster designation" (CD) markers (Bruder, et al.,
1998 J. Orthop. Res. 16:155-162; Conget, et al., 1999 3. Cell
Physiol. 181:67-73; Pittenger, et al., 1999 Science 284:143-147).
This study confirms that a unique combination of cell surface
proteins is expressed on PLA cells. Moreover, both PLA and MSC
populations show similar expression profiles. Like MSCs, PLA cells
expressed CD29, CD44, CD71, CD90, CD105/SH2, SH3 and STRO-1 as
shown by IF, in addition to CD13 as confirmed by FC (FIG. 24). Like
MSCs, PLA cells did not express CDs 4, 8, 11, 14, 16, 19, 31, 33,
34, 45, 56, and 62E on the cell surface (FIG. 24). The similar CD
profiles suggest that PLA cells may be a stem cell population like
MSCs. However, the degree of similarity may indicate that PLA cells
are simply an MSC population located within or contaminating the
adipose compartment. Lipoplasty results in the rupture of multiple
blood vessels and while vasoconstrictors are used to minimize blood
loss, the processed PLA pellet may be MSCs obtained from the
peripheral blood supply (Zvaifler, et al., 2000 Arthritis Res.
2:477-488). However, there appear to be a few subtle distinctions
between PLA and MSC populations. In contrast to MSCs, no expression
of CD58 could be detected on PLA cells using IF, while expression
was seen on MSCs (FIG. 23). Furthermore, MSCs have also been
reported to express CD104, CD106 and CD140a (Bruder, et al., 1998
J. Orthop. Res. 16:155-162; Conget, et al., 1999 J. Cell Physiol.
181:67-73; Pittenger, et al., 1999 Science 284:143-147). No
expression of these CD antigens were detected on PLA cells using IF
or FC (Table 12). These differences may indicate that the PLA
population is a distinct population of stem cells. However, the
possibility that PLA cells are a clonal variant of MSCs cannot be
ruled out.
PLA Cells Undergo Osteogenesis
[0339] The mesengenic process involves: 1) proliferation of
progenitor cells, 2) commitment of these cells via the action of
specific growth factors and cytokines, 3) lineage progression into
transitory cell types expressing specific genes and 4) terminal
differentiation characterized by the cessation of proliferation and
biosynthesis of tissue-specific products (Bruder, et al., 1997 J.
Cell Biochem. 64:278-294; Caplan 1994 Clin. Plas. Surg. 21:429-435;
Jaiswal, et al., 1997 J. Cell Biochem. 64:295-312). Osteogenesis
follows this pattern closely with osteogenic precursors developing
into mitotic pre-osteoblasts and secretory osteoblasts, which lose
their mitotic potential and form the mature osteocyte (Owen, et
al., 1990 J. Cell Physiol. 143:420-430; Stein, et al., 1989
Conncet. Tissue. Res. 20:3-13). Therefore, osteogenic
differentiation is characterized by distinct phases of
proliferation, matrix synthesis/maturation and mineralization
(Owen, et al., 1990 J. Cell Physiol. 143:420-430). Consistent with
this, distinct phases were observed upon osteogenic differentiation
of PLA cells. A relatively linear growth rate was measured within
the first week of induction, a period characterized by negligible
AP activity and Ca.sup.2+ deposition. Proliferation rates increased
between day 9 and day 13 and were accompanied by the appearance of
AP by day 13. Proliferation ceased temporarily between day 13 and
day 15 and no significant increase in AP staining was observed
during this time point . An increase in cell number and enhanced AP
staining was observed beyond 2 weeks induction. These findings are
similar to the sequence of events in reported calvarial cultures in
which cells first proliferate and then show elevated levels of AP
(Aronow, et al., J. Cell. Physiol. 143:213-221; Owen, et al., 1990
J. Cell Physiol. 143:420-430). Moreover, glucocorticoids have been
postulated to stimulate the proliferation of osteogenic progenitors
(Shalhoub, et al., 1989 Biochem. 28:5318-5322; Tenenbaum, et al.,
1985 Endo 117:2211-2217. In addition to AP activity, significant
levels of calcium were seen by 3 weeks and marked the onset of the
mineralization phase in PLA cells. Increased matrix mineralization
was accompanied by a dramatic increase in AP staining and was
consistent with results found in rat calvarial cultures (Collin et
al., 1992 Calcif. Tiss. Int. 50:175-183; Shalhoub, et al., 1989
Biochem. 28:5318-5322). Increased mineralization was also
accompanied by the cessation of proliferation (day 25), followed by
a reduction in PLA cell number. This reduction was likely due to
the increase in mineral deposition and coincided with the increased
appearance of ECM and the formation of cell-free intemodular zones
. In support of this, ECM formation has been suggested to
contribute to the shutdown of proliferation by rat osteoblasts
(Owen, et al., 1990 J. Cell Physiol. 143:420-430) and rat marrow
stromal cells (Malaval, et al., 1994 J. Cell. Physiol.
158:555-572). Taken together, the results suggest that PLA cells
possess distinct proliferative, synthetic and mineralization phases
during osteogenic differentiation.
[0340] Glucocorticoid excess and/or prolonged treatment in vivo is
associated with decreased bone formation (Baylink 1983 N. Engl. J.
Med. 309:306-308), possibly through a reduction of progenitor
conversion to osteoblasts (Chyun, et al., 1984 Endo. 114:477-480).
In contrast to dexamethasone, treatment with vitamin D metabolites
restores bone mineralization and bone formation by bone-derived
cells in vitro (Beresford, et al., 1986 Endo. 119:1776-1785; Kanis,
et al., 1982 in Endocrinology of Calcium Metabolism, ed. J A
Parsons, New York: Raven Press, pp 321). Therefore, the effects of
dexamethasone and 1,25-dihydroxyvitamin D3 (VD) on PLA osteogenesis
were examined. The bone/kidney/liver isoform of AP catalyzes the
cleavage of inorganic and organic phosphates at alkaline pH. While
its precise function during in vivo osteogenesis is unclear, AP
expression levels in pre-osteoblasts and MSCs are upregulated upon
the onset of osteogenic differentiation and this enzyme thought to
play a key role in matrix mineralization through its
pyrophosphatase activity (McComb, et al., 1979 Alkaline
Phosphatase, New York: Plenum Press; Robison 1923 Biochem. J.
17:286-293; Siffert 1951 J. Exp. Med. 93:415-426). Therefore,
analysis of AP levels and matrix mineralization are important
indicators of osteogenesis. Based on this, these parameters were
measured in induced PLA samples and compared to similarly treated
MSCs and human osteoblasts as controls.
[0341] While, the overall effect of osteogenic differentiation on
AP activity and matrix mineralization appeared to be similar in PLA
cells and MSCs, the kinetics of enzyme activity and the response to
induction conditions differed depending on differentiation stage,
suggesting that these two populations may possess distinct
phenotypes. AP activity appeared in both PLA and MSC populations
between 2 and 3 weeks induction. VD treatment of PLA cells resulted
in a significantly higher level of AP activity at 3 weeks versus
Dex induction and a greater level of enzyme induction from 2 and 3
weeks (17.2 fold/Dex vs. 71.3 fold/VD). This VD effect was seen at
each differentiation stage. In contrast, the effect of induction
condition was reversed in MSCs, with Dex producing greater AP
activities at each differentiation stage. In addition to
differences in measured enzyme activity and induction level, the
kinetics of AP activity differed between PLA cells and MSCs. AP
activity in both Dex and VD-induced PLA cells was bi-phasic.
Specifically, peak AP levels were measured under both induction
conditions at 3 and 6 weeks and a decreased level detected at 5
weeks. Like PLA cells, a bi-phasic response was also observed in
Dex-treated MSCs. However, the kinetics of AP activity appeared to
be accelerated in Dex-treated MSCs with peaks detected at 3 and 5
weeks and a decrease in enzyme at 4 weeks. Moreover, a distinct
bi-phasic pattern was not observed upon VD stimulation of MSCs,
lending further support to a putative distinction between these two
cell populations.
[0342] The reason for the biphasic response in PLA and MSC
populations is unclear. Time course studies using rat calvarial
models have shown that AP activity peaks early, during the
deposition of the bony ECM, and is subsequently downregulated
(Owen, et al., 1990 J. Cell Physiol. 143:420-430; Rodan and Rodan
1984 in Bone and Mineral Research, ed. W. A., Amsterdam: Elsevier
Science Publishers, pp. 244-285; Stein, et al., 1990 FASEB J.
4:3111-3123). A similar pattern is observed in marrow stromal cell
cultures and correlates with advanced matrix mineralization and
terminal osteogenic differentiation into osteocytes (Bruder and
Caplan 1990 Bone 11:189-198; Jaiswal, et al., 1997 J. Cell Biochem.
64:295-312; Malaval, et al., 1994 J. Cell. Physiol. 158:555-572).
The drop in PLA AP activity observed from 4 to 5 weeks correlates
to increasing calcium phosphate levels within the matrix. However,
we know of no studies in which AP levels are quantitated beyond
this matrix synthesis phase. Therefore, this study may be the first
to examine AP activity in stem cells over an extended time period.
It is possible that the pattern of PLA and MSC AP activity
represents a stage-specific response to osteogenic induction. In
support of this, increases in AP have been observed in VD-treated
immature osteosarcoma cultures (Majeska, et al., 1982 J. Biol.
Chem. 257:3362) whereas a dose-dependent inhibition was detected in
more mature cells, an effect thought to represent the return of a
cell fraction to the osteoprogenitor pool or their differentiation
to osteocytes, a cell population with low AP activity. Therefore,
the decrease in AP levels in PLA and MSC samples may be due to the
terminal differentiation of a cell fraction whereas the second AP
peak could be due to the delayed development of a fraction of
osteogenic progenitor cells.
[0343] Consistent with the AP results, induction of PLA cells with
VD produced a greater overall increase in calcium levels compared
to dexamethasone. Like AP activity subtle distinctions in calcium
accumulation could be observed between PLA cells and MSCs. In
support of the AP data, matrix mineralization by PLA cells was not
observed until 3 weeks induction. Beyond this time point, Dex
stimulation did not appear to significantly affect the rate of
matrix mineralization. However, a dramatic increase was detected in
VD-treated PLA samples, with 6 week samples containing
significantly more calcium phosphate. This increased mineralization
rate occurred despite the fact that AP did not differ dramatically
between 3 and 6 week VD samples. Moreover, the decrease in AP
activity observed between 4 and 5 weeks in PLA cells did not
translate into decreases in calcium level. Rather, mineral
accumulation continued to increase in these cells. This pattern has
previously been observed in human MSCs (Jaiswal, et al., 1997 J.
Cell Biochem. 64:295-312). Like PLA cells, a time dependent
increase in mineralization was observed in MSCs with a greater
overall increase observed in VD-treated samples. The pattern of
matrix mineralization in these cells correlated well with AP
activity within the first 4 weeks of induction. Specifically,
higher AP levels in Dex-treated MSCs resulted in greater calcium
accumulation. However, between 4 and 5 weeks induction a dramatic
shift takes place, with small increases in AP activity in
VD-treated MSCs producing dramatic increases in calcium level.
Moreover, AP activity in VD-induced MSCs were significantly lower
than Dex-treated cells, yet VD treatment resulted in dramatically
more calcium, suggesting that MSCs became more sensitive to VD
induction over time. Taken together, the appearance of AP upon
osteogenic induction and the accumulation of a mineralized ECM
support the osteogenic phenotype of PLA cells. In addition,
differences observed in the kinetics and pattern of these two
markers indicates that the PLA population may be distinct from
MSCs.
[0344] During osteogenesis, osteoblasts synthesize a wide
repertoire of no proteins that are incorporated into a surrounding
ECM scaffold. The composition of the matrix, together with the
kinetics of secretion, help define the unique properties of bone
tissue and can be used to confirm osteogenic differentiation.
However, with few exceptions, the actual matrix proteins are not
unique to bone. One of these exceptions is the protein osteocalcin
(OC). A highly conserved protein containing three
.gamma.-carboxyglutamic acid residues, OC is an inhibitor of
hydroxyapatite formation in vitro, suggesting that this protein
participates in mineralization (Boskey, et al., 1985 Calc. Tiss.
Int. 37:75; Price, et al., 1976 Proc. Natl. Acad. Sci. USA
73:1447-1451). In support of this, OC is expressed by mature
osteoblasts and its expression level rises dramatically during the
mineralization phase (Collin, et al., 1992 Calcif. Tiss. Int.
50:175-183; Malaval, et al., 1994 J. Cell. Physiol. 158:555-572;
Owen, et al., 1990 J. Cell Physiol. 143:420-430; Shalhoub, et al.,
1992 J. Cell. Biochem. 50:425-440; Stein, et al., 1990 FASEB J.
4:3111-3123). While OC is considered a relatively late marker of
osteoblast differentiation, it is expressed early in bone formation
in marrow stromal cell cultures before large amounts of matrix are
synthesized (Malaval, et al., 1994 J. Cell. Physiol. 158:555-572).
Consistent with osteogenic differentiation, osteo-induced PLA cells
expressed OC. However, its expression was dependent upon the
composition of the osteoinductive medium. Specifically, osteocalcin
expression was not observed in non-induced PLA cells nor in PLA
cells induced with OM containing dexamethasone. The lack of OC
expression in Dex-treated PLA cells may be due to an inhibitory
effect associated with glucocorticoids (Cooper, et al., 1999 J.
Endocrinol. 163:159-164). In support of this, negligible levels of
OC have been observed in rat MSCs and human bone cell cultures
induced with dexamethasone (Beresford, et al., 1986 Endo.
119:1776-1785; Leboy, et al., 1991 J. Cell Physiol. 146:370-378).
Furthermore, OC was not observed upon induction of a human
osteoblast cell line, NHOst, in this study (FIG. 27). In contrast
to dexamethasone induction, OC expression was seen only upon VD
stimulation and is consistent with studies confirming VD-dependent
increases in OC expression by osteosarcoma cells (Price, et al.,
1980 J. Biol. Chem. 225:11660-11663) and its stimulation of the OC
promoter (Lian, et al., 1988 Clin. Orthop. Rel. Res. 226:276-291;
Yoon, et al., 1988 Biochem. 27:8521-8526). In addition to its
appearance upon VD induction, a distinct bi-phasic expression
pattern of OC was observed. Consistent with bone marrow MSCs, the
appearance of OC was associated with an initial stage of
differentiation, appearing as early as 4 days induction. A dramatic
increase in OC level was detected after one week induction.
Induction for 2 weeks resulted in an apparent inhibition of OC
expression and was followed by increased expression beyond three
weeks. The reappearance of OC at three weeks was coincident with
the synthesis and mineralization of the surrounding ECM and may be
supportive of the proposed role for OC in matrix calcification.
With regards to OC's biphasic pattern, a similar effect to that
observed in AP expression may be occuring: i.e. a developmental
stage-specific response to VD. In addition to VD, several other
induction agents also exert stage-specific effects on osteogenesis,
including TGF.beta. (Breen, et al., 1994 J. Cell. Biochem.
160:323-335). Similar to VD-induced PLA cells, OC was also detected
in MSCs with several differences observed in OC expression pattern
observed in these cells. First, in contrast to PLA cultures, a low
level of OC expression was observed in non-induced MSCs. The basal
level of OC expression in control MSCs was extremely low and is
consistent with reports of constitutive OC expression in cultures
of rat MSCs (Malaval, et al., 1994 J. Cell. Physiol. 158:555-572).
Second, OC expression was observed in Dex-treated MSCs. Finally,
while VD-induction increased the expression of OC in MSCs, no
apparent biphasic pattern was observed. Taken together, the
expression of bone-specific OC by osteo-induced PLA cells supports
their osteogenic capacity. In addition, the distinct pattern of OC
expression and the differential response to induction factors
observed between MSCs and PLA cells further suggests that these two
populations may possess unique phenotypes.
[0345] In addition to OC, osteo-induced PLA cells also expressed
several other genes characteristic of the osteogenic lineage,
including OP, ON, CBFA1, AP and CNI. Cbfa-1 (core binding factor-1
or Osf-2) is a transcriptional regulatory factor encoded by the
gene, CBFA1, a member of the runt domain gene family (Kania, et
al., 1990 Genes Dev. 4:1701-1713). Isolated from the nuclear
extracts of primary osteoblasts, the Cbfa1 factor has been shown to
bind to the promoters of several osteogenic genes, including OC, OP
BSP and CN type I, thus acting as a master regulator of osteoblast
differentiation (Ducy, et al., 1997 Cell 89:747-754). Moreover,
mutations to the C-terminal region of human CBFA1 is associated
with Cleidocranial dysplasia (CCD), an autosomal-dominant condition
characterized by deformities in skeletal patterning (Jones, et al.,
1997 Smith's Recongizable Patterns of Human Malformation, 5.sup.th
edition, Philadelphia: W B Saunders Company; Mondlos, et al., 1997
Cell 89:773-779; Otto, et al., 1997 Cell 89:765-771; . Consistent
with its proposed role, both Dex and VD-induced PLA cells expressed
CBFA1 at all induction points and no significant difference in
expression level was observed between the two induction conditions.
In support of the PLA results, CBFA1 was also expressed in
osteo-induced MSCs and was restricted to a late differentiation
stage in NHOst cells. Finally, both undifferentiated PLA cells and
MSCs expressed low levels of this growth factor. However,
osteogenic induction of PLA cells resulted in an approximate 2-fold
increase in CBFA1 expression as confirmed using gene arrays.
Moreover, recent studies in developing mice have suggested that
Cbfa1 is expressed in progenitors of both the osteogenic and
chondrogenic lineages (Ducy, et al., 1997 Cell 89:747-754).
Therefore, the expression of CBFA1 in control PLA cells may
represent basal gene expression in cells with a progenitor
phenotype.
[0346] Like CBFA1, the expression of OP, ON, AP and CNI was
observed in control and osteo-induced PLA cells, MSCs and NHOsts
throughout differentiation. Expression of CNI in these cell types
appeared to be equivalent under each induction condition using
RT-PCR. However, decreased expression of this gene was detected in
osteo-induced PLA cells using gene arrays and is consistent with
the proposed inhibitory effect of glucocorticoids on collagen
expression (as reviewed in Cooper, et al., 1999 J. Endocrinol.
163:159-164). As with other genes, ON and OP expression did not
appear to be affected by induction condition. Moreover, osteogenic
induction resulted in significant decreases in OP level, as
measured by microarrays, and was consistent with decreases observed
upon induction of rat bone marrow stromal cells (Malaval, et al.,
1994 J. Cell. Physiol. 158:555-572). While not restricted to
osteogenic cells, both OP and ON are found in high amounts in bone
tissue. Therefore, their expression, together with
osteoblast-specific genes like OC, supports the osteogenic capacity
of PLA cells. In addition to these osteogenic genes, osteo-induced
PLA cells expressed several other genes, including the
proteoglycans decorin and biglycan and the transcription factors
PPAR.gamma.1 and PPAR.delta..
[0347] In addition to the RT-PCR results, expression of several
proteins characteristic of osteogenic differentiation was also
observed using both IF and Western blotting. In support of the
RT-PCR data, control and osteo-induced PLA cells expressed several
proteins consistent with an osteogenic phenotype, including CNI,
decorin, biglycan, OP and ON. Significant differences in CNI,
decorin, biglycan and ON expression were not observed upon
osteogenic induction and an increase in OP expression was seen
after 3 weeks induction. Expression of ON and OP was also observed
in control and osteo-induced PLA cells using IF with differences in
intracellular expression pattern detected between the two cell
populations. Specifically, OP expression in both control and
induced PLA cells concentrated to a perinuclear location, while its
distribution appeared to be more uniform in MSC samples. This
perinuclear concentration has been observed in MSCs during
osteogenesis and is a characteristic of secreted proteins (Zohar,
et al., 1998 Eur. J. Oral Sci. 106:401-407). However, contrary to
this study, a defined perinuclear concentration of OP was not
observed in our MSC populations and may represent a clonal variant
or specific culture conditions. Rather, OP in the MSCs concentrated
to the cell surface and at cell processes. This focal distribution
has also been observed in MSCs and may indicate cell migration by
these cells during differentiation (Zohar, et al., 1998 Eur. J.
Oral Sci. 106:401-407). Similar intracellular patterns were
observed for ON in control PLA and MSC samples. In these cells, ON
was distributed throughout the cell in a fine punctate pattern and
a low level was also found in the nucleus. Osteogenic induction did
not alter this pattern in MSCs. However, the nuclear expression was
lost upon differentiation of PLA cells. Furthermore, while ON was
found in virtually all control PLA cells, not all osteo-induced PLA
cells were ON-positive. Rather, expression of this protein was
found in regions of high cell density. Finally, no expression of OC
was observed in control PLA cells, whereas a very low level was
detected in undifferentiated MSCs, consistent with the RT-PCR
findings. Osteogenic induction resulted in OC expression by a small
percentage of the osteogenic PLA cells, while a larger percentage
of osteogenic MSCs expressed this protein. The expression pattern
of OC was similar in both osteogenic PLA and MSCs: distributed
throughout the cell and concentrated at defined regions along the
cell surface. Together, with CNI, OP and ON, the expression of OC
is supportive of the RT-PCR data and further confirms the
osteogenic capacity of PLA cells in vitro.
PLA Cells Undergo Adipogenic Differentiation
[0348] The differentiation of adipocytes in culture is dependent
upon many factors, including serum, hormonal supplementation
(insulin) and pharmacologic agents (indomethacin, IBMX) (Green, et
al., 1974 Cell 3:127-133; Russell, TR 1976 Proc. Natl. Acad. Sci.
USA 73:4516-4520; Williams and Polakis 1977 Biochem. Biophys. Res.
Commun. 77:175-186). However, initiation of the adipogenic program,
in contrast to terminal differentiation, does not require such
adipogenic agents but may be dependent upon increased culture
confluence. Moreover, it is known that reversible growth arrest at
confluence must occur before most pre-adipocytes can commit to the
adipogenic lineage (Scott, et al., 1982, J. Cell Biol. 94:400-405;
Speigelman and Farmer 1982 Cell 29:53-60; Trayhurn and Ashwell 1987
Proc. Nutr. Soc. 46:135-142). As adipogenic differentiation
proceeds, a loss of proliferative potential is observed and the
irreversible loss of replication potential is a characteristic of
terminal adipocyte differentiation. To investigate if PLA cells
exhibit the same characteristics, PLA proliferation was correlated
to adipogenesis, as measured by Oil Red O accumulation. Consistent
with studies on pre-adipocyte cell lines, high levels of
differentiation occurred in confluent PLA cultures. Differentiating
PLA cells assumed a more expanded morphology and began to
accumulate intracellular lipid droplets as early as 2 weeks
induction . Differentiation proceeded with no significant increase
in PLA cell number, suggesting that cell number and growth kinetics
are linked to PLA adipogenesis (FIG. 29).
PLA--Cd Markers and ECM--Supplements
[0349] Adipogenic differentiation is accompanied by several
molecular and biochemical events, including the increase in
lipogenic enzymes that catalyze the conversion of glucose into
fatty acids and triglycerides. Glycerol-3-phosphate (G3P) is the
primary substrate for triglyceride synthesis in adipose tissue and
the adipose conversion of 3T3 cells is characterized by a dramatic
increase in the enzymatic source of G3P, glycerophosphate
dehydrogenase (GPDH) (Kuri-Harcuch, et al., 1978 J. Biol Chem.
252:2158-2160; Pairault Greem 1979 J. Biol. Chem. 76). Based on
this, GPDH activity was measured in adipo-induced PLA and 3T3-L1
cells. No significant difference in GPDH levels was detected
between differentiated cells and non-induced controls until 3 weeks
differentiation. Moreover, the initial period of differentiation
was associated with higher basal GPDH levels. The increased level
of GPDH in adipo-induced PLA cells was associated with the
appearance of Oil Red O staining (FIG. 29). Induction from 3 to 4
weeks resulted in a significant increase in GPDH in both
differentiated PLA cells and 3T3-L1 controls and coincided with
increased lipid accumulation. Continued differentiation for an
additional week did not significantly change enzyme levels in these
cell populations. A similar pattern of GPDH activity was also
observed in adipo-induced MSCs. Therefore, the increase in GPDH
enzyme activity in PLA cells induced toward the adipogenic lineage
indicates that these cells may be undergoing adipogenic
differentiation.
[0350] Like osteogenesis, adipogenesis is characterized by the
expression of a distinct set of genes that are involved in lipid
synthesis and storage. One of these genes, PPAR.gamma.2, is a
member of the PPAR nuclear hormone receptor superfamily, together
with PPAR.gamma.1 and PPAR.delta. (reviewed in (Fajas, et al., 1998
Curr. Biol. 10:165-173). PPAR.gamma.2 has been identified as part
of a heterodimeric complex (with ARF6 and the retinoid X receptor)
that acts as a key transcriptional regulator of the tissue-specific
aP2 gene (Totonoz, et al., 1995 Nucl. Acid Res.). Moreover,
PPAR.gamma.2 is expressed at high levels specifically in fat and is
induced early in the differentiation of cultured adipocyte cell
lines (Totonoz, et al., 1994 Genes Dev. 8:1224-1234; Totonoz, et
al., 1994 Cell 79:1147-1156). Consistent with this, PPAR.gamma.2
was specifically detected in adipo-induced PLA and MSC samples.
Initial differentiation (i.e. 4 days) of these cell populations was
characterized by the absence of this transcription factor and is
agreement with previous results from differentiating 3T3 adipocytes
(Totonoz, et al., 1994 Genes Dev. 8:1224-1234; Totonoz, et al.,
1994 Cell 79:1147-1156). Detectable levels of PPAR.gamma.2 were
observed after one week induction. However, expression levels were
significantly higher at this time point in adipo-induced PLA cells,
suggesting that the kinetics of PPAR.gamma.2 expression may differ
slightly between MSC and PLA populations. Distinctions in
PPAR.gamma.1 expression were also observed between PLA cells and
MSCs. A similar time-dependent increase in PPAR.gamma.1 expression
was observed in PLA cells and MSCs. However, early differentiation
(i.e. 4 days) of MSCs was associated with an absence of this
transcription factor while low levels were observed in induced PLA
cells. Moreover, no PPAR.gamma.1 was detected in control MSCs.
Finally, while detectable levels of PPAR.gamma.1 were seen in
non-induced PLA cells, adipogenic induction was associated with a
significant increase in expression, consistent with adipogenic
differentiation. PPAR.gamma.2 is associated with growth arrest and
early commitment of pre-adipose cells to the adipogenic lineage.
This period of differentiation also marks the point at which the
gene LPL is expressed (Ailhaud, et al., 1992 Annu. Rev. Nutr.
12:207-233; Fajas, et al., 1998 Curr. Biol. 10:165-173). LPL
(Lipoprotein Lipase) is ubiquitously expressed but is significantly
upregulated in adipose tissue. Through its hydrolysis of
triglycerides, LPL promotes the exchange of lipids and affects the
metabolism of several triglyceride-rich lipoproteins, including HDL
and LDL (Eisenberg, et al., 1984 J. Lipid Res. 25:1017-1058).
Consistent with its ubiquitous expression, non-induced PLA and MSC
controls expressed a low level of LPL. However, adipogenic
induction of both PLA cells and MSCs was associated with a
significant increase in the expression of this gene. This increase
was observed after one week induction and levels remained
equivalent throughout the remaining differentiation period.
Finally, extended differentiation of preadipocytes results in the
expression of the late adipogenic markers and is associated with
the accumulation of lipid within the maturing adipocyte (Ailhaud,
et al., 1992 Annu. Rev. Nutr. 12:207-233; Fajas, et al., 1998 Curr.
Biol. 10:165-173). One such late marker is the fatty acid binding
protein, aP2 (Bemlohr, et al., 1984 Proc. Natl. Acad. Sci. USA
81:468-472; Bernlohr, et al., 1985 Biochem. Biophys. Res. Comun.
132:850-855). Consistent with previous results (Bemlohr, et al.,
1985 Biochem. Biophys. Res. Comun. 132:850-855), aP2 was detected
in 3T3-L1 controls, along with LPL and PPAR.gamma.2. However,
despite its classification as a late marker in adipocytes, aP2
expression was observed throughout adipogenic induction in both PLA
cells and MSCs and levels appeared to be equivalent at each
induction point. Moreover, aP2 expression preceded that of
PPAR.gamma.2, in direct contrast to the pattern of expression
observed in pre-adipocyte differentiation ((Totonoz, et al., 1994
Genes Dev. 8:1224-1234; Totonoz, et al., 1994 Cell 79:1147-1156).
Consistent with its function in adipogenesis, extremely low levels
of aP2 were found in non-induced controls. This constitutive
expression was in agreement with the expression of aP2 in tissues
other than fat (Zezulak and Green 1985 1985 Mol. Cell Biol.
5:419-421) and is similar to the LPL results. Taken together, the
adipogenic-specific expression of PPAR.gamma.2 in adipo-induced PLA
cells, together with the upregulated expression of LPL and aP2 is
supportive of the adipogenic capacity of these cells.
[0351] Furthermore, the adipogenic capacity in combination with the
osteogenic potential of these cells suggests that PLA cells may
possess multi-lineage potential.
PLA Cells Undergo Chondrogenesis
[0352] Chondrogenic differentiation of cell lines requires high
density culture (Johnstone, et al., 1998 Exp. Cell Res.
238:265-272), duplicating the process of cellular condensation
(Fell1925 J. Morphol. Physiol. 40), in addition, to supplementation
with specific growth factors, such as TGF.beta.1, TGF.beta.3 or
BMP2 (Johnstone, et al., 1998 Exp. Cell Res. 238:265-272; Mackay,
et al., 1998 Tissue Eng. 4:415-428). Consistent with this,
aggregate culture of PLA cells in CM, containing TGF-62 1, resulted
in the formation of small, compact micromass nodules as early as 24
hours induction . Induced PLA nodules stained positively using the
stain Alcian Blue, consistent with the presence of sulfated
proteoglycans within the nodule ECM and in agreement with the
results described in Example 7 above. Alcian blue staining appeared
to concentrate more in the interior of the nodule and was apparent
as early as 3 days induction. Consistent with Alcian Blue staining,
PLA nodules also contained keratan- and chondroitin-4-sulfate, two
proteoglycans expressed in high amounts in cartilage. In support of
these results, the expression of KS and CS has also been observed
in human bone marrow MSCs induced toward the chondrogenic lineage
(Yoo, et al., 1998 J. Bone Joint Surg. Am 80:1745-1757: Yoo, et
al., 1998 Clin. Orthop. S73-81). In addition to sulfated
proteoglycans, PLA nodules also expressed collagen type II, a
collagen isoform characteristic of cartilage tissue. Finally, PLA
nodules cultured under high-density conditions and maintained in
non-inductive control medium did not form nodules and failed to
stain for any cartilage-specific histologic marker, thus confirming
the specificity of our induction conditions.
[0353] Quantitation of sulfated proteoglycans can be accomplished
using a metachromatic dimethyldimethylene blue assay (Famdale, et
al., 1986 Biochimica et Biophysica Acta 883:173-177). Consistent
with our immunohistochemical results, the DMMB assay confirmed the
presence of sulfated proteoglycans in the differentiated PLA
samples.
[0354] Moreover, a time-dependent increase in KS and CS within
chondrogenic PLA nodules was observed up to 2 weeks of induction. A
similar increase has also been observed in induced MSC cultures
(Yoo, et al., 1998 J. Bone Joint Surg. Am 80:1745-1757: Yoo, et
al., 1998 Clin. Orthop. S73-81) and suggests that PLA cells have
accumulated an ECM characteristic of cartilage tissues. PG levels
decreased slightly beyond 2 weeks induction and may represent
remodeling of the cartilagenous ECM. Non-induced PLA cells,
maintained under high-density conditions, were also associated with
an ECM containing these proteoglycans. Moreover, basal PG levels
were greater than induced PLA sample at 4 and 7 days. However,
significantly more proteoglycan accumulation was observed in
induced PLA nodules at days 14 and 21. The significant accumulation
of KS and CS within the ECM of induced PLA nodules, together with
the histological results suggests that PLA cells also possess in
vitro chondrogenic capacity when cultured under high-density
conditions.
[0355] Induction of PLA cells in CM resulted in the expression of
several genes consistent with chondrogenesis. CNII expression was
observed specifically in induced PLA cells and was restricted to
day 7 and 10 and supported our immunohistochemical results. A
restricted expression pattern similar to CNII was observed in PLA
nodules using primers designed to the amino terminus of aggrecan
(AG), a large proteoglycan expressed in high amounts in cartilage.
Expression of aggrecan was also observed in PLA samples using
primers to the carboxy terminus (PG). However, in addition to
expression at days 7 and 10, PG was also detected at day 14 in
these nodules. In support of the PLA results, expression of
aggrecan in induced NHCK nodules was detected using both amino and
carboxy primer sets. Like CNII, the expression of aggrecan was
specific to induced PLA and NHCK nodules. In addition to CNII,
chondrogenic induction of PLA cells resulted in the restricted
expression of CNX, a marker of hypertrophic chondrocytes.
Expression of CNX was detected at day 14 and suggests that PLA
nodules undergo hypertrophy over time. Induced NHCK samples also
expressed CNX, although at a lower level. PLA nodules were also
associated with additional collagen types, including CNI and CNIII.
While the majority of PLA samples examined exhibited a restricted
collagen pattern (day 4 only), CNI and was detected in a few PLA
samples up to day 14. The expression of CNI has also been observed
in human MSC nodules by fibroblastic cells located in the outer
nodule, leading researchers to suggest that this region is
comprised of perichondrium-like cells involved in the
differentiation process (Yoo, et al., 1998 J. Bone Joint Surg. Am
80:1745-1757: Yoo, et al., 1998 Clin. Orthop. S73-81). In support
of this, perichondrium-like cells have also been observed in
high-density embryonic chick limb-bud cell cultures and cell
aggregates (Osdoby and Caplan 1979 Devel. Biol. 73:84-102;
Tachetti, et al., 1987 J. Cell Biol. 106:999-1006). Therefore, the
continued expression of CNI in select PLA samples may be due to the
presence of a similar cell population.
[0356] Induced and control PLA cells also expressed the
proteoglycans, decorin and biglycan and the gene CBFA1. Decorin and
biglycan make up the majority of the small leucine-rich
proteoglycans within the cartilagenous ECM and their expression
within PLA nodules further supports the chondrogenic phenotype. In
addition to its expression during osteogenesis, a role for CBFA-1
in the hypertrophy and terminal differentiation of chondrocytes has
recently been confirmed (Enomoto, et al., 2000 J. Biol. Chem.
275:8695-8702. Therefore, the expression of CBFA-1, together with
CNX, may indicate terminal differentiation of PLA cells within the
nodule. Chondrocyte hypertrophy may also precede the ossification
of cartilagenous tissue. However, expression of bone-specific OC by
chondrogenic PLA or NHCK cells was not seen at any time point,
confirming the absence of osteogenic differentiation within the PLA
nodule. Interestingly, micromass culture of MSCs in CM did not
result in the formation of nodules and was not examined. Taken
together, the specific expression of CNII, aggrecan and CNX in
induced PLA nodules, in addition to the presence of keratan- and
chondroitin-4-sulfate within the ECM supports the chondrogenic
phenotype of these cells. Moreover, the chondrogenic capacity of
PLA cells, together with their osteogenic and adipogenic potential,
further supports the multi-lineage capacity of these putative stem
cells.
PLA Cells Undergo Myogenic Differentiation
[0357] RT-PCR analysis of PLA cells induced toward the myogenic
lineage confirmed the expression of several myogenic genes,
including the transcription factors MyoD1, myogenin and myf5, in
addition to the muscle-specific protein, the myosin heavy chain.
Determination of the myogenic lineage is thought to be controlled
at the transcriptional level by MyoD1 and myf-5, which are
expressed in proliferating myoblasts (Atchley, et al., Proc. Natl.
Acad. Sci. 91:11522-11526; Lassar, et al., 1994 Curr. Opin. Cell
Biol. 6:432-442; Weintraub, et al., 1994 Genes Dev. 15:2203-2211),
whereas execution of the myogenic differentiation program is
controlled by myogenin and MRF4 expression (emerson, et al., 1993
Curr. Opin. Genet. Dev. 3:265-274; Olson, et al., 1996 Cell 5:1-4).
Finally, terminal differentiation of myoblasts can be confirmed
through the expression of the myosin heavy chain. Consistent with
these findings, the expression of myf5 was restricted to the first
3 weeks of myogenic PLA induction while increased MyoD1 expression
was detected within the first week relative to the remainder of the
differentiation period. Myo-induced PLA cells also expressed
myogenin at relatively equivalent levels throughout the 6 week
induction period. Finally, increased expression of the myosin heavy
chain was detected at 6 weeks induction and suggests that PLA cells
underwent of terminal differentiation. The expression of myf5 and
myogenesis further supports this potential and, together with the
osteogenic, adipogenic and chondrogenic capacity of PLA cells,
indicates their potential for differentiation to multiple
mesodermal lineages.
PLA Cells May Possess Neurogenic Potential
[0358] True pluripotency of a stem cell is achieved upon
differentiation to cells from distinct embryologic lineages. Recent
reports have documented the differentiation of MSCs to neural cells
(Deng, et al., 1994 Genes Devel. 8:3045-3057; Kopen, et al., 1999
Proc. Natl. Acad. Sci. USA 95:3908-3913; Sanchez-Ramos, et al. Exp.
Neurol. 164:247-256; Woodbury, et al., 2000 J. Neurosci. Res.
61:364-370) and neural stem cells (NSCs) to haematopoietic cells
(Bjornson, et al., 1999 Science 283:534-537), suggesting that stem
cell populations may not be as restricted as previously thought.
Based on these findings, we investigated if PLA cells could be
induced beyond their putative multilineage mesodermal capacity. To
this end, PLA cells were cultured in a medium known to induce
neurogenic differentiation (Vescovi, et al., 1999 Exp. Neurol.
156:71-83; Woodbury, et al., 2000 J. Neurosci. Res. 61:364-370) and
differentiation assessed by staining for neural markers, including
NSE, trk-a and MAP-2 or for the expression of GFAP and GaIC,
markers of astrocytes and oligodendrocytes, respectively. The
morphologic and histologic data suggest that PLA cells, like MSCs,
possess neurogenic potential in vitro. Induction of PLA cells in NM
for a minimum of 30 minutes resulted in a dramatic change in
morphology with cells assuming a neuronal-like phenotype.
NM-induced PLA cells underwent retraction, forming compact cells
bodies with multiple extensions. Cell bodies became more spherical
and cell processes exhibited secondary branches with increasing
induction time. A time-dependent increase in the proportion of PLA
cells with this phenotype was observed in all induced PLA cultures.
Similar morphologic changes have been observed upon neurogenic
induction of MSCs from both rodents and human (Woodbury, et al.,
2000 J. Neurosci. Res. 61:364-370). Moreover, this PLA morphology
was similar to that observed upon NGF stimulation of PC 12 cells, a
neuroendocrine cell line similar to primary sympathetic neurons
.
[0359] The observed morphologic changes in neuro-induced PLA cells
were accompanied by the increased expression of neuron-specific
markers, such as NSE, trk-a and NeuN, and did not result in
expression of markers for astrocytes and oligodendrocytes.
Furthermore, expression of these markers was also observed in PC12
cultures, suggesting that PLA cells may be assuming a neuronal-like
phenotype. In support of the PLA results, increased expression of
NSE, a neuron-specific enolase, and trk-a has been observed upon
induction of MSCs with .beta.-ME, with approximately 100% of the
neuronal-like MSCs positive for these markers (Woodbury, et al.,
2000 J. Neurosci. Res. 61:364-370). Like the MSC studies, all PLA
cells exhibiting a neuronal phenotype expressed significant levels
of NSE and trk-a. In addition to NSE, expression of NeuN has also
been used to identify neuronal development in neurogenic precursors
and MSCs (Sanchez-Ramos, et al., 2000 Exp. Neurol. 164:247-256).
Specifically, NeuN is expressed in post-mitotic neurons (Sarnat, et
al., 1998 Brain Res. 20:88-94) and its appearance is thought to
coincide with the withdrawl of the developing neuron from the cell
cycle and/or the initiation of terminal differentiation (Mullen, et
al., 1992 Development 116:210-211). The expression of NeuN within
the neuronal-like PLA cells, together with the presence of NSE and
trk-a, further supports the development of a neuronal phenotype in
PLA cells. Moreover, the expression of NeuN may indicate the
development of a post-mitotic neuronal phenotype. In contrast to
NSE, trk-a and NeuN, expression of the mature neuronal markers,
tau, MAP-2 and NF-70, was not observed , suggesting that induced
PLA cells represent an early developmental stage. Consistent with
this, MAP-2 expression in induced MSC cultures has not been
observed by several groups and may reflect the induction conditions
used or the need for prolonged induction time (Deng, et al., 2001
Biochem. Biophys. Res. Commun. 282:148-152; Sanchez-Ramos, et al.,
2000 Exp. Neurol. 164:247-256).
[0360] Finally, the putative neuronal potential of PLA cells was
confirmed using RT-PCR. Consistent with the immunohistochemistry
results, no expression of GFAP could be detected, supporting the
restriction of induced PLA cells to the neuronal lineage. In
addition, PLA cells were examined for the expression of the gene
nestin. Nestin, an intermediate filament protein, has been detected
in high amounts in CNS stem cells (Lendahl, et al., 1990 Cell
60:585-595), within the developing neural tubes of mice
(Frederikson and McKay 1988 J. Neurosci. 8:1144-1151) and in MSCs
induced toward the neurogenic lineage (Sanchez-Ramos, et al., 2000
Exp. Neurol. 164:247-256). Differentiation of neural precursors
results in a decrease in nestin expression levels, indicating that
this protein can be used as a marker of a progenitor phenotype
(Johe, et al., 1996 Genes Dev. 10:3129-3140; Lendahl, et al., 1990
Cell 60:585-595). The expression of nestin in control PLA cultures
is supportive of the presence of neurogenic precursors within the
PLA. However, differentiation of PLA cells did not result in an
appreciable decrease in nestin expression. This may be due to two
possibilities: 1) the differentiation of PLA cells into a
neurogenic progenitor population only or 2) the differentiation of
PLA cells into an early neuronal-like cell that retains nestin
expression. In support of the latter, nestin expression was also
detected in NGF-treated PC12 controls. Based on this, together with
the expression of NeuN, NSE and trk-a in induced PLA cells, leads
us to favor the latter possibility and further studies are
warranted. Like our IH results, RT-PCR analysis failed to detect
expression of a mature neuronal marker (GAD65), a marker detected
in PC12 controls and brain. It is possible that additional growth
factors or a prolonged induction period may be required to induce
PLA cells into a more mature stage. Finally, induction of PLA cells
with NM appeared to restrict their development to cells
characteristic of the CNS, as the cells did not express ChaT, a
specific marker of peripheral nerves. Nestin expression has also
been observed in non-induced MSCs, in addition to myogenic cells,
newly formed endothelial cells, epithelial cells of the developing
lens and hepatic stellate cells. This broad distribution indicates
that nestin cannot be used as a neurogenic precursor marker per se.
However, combined with the expression of additional neuronal
markers, such as NeuN, the possibility that PLA cells are forming
precursors of the neuroectodermal lineage is strengthened.
ADSC Clonal Isolates Demonstrate Multi-Lineage Capacity
[0361] Multi-lineage differentiation by PLA cells may result from
the commitment of multiple lineage-specific precursors rather than
the presence of a pluripotent stem cell population within adipose
tissue. Therefore, multi-lineage differentiation by clonal isolates
derived from single PLA cells is critical to the classification of
PLA cells as a source of stem cells. In support of this, single PLA
cell isolates expanded in culture exhibited multi-lineage capacity
in vitro, staining positively for alkaline phosphatase
(osteogenesis), Oil Red O (adipogenesis) and Alcian Blue
(chondrogenesis). Clonal analysis resulted in the isolation of
several lineage combinations, including tri-lineage (osteogenic,
adipogenic and chondrogenic), dual-lineage (osteogenic/adipogenic,
osteogenic/chondrogenic) and single lineage (adipogenic only). The
tri-lineage clones were subsequently termed Adipose Derived Stem
cells (ADSCs) and were analyzed for multilineage potential using
RT-PCR. Consistent with multilineage capacity ADSCs expressed
several genes characteristic of osteogenesis (OC, ON, OP, CNI and
AP), adipogenesis (PPAR.gamma.2, aP2 and LPL) and chondrogenesis
(AGG, CNX, decorin and biglycan). Furthermore, several tri-lineage
ADSCs also expressed the neuronal marker trk-a using IH (FIG. 39).
Based on these results, the expression of multiple lineage-specific
mesodermal genes by ADSCs suggests that these isolated clones
possess multipotentiality and may be considered stem cells.
EXAMPLE 12
[0362] The following provides a description of molecular and
biochemical characterization of adipose-derived stem cells.
Materials and Methods
[0363] All materials were purchased from Sigma (St. Louis, Mo.),
VWR (San Dimas, Calif.) and Fisher Scientific (Pittsburgh, Pa.)
unless otherwise stated. All tissue culture reagents were purchased
from Life Technologies (New York, N.Y.). Fetal Bovine Serum (FBS)
and Horse Serum (HS) were purchased from Hyclone (Logan, Utah) and
Life Technologies, respectively.
Antibodies
[0364] Monoclonal antibodies to CD29, CD34, CD44, CD45, CD58, CD90,
CD104, CD105 and CD140a were purchased from Pharmingen (Bedford,
Mass.). Monoclonal antibodies to CD31 and CD71 were obtained from
R&D Systems (Minneapolis, Mass.) and Zymed (S. San Francisco,
Calif.), respectively. FITC and PE-conjugated anti-CD antibodies
used for flow cytometry (FC) were purchased from Pharmingen. A
monoclonal antibody to the SH3 antigen was produced from the SH3
hybridoma (ATCC). The Stro-1 hybridoma supernatant was the generous
gift of Dr. John Fraser (UCLA). Monoclonal antibodies to the human
collagens 1 (.alpha.CNI) and 4 (.alpha.CNIV) were purchased from
Sigma. Monoclonal antibodies to human collagen 3 (.alpha.CNIII) and
collagen 5 (.alpha.CNV) was purchased from Biogenesis (Kingston,
N.H.).
Cell Harvest Culture and Differentiation Conditions
[0365] Processed lipoaspirate (PLA) cells were obtained from raw
lipoaspirates and cultured as described previously (Zuk, P. et al.,
2001 Tissue Engineering 7:209-226). PLA cells were maintained in
non-inductive Control medium (Table 13) while MSCs were maintained
in specialized Control medium (Clonetics). PLA cells were induced
toward the desired mesenchymal lineages using the induction media
outlined in Table 13. MSCs were induced using the commerical
control medium supplemented with the same growth factors as
outlined in Table 13.
PLA Clonal Isolation and Analysis: Adipose-Derived Stem Cells
(ADSCs)
[0366] ADSC Isolation: PLA cells were plated at extremely low
confluence in order to result in isolated single cells. Cultures
were maintained in Control medium until proliferation of single PLA
cells resulted in the formation of well-defined colonies. The
single PLA-cell derived colonies were termed Adipose Derived Stem
Cells (ADSCs). ADSCs were harvested using sterile cloning rings and
0.25% trypsin/EDTA. The harvested ADSCs were amplified in Cloning
Medium (15% FBS, 1% antibiotic/antimycotic in F12/DMEM (1:1)).
Indirect Immunofluorescence
[0367] Indirect Immunofluorescence (IF): PLA cells, ADSCs and MSCs
were processed for IF as described previously (Zuk, P. et al.,
2001, Tissue Engineering, 7:209-226) using the anti-CD marker
antibodies outlined in Table 14. In addition, PLA cells were
incubated with supernatants produced from the STRO-1 and SH3
hybridoma cell lines. To determine the cell characteristics of
differentiated PLA cells and MSCs, cells were induced toward either
the osteogenic lineage for 3 weeks or the adipogenic lineage for 2
weeks and incubated with anti-CD antibodies. The differentiated
cells were also analyzed using antibodies to human collagens 1, 4
and 5.
Flow Cytometry
[0368] PLA cells from multiple donors, in addition to MSCs, were
cultured for 3 weeks in Control medium and analyzed for the
expression of CD antigens by flow cytometry (FC) as described
previously (Zuk, P. et al., 2001, Tissue Engineering, 7:209-226).
PLA cells were also induced in either OM or AM for 2 weeks prior to
analysis. Briefly, cells were harvested a 80% confluence with
trypsin/EDTA, washed and resuspended in Flow Cytometry Buffer (FCB)
at a concentration of 1.times.10.sup.6 cells/ml. One hundred
microliters of the cell preparation (1.times.10.sup.5 cells) were
stained with saturating concentrations of FITC-conjugated (anti-CD
14, 44, 45 61, 71, 90 and 105) or PE-conjugated (anti-CD 13, 16,
31, 34, 44, 49d, 56, 62E and 106) antibodies for 1 hour at
4.degree. C. Cells were also incubated with isotype-matched IgG's
as a control to assess autofluorescence. After incubation, the
cells were washed three times with FCB and resuspended for
analysis. Flow cytometry was performed on a FACStar flow cytometer
(Becton Dickson). The geometric means, calculated from the absolute
numbers of cells per 10,000 events are shown in Table 12.
Results
PLA Cells Share Many Similarities with MSCs
[0369] The results described in this example demonstrate the
mutli-lineage potential of adipose-derived stem cells and their
clonal isolates. In order to characterize the PLA population
further, cells were examined using indirect IF and FC and compared
to a commercial population of human MSCs. MSCs have been shown to
express a unique set of cell surface markers that can be used to
help identify this stem cell population (Table 14) (Bruder, S. P.
et al., 1998, Clin. Orthop., S247-256; Conget, P. A. et al., 1999,
J. Cell Physiol, 181:67-73; Pittenger, M. F et al., 1999, Science,
284:143-147.) Like MSCs, PLA cells expressed several of these
proteins (FIGS. 23 and 24), supporting the characterization of
these cells as stem cells. Approximately 100% of the PLA and MSC
cultures were positive for the expression of CD29, CD44, CD90 and
CD105/SH2 with high expression levels for each of these markers
being observed in both cell populations. Both cell populations also
expressed the SH3 antigen, which, together with SH2, is considered
a specific marker for MSCs (Haynesworth, S. E. et al., 1992, Bone,
13:69-80.)
[0370] In addition, the majority of PLA cells and MSCs were also
positive for the transferrin receptor, CD71, indicating that a
fraction of these cell populations were replicating. PLA and MSCs
did not express the haematopoietic lineage markers, CD31 and CD34.
A small number of PLA samples did show negligible staining for
CD45, although the number of CD45-positive cells did not exceed 5%
of the total PLA cell number. Unlike MSCs, no staining for the
adhesion molecule CD58 was observed in PLA cells. The IF results
were subsequently confirmed by FC (FIG. 24, Panel B). Both MSC and
PLA cells showed similar profiles, comprised mainly of a population
of relatively small, agranular cells (FIG. 24, Panel A). However, a
greater proportion of the PLA population did appear to contain
larger, granular cells (see upper right corner), while a larger
proportion of the MSC population contained smaller agranular cells.
FC confirmed the expression of CD44, CD71, CD90 and CD105 on both
PLA and MSCs and did not detect significant levels of CD3 1, CD34,
CD45 and CD104. In addition to these markers, FC also measured
expression of CD13, CD49d, SH3 and STRO-1 on PLA cells yet did not
detect expression of CDs 4, 8, 11, 14, 16, 19, 33, 56, 62E and 106
(Table 12). Taken together, the immunofluorescent and flow results
demonstrate several similarities in CD expression profiles between
PLA populations and bone marrow-derived MSCs.
Phenotypic Characterization of Differentiated PLA Cells
[0371] Differentiation of stem cells may alter the expression of
several cell surface and intracellular proteins. In order to
characterize differentiated PLA cells, cells from the same patient
were maintained in non-inductive Control medium or were induced
toward the osteogenic and adipogenic lineages. Control and
differentiated PLA cells were subsequently analyzed by IF and
compared to MSCs as a control. The results are presented in Table
15. PLA cells induced for 3 weeks in OM underwent increased
proliferation and did not show any significant differences in CD
marker profile when compared to undifferentiated PLA cells. Like
control PLA cells, expression of CD45 was not observed in
osteogenic PLA cells while significant expression of CD44 and CD90
was detected (FIG. 40, Panel A: PLA--Bone).However, in contrast to
control cells, osteogenic differentiation resulted in localized
areas of CD34 expression. Like PLA cells, the CD marker profile of
control and osteo-induced MSCs was similar, with the exception of
CD34 and CD45. As shown in FIG. 40, Panel B, expression of CD34 and
CD45 was not observed in control MSCs. However, a slight increase
in CD34 expression level was observed upon induction in OM while an
increased number of CD45-positive cells were detected.
[0372] To induce adipogenic differentiation, PLA cells were
maintained for a minimum of 2 weeks in AM. In order to correlate CD
marker expression to cell morphology, fluorescent micrographs were
overlaid with light micrographs (inset pictures). Induction of PLA
cells with AM resulted in an expanded cellular morphology and the
accumulation of multiple, intracellular lipid vacuoles, consistent
with adipogenesis. These lipid-containing PLA cells were considered
to be mature adipocytes (white arrows) (FIG. 41, Panel A:
PLA--Fat). Like osteogenesis, adipogenic differentiation appeared
to result in slightly increased CD34 levels in both fibroblastic
and lipid-containing PLA cells (FIG. 41, Panel A). In addition, a
negligible fraction of the adipogenic PLA cultures contained
CD45-positive cells. However, these cells did not contain the lipid
vacuoles characteristic of mature adipocytes (CD45--inset). A
significant level of CD44 was also detected in adipogenic PLA
cultures. However, lipid-filled PLA cells appeared to express lower
levels of CD44 in comparison to their fibroblastic counterparts
(open arrows--CD44-ve adipocytes, filled arrows--CD44+ve cell).
Furthermore, CD44 staining levels varied among the fibroblasts,
ranging from intense to little or no CD44 expression. A similar
restriction was also observed for CD90 with all fibroblasts
expressing this protein at comparable levels. Like PLA cells,
adipo-induced MSCs expressed CD44 and CD90 and showed increased
staining for CD34 and CD45 (FIG. 41, Panel B and Table 16).
However, unlike adipo-induced PLA cells, both fibroblastic and
lipid-filled MSCs (filled vs. open arrows, respectively) appeared
to express CD44 and CD90 at similar levels.
[0373] In order to confirm the immunofluorescent results, FC was
performed on non-induced and differentiated PLA cells and the
geometric means calculated for each CD marker protein (FIG. 42)
(Table 16). Osteogenic differentiation did not appreciably change
the size and granularity of the PLA populations (FIG. 42, Panel A).
Adipogenesis, however, resulted in a significant increase in the
size and granularity of the PLA population, likely a reflection of
the expanded cellular morphology and the formation of intracellular
lipid vacuoles. Consistent with the immunofluorescent results,
control PLA cells were negative for CD34 and CD45 expression (Panel
B), nor could expression of CD14, CD16, CD31, CD34, CD45, CD56,
CD61, CD62, CD105 or CD106 be measured in these cells (Panel B).
Differentiation appeared to increase CD34 and CD61 expression with
a greater increase being observed upon osteogenic differentiation.
The increased CD34 expression was consistent with the increased
staining observed upon IF processing (FIG. 40). Slight increases in
CD56 and CD49d were detected specifically in osteo-induced PLA
cells. The expression of CD56 in adipo-induced PLA cells did not
differ significantly from controls, while a decrease in CD49d
expression was detected upon adipogenesis. FC also confirmed the
expression of CD13, CD44 and CD90 in control cells (FIG. 42, Panel
C.). Osteogenesis significantly increased expression of these
markers and a further increase in CD90 was measured in
adipo-induced PLA cells. Finally, adipogenic differentiation
resulted in a decreased expression of CD13 and CD44 to below that
of undifferentiated PLA cells. The decrease in CD44 was consistent
with the IF results, in which lower expression levels were seen in
lipid-containing PLA cells.
[0374] Mesodermally-derived cells, such as adipocytes and
osteoblasts are associated with extensive extracellular matrices
(ECMs). To assess the expression of ECM proteins in differentiated
PLA cells, adipogenic and osteogenic PLA cells were analyzed by IF
for the expression of ECM collagens. The results are summarized in
Table 17. The majority of undifferentiated PLA cells expressed
collagen types 1 and 3 (CNI, CNIII) (FIG. 43, Panel A:
PLA--Control). CNI and CNIII in fibroblastic PLA cells were
restricted to a defined perinuclear concentration and was evenly
distributed throughout while cells with an expanded morphology.
Contrary to CNI and CNIII, the expression of collagen types 4 and 5
(CNIV, CNV) was restricted to defined culture regions of
concentrated PLA cells and matrix formation. The expression
patterns of CNIV and CNV were fibrillar in nature, consistent with
the secretion of these proteins into the extracellular space
surrounding these cells.
[0375] Adipogenic differentiation of PLA cells inhibited both CNI
and CNV expression (FIG. 43, Panel A: PLA--Fat). Differentiation
also appeared to alter the intracellular expression pattern of
CNIII, redistributing it evenly throughout the mature PLA
adipocyte. In addition, a lower level of CNIV expression was
observed in adipogenic PLA samples, with the majority of the CNIV
fluorescence observed in lipid-filled PLA cells (see arrows;
inset). While an inhibition of CNV expression was also observed
upon osteogenic induction, no significant difference in CNI
expression pattern could be detected for this lineage (FIG. 43,
Panel A; PLA--Bone). Finally, osteogenesis appeared to increase
CNIV expression levels and resulted in a more widespread
distribution of this collagen within the PLA culture. The
expression patterns of CNI, CNIII, CNIV and CNV in control MSCs
were found to be similar to control PLA cells (FIG. 43, Panel B:
MSC--Control). Like PLA cultures, CNIII and CNIV were detected in
adipo-induced MSC samples. However, CNIV appeared to be restricted
to extracellular fibrils rather than an intracellular distribution
(FIG. 43, Panel B: MSC--Fat, see arrows). In contrast to adipogenic
PLA cultures, induction toward this lineage did not inhibit
synthesis of CNI and CNV by MSCs. Rather, these collagens could be
detected within extracellular fibrils and weak CNI expression could
also be observed within lipid-filled MSCs. Finally, in contrast to
osteo-induced PLA cells, osteogenic induction of MSCs did not alter
the intracellular expression pattern of CNI or the synthesis and
extracellular deposition of CNIV and CNV (FIG. 43, Panel B;
MSC--Bone). Taken together, the immunofluorescent data suggests
that both adipogenic and osteogenic differentiation of PLA cells
leads to a remodeling of the associated ECM, resulting in a matrix
that appears to be distinct from those of MSCs.
PLA Clonal Isolates (ADSCs) Express a Similar Complement of CD
Marker Proteins
[0376] Multi-lineage differentiation by PLA cells may be the result
of the commitment of multiple lineage-specific precursors rather
than the presence of a pluripotent stem cell population within
adipose tissue. Therefore, multi-lineage differentiation by clonal
isolates derived from single PLA cells is critical to the
classification of PLA cells as a source of stem cells. In support
of this, single PLA cells colonies, termed Adipose-Derived Stem
Cells (ADSCs), exhibited multi-lineage capacity in vitro (FIG. 35).
Analysis of 500 ADSC isolates confirmed differentiation potential
in approximately 6% of the total number of clones examined. Seven
ADSC isolates exhibited tri-lineage potential, differentiating into
cells of the osteogenic, adipogenic and chondrogenic lineages,
approximately 24% of the total number of ADSCs positive for
differentiation potential (Table 11). Furthermore, a qualitative
increase in differentiation level, as measured by histologic
staining, was also observed in these tri-lineage ADSC populations.
In addition to tri-lineage ADSCs, several dual-lineage clones (O/A,
O/C and A/O) and single adipogenic lineage clones were also
isolated. Isolation and expansion of ADSCs did not alter the CD
expression profile of the clones, as no difference in CD expression
could be detected by IF. Furthermore, no difference was also
observed between tri- and dual lineage ADSC isolates (FIG. 36).
Like the heterogenous PLA populations, ADSCs were positive for
CD29, CD44, CD71 and CD90 expression, while no expression of CD31,
34, 45 and 104 was observed. Therefore, the presence of
multi-lineage ADSC isolates within the heterogenous PLA cell
population and their identical CD marker profile to PLA cells
further supports the theory that the adipose compartment is a
source of multi-potential stem cells.
14TABLE 13 Lineage-specific differentiation induced by media
supplementation Medium Media Serum Supplementation Control DMEM 10%
FBS None Adipogenic DMEM 10% FBS 0.5 mM isobutyl-methylxanthine
(AM) (IBMX), 1 .mu.M dexamethasone, 10 .mu.M insulin, 200 .mu.M
indomethacin, 1% antibiotic/antimycotic Osteogenic DMEM 10% FBS 0.1
.mu.M dexamethasone, 50 .mu.M (OM) ascorbate-2-phosphate, 10 mM
.beta.- glycerophosphate, 1% antibiotic/ antimycotic
[0377]
15TABLE 14 Monoclonal antibodies to CD antigens: Reported cell
specificity and distribution CD Antigen Clone Cell Specificity 29
Integrin .beta.1 MAR4 broad distribution--lymphocytes, monocytes,
granulocytes NOT on erythrocytes 31 PECAM-1 9G11 endothelial cells,
platelets, monocytes, granulocytes, haematopoietic precursors 34 --
581 endothelial cells, some tissue fibroblasts, haematopoietic
precursors 44 Pgp-1 G44-26 leucocytes, erythrocytes, epithelial
cells, platelets 45 LCA HI30 leucocytes, haematopoietic cells 58
LFA-3 L306.4 wide distribution--haematopoietic cells, endothelial
cells, fibroblasts 71 TfR H68.4 most dividing cells 90 Thy-1 5E10
immature CD34+ cells, cells capable of long term culture, primitive
progenitor cells 104 105 Endoglin -- endothelial cells, B cell
precursors, MSCs SH3 -- -- mesenchymal stem cells
[0378]
16TABLE 15 Immunofluorescent analysis of differentiated PLA and MSC
populations CD marker PLA PLA-OM PLA-AM MSC MSC-OM MSC-AM CD29 +ve
+ve +ve +ve +ve +ve CD31 -ve -ve -ve -ve -ve -ve CD34 -ve +/-,
restricted +ve (weak) -ve +ve (weak) +ve (weak) CD44 +ve +ve +ve,
+ve +ve +ve fibroblastic cells CD45 -ve -ve +/-, -ve +ve +ve (weak)
fibroblastic cells CD58 -ve -ve -ve +ve +ve +ve CD71 +ve +ve +ve
+ve +ve +ve CD90 +ve +ve +ve +ve +ve +ve fibroblastic cells CD105
+ve +ve +ve +ve +ve +ve - veno staining observed +/- minimal
staining observed (less than 10% of the population) +ve staining
observed
[0379]
17TABLE 16 Flow cytometric analysis of CD marker expression in
osteogenic, adipogenic and control PLA cells. CD Antigen PLA-CM
PLA-OM PLA-AM CD13 148.88 924.79 134.34 CD14 2.43 3.54 3.08 CD16
2.38 3.43 2.70 CD31 2.22 2.92 2.53 CD34 3.55 9.10 5.27 CD44 16.92
64.62 8.76 CD45 2.52 3.85 3.47 CD49d 5.33 13.05 4.27 CD56 2.66 4.86
2.72 CD61 3.68 7.55 4.12 CD62E 2.30 2.89 2.38 CD90 25.96 45.32
46.53 CD105 8.39 16.70 11.53 CD106 2.45 3.27 2.51 SH3 8.95 25.15
14.65 -ve 2.59 3.57 3.08
[0380]
18TABLE 17 Immunuofluorescent staining patterns of extracellular
matrix collagens: Effect of differentiation. Collagen
Immunofluorescent Staining Pattern type PLA MSC PLA-Fat MSC-Fat
PLA-Bone MSC-Bone 1 punctate + punctate + no expression weak
cellular expression + punctate + punctate + perinuclear perinuclear
fibrillar pattern perinuclear perinuclear concentration
concentration concentration concentration 4 fibrillar, localized
fibrillar pattern cellular distribution, fibrillar, decreased
fibrillar, weak fibrillar pattern to defined regions lipid-filled
cells only expression expression 5 fibrillar, localized fibrillar
pattern no expression fibrillar pattern no expression cellular +
fibrillar to defined regions pattern
Discussion
[0381] In this study, a more comprehensive characterization of the
PLA and ADSC populations was performed using a combination of
immunofluorescence and flow cytometry. While PLA cells expressed a
similar complement of CD antigens with MSCs (positive: CD29, CD44,
CD71, CD90, CD105 SH3, negative: CD31, CD34, CD45), the expression
of CD58, CD104 and CD140a differed on PLA cells when examined by
immunofluorescence. Flow cytometry also confirmed the expression of
CD13 and the absence of CD14, CD16, CD56 and CD62E. Subtle
distinctions between non-induced and differentiation PLA cells
could be determined using flow cytometry. Specifically, increases
in CD13, CD44 and CD90 were observed upon osteogenic induction,
whereas CD13 and CD44 levels in adipogenic cultures were found to
be lower. Consistent with this, IF analysis indicated a lower level
of CD44 expression within lipid-filled PLA cells (i.e. mature
adipocytes). Osteogenic differentiation also resulted in slight
increases in CD34, CD49d, CD56 and CD61. CD34 expression was
confirmed using immunofluorescence, with CD34-positive regions
being observed in osteogenic PLA cultures. ADSC clonal populations
also expressed a similar complement of CD antigens to that observed
in the heterogenous PLA population, suggesting that clonal
isolation and expansion of these cells does not affect cell surface
protein expression. Finally, differentiation of PLA cells also
resulted in changes to the associated ECM and differences in the
expression patterns and levels of collagen types 1, 4 and 5 were
found between differentiated PLA and MSC cultures. Taken together,
this data suggests that PLA cells may represent a stem cell
population within adipose tissue but is a population that possesses
subtle distinctions from MSCs.
[0382] While PLA cells expressed a similar complement of CD
antigens with MSCs, an established mesenchymal stem cell
population, PLA cells did show subtle differences in the expression
of CD58, CD104 and CD140a. The CD marker profile on PLA cells was
further confirmed using flow cytometry. Osteogenic and adipogenic
differentiation did not significantly change the CD profile, but,
as with control cells, subtle distinctions could be determined
using flow cytometry. Differentiation also resulted in changes to
the associated ECM. Finally, both ADSC clonal populations expressed
a similar complement of CD antigens to that observed in the
heterogenous PLA population, suggesting that clonal isolation of a
multi-lineage population from the PLA does not affect the
expression of cell surface proteins.
[0383] Characterization of a cell population can be performed
through identification of unique proteins expressed on the cell
surface. Several groups have subsequently characterized MSCs based
on their expression of cell-specific proteins (e.g. STRO-1, SH2,
SH3, SH4) and "cluster designation" (CD) marker profiles (Bruder,
S. P. et al., 1998, Clin. Orthop., S247-256; Conget, P. A. et al.,
1999, J. Cell Physiol, 181:67-73; Pittenger, M. F et al., 1999,
Science, 284:143-147.) This study confirms that, like MSCs, a
unique combination of cell surface proteins are expressed on PLA
cells with the two populations showing similar expression profiles.
Like MSCs, PLA cells expressed CD13, CD29, CD44, CD71, CD90,
CD105/SH2 and SH3 as shown by a combination of IF and FC. In
addition, PLA cells did not express CD14, CD16, CD31, CD34, CD45,
CD56, and CD62E on the cell surface. The similarity in CD profiles
to MSCs lends support to the theory that PLA cells are a stem cell
population. However, the degree of similarity may indicate that PLA
cells are simply an MSC population located within or contaminating
the adipose compartment. Lipoplasty results in the rupture of
multiple blood vessels and while vasoconstrictors are used to
minimize blood loss, the processed PLA pellet may be MSCs obtained
from the peripheral blood supply (Zvaifler, N.J. et al., 2000,
Arthritis Res., 2:477-488.) However, there appear to be a few
subtle distinctions between PLA and MSC populations. In contrast to
MSCs, no expression of CD58 could be detected on PLA cells using
IF, while expression was seen on MSCs (FIG. 23). Furthermore, MSCs
have also been reported to express CD104, CD106 and CD140a (Bruder,
S. P. et al., 1998, Clin. Orthop., S247-256; Conget, P. A. et al.,
1999, J. Cell Physiol, 181:67-73; Pittenger, M. F et al., 1999,
Science, 284:143-147.) No expression of these CD antigens were
detected on PLA cells using IF or FC (FIGS. 23 and 24). These
differences may indicate that the PLA population represents a
distinct population of stem cells. However, the possibility that
PLA cells are a clonal variation of MSCs cannot be completely ruled
out.
[0384] Multi-lineage differentiation by PLA cells may result from
the commitment of multiple lineage-specific precursors rather than
the presence of a pluripotent stem cell population within adipose
tissue. Therefore, multi-lineage differentiation by clonal isolates
derived from single PLA cells is critical to the classification of
PLA cells as a source of stem cells. In support of this, ADSC
isolated exhibited multi-lineage capacity in vitro staining
positively using the histologic assays alkaline phosphatase
(osteogenesis), Oil Red O (adipogenesis) and Alcian Blue
(chondrogenesis). Several lineage combinations were observed,
including tri-lineage (osteogenic, adipogenic and chondrogenic),
dual-lineage (osteogenic/adipogenic, osteogenic/chondrogenic) and
single lineage (adipogenic only). Isolation and expansion of ADSCs
did not alter the CD expression profile and no difference in CD
expression could be detected between any tri-lineage and
dual-lineage ADSC population. Therefore, the presence of
multi-lineage ADSC isolates and their identical CD marker profile
to heterogenous PLA cells further supports the theory that the
adipose compartment is a source of multi-potential stem cells.
[0385] Differentiation of mesenchymal precursors and stem cells may
lead to changes in the expression of several cell surface and
intracellular proteins as these cells acquire a new fate and
function. To assess this, undifferentiated PLA cells and cells
induced toward the osteogenic and adipogenic lineages were examined
by IF and FC for any changes in CD marker profile. Osteogenic
differentiation did not significantly alter the CD profiles of PLA
cells (FIG. 2 and Tables 16 and 17). Indirect IF confirmed the
expression of CD44 and CD90 and did not detect expression of CD34
and CD45 in both osteogenic PLA and MSC cultures. In addition, both
osteogenic PLA and MSC cultures were positive for CD29, CD71, CD105
and SH3 expression, whereas no expression of CD31 could be
detected. However, further analysis of osteogenic PLA cultures by
FC revealed subtle changes to the CD profile. Specifically,
osteo-induction resulted in a 1.8-fold and 3.8-fold increase in
CD90 and CD44 expression levels, respectively. The increased
expression of CD44, the hyaluronan receptor, is likely the result
of increased matrix synthesis and cell-matrix interaction by PLA
cells upon osteogenesis. Recent work has also confirmed the
expression of Thy-1/CD90 on osteoblasts and osteoblast-like cells
derived from mice, rats and human. Expression of this protein
increased markedly during the earliest stages of maturation
(proliferative phase) and decreased as the osteoblasts matured. The
increased expression of CD90 upon osteogenic induction of PLA cells
may, therefore, reflect the increased expression of this protein as
the osteogenic PLA cells proliferate during the earliest phases of
differentiation. In addition to CD44 and CD90, a dramatic increase
(6.2-fold) in the metalloprotease, CD13/aminopeptidaseN, was also
observed in osteogenic PLA cells. In addition to its expression on
committed progenitors of granulocytes and monocytes [Kishimoto,
1997 #1082], CD13 has also been identified on fibroblasts, bone
marrow stromal cells and osteoclasts (Syrjala, M. et al., 1994, Br.
J. Haematol., 88:679-684). Recent work has identified an increase
in CD13 mRNA levels upon cell-cell contact (Kehlen, A. et al.,
2000, J. Cell Biochem, 80:115-123; Reimann, D. et al., 1997, J.
Immunol., 158:33425-3432.) The dramatic increase in CD13 on PLA
cells may therefore be due to the increased cell to cell contact
within osteogenic PLA cultures. Additionally, increased expression
of proteases, such as CD13, on stem cells may also participate in
differentiation by degrading regulatory peptides and proliferation
agents that may affect the development of these cells (Young, H. E.
et al, 1998, Wound Repair Regen, 6:66-75; Young, H. E. et al.,
1999, Proc. Soc. Exp. Biol. Med., 221:63-71.)
[0386] Interestingly, FC measured slight increases in CD34, CD56,
CD49d, CD61 and CD105 expression upon osteogenic induction. With
the exception of CD105, these markers were not expressed on
undifferentiated PLA cells and MSCs and their increase is likely
the result of differentiation. A 2.6-fold increase in CD34
expression was detected in osteo-induced PLA cultures. This
increase was consistent with the appearance of CD34-positive
regions within 6steogenic PLA cultures as shown by IF (FIG. 23). A
slight increase in CD34 was also observed upon IF analysis of
osteogenic MSCs (FIG. 40). However, this increase appeared to be
the result of an overall enhanced expression level by all MSCs.
Osteogenic induction also resulted in a 1.8-fold increase in CD56
expression. Identified as neural cell adhesion molecule (NCAM),
CD56 is expressed on haematopoietic stem cells (Kishimoto, T. et
al, 1997, Leucocyte Typing VI. White Cell Differentiation Antigens.
(Hamden, Conn.: Garland Publishing), mediating their adhesion with
adjacent cells and the surrounding matrix (Lanier, L. L. et al,
1991, J. Immunol, 146:4421-4426; Lanier, L. L. et al., 1989, J.
Exp. Med., 183:681-689.) Although its function has not been
confirmed, CD56 may act in a similar manner in non-haematopoietic
cells. In support of this, osteoblasts express NCAM, using this
adhesion molecule to mediate cell and matrix interactions and
leading to their differentiation (Lee, Y. A. et al., 1992, J. Bone
Miner. Res., 7:1435:1466.) The osteogenic differentiation of PLA
cells, therefore, may induce elevated levels of this CD protein in
order to regulate the increasing cell-cell and cell-matrix
interactions during differentiation. The same explanation can
likely be applied to the observed 3-fold increase in the .alpha.4
integrin, CD49d. Finally, a small increase in CD105 expression was
measured on osteogenic PLA cells. Classified as a type III
TGF.beta.3 receptor (Cheifetz, S. et al., 1992, J. Biol. Chem.,
267:19027-19030), CD105 is expressed on a wide variety of cells,
including endothelial cells, B-lineage precursors, MSCs and a
subset of CD34+ cells isolated from peripheral blood (Rokhlin, O.
W. et al., 1995, J. Immunol., 154:4456-4465; Majumdar, M. K. et
al., 1998, J. Cell Physiol., 176:57-66; Barry, F. P. et al., 1999,
Biochem. Biophys. Res. Commun., 265:134-139; Pierelli, L. et al.,
2000, Br. J. Hematol., 108:610-620.) While little is know of this
protein during bone development, expression of CD1 05 is thought to
decrease as osteogenic precursors proceed toward terminal
differentiation, disappearing on mature osteoblasts (Haynesworth,
S. E. et al., 1992, Bone, 13:69-80.) Therefore, the expression of
CD105 on osteogenic PLA and MSCs, as shown by IF, may indicate that
these cells represent an early stage in differentiation and have
not reached their final differentiation stage. Furthermore, the
slight increase in CD105 expression on PLA cells, as measured by
FC, correlates to the increase in CD34 and may reflect the increase
in a CD34.sup.+ subset within the osteogenic culture.
[0387] Adipogenic differentiation of PLA cells has been shown to
result in an expanded morphology, together with the accumulation of
multiple lipid-filled intracellular vacuoles (Zuk, P. et al., 2001,
Tissue Engineering, 7:209-226.) As a result, adipo-induced PLA
cultures are a heterogenous mixture of lipid-filled cells (i.e.
mature PLA adipocytes) and more immature fibroblastic cells.
Consistent with this, FC characterization of adipogenic PLA
cultures demonstrated a shift toward a population of larger, more
granular cells. IF analysis confirmed the expression of CD29, CD44,
CD71, CD90 and CD105 on adipogenic PLAs and MSCs (FIG. 41). While
equivalent levels of CD29, CD71 and CD105 were found on both
fibroblastic and lipid-filled cells, lower levels of CD44 and CD90
were observed in the mature PLA adipocytes. Contrary to PLA
cultures, no such restriction could be detected by IF in
adipo-induced MSCs. While expression of CD90 appeared to be
decreased in lipid-filled PLA cells, virtually 100% of the PLA
fibroblasts stained brightly for CD90 and a 1.8-fold increase in
this protein was measured using FC, a level comparable to that
measured in osteogenic cultures (1.75-fold). Contrary to CD90,
expression levels of the CD44-positive PLA fibroblasts appeared to
vary, with cell staining ranging from intense to little or no CD44.
In support of this, FC confirmed a 48% decrease in CD44 expression
in adipogenic PLA samples. A decrease was also measured for
CD13/aminopeptidase N and the decrease of these two proteins is
likely a reflection of the remodeling of the ECM to one more
consistent with adipogenic tissue.
[0388] Like osteogenic PLA cells, FC confirmed the absence of CD14,
CD16, CD31, CD45, CD62E and CD106 in adipogenic PLA cultures while
expression of CD34, CD49d and CD61 were slightly elevated in these
cells. While FC did not detect a significant increase in CD45 upon
adipogenic induction, a small percentage of PLA cells positive for
this protein was observed upon IF analysis. The increased
expression of CD34 on adipogenic PLA cells was not as large as that
measured upon osteogenesis and IF analysis confirmed CD34
expression by all PLA morphologies. However, expression was
restricted to cells with a fibroblastic morphology. Weak expression
of both CD34 and CD45 were also detected upon IF analysis of
adipogenic MSCs with expression observed in both fibroblastic and
lipid-filled cells.
[0389] Differentiation of mesenchymal precursors to their
lineage-committed cell types (i.e. osteoblasts, adipocytes) is
accompanied by synthesis and remodelling of an ECM. Variation of
ECM composition and organization gives each tissue its specific
characteristics and participates in the differentiation and growth
of the constituent cell types. For example, bone matrix consists of
inorganic hydroxyapatite together with an organic fraction
comprised of proteoglycans and collagens, with collagen type 1
making up the majority (approx. 90% of the organic fraction).
Cartilage matrix consists mainly of collagens type 2 and 10 and
multiple sulfated proteoglycans. Adipogenic ECMs are comprised of
multiple collagen subtypes (1 through 6), laminin and fibronectin.
Together, these collagens are a part of the unique extracellular
environment of each tissue and are crucial to the survival and
function of the component cells. Based on this, the expression of
ECM collagens were examined in both control and induced PLA cells
and MSCs.
[0390] Non-induced PLA cells and MSCs expressed CNI, CNIV and CNV
(FIG. 43), in addition to CNIII. Both CNI and CNIII exhibited
similar staining patterns in both cell populations and osteogenic
induction did not alter the intracellular distribution of these
collagens. Furthermore, a qualitative increase in CNI was observed
in several PLA and MSC samples. A large volume of work confirms the
role of collagen type 1 in osteogenic differentiation. For example,
CNI levels increase during the early stages of rat calvarial
osteoblast differentiation and inhibition of this collagen totally
blocks osteogenic differentiation (Stein, G. S. et al., 1990, Faseb
J., 4:3111-3123; Lynch, et al., 1995, Exp. Cell Res., 216:35-45.)
Factors that are known to affect osteogenesis, such as
dexamethasone, vitamin D and the parathyroid hormone, can directly
affect levels of CNI. Furthermore, bone marrow stromal cells
maintained on CNI matrices differentiate into osteoblasts in vitro
and induce bone formation in vivo, an effect that is not seen on
CNII, CNIII or CNV matrices. Therefore, the synthesis of CNI in
pre-induced and osteo-induced PLA cultures in consistent with the
role of this collagen in osteogenesis. Moreover, the similarities
in CNI expression observed in both osteo-induced PLA cells and MSCs
suggests that similar mechanisms may function in the osteogenic
differentiation of these cell types.
[0391] In addition to CNI, expression of CNIV and CNV were also
observed in both control PLA and MSC cultures, distributed in a
fibrillar pattern consistent their secretion into the extracellular
environment. The presence of these collagens is a vital component
of the osteogenic ECM as expression of these collagens is observed
in whole bone marrow stroma, the osteoblasts of newly forming bone
and in STRO-1-positive colony derived stromal cell lines. In
contrast to induced MSC cultures, osteogenic induction of PLA cells
appeared to significantly decrease CNIV synthesis and completely
inhibited CNV expression.
[0392] Adipogenic differentiation resulted in additional
distinctions between PLA and MSC populations. Like osteogenic
cultures, adipogenic induction of PLA cells resulted in an
inhibition of CNV expression. Moreover, adipogenesis also resulted
in the inhibition of CNI. No such inhibition was seen in
adipo-induced MSCs. Rather, a reduced level of CNIV synthesis was
observed in adipogenic MSC populations with all three collagen
types (I, IV and V) exhibiting a fibrillar, extracellular
expression pattern. While weak cellular expression of CNI was also
observed in lipid-filled MSCs, the expression of CNIV and CNV
appeared to remain extracellular. Like MSC samples, adipo-induced
PLA cells also expressed CNIV. However, CNIV expression in these
cells remained intracellular and appeared to be expressed
exclusively in lipid-filled PLAs.
[0393] Like osteogenesis, several lines of evidence suggest that
ECM components, such as collagens, participate in adipogenesis.
First, changes in the ECM lead to morphologic and cytoskeletal
alterations that are required for the expression of lipogenic
enzymes (Kuri-Haruch, W. et al., 1984, Differentiation, 28;
Spiegelman, B. M. et al., 1983, Cell, 357-666.) Second, expression
of CNI, CNIII and CNIV varies dramatically upon differentiation of
3T3-L1 cells (Weiner, F. R. et al., 1989, Biochem., 28:4094-4099.)
Lastly, the ECM of developing adipose tissue is organized during
differentiation, an event thought to be mediated by the adipocytes
themselves (Nakajima, I. et al., 1998, Differentiation,
63:193-200.) Fibroblasts and adipocyte precursors with a
fibroblastic morphology synthesize and secrete type I and III
collagens, in addition to small amounts of the basement membrane
collagen, type IV (Goldberg, B., 1977, PNAS, 74:3322-3325; Alitano,
K. et al., 1982, J. Cell Biol., 94:497-505; Cryer, A. et al., 1982,
Eur. J. Clin., Invest., 12:235-238; Kuri-Harcuch, W. et al., 1984,
Differentation, 28; Liau, G. et al., 1985, J. Biol., Chem.,
260:531-536.) As these cells begin to differentiate changes occur
in cell morphology, cytoskeleton and the level and type of ECM
secreted (Napolitano, L., 1963, J. Cell Biol., 18:663-679; Aratani,
Y. et al., 1988, J. Biol. Chem., 263:16163-16169; Weiner, F. R. et
al., 1989, Biochem., 28:4094-4099.) These changes, in turn, may be
a requirement for their terminal differentiation into
adipocytes.
[0394] To study the synthesis and distribution of ECM components
upon adipogenesis, several preadipocyte cell lines have been
developed, including several 3T3 variants (Green, H. et al., 1974,
Cell, 3:127-133) and a clonal preadipocyte cell line from Japanese
cattle (BIP cells) (Aso, H. et al., 1995, Biochem. Biophys. Res.
Commun., 213:369-374.) Adipose conversion of BIP cells results in
production of an ECM similar to adipose tissue in which adipocytes
are interconnected by a fibrillar network of collagens I, II, IV, V
and VI together with an intracellular expression of CNIII
(Nakajima, I. et al., 1998, Differentiation, 63:193-200.) Like BIP
cells, adipo-induction of both PLA cells and MSCs resulted in a
similar intracellular distribution of CNIII. Furthermore, fibrils
of CNI, CNIV and CNV were also associated with adipogenic MSCs and
appeared to be organized randomly. The expression of similar
collagens and their random organization in adipogenic MSCs is
consistent with that observed upon differentiation of preadipocytes
and suggests that comparable ECM synthesis and remodeling may occur
upon differentiation of these stem cells.
[0395] However, the adipogenic differentiation of PLA cells
presents several differences to several preadipocyte cell lines and
MSCs. Like preadipocyte cells, including BIP cells from cattle, and
3T3 cells from mice, pre-differentiated PLA cells synthesize CNI
and CNV. However, these collagens are no longer observed upon
differentiation. The disappearance of CNI and CNV in adipogenic PLA
cultures may represent a specific remodelling pathway unique to
these cells. In support of this, changes in the pericellular
environment that occur during differentiation can change the
intracellular environment and the secretion of MMPs that degrade
the surrounding ECM. Low levels of CNIV are also produced by
preadipocytes and a dramatic increase is observed upon adipogenesis
(Aratani, Y. et al., 1988, J. Biol. Chem., 263:16163-16169;
Nakajima, I. et al., 1998, Differentiation, 63:193-200.) While a
qualitative increase in CNIV is observed in adipogenic PLA
cultures, its fibrillar distribution is lost and the collagen is
restricted to lipid-filled PLA cells. The change in CNIV expression
pattern in comparison to preadipocytes and MSCs remains
unclear.
[0396] While EM observations of mature fat cells have identified a
CNIV-rich basement membrane associated with several other fibrillar
collagens (Chase, W. H., 1959, J. Ultrastruc. Res., 2:283-287;
Barnett, R. J., 1962, L. W. Kinsell, ed., (Springfield, Ill.:
Charles C. Thomas); Angel., A. et al., 1970, B. Jeanrenaud and
Hepp, D., et ed. (Thiene, Stuttgard: Academic Press), mature fat
cells do not synthesize collagens. Moreover, adipogenic precursors
lose the capacity for collagen synthesis in vitro during the
post-confluent differentiation stage. However, collagen synthesis
is critical for terminal adipocyte differentiation and
triacylglyerol accumulation indicating that the predifferentiation
expression of an ECM determines their ultimate phenotype.
Therefore, the predifferentiation expression of CNI, CNIII, CNIV
and CNV by PLA cells and MSCs may serve to initiate their
differentiation program. As differentiation proceeds and the
appearance of lipid-filled cells (i.e. mature adipocytes)
increases, the synthesis of these collagens ceases, resulting in a
collagenous ECM unique to adipose tissue. This is likely the case
with the MSC population. However, the absence of CNI and CNV in PLA
cultures may be the result of a direct inhibition of synthesis or a
dramatic remodeling of the ECM. The precise time of collagen
inhibition upon PLA adipogenesis and/or the possible existence of
agents involved in collagen degradation remains unknown.
* * * * *