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TISSUE-SPECIFIC STEM CELLS |
Institute of Transfusion Medicine and Immunology, German Red Cross Blood Service of Baden-Württemberg – Hessen, Medical Faculty Mannheim, University of Heidelberg, Heidelberg, Germany
Key Words. Mesenchymal stem cells • Fetal calf serum • Platelet-rich plasma • AB serum • Adipose tissue
Correspondence: Karen Bieback, Ph.D., Institute of Transfusion Medicine and Immunology, German Red Cross Blood Service of Baden-Württemberg – Hessen, Medical Faculty Mannheim, University of Heidelberg, Friedrich-Ebert-Str. 107, 68167 Mannheim, Germany. Telephone: +49-621-3706-8216; Fax: +49-621-3706-851; e-mail: k.bieback{at}iti-ma.blutspende.de
Received January 12, 2007;
First published online in STEM CELLS EXPRESS January 25, 2007.
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ABSTRACT |
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Disclosure of potential conflicts of interest is found at the end of this article.
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosures of Potential...AcknowledgmentsReferences |
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MATERIALS AND METHODS |
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AB-HS
Human serum was derived from whole-blood donations of 10 AB-blood-group-typed donors. Blood donations were drained into blood bags without anticoagulants and allowed to clot overnight at 4°C. Subsequently, the serum was separated by centrifugation at 2,000g for 15 minutes. The serum was aliquoted into 15-ml sterile tubes (Greiner Bio-One, Frickenhausen, Germany, http://www.gbo.com/en) and frozen at –30°C. After thawing, AB-HS of at least five donors was pooled and sterile filtered through 0.2-µm pores (Nalgene filtration device; Nalge Nunc International, Rochester, NY, http://www.nuncbrand.com).
Preparation of Platelet-Rich Plasma and Platelet Activation
Whole-blood donations of four AB-blood-group-typed donors were used to prepare one pooled platelet concentrate out of buffy coats as described in Janetzko et al. [24]. The protocol was slightly modified by suspending the pooled platelet concentrate in AB plasma of one donor instead of using a platelet stabilizer such as T-sol. Platelet count was between 2x 106 and 3 x 106 platelets per microliter for all platelet concentrates as determined by CellDyn (3200; Abbott, Wiesbaden, Germany, http://www.abbott.de). The resulting pool of platelets in plasma was termed platelet-rich plasma (PRP). Two different protocols to derive platelet factor rich supernatants were tested. First, a platelet lysate protocol derived from Doucet et al. [17] in which PRP was shock frozen twice in liquid nitrogen was developed. After 30 minutes centrifugation at 3,000g, the platelet-factor rich supernatant was sterile filtered through 0.2-µm pores and stored in aliquots of 5 ml at –80°C. Alternatively, platelets were activated by (a) 1 unit of human thrombin [19] (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com) per milliliter of cell suspension, (b) 5 µM adenosinediphosphate (ADP) (Sigma-Aldrich)/5 µM epinephrine [25] (Sigma-Aldrich), or (c) 1 mM thrombin receptor activated peptide (TRAP)-6 [26] (Sigma-Aldrich) for 45 minutes at room temperature with gentle agitation. Calcium was not used in the activation protocols to prevent increased osteogenic differentiation [18]. This was followed by centrifugation and sterile filtration as described earlier herein. After thawing, the aliquot was centrifuged again for 5 minutes at 1,500g to remove any developing clots. To prevent in vitro gel formation, 2 units of heparin (Liquemin 5000; Roche, Basel, Switzerland, http://www.roche.com) per milliliter of medium were added to the basal medium before adding the platelet factors. We selected tPRP as the best option of all platelet-activation protocols (see Results), and therefore used this for MSC isolation and expansion in repetitive experiments.
Isolation and Culture of Adipose Tissue MSCs
AT-MSCs were obtained from lipoaspirates of 10 donors (two males and eight females) ranging in age from 25 to 64 years, with a median age of 31.5 years. Lipoaspirates were obtained in accordance with the ethical standards of the local ethical committee. Two hundred milliliters of each raw lipoaspirate were processed as described previously [3]. Briefly, lipoaspirates were washed with phosphate buffered saline (PBS; Biochrom AG, Berlin, Germany, http://www.biochrom.de) and digested with 0.075% collagenase type I (Sigma-Aldrich) for 30–45 minutes at 37°C. After neutralization of collagenase activity with basal medium and supplements, the stromal vascular fraction (SVF) was derived by centrifugation at 1,200g for 10 minutes. The pellet was resuspended and filtered through a 100-µm nylon cell strainer (Falcon; Becton Dickinson, Heidelberg, Germany, http://www.bd.com). After one more centrifugation at 1,200g for 10 minutes, the resuspended SVF cells were plated into T175 cell culture flasks (Greiner Bio-One) using basal medium with (a) 10% FCS, (b) 10% AB-HS, or (c) 10% tPRP. All flasks were cultivated at 37°C in a humidified atmosphere containing 5% CO2. Nonadherent cells were removed 12–18 hours after plating by extensive washing. The resulting plastic-adherent cell population was termed AT-MSCs.
AT-MSCs were maintained under the three culture conditions described in the preceding paragraph with twice-weekly media changes until a confluence of 70%–80% was reached. At this time, cells were passaged using Trypsin/EDTA (Promocell, Heidelberg, Germany, http://www.promocell.com) and replated at a density of 200 cells per square centimeter for all passages. Viability was determined by Trypan blue exclusion (Sigma-Aldrich), and cells were counted using a Neubauer chamber (Marienfeld GmbH, Lauda-Königshofen, Germany, http://www.superior.de).
Proliferation Kinetics
AT-MSCs of all 10 donors were expanded using three different supplements. Cells were counted and passaged at a confluence of 70%–80%. If cells did not reach a minimum confluence of 50% after 21 days, the culture was discontinued. At each passage, the population doubling rate was determined by using the formula x = [log10(NH) – log10(N1)]/log10(2) [3], where N1 is the plated cell number and NH is the cell number at harvest. To calculate the cumulative population doubling rate, the population doublings for each passage were determined and then added to the previous passages. Because the cell number of plastic-adherent cells could first be counted at passage 1, the cumulative doubling rates were calculated starting at passage 2. Fold expansion rates were also determined for passages 2–6 by dividing the cell number at harvest by the plated cell number (NH/N1).
At the same time, a growth curve of AT-MSCs from three different lipoaspirates was started at passage 3. Two hundred cells per square centimeter were plated in five T25 flasks (Greiner). At days 3, 5, 7, 9, and 11, cells from one T25 flask were harvested and counted. The fold expansion from seeding at day 1 was calculated and compared for all supplements.
The use of heparin and thrombin in tPRP might have an additional proliferative effect; therefore, separate heparin and thrombin controls were started at passage 2. AT-MSCs from four lipoaspirates were cultured in basal medium with 10% FCS until passage 2, and then plated at a density of 200 cells per square centimeter in three different T75 flasks (Nunc) using basal medium with either 10% FCS, 10% FCS + 2 units of heparin per milliliter, or 10% FCS + 1 unit of thrombin per milliliter. Cells were counted and passaged at days 8, 16, 26, and 36.
Fibroblastoid Colony-Forming Unit Assays
AT-MSC frequency in primary culture at P0 was determined for all 10 donors by counting fibroblastoid colony-forming units (CFU-Fs). After isolation from raw lipoaspirates, SVF cells were plated in six-well plates (Falcon) at low densities of 1 x 103, 5 x 103, and 1 x 104 per well using the three different culture conditions. The medium was replaced twice weekly, and cultures were stopped on day 10. Then, the cell layer was fixed with methanol and stained with a Giemsa solution (Merck, Darmstadt, Germany, http://www.merck.de). AT-MSC frequency was calculated as the mean number of colonies per 1,000 cells seeded.
In Vitro Differentiation Assays
Differentiation assays into the adipogenic and osteogenic lineages were performed for AT-MSCs of all 10 donors and of all culture conditions at passage 1 and then again at passage 7 to assess whether differentiation capacities were retained throughout long-term culture. Osteogenic differentiation was confirmed by the increase of alkaline phosphatase (AP) expression by histochemical staining following manufacturer's instructions (leukocyte alkaline phosphatase kit 85L-3R; Sigma-Aldrich). The formation of a hydroxyapatite matrix was confirmed by the van Kossa stain as described in Kern et al. [3].
Adipogenic differentiation was induced in 100% confluent AT-MSC cultures with three cycles of alternating induction and maintenance medium. The developing lipid vacuoles were stained with Oil Red. All osteogenic and adipogenic media consisted of bullet kits by Cambrex, which included osteogenic and adipogenic basal media in addition to the supplements.
Flow Cytometry Analysis
Immunophenotypic analyses with flow cytometry were performed for AT-MSCs at passage 1 for four donors and again at passage 7 for three of the same donors. The cells used for analyses at passage 1 were cryopreserved in FCS with 10% dimethyl sulfoxide (DMSO) for cells cultured in FCS, or in 4% Albumin-isotonic sodium-chloride solution and 10% DMSO for all cells cultured in either AB-HS or tPRP. Before flow cytometry analysis, cells were thawed and cultured for at least 1 week.
At passage 1, the following mouse anti-human antibodies were used: CD31-phycoerythrin (PE), CD34-PE, CD29-fluorescein isothiocyanate (FITC; Becton Dickinson), CD45-PE (Diatec, Oslo, Norway, http://www.diatec.com), CD144-PE (Beckman Coulter, Krefeld, Germany, http://www.beckmancoulter.com), CD14-PE, CD73-PE, CD90-Cy5, CD44-FITC, human leukocyte antigen (HLA)-ABC-FITC, HLA-DR-FITC (Immunotech-Coulter, Marseille, France, http://www.beckmancoulter.com/products/pr_immunology.asp), CD133/1-PE (Miltenyi Biotech, Bergisch Gladbach, Germany,http://www.miltenyibiotec.com), and vascular endothelial growth factor receptor (VEGFR)-2-PE (R&D Systems, Minneapolis, http://rndsystems.com). CD14-PE, CD31-PE, CD44-FITC, CD45-PE, CD73-PE, CD90-Cy5, HLA-ABC-FITC, and HLA-DR-FITC were again reassessed at passage 7. Mouse isotype antibodies served as control (Becton Dickinson; Beckman Coulter). Ten thousand labeled cells were acquired and analyzed using a FACScan flow cytometer running CellQuest software (Becton Dickinson).
Human Cytokine Antibody Array
Human AB-HS of all 10 donors and tPRP of all preparations were pooled, respectively, and their cytokine profiles were analyzed with a semiquantitative human cytokine antibody array that detects 174 cytokines in one experiment (RayBio Human Cytokine Antibody Array G series 2000; Tebu-bio GmbH, Berlin, http://www.tebu-bio.com). Total protein content of all supplements was determined with the Bradford reaction. The array consisted of three glass slides (array 6, 7, and 8) that were pretreated according to the manufacturer's instructions and incubated with 10-fold diluted AB-HS or tPRP for 2 hours. All sample measurements were performed in duplicate. Because of the human specificity of the array, FCS was not tested. The array glass slides were washed, incubated with a biotin-conjugated anti-cytokine mix for 2 hours, washed again, and developed for 2 hours with Cy3-conjugated streptavidin. The signals were detected using a laser scanner (GMS 418 array scanner; Affymetrix, Santa Clara, CA, http://www.affymetrix.com) and analyzed with ArrayVision version 7 (Imaging Research Inc., St. Catharines, ON, Canada, http://www.imagingresearch.com). Signals were normalized using positive, negative, and internal controls included on the array.
Statistical Analysis
Data are presented as arithmetic mean ± SD. A two-sided paired Student's t test was used to compare the mean values of cumulative population doublings and cell numbers, as well as to analyze flow cytometry data. Differences were considered to be significant at p < .05. All statistical tests were performed using the SAS software package (version 9; SAS Institute, Cary, NC, http://www.sas.com).
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RESULTS |
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Morphology and Growth Pattern
AT-MSCs of all culture conditions displayed a characteristic fibroblast-like morphology (Fig. 1A–1C). However, cells cultured with AB-HS and tPRP appeared smaller and more spindle-shaped with fewer cytoplasmic processes than did cells cultured with FCS, and had a pronounced clustering of vesicles around the nucleus. In primary culture, AB-HS (Fig. 1B) and tPRP (Fig. 1C) cells had a much denser growth on day 6 compared with FCS cells (Fig. 1A). This resulted in higher cell numbers harvested from the same culture area compared with FCS. Starting at passage 3–4, AT-MSCs cultured in AB-HS and tPRP showed a distinct mesh-like growth pattern (Fig. 1E, 1F), which was even more pronounced in later passages, as seen in Figure 1H and 1I. This mesh-like growth pattern was most evident for tPRP cells and prevented the formation of a confluent monolayer.
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Furthermore, AT-MSCs cultured with AB-HS and tPRP had decreased adhesion potential compared with AT-MSCs cultured with FCS. In the FCS condition, cells detached from plastic after 5–6 minutes of trypsination, whereas AB-HS and tPRP cells had already detached after 3–4 minutes. Even without trypsination, tPRP cells tended to detach from the middle of the cell clusters on reaching higher cell densities (Fig. 1I).
Isolation and Expansion Potential
The isolation of AT-MSCs from lipoaspirates was successful for all donors (n = 10) irrespective of culture condition. Cumulative population doublings were calculated for all passages to compare the effect of different supplements on the proliferative capacity of AT-MSCs. We showed that cumulative expansion rates were significantly higher when MSCs were cultured in AB-HS or tPRP rather than in FCS (Fig. 2A; p < .001). However, we did not observe a significant difference between AB-HS and tPRP (p = .37). AB-HS and tPRP exerted an early and strong proliferative effect on AT-MSC growth that leveled off after 40 ± 2.5 population doublings at passage 8 and proceeded at a slower rate until a maximum population doubling of 54 for tPRP and 50 for AB-HS, both at passage 11. On the contrary, AT-MSCs cultured in FCS exhibited a slow but continuous proliferation, reaching 31 ± 1.3 population doublings only at passage 8 and proceeding to the same maximal population doubling of 54 at passage 14.
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Colony Frequency
The CFU-F frequency of AB-HS and tPRP determined in primary culture did not differ significantly from that of FCS. We calculated the CFU-F frequency to be 1 in 1,000 seeded cells, independent of the supplement used (FCS, 0.7 ± 0.5; AB-HS, 0.7 ± 0.6; tPRP, 0.6 ± 0.5). However, the colonies differed in size and cell densities instead of number, indicating an increased proliferation of cells within the colonies (Fig. 3S–3U).
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Differentiation Capacity
The differentiation potential of AT-MSCs from all donors and culture conditions was tested for the adipogenic and osteogenic lineages at passages 1 and 7. Most importantly, we demonstrated that AB-HS and tPRP did not adversely affect the differentiation capacity of AT-MSCs.
Adipogenic differentiation was confirmed by the formation of lipid vacuoles that were stained by the oil red O (Fig. 3A–3C). Osteogenic differentiation was indicated by deposition of a mineralized matrix stained with van Kossa (Fig. 3G–3I) and the increased expression of alkaline phosphatase (Fig. 3M–3O).
We showed that all samples within the adipogenic and osteogenic differentiation assays were consistently positive defined by the appropriate stain. The differentiation capacities for FCS, AB-HS, and tPRP were comparable and was maintained throughout long-term culture.
Characterization of Surface Marker Expression
The expression of cell surface markers was analyzed by flow cytometry at passages 3 (n = 4) and 7 (n = 3) for all culture conditions (Table 2). All AT-MSCs were highly positive for CD29, CD44, CD73, and CD90, although FCS cells had a significantly higher CD44 expression intensity in passage 3 compared with AB-HS and tPRP (p < .05). The mean fluorescence intensity (MFI) for FCS was 843.4 ± 245.7, whereas AB-HS had an MFI of 317.7 ± 61.2 and tPRP an MFI of 416 ± 98.4. In addition, AT-MSCs of all culture conditions stained positive for HLA-class I and negative for HLA-class II, CD133, and endothelial markers CD31, CD144, and VEGFR-2. All AT-MSCs were negative for the hematopoietic marker CD34; however, there was a marked side population of CD45- and CD14-positive cells in passage 3 (Fig. 4) of AB-HS (CD45, 61% ± 2.1%; CD14, 56.2% ± 9.9%) and tPRP cells (CD45, 30.5% ± 8.3%; CD14, 30.6% ± 13%), compared with FCS (CD45, 11.6% ± 11.9%; CD14, 9.5% ± 12.1; p < .05). This CD45-/CD14-positive population was extremely heterogeneous, varying widely between AT-MSCs of different donors (also for FCS cultures). Subsequent passaging tended to diminish this population (p < .05). At passage 7, we found 3.0% ± 1.3% CD45- and 2.8% ± 3.6% CD14-positive cells for FCS, but still 5.0% ± 4.1% CD45- and 12.4% ± 15.1% CD14-positive cells for AB-HS and 10.5% ± 10.0% CD45- and 14.1% ± 14.4% CD14-positive cells for tPRP.
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Cytokine Profile
To characterize the cytokine and growth factor profile of AB-HS and tPRP, we used a semiquantitative human cytokine antibody array that detects 174 cytokines in one experiment. First, we determined total protein content of all culture supplements with the Bradford reaction. AB-HS contained 90 mg/ml total protein, tPRP 97.2 mg/ml, and FCS 87 mg/ml. Only AB-HS and tPRP were used in the cytokine antibody array because of the anti-human antibody specificities. For analysis, the internal negative controls were used to determine the cut-off rate for a positive signal. Intensities up to 20,000 were regarded as a weak signal, 20,000–40,000 as a moderate signal, and > 40,000 as a strong signal. If the difference of signal intensity between a factor in AB-HS and tPRP differed by more than twofold, it was considered to be a significant difference. In total, 43 of 174 chemokines were not detectable in AB-HS or tPRP (for a complete list, see supplemental online data). Another 83 factors had a weak signal intensity for both AB-HS and tPRP. Thirty-two factors were either negative or weakly positive in one or the other. There were 16 factors that had a moderate or strong signaling intensity. Of all 174 factors tested, only four chemokines had a significantly higher signal in AB-HS than in tPRP: angiogenin, PARC (pulmonary and activation-regulated chemokine), HCC-4 (hemofiltrate CC chemokine 4), and IL (interleukin)-13 receptor 
2. Conversely, another five factors had a higher signaling intensity in tPRP compared with AB-HS: MCP-2 (monocyte chemoattractant protein 2), GRO (growth related oncogene), uPAR (urokinase plasminogen activator receptor), ICAM-2 (intercellular adhesion molecule 2), and LAP (liver activator protein).
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DISCUSSION |
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This is the first study that compared the in vitro effects of pooled human AB serum and thrombin-activated PRP on AT-MSCs with FCS. Isolation and expansion of AT-MSCs were successful for all donors independent of medium supplement, and morphology was comparable, although AB-HS and tPRP exhibited a more spindle-shaped morphology. Kobayashi et al. [21] have shown that this could be an indication of increased motility.
We show that human AB serum as well as tPRP have a significantly higher proliferative effect on AT-MSC than FCS providing a 1,320–1,400-fold expansion in 11 days compared with only 254-fold expansion with FCS. The fact that both alternative supplements trigger such an early explosive growth rate makes them attractive candidates for MSC expansion in clinical settings, since the large number of MSC required for transplantation [31] can be achieved in a shorter period of time.
We also demonstrated that AT-MSC cultured with AB-HS or tPRP consistently differentiated into the adipogenic and osteogenic lineages and retained their differentiation capacity throughout long-term culture. Gruber et al. observed that thrombin-activated platelets decreased osteogenic differentiation of bone marrow derived MSC [19], whereas Doucet et al. postulated that platelet lysate stimulated osteogenic differentiation [17]. We did not find a decrease in the number of samples displaying osteogenic differentiation. Van Kossa and AP staining seemed to be even more intense for AB-HS and tPRP, however, this could be due to the increased proliferation of AT-MSC rather than a true increase in differentiation. Future studies will have to quantify whether human substitutes affect the number and intensity of differentiation or bias differentiation into one direction or the other.
Surface marker expression did not differ markedly between the different cell culture conditions and remained stable throughout long-term culture with the exception of a distinct CD45-/CD14-positive side population for AB-HS and tPRP cells in comparison to FCS which contained a much smaller number of CD45-/CD14-positive cells. The increased CD45-/CD14-positive population in AB-HS and tPRP could be due to cytokines such as HCC-4 and Flt-3-ligand which are present in serum/plasma and are chemoattractants for monocytes. The presence of this population was extremely donor dependent for all culture conditions and could relate to the AT-MSC isolation technique. Yoshimura et al. have shown that the fluid portion of the lipoaspirate contains a higher proportion of CD45+ cells than the fatty portion [32]. Although this CD45-/CD14-positive population diminishes with increased passaging time, we consider it essential to eliminate these contaminating cells before clinical application.
We also observed that the mean fluorescence activity of CD44 was decreased for AB-HS and tPRP in the early passages, possibly due to the presence of bFGF in serum/plasma which has been shown to downregulate CD44 [33]. CD44 is involved in cell-cell and cell-matrix interactions, therefore being involved in extravasation and homing [34]. CD44 expression may relate to the proliferative strength as when expansion of AT-MSC in HS-AB and tPRP ceases after long-term culture, levels of CD44 expression are similar across culture conditions.
There has been much controversy about what constitutes a suitable FCS supplement. Most studies have concluded that serum-free media cannot promote MSC growth without the addition of cytokines [35, 36]. This is possibly due to the fact that serum induces intracellular Ca2+ oscillations which are vital to MSC proliferation and differentiation [37]. Recently, several studies have concentrated on the use of autologous serum [11, 16, 23, 36]. Yet, this approach is limited by the amount of autologous serum necessary to expand MSC for clinical use [38] and the variability of serum, especially for patients receiving prior chemotherapy. Some studies have been successful in isolating and expanding MSC using AB serum [23, 39]; however, others have reported growth arrest of MSCs after the first passage [11, 22]. We demonstrate that AT-MSC can be efficiently kept throughout long-term culture, with AB serum reaching the same final cumulative population doublings as FCS. In our study, we used serum donors of the German Red Cross Blood Service who were all carefully screened for the absence of HLA allo-antibodies. Possibly, this quality control contributed to the effectiveness of AB-HS in our study. Alternatively, there is evidence that tissue source (i.e., adipose tissue vs. bone marrow) may play a role as well (A. Hecker, H. Lannert, unpublished data).
Besides human serum, platelet-factor rich supernatant has been described as a potential substitute for FCS [17–19]. On the basis of our study results, we conclude that thrombin-activated PRP is the optimal way to achieve a platelet-factor-rich supernatant. Platelet activation with thrombin closely imitates the physiologic activation of platelets, ensuring the bioactivity of secreted growth factors [40]. Also the release of small-membrane vesicles, termed microparticles [41], has an additional mitogenic effect [42]. Furthermore, the presence of thrombin-cleaved osteopontin could have an effect as well, because it promotes cell attachment and spreading [43]. In comparison, platelet lysate does not provide physiologic activation and contains aggregates of platelet membranes that are difficult to remove without compromising the proliferative effect. Furthermore, membrane-associated platelet antigens pose the threat of immunological responses [44]. However, clearing AT-MSC of platelet membranes before clinical administration seems to be technically difficult. Therefore, we favor the use of tPRP even though there is evidence that platelet lysate promotes MSC growth as well [17].
In conclusion, we show that pooled human AB-HS as well as tPRP enhance MSC proliferation without compromising differentiation capacity or immunophenotype. Our results provide a further step in the direction of producing an off-the-shelf animal-free medium supplement for that is widely available and prepared in as few processing steps as possible. We promote pooled AB serum as the optimal choice on the basis of the ease of preparation and the fact that further additives such as heparin and thrombin in tPRP are not included (Table 3). In addition, AB-HS in and of itself provides a good enough proliferative stimulus such that further addition of growth factors such as fibroblast growth factor-2 does not seem necessary. Furthermore, the decreased adhesion potential of tPRP cells may have an impact on the clinical application of platelet-derived alternatives. The premature culture surface detachment of tPRP cells could possibly be a sign of a change in the in vivo homing and docking ability of these cells. Therefore, we are currently addressing this issue in molecular and functional experiments. Although we consider it necessary to conduct further studies, our study is a first step toward a GMP-compliant manufacturing protocol of AT-MSCs.
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DISCLOSURES OF POTENTIAL CONFLICTS OF INTEREST |
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ACKNOWLEDGMENTS |
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REFERENCES |
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