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Abstract

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. REFERENCES

Mesenchymal stem cells (MSCs) residing in bone marrow (BM) are the progenitors for osteoblasts and for several other cell types. In humans, the age-related decrease in bone mass could reflect decreased osteoblasts secondary to an age-related loss of osteoprogenitors. To test this hypothesis, BM cells were isolated from vertebral bodies of thoracic and lumbar spine (T1–L5) from 41 donors (16 women and 25 men) of various ages (3–70 years old) after death from traumatic injury. Primary cultures were grown in alpha modified essential medium with fetal bovine serum for 13 days until adherent cells formed colonies (CFU-Fs). Colonies that stained positive for alkaline phosphatase activity (CFU-F/ALP+) were considered to have osteogenic potential. BM nucleated cells were plated (0.5, 1, 2.5, 5, or 10 × 106 cells/10-cm dish) and grown in dexamethasone (Dex), which promotes osteoblastic differentiation. The optimal plating efficiency using BM-derived cells from donors of various ages was 5 × 106 cells/10-cm dish. BM-derived cells were also grown in the absence of Dex at this plating density. At the optimal plating density, in the presence of Dex, the number of CFU-F/ALP+ present in the BM of the younger donors (3–36 years old) was 66.2 ± 9.6 per 106 cells (mean ± SEM), but only 14.7 ± 2.6 per 106 cells in the older donors (41–70 years old). With longer-term culture (4–5 weeks) of these BM cells in medium containing 10 mM β-glycerophosphate and 100 μg/ml ascorbic acid, the extracellular matrix mineralized, a result consistent with mature osteoblastic function. These results demonstrate that the number of MSCs with osteogenic potential (CFU-F/ALP+) decreases early during aging in humans and may be responsible for the age-related reduction in osteoblast number. Our results are particularly important in that the vertebrae are a site of high turnover osteoporosis and, possibly, the earliest site of bone loss in age-related osteoporosis.


INTRODUCTION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. REFERENCES

Osteoblasts originate from mesenchymal stem cells (MSCs),(1–5) which reside in bone marrow (BM) together with hematopoietic stem cells.(5) These two stem cell types cooperate through direct cell-to-cell interactions and release of cytokines and growth factors.(6–9) Generally, MSCs in BM are arrested in G0(10); however, when BM cells are plated in vitro, MSCs exit their quiescent state and start to proliferate, forming individual colonies (colony forming unit-fibroblasts, CFU-Fs), each derived from a single stem cell.(1,5) In vitro MSCs appear to be multipotent for differentiating into osteoblasts, chondroblasts, adipocytes, fibroblasts, myoblasts, and reticular cells.(1–5,11,12) Although CFU-Fs are a heterogeneous population of stem and progenitor cells,(2,13) colonies with alkaline phosphatase activity (CFU-F/ALP+) are considered to have osteogenic potential(13) and thus are considered osteoprogenitors.

The number of MSCs in BM and how this number varies with donor age have not been clearly established in humans, nor has a method been validated to identify MSCs before they become functional. Several animal studies on the influence of aging on CFU-F/ALP+ number suggest that the number of osteoprogenitor cells decreases with aging.(5,14–17) However, there is a discrepancy among studies using human cells, with some reports of decreasing numbers with age(18) and other reports that find no change with donor age.(19,20) Moreover, there have been several reports of a decrease in skeletal mass at a relatively early age—substantially before menopause—which would correlate with an age-related decrease in osteoprogenitor cell number.(21–23)

We hypothesize that a decrease in the number of osteoblasts and the age-related reduction in bone formation and in the mechanical properties and integrity of bone in humans reflect decreased osteoprogenitor generation during aging. Accordingly, we have investigated the number of CFU-F/ALP+ in cultures of BM from thoracolumbar vertebrae obtained from persons 3–70 years old at death from traumatic injury.

MATERIALS AND METHODS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. REFERENCES

Human BM cells

Human BM was isolated from postmortem thoracolumbar (T1–L5) vertebral bodies of 41 donors (25 men and 16 women) immediately after death from traumatic injuries.(24) Hepatitis and HIV were ruled out by history and lab tests on BM specimens. Nothing was known about the osteoporosis history of the donors. The BM isolation was performed in a class-1000 clean room facility equipped with class-100 safety cabinets. After harvesting the vertebral column from the cadaver, all the adherent soft tissue, the intervertebral disc, and the periosteum were removed from each single vertebra. The vertebral bodies were divided along the sagittal midline craniocaudal axis using a sterile hammer and chisel; a Rongeur was used to obtain small bone chips (∼5 mm3). The bone chips were placed at room temperature in 1000 ml of processing medium (X-Vivo 10; Biowhittaker, Inc., Walkersville, MD, U.S.A.), containing 100 ml of human serum albumin 25% (Calbiochem, La Jolla, CA, U.S.A.), 10 ml of bacitracin (5 × 103 U/ml), 10 ml of polymyxin B (5 × 104 U/ml), 10 ml of heparin (1000 U/ml), and 2 ml of 1 M HEPES buffer (pH 7.2). All the bone chips were separated from the cell suspension by filtering through two consecutive stainless steel screens (pore diameter 450 μm and 180 μm, respectively). The filtered cell suspension was centrifuged at 300g for 10 minutes at 4°C. The cell pellet was resuspended in alpha modified essential medium (α-MEM) (GIBCO-BRL, Grand Island, NY, U.S.A.) with 10% heat-inactivated fetal bovine serum (FBS; HyClone Laboratories, Inc., Logan, UT, U.S.A.). Remaining BM cells trapped within bone trabeculae were released by two additional 30-minute cycles, at room temperature, of gentle rocking agitation (∼40 cycles/minute) of the bone fragments in resuspension medium (1000 ml RPMI 1640 [GIBCO-BRL], 100 ml human serum albumin [25%], 10 ml gentamicin [50 mg/ml], 10 ml of heparin [1000 U/ml], and 2 ml of 1 M HEPES buffer, pH 7.2). All cells from all vertebrae (T1–L5) of a given donor were pooled, centrifuged at 300g for 10 minutes at 4°C, and resuspended in α-MEM with 10% FBS. The pooled suspension of BM cells was filtered through a Y-Type Blood Set (McGaw, Irvine, CA, U.S.A.) to remove any bone fragments that might have been trapped in the cell pellet and to separate any clumps of cells. An aliquot of cells from the final filtered suspension was treated with 4% acetic acid (final concentration; to lyse erythrocytes) and trypan blue and then counted with a hemacytometer.

Cell culture

Nucleated BM cells were counted and plated into 10-cm dishes (Costar, Cambridge, MA, U.S.A.) in 10 ml of α-MEM/10% FBS with 100 U/ml penicillin (GIBCO-BRL), 1 mg/ml streptomycin (GIBCO-BRL), and 100 μg/ml ascorbic acid. The next day, dexamethasone (Dex; Sigma, St. Louis, MO, U.S.A.) dissolved in dimethylsulfoxide was added to a final concentration of 10 nM. Dimethylsulfoxide at the final concentration of 0.001% was used as vehicle. After 1 week in culture, half of the growth medium was removed and replaced with fresh. Cells were incubated for 13 days in a 100% humidified atmosphere of 95% air, 5% CO2 at 37°C. For studies involving production of mineralized matrix, cells were grown for 4–5 weeks in the same medium supplemented with 10 mM β-glycerophosphate.

Alkaline phosphatase activity

At the indicated times, cells were fixed with cold 10% neutral-buffered formalin (30 minutes at 4°C) and then assayed for ALP activity as previously described.(25) Briefly, cells were incubated with fresh substrate at 37°C for 30 minutes, then rinsed extensively with distilled H2O and photographed. Substrate solution was prepared by dissolving 8 mg of naphthol AS-TR phosphate (Sigma) in 0.3 ml of N,N′-dimethylformamide (Sigma) while separately dissolving 24 mg of fast blue BB salt (Sigma) in 30 ml of 100 mM Tris-HCl (pH 9.6). The above solutions were mixed, 10 mg of MgCl2 was added and dissolved, and the pH was adjusted to 9.0 using 1 N HCl. The final substrate solution was filtered through a 0.2-μm filter and used immediately.

Colony size and number

After assaying for ALP activity on day 13, CFU-F colonies with 50 or more cells (the conventional value for defining a colony)(5) were scored visually as positive (i.e., blue stain, CFU-F/ALP+) using a Nikon SMZ-U (Nikon, Inc., Melville, NY, U.S.A.) dissecting microscope. ALP-positive and ALP-negative colonies were counted.

Colony size (area) was determined by image analysis of 20 randomly chosen colonies per plate from a defined area of 15 cm2 on each plate, using Sigma Scan Pro 4.0 for Windows (SPSS, Inc., Chicago, IL, U.S.A.). Only four donors (41- and 47-year-old females and 51- and 59-year-old males) had fewer than 20 colonies in that specific area.

Matrix Ca+2 content/mineralization

At the times indicated, mineralization was determined as described by Stanford et al.(26) Briefly, cells were washed thrice with PBS at room temperature and fixed with ice-cold 70% ethanol for 60 minutes at 4°C. Fixed cultures were incubated with 40 mM alizarin red-S (AR-S; Sigma), pH 4.2, for 10 minutes at room temperature with agitation via an orbital shaker (100 rpm). To minimize nonspecifically bound stain, dishes were rinsed briefly with five changes of deionized water and one extended rinse (15 minutes) with PBS at room temperature. Extracellular matrix (ECM) mineral-bound stain was photographed under light microscopy.

Solubilized AR-S was quantitated spectrophotometrically at 562 nm; AR-S was solubilized by 15 minutes of agitated (orbital shaker) incubation in 5 ml of 10% cetylpyridinium chloride buffered with 10 mM sodium phosphate, pH 7.0.

Statistical analysis

Student's t-test and regression analyses were used for assessing differences and correlations. Three 10-cm dishes of cells were analyzed for each donor; age group data are shown as the mean ± SEM.

RESULTS

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. REFERENCES

Characteristics of primary cultures of human vertebra-derived BM cells

After 2–4 days in culture, a heterogeneous population of mononuclear cells (with adherence capacity) including fibroblast-like spindle-shape cells, monocytes, macrophages, endothelial cells, and multinucleated osteoclasts attached to the surface of the dishes. As reported by others,(1–5,13) serial observation of cell shape and size showed that only MSC-derived fibroblast-like cells proliferate to form CFU-F colonies (Fig. 1). The other cell types that adhere to the dish surface do not proliferate and appear eventually to die, perhaps because the culture conditions are not optimal. The floating hematopoietic cells, also present in the cultures, were not initially removed because they may provide necessary factors for the MSCs.(9)

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Figure FIG. 1.. Production of CFU-Fs from human BM cells of the same donor. 5 × 106 nucleated BM cells, derived from vertebral column (T1–L5), were plated in 10-cm dishes in α-MEM with 10% FBS heat-inactivated and 100 μg/ml ascorbic acid and grown for 13 days in the presence (A) or absence (B) of 10 nM Dex. Cells were then fixed and stained for ALP activity (see Materials and Methods).

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Small fibroblastoid colonies formed at 5–7 days. At the end of the incubation period (13 days), colonies were clearly formed. CFU-Fs consisted (by visual inspection) of mainly two types of cells, elongated and cuboidal (not apparent from the magnification used for Fig. 1). Many CFU-Fs showed alkaline phosphatase activity (ALP+) expressed at different levels (Fig. 1). Dex increased the total number of colonies 20–40% regardless of donor age. Moreover, in the presence of Dex, >90% of the colonies were ALP-positive compared with <50% in its absence. ALP activity appeared visually to be consistently higher in cells grown in the presence of Dex (Fig. 1).

Effect of plating density

To determine the optimal cell number for colony formation (i.e., colony = aggregates of more than 50 cells), nucleated BM cells (0.5, 1, 2.5, 5, or 10 × 106 per 10-cm dish) were plated with Dex. After 13 days in culture, the highest number of ALP-positive colonies (CFU-F/ALP+) was obtained when 5 × 106 nucleated cells were plated per dish (Fig. 2). Plating a larger number of cells did not significantly increase the number of CFU-F/ALP+ (Fig. 2). Similar results were obtained when the total number of colonies was counted (data not shown).

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Figure FIG. 2.. Number of CFU-F/ALP+ at various plating densities. Whole BM cells were plated at 0.5, 1, 2.5, 5, and 10 × 106 per 10-cm dish and allowed to proliferate for 13 days. Young donors (3–36 years old) (filled square), older donors (41–70 years old) (filled circle). All donors (3–70 years old) (filled triangle). ***Significant difference, young versus older donors, p < 0.001.

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Age-related changes in the number of CFU-F/ALP+

The number of CFU-F/ALP+ obtained from all 41 donors when 5 × 106 cells were plated in the presence of Dex averaged 38.6 per 106 BM cells plated. Using only younger donors (ages 3–36; n = 19, mean = 20.6 ± 1.9 [mean ± SEM]); 5 females and 14 males), we observed 66.2 per 106 BM cells. There were only 14.7 per 106 BM cells from older donors (ages 41–70; n = 22, mean = 45.3 ± 2.8 [mean ± SEM]; 11 females and 11 males).

The number of CFU-F/ALP+ obtained after the 13-day culture period decreased with donor age at all plating densities in the presence or absence of Dex. Figure 3 shows the results obtained when BM cells were plated at 5 × 106 cells/dish. In the presence of Dex (Fig. 3A), ΔCFU-F/ALP+ ÷ Δdonor age (i.e., slope) = –7.17 ± 1.27 CFU-F/ALP+ per donor per year of age; significantly different from 0 (r2 = 0.45, p < 0.001). We observed an age-related decrease in the number of CFU-F/ALP+ from donors up to age 40 years (Fig. 4, broken line). Moreover, the mean value dropped from 331 ± 48 (mean ± SEM) for donors 3–36 years old (n = 19) to 73.8 ± 13 CFU-F/ALP+ for donors 41–70 years old (n = 22) (p < 0.001). In the absence of Dex, the slope is less steep (slope = –4.34 ± 1.08; r2 = 0.36, p < 0.001) (Fig. 3B).

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Figure FIG. 3.. Effect of Dex on the number of CFU-F/ALP+ from human vertebral BM. Whole BM cells were plated at 5 × 106 cells/10-cm dish and allowed to proliferate for 13 days (A) in the presence of 10 nM Dex, n = 41 (25 men, 16 women). r2 = 0.45; p < 0.001, slope = –7.17 ± 1.27, significantly different from 0; or (B) in the absence of Dex, n = 30 (17 men, 13 women). r2 = 0.36, p < 0.001, slope = –4.34 ± 1.08, significantly different from 0. Solid line in each panel indicates linear regression of data points shown; dotted lines indicate 95% confidence limits.

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Figure FIG. 4.. Regression analyses of data from the younger and older age groups with respect to the number of CFU-F/ALP+ obtained from Dex-treated cells. Data are from the experiment described in the legend to Fig. 3A. For the polynomial regression (solid line), r2 = 0.47, t1 = −2.55, t2 = 1.36; for the linear regressions (broken lines) of the younger age group, r2 = 0.12 (p < 0.05), and of the older age group, r2 = 0.002 (p < 0.84).

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The second-order polynomial regression of Dex-treated cultures confirms the slope change between ages 39 and 41 years, r2 = 0.47, t1 = –2.55, t2 = 1.36 (Fig. 4). However, if we draw two separate linear regressions, one for each age group, younger and older (Fig. 4), there is no further decrease in the number of CFU-F/ALP+ after 40 years, while a dramatic age-related reduction is evident in the younger donors at earlier ages. Further, there is no significant difference in colony number between male and female donors in either age group (data not shown).

Colony size changes with age

In the presence of Dex, colony size was significantly different between young (3–36 years old) and old (41–70 years old) donors, p < 0.001; in the young group, the average colony size was 10.23 ± 0.33 mm2, vs. 8.64 ± 0.23 mm2 in the older group (Fig. 5A). In the untreated cells (i.e., no Dex), the two groups (young and old) did not show any significant difference (p = 0.40). However, the colonies observed in the absence of Dex were significantly larger than those seen in Dex-treated cultures: 11.65 ± 0.37 mm2 vs. 9.94 ± 0.25 mm2, p < 0.001 (Fig. 5B). A difference was also seen between the young group not treated with Dex and the young group treated with Dex (p < 0.05) and between the untreated old group and the old Dex-treated group, p < 0.01 (Fig. 5B).

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Figure FIG. 5.. The effect of Dex and donor age on colony size. Whole BM cells were plated and treated as described in the legend to Fig. 3. (A) young donors (3–36 years old); older donors (41–70 years old). ***Significantly different only when Dex-treated, p < 0.001. (B) control, Dex-treated. In all donors, ***p < 0.001; in young donors, *p < 0.05; in older donors, ***p < 0.001.

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Osteogenic potential of MSCs

To verify the osteogenic potential (i.e., mineralizing capability) of the MSCs isolated from vertebral bodies, primary cultures of cells were grown for 4–5 weeks in α-MEM medium containing β-glycerophosphate, in the presence or absence of Dex from day 1. Mineralization occurred only in the Dex-treated cultures (Fig. 6).

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Figure FIG. 6.. Mineralization by BM-derived cells in culture. Whole BM cells were incubated for 4–5 weeks in α-MEM with 10% FBS, 100 μg/ml ascorbic acid, and 10 nM β-glycerophosphate in the absence (A) or in the presence (B) of 10 nM Dex. Cultures were stained for Ca2+ mineralization with AR-S.

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DISCUSSION

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. REFERENCES

In humans, bone mass increases, plateaus, then decreases with aging.(21,27) While resorption can increase, formation decreases, possibly because osteoblasts decrease with age.(28) Since osteoblast number might relate to progenitor number, the goal of this study was to determine whether the number of MSCs (with osteogenic potential) residing in the BM of human thoracic/lumbar vertebrae, a skeletal site of high turnover in bone, could be associated with age-related osteoporosis.(27,29,30)

BM cells derived from thoracic/lumbar vertebral bodies have the same fibroblastic spindle shape as reported for those isolated from ribs, femoral head, and iliac crest aspirates of both animals and humans.(5,9,13,31–34) Cultures of these nucleated BM cells in the presence of Dex showed high levels of ALP activity and a higher number of CFU-F/ALP+ than the nucleated cells cultured in the absence of Dex. These results support the concept that glucocorticoids stimulate recruitment of uncommitted stem cells toward the osteogenic lineage.(35,36) BM cells cultured in the absence of Dex resulted in CFU-F with lower ALP activity (Fig. 1B), suggesting that these cells are nonosteogenic progenitor cells or cells at a more immature stage along the osteogenic pathway.(13) Nonetheless, the colonies in the presence of Dex were significantly smaller in size (Fig. 6B), presumably because glucocorticoids inhibit bone cell proliferation by slowing growth and promoting differentiation.(13,31,34,36)

As reported by others, the morphology of the cells cultured with Dex was more osteoblastic (cuboidal shaped) than the fibroblastic morphology seen for those cultured without glucocorticoid.(32,33) Moreover, the number of colonies formed in the presence of Dex was directly proportional to the number of cells plated, as previously reported in animal studies.(9,13,17,37)

The mineralization potential of vertebra-derived human osteogenic MSCs was demonstrated in long-term cultures using β-glycerophosphate–containing medium. As reported in other studies using human BM aspirate from iliac crest, mineralization occurred only in the presence of Dex.(30,31) Moreover, individual CFU-F/ALP+ colonies can be subcultured and the subsequently derived colonies shown to mineralize.(38)

Regardless of the presence or absence of Dex, MSCs (total and those with osteogenic potential, i.e., CFU-F/ALP+) decreased with age (Fig. 3). Our results agree with findings from various animal studies(5,14–17) and with a report of human cells,(18) although they conflict with the data of others(19,20) who failed to note any age-related changes in the number of osteogenic precursors. However, one of those discrepant studies(19) used cells only from donors 40 years of age and older. Our data from donors older than 40 are consistent with the findings of that study, in that no further fall in the number of cells with osteogenic potential was found after age 40 (Fig. 4). Indeed, we can postulate that what is being observed is not so much a linear age-related decline in CFU-F/ALP+ over the entire age range of 3–70 years, but rather a sharp drop after the rapid juvenile bone growth period to a lower rate in mature adults. As indicated in the Introduction, several studies have reported a substantial premenopausal bone loss in the lumbar spine.(21–23) These data correlate well with our observation of a reduction in osteoprogenitor cells derived from thoracolumbar spine of donors substantially younger than age 40. In fact, most skeletal mass is accumulated by age 18, and some skeletal sites begin to lose bone immediately after that age (including trabecular bone in the vertebrae) (see Matkovic and references therein(39)).

Our results support the hypothesis that MSCs, and in particular MSCs with osteogenic potential, decrease during aging. We have not yet explored the mechanism, but there are several possibilities. There may be an age-related reduction in the number of MSCs in the BM as a result of cellular senescence, by which pluripotent cells could replicate only a limited number of times before undergoing apoptosis. Also, growth factors and hormones and MSC growth responsiveness to these factors could change.(10,21) Nevertheless, it is well known that the replication of cells in vitro is limited(40–42) and that with increasing donor age comes a decreased response to mitogens or a decrease in the number of mitogens elaborated in the medium. Further, the significant difference in colony size between the two groups treated with Dex (Fig. 5A) suggests that cells from elderly donors proliferate slower.

The BM microenvironment changes with age, resulting in cell-to-cell and cell-to-matrix interactions that may be unfavorable for MSC proliferation or that may favor MSC maturation toward a different lineage (e.g., adipogenic). Total marrow fat increases with age, and there is an inverse relationship between marrow adipocytes and osteoblasts with aging.(43–45)

Recently Ducy et al.(46) cloned a gene encoding a factor that binds to an osteoblast-specific cis-acting element (OSE-2). OSE-2–like elements are found in the promoter of a large number of genes in osteoblasts: type(I) collagen, osteocalcin, bone sialoprotein, osteopontin, etc. This factor, designated Osf2/Cbfa1, has been identified as an osteoblast-specific transcription factor involved in regulation of osteoblast differentiation and bone development.(47,48) These studies suggest a critical role for Osf2/Cbfa1 in the control of lineage-specific differentiation in MSCs. It would be interesting to determine at what stage in the maturation of MSCs toward the osteoblastic phenotype expression of Osf2/Cbfa1 is induced and by which growth factor(s) or hormone(s). In addition, we plan to determine whether there is an age-related decrease or delay in the expression of Osf2/Cbfa1 or in the capacity of the promoters of osteoblast-specific genes to respond to Osf2/Cbfa1.

In conclusion, we have demonstrated that human MSCs derived from vertebral bodies have characteristics in common with cells derived from other skeletal sites and can produce a mineralized ECM in vitro and that the total and osteoprogenitor (CFU-F/ALP+) colonies of MSCs decrease with donor age.

Acknowledgements

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. REFERENCES

We thank Mr. David Vazquez and Ms. Blanca Rodriguez for expert technical assistance in cell culture and cell assays, Mr. James McMannis and Ms. Topaz Kirlew from the Cell Transplantation Center (Diabetes Research Institute, Miami) for whole marrow isolation, and Ms. Ginnie Roos for help in preparing the manuscript. We thank Dr. Robert Morgan for assistance with statistical analysis of the data. Dr. Howard is the recipient of a VA Research Career Scientist award. Dr. Roos is supported by a Department of Defense grant (PC 970565).

REFERENCES

  1. Top of page
  2. Abstract
  3. INTRODUCTION
  4. MATERIALS AND METHODS
  5. RESULTS
  6. DISCUSSION
  7. Acknowledgements
  8. REFERENCES
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