設計熱敏感表面塗佈細胞外間質用於人羊水幹細胞的分化

簡易檢索 / 詳目顯示

回結果列表

研究生：	黃于茹 Yu-Ru Huang
論文名稱：	設計熱敏感表面塗佈細胞外間質用於人羊水幹細胞的分化 Design of Thermoresponsive Surface Immobilized with ECM for Differentiation of Human Amniotic Fluid Stem Cells
指導教授：	樋口亞紺 Akon Higuchi
口試委員:
學位類別：	碩士 Master
系所名稱：	工學院 - 化學工程與材料工程學系 Department of Chemical & Materials Engineering
論文出版年：	2019
畢業學年度：	107
語文別：	英文
論文頁數：	171
中文關鍵詞：	熱敏感、羊水幹細胞、成骨分化
外文關鍵詞：	thermoresponsive polymer, hAFSCs
相關次數：	點閱：7 下載：0
分享至:	分享至facebook 分享至twitter

查詢本校圖書館目錄查詢臺灣博碩士論文知識加值系統勘誤回報

人羊水幹細胞（hAFSCs）是多能胚胎細胞，它能夠分化成多個譜系。分離的hAFSCs貼附和增殖取決於所使用的基質。微環境在hAFSCs的分化和基因表達中扮演著關鍵的角色。因此，在本研究中，我們設計了hAFSCs的培養方法，其中將不同的基質（細胞外間質或熱敏感聚合物）塗覆被在組織培養聚苯乙烯（TCPS）培養皿上，可以在成骨細胞和軟骨細胞獲得更高的細胞增殖和分化能力。我們培養了使用不同基質的hAFSCs，例如TCPS以及TCPS塗有（a）基質膠，（b）Synthemax II，（c）人類重組-玻璃粘連蛋白（rVN），（d）第一型膠原蛋白，（e）纖連蛋白（f）聚（N-異丙基丙烯酰胺）（PolyNIPAAm），（g）聚（N-異丙基丙烯酰胺-丙烯酸丁酯共聚物）（PolyNIPAAm-BA）和（h）用ECM固定的熱敏感聚合物（NIPAAm-ECMs）。
在這些材料中，在rVN，Synthemax II和NIPAAm-ECM上培養的hAFSCs呈現出更高的增殖和分化能力。
此外，我們研究了使用間充質幹細胞（MSCs）衍生為視網膜色素上皮細胞（RPE）的潛力和可行性。先前的研究已經使用人類胚胎幹細胞（hESC）或人類誘導性多能幹細胞（hiPSC）來分化成RPE。然而，這可能導致免疫排斥和畸胎瘤產生的問題。對於使用間充質乾細胞（MSCs），例如hAFSC衍生的自RPE，應該更具前瞻性。
而且我們開發了一種新的細胞培養基，它使用人血小板裂解液（hPL）代替胎牛血清（FBS）作為補充劑，以提高hAFSCs的分化率，並建立更完整的hAFSCs無異種培養條件作為未來的臨床應用。

Human amniotic fluid stem cells (hAFSCs) are pluripotent fetal cells, which are capable to differentiate into multiple lineages. The isolated hAFSCs adhesion and proliferation depending on the substrates used. Microenvironment plays a key role in differentiation and gene expression for hAFSCs. Therefore, in this study, we designed a culture method of hAFSCs where the different substrates (extracellular matrices or thermoresponsive polymers) were coated on tissue culture polystyrene (TCPS) dishes could get higher cell proliferation and differentiation ability into osteoblasts and chondrocytes. We cultivated the hAFSCs where on different substrates were used such as TCPS and TCPS coated with (a) Matrigel, (b) Synthemax II, (c) human recombinant-vitronectin (rVN), (d) collagen type I, (e) fibronectin, (f) poly(N-isopropylacrylamide) (PNIPAAm), (g) poly(N-isopropylacrylamide-co-butylacrylate) (PNIPAAm-BA) and (h) thermoresponsive polymers immobilized with ECMs (PNIPAAm-ECMs).
Among these materials hAFSCs cultured on rVN, Synthemax II and PNIPAAm-ECMs presented the higher proliferation and differentiation abilities.
Moreover, we investigated the potential and feasibility of mesenchymal stem cells (MSCs) derived into retinal pigment epithelium (RPE). Previous studies have used human embryonic stem cells (hESCs) or human induced pluripotent stem cells (hiPSCs) to differentiate into RPE. However, this may cause immune rejection and teratoma production problems. It should be more prospective for mesenchymal stem cells (MSCs) such as hAFSCs derived into RPE.
Furthermore, a new cell culture medium was developed, which used human platelet lysate (hPL) instead of fetal bovine serum (FBS) as a supplement to improve the differentiation ratio of hAFSCs and built up a more complete hAFSCs with xeno-free culture conditions for clinical application in future.

Index of content
Chapter 1.    INTRODUCITON    1
1-1    Regenerative medicine for cell therapy    1
1-2    Stem cells    2
1-2.1    Totipotent stem cells    3
1-2.2    Pluripotent stem cells    3
1-2.3    Multipotent stem cells    5
1-2.4    Unipotent stem cells    7
1-3    Amniotic fluid    7
1-3.1    Definition    7
1-3.2    Developmental diagnosis using amniotic fluid    8
1-3.3    Content of amniotic fluid    8
1-3.4    Clinical significance and potential of amniotic fluid    9
1-3.5    Amniotic fluid cells (AFs)    10
1-3.5.1    AF-type cells    11
1-3.5.2    F-type cells    11
1-3.5.3    Epithelioid E-type cells    12
1-4    Amniotic fluid stem cells (AFSCs)    14
1-4.1    Isolation of amniotic fluid stem cells    14
1-4.2    Characterization of amniotic fluid stem cells    15
1-4.3    Pluripotency of Amniotic fluid stem cells    17
1-5    Stem cell microenvironments    19
1-5.1    Cell-soluble factor interactions    20
1-5.2    Cell-cell interactions    22
1-5.3    Cell-biomaterials interactions    22
1-5.4    Physical factors    23
1-6    ECMs    24
1-6.1    What is ECM?    24
1-6.2    The function of ECM    25
1-6.3    The component of ECM    26
1-6.4    ECM receptors    26
1-6.4.1    Matrigel (M)    29
1-6.4.2    Collagen (Col)    30
1-6.4.3    Fibronectin    31
1-6.4.4    Vitronectin    32
1-7    Human platelet lysate (hPL)    33
1-7.1    Preparation of human platelet lysate    36
1-7.2    Processing of platelet concentrates into hPL    37
1-7.3    Components within human platelet lysate    39
1-8    Thermoresponsive polymer and characteristics    42
1-9    Goal of the study    45
Chapter 2.    MATERIALS AND METHODS    46
2-1    Experimental materials    46
2-1.1    Cell source for cultivation    46
2-1.2    Cell culture dishes coated with ECM    47
2-1.3    Materials for thermoresponsive surface    48
2-1.4    Differentiation of hAFSCs    48
2-1.5    Characteristic evaluation of hAFSCs    50
2-2    Experimental instrument    52
2-3    Experimental methods    52
2-3.1    Preparation of the cell culture medium    52
2-3.2    Cell cultivation    53
2-3.3    Passage of the hAFSCs    54
2-3.4    Cell density measurement    55
2-3.5    Preparation of extracellular matrix (EMC) coated dishes    56
2-3.6    Preparation of thermoresponsive polymer coated with extracellular matrix (ECM) dishes    56
2-3.7    Pluripotent gene expression analysis    57
2-3.8    Immunofluorescence staining    59
2-3.9    Flow cytometry analysis    60
2-3.10    Osteogenic differentiation    60
2-3.11    Adipogenic differentiation of hAFSCs    62
2-3.12    Chondrogenic differentiation of hAFSCs    63
2-3.13    Quantitative analysis of differentiation    63
2-3.14    Retinal pigment epithelium (RPE) differentiation of hAFSCs    64
Chapter 3.    RESULT AND DISSCUSION    67
3-1    Cultivation of hAFSCs    67
3-1.1    The morphology of hAFSCs on ECMs-coating, polymer-coating, and polymer-ECM-coating dishes    68
3-1.2    The effect of human platelet lysate on hAFSCs culture    76
3-2.1    Osteogenic differentiation of hAFSCs cultured on ECMs, thermoresponsive polymers coating dishes    86
3-2.2    Development of home-made differentiation medium for inducing hAFSCs    94
3-2.3    Materials surface analysis - AFM (atomic force microscope)    108
3-2.4    Retinal pigment epithelium differentiation of hAFSCs    110
Chapter 4.    Conclusion    131
Supplementary data    135
S3-2.5    Neural differentiation of hAFSCs cultured on ECMs, thermoresponsive polymers coating dishes    135
S3-2.6    Adipocyte differentiation of hAFSCs cultured on ECMs, thermoresponsive polymers coating dishes    139

 
Index of figure
Figure 1-1    Outline of Bone Marrow Transplant.    1
Figure 1-2    Classification of stem cells on basis of their potency.    2
Figure 1-3    Derivation and differentiation potential of ESCs.    4
Figure 1-4    Developmental model of cell differentiation and dedifferentiation.    4
Figure 1-5    The mesengenic process. Mesenchymal stem cells are multipotent and possess the ability to proliferate and commit to different cell types based on the environmental conditions. They also may be redirected from one lineage to another. (https://www.frontiersin.org/articles/10.3389/fimmu.2013.00201/full).    7
Figure 1-6    The fetus is surrounded by amniotic fluid.    8
Figure 1-7    Morphological differences in the primary cells. A – The morphology of the AF-type cells. i. A phase-contrast image of a primary AF-type cell colony on the fifth day after seeding the amniotic fluid (magnification: 100x). ii. A typical AF-type cell colony on the eighth day after seeding (magnification: 40x). B – The morphology of the F-type cells. i. A phase-contrast image of a primary F-type cell colony on the seventh day after seeding (magnification: 100x). ii. A typical AF-type cell colony on the ninth day after seeding (magnification: 40x). C – The morphology of the E-type cells. i. A phase-contrast image of a primary E-type cell colony on the fifth day after seeding (magnification: 40x). ii. An enlarged photo of Fig. 1Ci (magnification: 100x). P = passage; D = day; AF = AF-type; F = F-type; E = E-type [58].    12
Figure 1-8    Extraction of amniotic fluid (Amniocentesis).    13
Figure 1-9    The time line of pregnant period.    14
Figure 1-10    Expression of MSC surface and postmitotic neuron markers by flow cytometry of hAFSCs [69].    16
Figure 1-11    Microenvironment of stem cells [86].    20
Figure 1-12    Effects of soluble factors on bone marrow mesenchymal stem cells (MSCs) trilineage differentiation [87].    21
Figure 1-13    Regulation of cell behavior by ECM [98].    23
Figure 1-14    Stem cells derive the tissues across that body that vary in stiffness of wide scales [100].    24
Figure 1-15    Extracellular matrix (ECM). Typical components include collagen, proteoglycans, bronectin, and laminin [101].    24
Figure 1-16    A model of adhesion between integrins and ECM.    26
Figure 1-17    Molecular structure of fibronectin. The various structural domains as well as binding sites for fibronectin (FN), fibrin, collagen, cells, and heparin are indicated. The amino acid sequences RGD and PHSRN constitute the major binding site for integrin α5β1 and second site that supports RGD-mediated adhesion, respectively [124].    32
Figure 1-18    The modular structure of vitronectin and its binding domains.    33
Figure 1-19    Platelet derived growth factors in wound healing [129].    34
Figure 1-20    Different modalities of platelet concentrate preparation [129].    37
Figure 1-21    Preparation of allogeneic pooled human platelet lysate (pHPL) from platelet concentrates [129].    38
Figure 1-22    Platelet granule cargo. The various types of platelet granules store a plethora of potent substances including lysosomal enzymes, coagulation factors, immunologic and adhesion molecules, chemokines and growth factors for hemostasis, host defense, angiogenesis and tissue repair [129].    40
Figure 1-23    Curves showing phase transition phenomenon. (a) Lower critical solution temperature (LCST) and (b) upper critical solution temperature (UCST) phase transition behaviors of thermo-responsive polymers in solution [159].    43
Figure 1-24    The structures of thermoresponsive polymers. (a) poly(N-isopropylacrylamide) (PNIPAAm). (b) poly (acrylic acid) (PAA). (c) polyacrylamide (PAAm). (d) poly (N-isopropylacrylamide-co-butylacrylate) (PNIPAAm-co-BA)    43
Figure 1-25    The mechanism for the hydrophilicity of PNIPAAm.    44
Figure 2-1    The counting grid pattern.    55
Figure 2-2    The thermoresponsive polymer surface dishes coating procedures    57
Figure 2-3    The timeline of RPE differentiation method I.    65
Figure 2-4    The time line of RPE differentiation method II.    66
Figure 3-1    Morphology of hAFSCs on different substrates-coating dishes at day 4. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm    71
Figure 3-2    The doubling time of hAFSCs on ECM or Polymer-coating dishes at Passage 7 for 4 days. (Seeding density: 2×104 /well).    72
Figure 3-3    Morphology of hAFSCs on different substrates-coating dishes at day 5. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm    74
Figure 3-4    The doubling time of hAFSCs on ECM or Polymer-coating dishes at Passage 7 for 5 days. (Seeding density: 2×104 /well).    75
Figure 3-5    Morphology of hAFSCs on different substrates-coating dishes at day 4. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm    80
Figure 3-6    The doubling time of hAFSCs on ECM or Polymer-coating dishes at Passage 4 for 4 days. (Seeding density: 2×104 /well).    81
Figure 3-7    Morphology of hAFSCs on different substrates-coating dishes at day 4. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm    83
Figure 3-8    The doubling time of hAFSCs on ECM or Polymer-coating dishes at Passage 5 for 4 days. (Seeding density: 2×104 /well).    83
Figure 3-9    Compare doubling time of 20% FBS and 10% hPL culture medium on thermoresponsive surface coated with ECMs    84
Figure 3-10    The mesengenic process. Mesenchymal stem cells are multipotent and possess the ability to proliferate and commit to different cell types based on the environmental conditions. They also may be redirected from one lineage to another (https://www.frontiersin.org/articles/10.3389/fimmu.2013.00201/full).    85
Figure 3-11    The time line of osteogenic differentiation of hAFSCs.    86
Figure 3-12    The morphology of hAFSCs after inducing 14 days with MODM (commercial product) inducing medium differentiation into osteoblast. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 7    88
Figure 3-13    ALP activity (early stage marker of osteoblasts) of hAFSCs on several substrates coating dishes after 14 days induction into osteoblast.    89
Figure 3-14    The morphology of hAFSCs after inducing 28 days with MODM (commercial product) inducing medium differentiation into osteoblast. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 7    90
Figure 3-15    Osteogenic differentiation of hAFSCs on various substrates coating dishes after 28 days induction into osteoblast. Upper row was top view and lower row was the morphology of the cells stained with Alizarin Red S under microscopy, Scale bar = 500μm. Red color sites indicate calcium deposition.    91
Figure 3-16    The quantification of staining cell ratio of Alizarin Red S staining.    92
Figure 3-17    Osteogenic differentiation of hAFSCs on various substrates coating dishes after 28 days induction into osteoblast. Upper row was top view and lower row was the morphology of the cells stained with von Kossa staining under microscopy, Scale bar = 500μm. Silver black color sites indicate calcium phosphate deposition.    93
Figure 3-18    The quantification of staining cell ratio of von Kossa staining.    94
Figure 3-19    The morphology of hAFSCs after inducing 14 days with 20% FBS (home-made) inducing medium differentiation into osteoblast. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 5, 80390.    97
Figure 3-20    ALP activity (early stage marker of osteoblasts) of hAFSCs on several substrates coating dishes after 14 days induction into osteoblast.    97
Figure 3-21    The morphology of hAFSCs after inducing 28 days with 20% FBS (home-made) inducing medium differentiation into osteoblast. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 5, 80390.    98
Figure 3-22    Osteogenic differentiation of hAFSCs on various substrates coating dishes after 28 days induction into osteoblast. Upper row was top view and lower row was the morphology of the cells stained with Alizarin Red S under microscopy, Scale bar = 500μm. Red color sites indicate calcium deposition.    99
Figure 3-23    The quantification of staining cell ratio of Alizarin Red S staining.    100
Figure 3-24    Osteogenic differentiation of hAFSCs on various substrates coating dishes after 28 days induction into osteoblast. The top view was the morphology of the cells stained with von Kossa staining.    100
Figure 3-25    The morphology of hAFSCs after inducing 14 days with 10% hPL (home-made) inducing medium differentiation into osteoblast. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 5, 80390.    102
Figure 3-26    ALP activity (early stage marker of osteoblasts) of hAFSCs on several substrates coating dishes after 14 days induction into osteoblast.    103
Figure 3-27    The morphology of hAFSCs after inducing 28 days with 10% hPL (home-made) inducing medium differentiation into osteoblast. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 5, 80390.    104
Figure 3-28    Osteogenic differentiation of hAFSCs on various substrates coating dishes after 28 days induction into osteoblast. Upper row was top view and lower row was the morphology of the cells stained with Alizarin Red S under microscopy, Scale bar = 500μm. Red color sites indicate calcium deposition.    105
Figure 3-29    The quantification of staining cell ratio of Alizarin Red S staining.    106
Figure 3-30    Osteogenic differentiation of hAFSCs on various substrates coating dishes after 28 days induction into osteoblast. The top view was the morphology of the cells stained with von Kossa staining.    106
Figure 3-31    ALP activity of hAFSCs induced into osteoblasts at 14 days of differentiation in commercial induction medium, home-made induction medium containing 20% FBS, and home-made induction medium containing 10% hPL after hAFSCs cultured on various substrates coating dishes.    107
Figure 3-32    Alizarin Red S staining of hAFSCs induced into osteoblasts at 28 days of differentiation in commercial induction medium, home-made induction medium containing 20% FBS, and home-made induction medium containing 10% hPL after hAFSCs cultured on various substrates coating dishes.    108
Figure 3-33    AFM analysis of ECMs, NIPAAm and NIPAAm-ECM coating dishes. AFM topography images.    109
Figure 3-34    The Rq value (roughness) on these coating substrates dishes.    109
Figure 3-35    The time line of RPE differentiation of hAFSCs (Schwartz., 2012).    110
Figure 3-36    The time line of RPE differentiation of hAFSCs (Osakada., 2009).    110
Figure 3-37    The morphology process of hAFSCs differentiation into RPE (Schwartz., 2012). 40X: Scale bar = 500μm    111
Figure 3-38    The morphology process of hAFSCs differentiation into RPE (Schwartz., 2012). 40X: Scale bar = 500μm    113
Figure 3-39    The morphology process of hAFSCs differentiation into RPE (Schwartz., 2012). 40X: Scale bar = 500μm    116
Figure 3-40    The morphology process of hAFSCs differentiation into RPE (Schwartz., 2012). 40X: Scale bar = 500μm    119
Figure 3-41    The morphology process of hAFSCs differentiation into RPE (Schwartz., 2012). 40X: Scale bar = 500μm    121
Figure 3-42    The morphology process of hAFSCs differentiation into RPE (Osakada., 2009). 40X: Scale bar = 500μm    122
Figure 3-43    RPE65 expression of hAFSCs-derived RPE analyzed by flow cytometry.    123
Figure 3-44    The morphology process of hAFSCs differentiation into RPE (Osakada., 2009). 40X: Scale bar = 500μm.    124
Figure 3-45    RPE65 expression of hAFSCs-derived RPE analyzed by flow cytometry.    124
Figure 3-46    The morphology process of hAFSCs differentiation into RPE (Zhang L et al., 2011) [170]. 40X: Scale bar = 500μm    125
Figure 3-47    RPE65 expression of hAFSCs-derived RPE analyzed by flow cytometry.    126
Figure 3-48    The morphology process of hAFSCs differentiation into RPE (Zhu, J., 2013 & Leach, L. L., 2016). 40X: Scale bar = 500μm    127
Figure 3-49    RPE65 expression of hAFSCs-derived RPE analyzed by flow cytometry.    127
Figure 3-50    The morphology process of hAFSCs differentiation into RPE (Leach LL., 2016) [173]. 40X: Scale bar = 500μm    128
Figure 3-51    RPE65 expression of hAFSCs-derived RPE analyzed by flow cytometry.    129
Figure 4-1.    The main idea of developing optimal hAFSCs culture and differentiation.    131
Figure 4-2.    The doubling time of hAFSCs cultivated with 20% FBS cultured on ECM, polymers and polymer immobilized with ECM-coating dishes.    132
Figure 4-3.    The doubling time of hAFSCs cultivated with 10% hPL cultured on ECMs, polymers and polymer immobilized with ECM-coating dishes.    132
Figure 4-4.    The osteogenic differentiation ratio of hAFSCs after culture on ECM-coating dishes, polymer-coating dishes and polymer immobilized with ECM-coating dishes with commercial (5%FBS), 20% FBS, and10% hPL contained in inducing medium.    134

 
Index of table
Table-1-1    Summary of hMSCs sources, cell surface markers and expansion media with serum supplements [11]    6
Table 1-2    In vitro growth potentials of colony-forming amniotic fluid cell types.    13
Table 1-3    Expression of some markers at varying stem cells.    17
Table 1-4    Lists of markers for AFSCs [12, 61, 81].    19
Table 1-5    Medium for induction of lineage-specific differentiation [84, 85].    19
Neurogenic differentiation (Ectoderm)    19
Table 1-6    Ligand-binding specificities of human integrins [111].    28
Table 1-7    Collagen types, forms and distribution [121].    31
Table 1-8    Advantages and disadvantages of FBS and hPL [128].    35
Table 2-1    Probes of pluripotent gene    51
Table 2-2    The formula of ECM-coating dishes.    56
Table 2-3    Composition of RNA/primer solution in PCR reaction    58
Table 2-4    Formula of qRT-PCR reaction.    58
Table 2-5    qRT-PCR step condition.    59
Table 2-6    Formula of home-made osteogenic differentiation medium.    61
Table 3-1    The doubling time of hAFSCs on ECM or Polymer-coating dishes at Passage 7.    72
Table 3-2    The doubling time of hAFSCs on substrates-coating dishes at Passage 7 for 5 days.    75
Table 3-3    The doubling time of hAFSCs on substrates-coating dishes at Passage 4 for 4 days.    81
Table 3-4    The doubling time of hAFSCs on substrates-coating dishes at Passage 5 for 4 days.    84
Table 3-5    The osteogenic induction medium developed in this study.    96
Table 3-6    The results sorting of RPE differentiation method I of hAFSCs.    121
Table 3-7    The results sorting of RPE differentiation method II of hAFSCs.    130


Supplementary data
Figure S3-52    The time line of neural differentiation of hAFSCs.    135
Figure S3-53    The morphology of hAFSCs expansion for 7 days before neural differentiation. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 3    136
Figure S3-54    The morphology of hAFSCs after inducing 21 days with HNEU.D. Media-450 (commercial product) inducing medium differentiation into neural. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 3    137
Figure S3-55    The immunofluorescence staining of pluripotent proteins detected on hAFSCs cultured on ECMs, thermoresponsive polymers coating dishes passage 4. (A) Hoechst (nuclei) expression, (B) Nestin expression, (C)β-III tubulin expression, and (D) merged pictures. Scale bar = 100μm.    138
Figure S3-56    The time line of adipocyte differentiation of hAFSCs.    139
Figure S3-57    The morphology of hAFSCs after inducing 21 days with StemPro™ Adipogenesis Differentiation Kit (No. A1007001, Gibco™) (commercial product) inducing medium differentiation into adipocyte cells. 40X: Scale bar = 500μm; 100X: Scale bar = 100μm, Passage 6.    140
Figure S3-58    The top view of Oil red staining detected the hAFSCs cultured on ECMs, thermoresponsive polymers coating dishes at passage 6.    141


                                

1. Mao, A.S. and D.J. Mooney, Regenerative medicine: Current therapies and future directions. Proceedings of the National Academy of Sciences of the United States of America, 2015. 112(47): p. 14452-14459.
2. Parveen, S., et al., New era in health care: tissue engineering. 2006. 1(1): p. 8.
3. Malard, F. and M.J.M.o.i. Mohty, New insight for the diagnosis of gastrointestinal acute graft-versus-host disease. 2014. 2014.
4. Irfan, A. and I.J.M.E.I. Ahmed, Stem Cells: The Future of Personalised Medicine? 2014. 5: p. MEI. S13177.
5. Dick, J.E.J.B., Stem cell concepts renew cancer research. 2008. 112(13): p. 4793-4807.
6. Papapetrou, E.P., et al., Stoichiometric and temporal requirements of Oct4, Sox2, Klf4, and c-Myc expression for efficient human iPSC induction and differentiation. Proceedings of the National Academy of Sciences of the United States of America, 2009. 106(31): p. 12759-12764.
7. Fischbach, G.D. and R.L. Fischbach, Stem cells: science, policy, and ethics. Journal of Clinical Investigation, 2004. 114(10): p. 1364-1370.
8. Yu, J.Y., et al., Induced pluripotent stem cell lines derived from human somatic cells. Science, 2007. 318(5858): p. 1917-1920.
9. Takahashi, K. and S. Yamanaka, Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell, 2006. 126(4): p. 663-676.
10. Bonfanti, P., et al., Microenvironmental reprogramming of thymic epithelial cells to skin multipotent stem cells. Nature, 2010. 466(7309): p. 978-U105.
11. Ullah, I., R.B. Subbarao, and G.J. Rho, Human mesenchymal stem cells - current trends and future prospective. Bioscience Reports, 2015. 35: p. 18.
12. Pittenger, M.F., et al., Multilineage potential of adult human mesenchymal stem cells. Science, 1999. 284(5411): p. 143-147.
13. in 'tAnker, P.S., et al., Amniotic fluid as a novel source of mesenchymal stem cells for therapeutic transplantation. Blood, 2003. 102(4): p. 1548-1549.
14. Tsai, M.S., et al., Isolation of human multipotent mesenchymal stem cells from second-trimester amniotic fluid using a novel two-stage culture protocol. Human Reproduction, 2004. 19(6): p. 1450-1456.
15. Cai, J.L., et al., Generation of Human Induced Pluripotent Stem Cells from Umbilical Cord Matrix and Amniotic Membrane Mesenchymal Cells. Journal of Biological Chemistry, 2010. 285(15): p. 11227-11234.
16. Wagner, W., et al., Comparative characteristics of mesenchymal stem cells from human bone marrow, adipose tissue, and umbilical cord blood. Experimental Hematology, 2005. 33(11): p. 1402-1416.
17. Zhang, X., et al., Runx2 overexpression enhances osteoblastic differentiation and mineralization in adipose - derived stem cells in vitro and in vivo. Calcified Tissue International, 2006. 79(3): p. 169-178.
18. Huang, G.T.J., S. Gronthos, and S. Shi, Mesenchymal Stem Cells Derived from Dental Tissues vs. Those from Other Sources: Their Biology and Role in Regenerative Medicine. Journal of Dental Research, 2009. 88(9): p. 792-806.
19. Seifrtova, M., et al., The response of human ectomesenchymal dental pulp stem cells to cisplatin treatment. International Endodontic Journal, 2012. 45(5): p. 401-412.
20. Schuring, A.N., et al., Characterization of endometrial mesenchymal stem-like cells obtained by endometrial biopsy during routine diagnostics. Fertility and Sterility, 2011. 95(1): p. 423-426.
21. Jiao, F., et al., Human Mesenchymal Stem Cells Derived From Limb Bud Can Differentiate into All Three Embryonic Germ Layers Lineages. Cellular Reprogramming, 2012. 14(4): p. 324-333.
22. Allickson, J.G., et al., Recent studies assessing the proliferative capability of a novel adult stem cell identified in menstrual blood. 2011. 3(2011): p. 4.
23. Ab Kadir, R., et al., Characterization of Mononucleated Human Peripheral Blood Cells. Scientific World Journal, 2012: p. 8.
24. Raynaud, C.M., et al., Comprehensive Characterization of Mesenchymal Stem Cells from Human Placenta and Fetal Membrane and Their Response to Osteoactivin Stimulation. Stem Cells International, 2012: p. 13.
25. Rotter, N., et al., Isolation and characterization of adult stem cells from human salivary glands. Stem Cells and Development, 2008. 17(3): p. 509-518.
26. Bartsch, G., et al., Propagation, expansion, and multilineage differentiation of human somatic stem cells from dermal progenitors. Stem Cells and Development, 2005. 14(3): p. 337-348.
27. Riekstina, U., et al., Characterization of human skin-derived mesenchymal stem cell proliferation rate in different growth conditions. Cytotechnology, 2008. 58(3): p. 153-162.
28. Kita, K., et al., Isolation and Characterization of Mesenchymal Stem Cells From the Sub-Amniotic Human Umbilical Cord Lining Membrane. Stem Cells and Development, 2010. 19(4): p. 491-501.
29. Morito, T., et al., Synovial fluid-derived mesenchymal stem cells increase after intra-articular ligament injury in humans. Rheumatology, 2008. 47(8): p. 1137-1143.
30. Wang, H.S., et al., Mesenchymal stem cells in the Wharton's jelly of the human umbilical cord. Stem Cells, 2004. 22(7): p. 1330-1337.
31. Hou, T.Y., et al., Umbilical Cord Wharton's Jelly: A New Potential Cell Source of Mesenchymal Stromal Cells for Bone Tissue Engineering. Tissue Engineering Part A, 2009. 15(9): p. 2325-2334.
32. Goff, L.A., et al., Differentiating human multipotent mesenchymal stromal cells regulate microRNAs: Prediction of microRNA regulation by PDGF during osteogenesis. Experimental Hematology, 2008. 36(10): p. 1354-1369.
33. Mamidi, M.K., et al., Comparative cellular and molecular analyses of pooled bone marrow multipotent mesenchymal stromal cells during continuous passaging and after successive cryopreservation. Journal of Cellular Biochemistry, 2012. 113(10): p. 3153-3164.
34. Otsuru, S., et al., Improved isolation and expansion of bone marrow mesenchymal stromal cells using a novel marrow filter device. Cytotherapy, 2013. 15(2): p. 146-153.
35. Gronthos, S., et al., THE STRO-1(+) FRACTION OF ADULT HUMAN BONE-MARROW CONTAINS THE OSTEOGENIC PRECURSORS. Blood, 1994. 84(12): p. 4164-4173.
36. Stewart, K., et al., Further characterization of cells expressing STRO-1 in cultures of adult human bone, marrow stromal cells. Journal of Bone and Mineral Research, 1999. 14(8): p. 1345-1356.
37. Pendleton, C., et al., Mesenchymal Stem Cells Derived from Adipose Tissue vs Bone Marrow: In Vitro Comparison of Their Tropism towards Gliomas. Plos One, 2013. 8(3): p. 7.
38. Zhang, X., et al., Isolation and Characterization of Mesenchymal Stem Cells From Human Umbilical Cord Blood: Reevaluation of Critical Factors for Successful Isolation and High Ability to Proliferate and Differentiate to Chondrocytes as Compared to Mesenchymal Stem Cells From Bone Marrow and Adipose Tissue. Journal of Cellular Biochemistry, 2011. 112(4): p. 1206-1218.
39. Gronthos, S., et al., Surface protein characterization of human adipose tissue-derived stromal cells. Journal of Cellular Physiology, 2001. 189(1): p. 54-63.
40. Baglioni, S., et al., Characterization of human adult stem-cell populations isolated from visceral and subcutaneous adipose tissue. Faseb Journal, 2009. 23(10): p. 3494-3505.
41. Kadar, K., et al., DIFFERENTIATION POTENTIAL OF STEM CELLS FROM HUMAN DENTAL ORIGIN - PROMISE FOR TISSUE ENGINEERING. Journal of Physiology and Pharmacology, 2009. 60: p. 167-175.
42. Moretti, P., et al., Mesenchymal Stromal Cells Derived from Human Umbilical Cord Tissues: Primitive Cells with Potential for Clinical and Tissue Engineering Applications, in Bioreactor Systems for Tissue Engineering Ii: Strategies for the Expanison and Directed Differentiation of Stem Cells, C. Kasper, M. VanGriensven, and R. Portner, Editors. 2010, Springer-Verlag Berlin: Berlin. p. 29-54.
43. Denu, R.A., et al., Fibroblasts and Mesenchymal Stromal/Stem Cells Are Phenotypically Indistinguishable. Acta Haematologica, 2016. 136(2): p. 85-97.
44. Van Keymeulen, A., et al., Distinct stem cells contribute to mammary gland development and maintenance. Nature, 2011. 479(7372): p. 189-U58.
45. Delo, D.M., et al., Amniotic fluid and placental stem cells, in Methods in Enzymology. 2006, Elsevier. p. 426-438.
46. Coady, A.M. and S. Bower, Twining's Textbook of Fetal Abnormalities E-Book: Expert Consult: Online and Print. 2014: Elsevier Health Sciences.
47. Underwood, M.A., W.M. Gilbert, and M.P.J.J.o.p. Sherman, Amniotic fluid: not just fetal urine anymore. 2005. 25(5): p. 341.
48. Eslaminejad, M.B. and S. Jahangir, Amniotic Fluid Stem Cells and Their Application in Cell-Based Tissue Regeneration. International Journal of Fertility & Sterility, 2012. 6(3): p. 147-156.
49. Jarzembowski, J., Genetic Disorders and the Fetus: Diagnosis, Prevention, and Treatment. Pediatric and Developmental Pathology, 2011. 14(1): p. 84-84.
50. Hoehn, H. and D. Salk, . Morphological and Biochemical Heterogeneity of Amniotic Fluid Cells in Culture, in Methods in cell biology. 1982, Elsevier. p. 11-34.
51. GOSDEN, C.M.J.B.m.b., Amniotic fluid cell types and culture. 1983. 39(4): p. 348-354.
52. Prusa, A.R., et al., Oct-4-expressing cells in human amniotic fluid: a new source for stem cell research? Human Reproduction, 2003. 18(7): p. 1489-1493.
53. Prusa, A.-R. and M.J.M.S.M. Hengstschlager, Amniotic fluid cells and human stem cell research: a new connection. 2002. 8(11): p. RA253-RA257.
54. Kim, J., et al., Human amniotic fluid-derived stem cells have characteristics of multipotent stem cells. Cell Proliferation, 2007. 40(1): p. 75-90.
55. Tsai, M.S., et al., Clonal amniotic fluid-derived stem cells express characteristics of both mesenchymal and neural stem cells. Biology of Reproduction, 2006. 74(3): p. 545-551.
56. Simoni, G. and R.J.J.o.p.m. Colognato, The amniotic fluid-derived cells: the biomedical challenge for the third millennium. 2009. 3(3): p. 34.
57. Fauza, D., Amniotic fluid and placental stem cells. Best Practice & Research Clinical Obstetrics & Gynaecology, 2004. 18(6): p. 877-891.
58. Zhang, S.L., et al., The heterogeneity of cell subtypes from a primary culture of human amniotic fluid. Cellular & Molecular Biology Letters, 2010. 15(3): p. 424-439.
59. Jiang, Y.H., et al., Pluripotency of mesenchymal stem cells derived from adult marrow (vol 418, pg 41, 2002). Nature, 2007. 447(7146): p. 879-880.
60. Perrone, L., et al., Postnatal weight change is influenced by mother-newborn pair leptin levels. Nutrition Research, 2000. 20(11): p. 1531-1536.
61. Zambotti, F., et al., Monoamine metabolites and related compounds in human amniotic fluid: Assay by gas chromatography and gas chromatography-mass spectrometry. 1975. 61(3): p. 247-256.
62. Loukogeorgakis, S.P. and P. De Coppi, Concise Review: Amniotic Fluid Stem Cells: The Known, the Unknown, and Potential Regenerative Medicine Applications. Stem Cells, 2017. 35(7): p. 1663-1673.
63. You, Q., et al., The Biological Characteristics of Human Third Trimester Amniotic Fluid Stem Cells. Journal of International Medical Research, 2009. 37(1): p. 105-112.
64. Nadri, S. and M. Soleimani, Comparative analysis of mesenchymal stromal cells from murine bone marrow and amniotic fluid. Cytotherapy, 2007. 9(8): p. 729-737.
65. Fuchs, J.R., et al., Diaphragmatic reconstruction with autologous tendon engineered from mesenchymal amniocytes. Journal of Pediatric Surgery, 2004. 39(6): p. 834-837.
66. Barry, F.P., et al., The monoclonal antibody SH-2, raised against human mesenchymal stem cells, recognizes an epitope on endoglin (CD105). Biochemical and Biophysical Research Communications, 1999. 265(1): p. 134-139.
67. Barry, F., et al., The SH-3 and SH-4 antibodies recognize distinct epitopes on CD73 from human mesenchymal stem cells. Biochemical and Biophysical Research Communications, 2001. 289(2): p. 519-524.
68. Kannagi, R., et al., Stage‐specific embryonic antigens (SSEA‐3 and‐4) are epitopes of a unique globo‐series ganglioside isolated from human teratocarcinoma cells. 1983. 2(12): p. 2355-2361.
69. Tan, S.M., A. Sharma, and J.B. de Haan, Oxidative stress and novel antioxidant approaches to reduce diabetic complications, in Oxidative Stress and Diseases. 2012, IntechOpen.
70. Barlow, S., et al., Comparison of Human Placenta- and Bone Marrow-Derived Multipotent Mesenchymal Stem Cells. Stem Cells and Development, 2008. 17(6): p. 1095-1107.
71. Klemmt, P.A.B., V. Vafaizadeh, and B. Groner, Murine amniotic fluid stem cells contribute mesenchymal but not epithelial components to reconstituted mammary ducts. Stem Cell Research & Therapy, 2010. 1: p. 13.
72. Sessarego, N., et al., Multipotent mesenchymal stromal cells from amniotic fluid: solid perspectives for clinical application. Haematologica-the Hematology Journal, 2008. 93(3): p. 339-346.
73. De Rosa, A., et al., Amniotic fluid-derived mesenchymal stem cells lead to bone differentiation when cocultured with dental pulp stem cells. 2010. 17(5-6): p. 645-653.
74. Suh, M.R., et al., Human embryonic stem cells express a unique set of microRNAs. Developmental Biology, 2004. 270(2): p. 488-498.
75. Draper, J.S., et al., Surface antigens of human embryonic stem cells: changes upon differentiation in culture. Journal of Anatomy, 2002. 200(3): p. 249-258.
76. Da Sacco, S., R.E. De Filippo, and L. Perin, Amniotic fluid as a source of pluripotent and multipotent stem cells for organ regeneration. Current Opinion in Organ Transplantation, 2011. 16(1): p. 101-105.
77. Kunisaki, S.M., et al., Tissue engineering from human mesenchymal amniocytes: a prelude to clinical trials. Journal of Pediatric Surgery, 2007. 42(6): p. 974-980.
78. Miki, T. and S.C. Strom, Amnion-derived pluripotent/multipotent stem cells. Stem Cell Reviews, 2006. 2(2): p. 133-141.
79. Baksh, D., L. Song, and R.S. Tuan, Adult mesenchymal stem cells: characterization, differentiation, and application in cell and gene therapy. Journal of Cellular and Molecular Medicine, 2004. 8(3): p. 301-316.
80. Ririe, K.M., R.P. Rasmussen, and C.T. Wittwer, Product differentiation by analysis of DNA melting curves during the polymerase chain reaction. Analytical Biochemistry, 1997. 245(2): p. 154-160.
81. Shamblott, M.J., et al., Human embryonic germ cell derivatives express a broad range of developmentally distinct markers and proliferate extensively in vitro. Proceedings of the National Academy of Sciences of the United States of America, 2001. 98(1): p. 113-118.
82. De Coppi, P., et al., Isolation of amniotic stem cell lines with potential for therapy. Nature Biotechnology, 2007. 25(1): p. 100-106.
83. Zheng, Y.B., et al., Characterization and hepatogenic differentiation of mesenchymal stem cells from human amniotic fluid and human bone marrow: A comparative study. Cell Biology International, 2008. 32(11): p. 1439-1448.
84. Zuk, P.A., et al., Human adipose tissue is a source of multipotent stem cells. Molecular Biology of the Cell, 2002. 13(12): p. 4279-4295.
85. Kubista, M., et al., Brca1 regulates in vitro differentiation of mammary epithelial cells. Oncogene, 2002. 21(31): p. 4747-4756.
86. Higuchi, A., et al., Biomimetic Cell Culture Proteins as Extracellular Matrices for Stem Cell Differentiation. Chemical Reviews, 2012. 112(8): p. 4507-4540.
87. Sobacchi, C., et al., Soluble factors on stage to direct mesenchymal stem cells fate. 2017. 5: p. 32.
88. Maes, C., et al., Soluble VEGF isoforms are essential for establishing epiphyseal vascularization and regulating chondrocyte development and survival. Journal of Clinical Investigation, 2004. 113(2): p. 188-199.
89. Zelzer, E., et al., VEGFA is necessary for chondrocyte survival during bone development. Development, 2004. 131(9): p. 2161-2171.
90. Le Blanc, S., et al., Fibroblast growth factors 1 and 2 inhibit adipogenesis of human bone marrow stromal cells in 3D collagen gels. Experimental Cell Research, 2015. 338(2): p. 136-148.
91. Simann, M., et al., Canonical FGFs Prevent Osteogenic Lineage Commitment and Differentiation of Human Bone Marrow Stromal Cells Via ERK1/2 Signaling. Journal of Cellular Biochemistry, 2017. 118(2): p. 263-275.
92. Choy, L. and R. Derynck, Transforming growth factor-beta inhibits adipocyte differentiation by Smad3 interacting with CCAAT/enhancer-binding protein (C/EBP) and repressing C/EBP transactivation function. Journal of Biological Chemistry, 2003. 278(11): p. 9609-9619.
93. van Zoelen, E.J., et al., TGFβ-induced switch from adipogenic to osteogenic differentiation of human mesenchymal stem cells: identification of drug targets for prevention of fat cell differentiation. 2016. 7(1): p. 123.
94. Majno, G. and I. Joris, Cells, tissues, and disease: principles of general pathology. 2004: Oxford University Press.
95. Cooper, G.M., R.E. Hausman, and R.E. Hausman, The cell: a molecular approach. Vol. 10. 2000: ASM press Washington, DC.
96. Gelain, F., et al., Designer Self-Assembling Peptide Nanofiber Scaffolds for Adult Mouse Neural Stem Cell 3-Dimensional Cultures. Plos One, 2006. 1(2): p. 11.
97. Ashraf, U., et al., Planting geometry and herbicides for weed control in rice: implications and challenges, in Grasses as Food and Feed. 2018, IntechOpen.
98. Saha, K., et al., Substrate Modulus Directs Neural Stem Cell Behavior. Biophysical Journal, 2008. 95(9): p. 4426-4438.
99. Engler, A.J., et al., Matrix elasticity directs stem cell lineage specification. Cell, 2006. 126(4): p. 677-689.
100. Smith, L.R., S. Cho, and D.E. Discher, Stem Cell Differentiation is Regulated by Extracellular Matrix Mechanics. Physiology, 2018. 33(1): p. 16-25.
101. E. V. Wong, P.D., Cells: Molecules and Mechanisms, in Cells: Molecules and Mechanisms, Chapter 13, ECM & Adhesion. 2009, Axolotl Academic Publishing Company, Louisville, KY. p. 198.
102. Gattazzo, F., A. Urciuolo, and P. Bonaldo, Extracellular matrix: A dynamic microenvironment for stem cell niche. Biochimica Et Biophysica Acta-General Subjects, 2014. 1840(8): p. 2506-2519.
103. He, N.N., et al., Extracellular Matrix can Recover the Downregulation of Adhesion Molecules after Cell Detachment and Enhance Endothelial Cell Engraftment. Scientific Reports, 2015. 5: p. 12.
104. Zhang, J.H., et al., Extracellular Matrix Promotes Highly Efficient Cardiac Differentiation of Human Pluripotent Stem Cells The Matrix Sandwich Method. Circulation Research, 2012. 111(9): p. 1125-1136.
105. Yao, X.P., et al., Nitric oxide releasing hydrogel enhances the therapeutic efficacy of mesenchymal stem cells for myocardial infarction. Biomaterials, 2015. 60: p. 130-140.
106. Taddei, M.L., et al., Anoikis: an emerging hallmark in health and diseases. Journal of Pathology, 2012. 226(2): p. 380-393.
107. Page-McCaw, A., A.J. Ewald, and Z. Werb, Matrix metalloproteinases and the regulation of tissue remodelling. Nature Reviews Molecular Cell Biology, 2007. 8(3): p. 221-233.
108. Lu, P., et al., Extracellular Matrix Degradation and Remodeling in Development and Disease. Cold Spring Harbor Perspectives in Biology, 2011. 3(12).
109. Theocharis, A.D., et al., Extracellular matrix structure. Advanced Drug Delivery Reviews, 2016. 97: p. 4-27.
110. Stendahl, J.C., D.B. Kaufman, and S.I. Stupp, Extracellular Matrix in Pancreatic Islets: Relevance to Scaffold Design and Transplantation. Cell Transplantation, 2009. 18(1): p. 1-12.
111. Takada, Y., X. Ye, and S. Simon, The integrins. Genome Biology, 2007. 8(5).
112. Langer, R. and J.P. Vacanti, TISSUE ENGINEERING. Science, 1993. 260(5110): p. 920-926.
113. Barbucci, R., Integrated Biomaterials Science. 2002.
114. Daley, W.P., S.B. Peters, and M. Larsen, Extracellular matrix dynamics in development and regenerative medicine. Journal of Cell Science, 2008. 121(3): p. 255-264.
115. Rozario, T. and D.W. DeSimone, The extracellular matrix in development and morphogenesis: A dynamic view. Developmental Biology, 2010. 341(1): p. 126-140.
116. Mullen, P., The Use of Matrigel to Facilitate the Establishment of Human Cancer Lines as Xenografts. Cancer Cell Culture, 2004: p. 287–292.
117. Hughes, C.S., L.M. Postovit, and G.A. Lajoie, Matrigel: A complex protein mixture required for optimal growth of cell culture. Proteomics, 2010. 10(9): p. 1886-1890.
118. Ramshaw, J.A.M., et al., Collagens as biomaterials. Journal of Materials Science-Materials in Medicine, 2009. 20: p. 3-8.
119. Muller, W.E.G., The origin of metazoan complexity: Porifera as integrated animals. Integrative and Comparative Biology, 2003. 43(1): p. 3-10.
120. Silvipriya, K., et al., Collagen: Animal Sources and Biomedical Application. Journal of Applied Pharmaceutical Science, 2015: p. 123-127.
121. Parenteau-Bareil, R., R. Gauvin, and F. Berthod, Collagen-Based Biomaterials for Tissue Engineering Applications. Materials, 2010. 3(3): p. 1863-1887.
122. Hsiao, C.T., et al., Fibronectin in cell adhesion and migration via N-glycosylation. Oncotarget, 2017. 8(41): p. 70653-70668.
123. Erickson, H.P., Stretching fibronectin. Journal of Muscle Research and Cell Motility, 2002. 23(5-6): p. 575-580.
124. Nishida, T., M. Inui, and M. Nomizu, Peptide therapies for ocular surface disturbances based on fibronectin-integrin interactions. Progress in Retinal and Eye Research, 2015. 47: p. 38-63.
125. Savage, B. and Z.M. Ruggeri, Platelet Thrombus Formation in Flowing Blood. PLATELETS, 2013: p. 399–423.
126. Boskey, A.L.R., P. G., The Regulatory Role of Matrix Proteins in Mineralization of Bone. Osteoporosis, 2013: p. 235–255.
127. Maeda, H., et al., Prospective potency of TGF-beta1 on maintenance and regeneration of periodontal tissue. Int Rev Cell Mol Biol, 2013. 304: p. 283-367.
128. Hemeda, H., B. Giebel, and W. Wagner, Evaluation of human platelet lysate versus fetal bovine serum for culture of mesenchymal stromal cells. Cytotherapy, 2014. 16(2): p. 170-180.
129. Burnouf, T., et al., Human platelet lysate: Replacing fetal bovine serum as a gold standard for human cell propagation? Biomaterials, 2016. 76: p. 371-387.
130. Bieback, K., et al., Human Alternatives to Fetal Bovine Serum for the Expansion of Mesenchymal Stromal Cells from Bone Marrow. Stem Cells, 2009. 27(9): p. 2331-2341.
131. Mannello, F. and G.A. Tonti, Concise review: no breakthroughs for human mesenchymal and embryonic stem cell culture: conditioned medium, feeder layer, or feeder-free; medium with fetal calf serum, human serum, or enriched plasma; serum-free, serum replacement nonconditioned medium, or ad hoc formula? All that glitters is not gold! Stem Cells, 2007. 25(7): p. 1603-9.
132. Shih, D.T.B. and T. Burnouf, Preparation, quality criteria, and properties of human blood platelet lysate supplements for ex vivo stem cell expansion. New Biotechnology, 2015. 32(1): p. 199-211.
133. Stute, N., et al., Autologous serum for isolation and expansion of human mesenchymal stem cells for clinical use. Experimental Hematology, 2004. 32(12): p. 1212-1225.
134. Shahdadfar, A., et al., In vitro expansion of human mesenchymal stem cells: Choice of serum is a determinant of cell proliferation, differentiation, gene expression, and transcriptome stability. Stem Cells, 2005. 23(9): p. 1357-1366.
135. Horn, P., et al., Impact of individual platelet lysates on isolation and growth of human mesenchymal stromal cells. Cytotherapy, 2010. 12(7): p. 888-898.
136. Kocaoemer, A., et al., Human AB serum and thrombin-activated platelet-rich plasma are suitable alternatives to fetal calf serum for the expansion of mesenchymal stem cells from adipose tissue. Stem Cells, 2007. 25(5): p. 1270-8.
137. Eriksson, L., et al., PLATELET CONCENTRATES IN AN ADDITIVE SOLUTION PREPARED FROM POOLED BUFFY COATS - INVIVO STUDIES. Vox Sanguinis, 1993. 64(3): p. 133-138.
138. Azuma, H., et al., Platelet additive solution - Electrolytes. Transfusion and Apheresis Science, 2011. 44(3): p. 277-281.
139. Doucet, C., et al., Platelet lysates promote mesenchymal stem cell expansion: A safety substitute for animal serum in cell-based therapy applications. Journal of Cellular Physiology, 2005. 205(2): p. 228-236.
140. Schallmoser, K., et al., Human platelet lysate can replace fetal bovine serum for clinical-scale expansion of functional mesenchymal stromal cells. Transfusion, 2007. 47(8): p. 1436-1446.
141. Kilian, O., et al., Effects of platelet growth factors on human mesenchymal stem cells and human endothelial cells in vitro. European Journal of Medical Research, 2004. 9(7): p. 337-344.
142. Van Pham, P., et al., Activated platelet-rich plasma improves adipose-derived stem cell transplantation efficiency in injured articular cartilage. Stem Cell Research & Therapy, 2013. 4: p. 11.
143. Cervelli, V., et al., Platelet-Rich Plasma Greatly Potentiates Insulin-Induced Adipogenic Differentiation of Human Adipose-Derived Stem Cells Through a Serine/Threonine Kinase Akt-Dependent Mechanism and Promotes Clinical Fat Graft Maintenance. Stem Cells Translational Medicine, 2012. 1(3): p. 206-220.
144. Hara, Y., M. Steiner, and M.G. Baldini, Platelets as a source of growth-promoting factor(s) for tumor cells. Cancer Res, 1980. 40(4): p. 1212-6.
145. Umeno, Y., A. Okuda, and G. Kimura, Proliferative behaviour of fibroblasts in plasma-rich culture medium. J Cell Sci, 1989. 94 ( Pt 3): p. 567-75.
146. King, G.L. and S. Buchwald, Characterization and partial purification of an endothelial cell growth factor from human platelets. J Clin Invest, 1984. 73(2): p. 392-6.
147. Bernardi, M., et al., Production of human platelet lysate by use of ultrasound for ex vivo expansion of human bone marrow-derived mesenchymal stromal cells. Cytotherapy, 2013. 15(8): p. 920-929.
148. Luttenberger, T., et al., Platelet-derived growth factors stimulate proliferation and extracellular matrix synthesis of pancreatic stellate cells: Implications in pathogenesis of pancreas fibrosis. Laboratory Investigation, 2000. 80(1): p. 47-55.
149. Schallmoser, K., et al., Rapid large-scale expansion of functional mesenchymal stem cells from unmanipulated bone marrow without animal serum. Tissue Engineering Part C-Methods, 2008. 14(3): p. 185-196.
150. Hemeda, H., et al., Heparin concentration is critical for cell culture with human platelet lysate. Cytotherapy, 2013. 15(9): p. 1174-1181.
151. Burnouf, T., et al., A chromatographically purified human TGF-beta1 fraction from virally inactivated platelet lysates. Vox Sang, 2011. 101(3): p. 215-20.
152. Burnouf, T., et al., A virally inactivated platelet-derived growth factor/vascular endothelial growth factor concentrate fractionated from human platelets. Transfusion, 2010. 50(8): p. 1702-1711.
153. Mekaj, Y.H., The roles of platelets in inflammation, immunity, wound healing and malignancy. International Journal of Clinical and Experimental Medicine, 2016. 9(3): p. 5347-5358.
154. Shimizu, K., H. Fujita, and E. Nagamori, Oxygen Plasma-Treated Thermoresponsive Polymer Surfaces for Cell Sheet Engineering. Biotechnology and Bioengineering, 2010. 106(2): p. 303-310.
155. Twaites, B.R., et al., Thermoresponsive polymers as gene delivery vectors: Cell viability, DNA transport and transfection studies. Journal of Controlled Release, 2005. 108(2-3): p. 472-483.
156. Doorty, K.B., et al., Poly(N-isopropylacrylamide) co-polymer films as potential vehicles for delivery of an antimitotic agent to vascular smooth muscle cells. Cardiovascular Pathology, 2003. 12(2): p. 105-110.
157. Stile, R.A. and K.E. Healy, Thermo-responsive peptide-modified hydrogels for tissue regeneration. Biomacromolecules, 2001. 2(1): p. 185-194.
158. Hacker, M.C., et al., Synthesis and Characterization of Injectable, Thermally and Chemically Gelable, Amphiphilic Poly(N-isopropylacrylamide)-Based Macromers. Biomacromolecules, 2008. 9(6): p. 1558-1570.
159. Teotia, A.K., Sami, H., & Kumar, A., Switchable and Responsive Surfaces and Materials for Biomedical Applications. 2015. 3–43.
160. Haq, M.A., Y.L. Su, and D.J. Wang, Mechanical properties of PNIPAM based hydrogels: A review. Materials Science & Engineering C-Materials for Biological Applications, 2017. 70: p. 842-855.
161. Mano, J.F., Stimuli-responsive polymeric systems for biomedical applications. Advanced Engineering Materials, 2008. 10(6): p. 515-527.
162. Janmey, P.A., et al., The hard life of soft cells. 2009. 66(8): p. 597-605.
163. Discher, D.E., P. Janmey, and Y.L. Wang, Tissue cells feel and respond to the stiffness of their substrate. Science, 2005. 310(5751): p. 1139-1143.
164. Xue, W., et al., Synthesis and characterization of hydrophobically modified polyacrylamides and some observations on rheological properties. European Polymer Journal, 2004. 40(1): p. 47-56.
165. Wanwipa Siriwatwechakul, N.T., Vatcharani Ngaotheppitak, and Sureeporn, Thermo-Sensitive Hydrogel: Control of Hydrophilic-Hydrophobic Transition. International Journal of Chemical & Biomolecular Engineering, 2008: p. 165-170.
166. Halperin, A. and M. Kroger, Thermoresponsive Cell Culture Substrates Based on PNIPAM Brushes Functionalized with Adhesion Peptides: Theoretical Considerations of Mechanism and Design. Langmuir, 2012. 28(48): p. 16623-16637.
167. Schwartz, S.D., et al., Embryonic stem cell trials for macular degeneration: a preliminary report. Lancet, 2012. 379(9817): p. 713-20.
168. Osakada, F., et al., In vitro differentiation of retinal cells from human pluripotent stem cells by small-molecule induction. J Cell Sci, 2009. 122(Pt 17): p. 3169-79.
169. Zaky, S.H., et al., Platelet lysate favours in vitro expansion of human bone marrow stromal cells for bone and cartilage engineering. J Tissue Eng Regen Med, 2008. 2(8): p. 472-81.
170. Zhang, L., et al., ROCK inhibitor Y-27632 suppresses dissociation-induced apoptosis of murine prostate stem/progenitor cells and increases their cloning efficiency. PloS one, 2011. 6(3): p. e18271-e18271.
171. Emre, N., et al., The ROCK Inhibitor Y-27632 Improves Recovery of Human Embryonic Stem Cells after Fluorescence-Activated Cell Sorting with Multiple Cell Surface Markers. Plos One, 2010. 5(8): p. 10.
172. Zhu, J., et al., Inhibition of RhoA/Rho-kinase pathway suppresses the expression of extracellular matrix induced by CTGF or TGF-β in ARPE-19. International journal of ophthalmology, 2013. 6(1): p. 8-14.
173. Leach, L.L., et al., Induced Pluripotent Stem Cell-Derived Retinal Pigmented Epithelium: A Comparative Study Between Cell Lines and Differentiation Methods. J Ocul Pharmacol Ther, 2016. 32(5): p. 317-30.

簡易檢索 / 詳目顯示

相關論文