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Are bFGF and/or EGF necessary in NSC expansion medium?

Are bFGF and/or EGF necessary in NSC expansion medium?



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My understanding is that it isn't necessary since it appears that the idea behind a feeder layer is that the stem cells produce their own bFGF & EGF along with other growth factors. Yet I've still seen some papers (e.g. this one which I am trying to reproduce), use it in mediums used for NSC maintenance (their neural progenitor culture contains both). This paper seems to suggest that a feeder layer for NSCs is unnecessary which would make sense but I wanted to know what is common practice and the most robust method and how it is related to these findings. Because of that I would like to hear from personal experience but references are also obviously useful.


A novel role for Gab2 in bFGF-mediated cell survival during retinoic acid–induced neuronal differentiation

Gab proteins amplify and integrate signals stimulated by many growth factors. In culture and animals, retinoic acid (RA) induces neuronal differentiation. We show that Gab2 expression is detected in neurons in three models of neuronal differentiation: embryonic carcinoma (EC) stem cells, embryonic stem cells, and primary neural stem cells (NSCs). RA treatment induces apoptosis, countered by basic FGF (bFGF). In EC cells, Gab2 silencing results in hypersensitivity to RA-induced apoptosis and abrogates the protection by bFGF. Gab2 suppression reduces bFGF-dependent activation of AKT but not ERK, and constitutively active AKT, but not constitutively active MEK1, reverses the hypersensitization. Thus, Gab2-mediated AKT activation is required for bFGF's protection. Moreover, Gab2 silencing impairs the differentiation of EC cells to neurons. Similarly, in NSCs, Gab2 suppression reduces bFGF-dependent proliferation as well as neuronal survival and production upon differentiation. Our findings provide the first evidence that Gab2 is an important player in neural differentiation, partly by acting downstream of bFGF to mediate survival through phosphoinositide 3 kinase𠄺KT.


Introduction

Neural stem cells (NSCs) hold the capacity of self-renewal and can differentiate into neurons, astrocytes and oligodendrocytes. They play a major role in the development of the embryonic central nervous system and continue to function throughout adulthood 1, 2, 3 . The proliferation and differentiation of NSCs depend on microenvironmental niche signals 4, 5 , including a number of growth factors such as vascular endothelial growth factor (VEGF) and basic fibroblast growth factor (bFGF) 6 .

VEGF was originally identified as a major mediator of angiogenesis 7, 8 . It plays an important role in mediating vascular permeability and tissue regeneration 9, 10 . In most occasions, VEGF exerts its action via its receptor, Flk-1, in endothelial cells, hematopoietic stem/progenitor cells and some tumor cells 11, 12, 13, 14 . In the central nervous system (CNS), for instance the hippocampus, VEGF stimulates the expansion of NSCs and neurogenesis in various animal models, resulting in improved learning ability 15, 16, 17, 18 . In adults, NSCs are found in close proximity to blood vessels and surrounded by glial cells in the hippocampus and the subventricular zone. Previous studies by others suggested that both vascular cells and glial cells may serve as a niche for NSCs 19, 20 . Since both cell types express VEGF and bFGF 21 , we hypothesize that these two growth factors may serve as niche signals for NSCs.

There are a number of observations suggesting that bFGF may also modulate neurogenesis, both in vivo and in vitro. First, bFGF and its main receptor, FGFR-1, are both present in the mouse CNS during corticogenesis 22 . Second, bFGF is secreted from cells within the brain through an energy-dependent exocytosis process 23 . Third, during the early phase of development, neural progenitor cells proliferate in response to bFGF during the neurogenic phase 24, 25 . It has been reported that knocking out bFGF during critical periods of brain development led to an overall reduction in progenitor proliferation and subsequent neuronal differentiation 26, 27 .

To assess the roles of VEGF and bFGF in regulation of NSCs, we used a nearly homogeneous population of NSCs derived from mouse embryonic stem (ES) cells to test the hypothesis that bFGF and VEGF coordinately regulate NSC proliferation.


Methods

Cell culture

Islet-depleted pancreatic cell aggregates from human donors (10 in total age 39넙 years body weight 72넓 kg) were kindly provided by Dr. Garth Warnock and Dr. Ziliang Ao at the Ike Barber Human Islet Transplant Laboratory (Vancouver, BC, Canada). Pancreata were obtained with the written informed consent of family members under the approval of the University of British Columbia Clinical Research Ethics Board. Cell clusters received on day 0 were washed twice in CMRL medium with 10% fetal bovine serum, 100 units/mL penicillin and 100 µg/mL streptomycin (all from Invitrogen, Carlsbad, CA), referred to as CMRL⬐% FBS medium. Clusters were then dispersed or seeded overnight in CMRL⬐% FBS medium at 1 µL packed cell volume (PCV)/cm 2 (PCV tubes, Techno Plastic Product, Trasadingen, Switzerland) in tissue culture-treated flasks (Sarstedt, Nümbrecht, Germany) before magnetic-activated cell sorting (MACS) on day 1. Unsorted or sorted cells were seeded at 1.25휐 5 cells/cm 2 in 0.32 mL/cm 2 CMRL⬐% FBS medium in plates containing 0.32 mL/cm 2 pre-incubated medium and cultured at 37ଌ, 5% CO2 and 90% humidity. Live cells were enumerated by trypan blue exclusion using a Cedex cell counter (Roche Innovatis, Bielefeld, Germany). The unsorted and MACS-sorted cultures were maintained in CMRL⬐% FBS medium for 8 days, with medium exchanges on days 2, 5 and 7. For serum-free assay development and screening, the cultures were shortened to 6 days with medium exchanges on days 2 and 5. The basal CMRL-0.1X ITS serum-free medium consisted of CMRL with 0.5 mg/L insulinʰ.5 mg/L transferrinʰ.5 µg/L selenite (i.e. 0.1 times I-1884 from Sigma), 10 mM nicotinamide and 0.2% BSA (STEMCELL, BC, Canada). The test solutions contained 0.1%, 1% or 10% FBS (Invitrogen), or 50% pancreatic fibroblast-conditioned medium diluted in CMRL-0.1X ITS. The following recombinant human growth factors were also tested at 20 ng/mL unless otherwise mentioned: basic fibroblast growth factor (bFGF, STEMCELL), epidermal growth factor (EGF, STEMCELL), hepatocyte growth factor (HGF, Sigma), keratinocyte growth factor (KGF, Sigma), vascular endothelial growth factor (VEGF, Sigma). One h after the last medium exchange, 10 µM of 5-Bromo-2′-deoxy-uridine was added (BrdU, Labelling and Detection Kit II, Roche, Basel, Switzerland) and incubated for 20 h prior to fixing, staining and Cellomics analysis.

Conditioned medium

Passaged (P3 to P8) CD90-enriched pancreatic cells cultured in CMRL⬐% FBS medium until reaching 90% confluency (𢏆휐 4 cells/cm 2 ) were treated with 10 µg/mL mitomycin c (Sigma) for 1 h, washed 3 times and incubated for 24 h in CMRL-0.1X ITS, then filter-sterilized and stored at �ଌ.

Cell dispersion

Cell clusters were washed twice with dispersion medium (1 mM EDTA from Invitrogen, 10 mM HEPES from Sigma and 0.5% bovine serum albumin from STEMCELL prepared in Ca 2+ and Mg 2+ -free HBSS from Invitrogen), then re-suspended at 0.05 mL PCV/mL in dispersion medium and kept at 37ଌ for 7 min with 75 rpm agitation. Clusters were digested for 10 min at 37ଌ by adding 25 µg/mL trypsin and 4 µg/mL DNase (Sigma). After adding CMRL⬐% FBS medium, the cells were triturated and filtered through a 40 µm nylon sieve (BD Biosciences). The total cell yield based on PCV was 43ଗ%, providing 168넹 million live dispersed cells/mL PCV initial tissue clusters.

Magnetic-activated cell sorting

Adherent cells from undispersed clusters were washed with MACS buffer consisting of Ca 2+ and Mg 2+ -free phosphate buffered saline (PBS) with 1% FBS, 2 mM EDTA, 100 units/mL penicillin and 100 µg/mL streptomycin. Cells were trypsinized, triturated after adding CMRL⬐% FBS medium and then filtered through a 40 µm cell strainer (BD Biosciences), yielding 0.17ଐ.05휐 6 cells/cm 2 on day 1. The cells were then washed twice with MACS buffer and re-suspended at 𢙂휐 7 cells/mL in � µL MACS buffer. An equal volume of primary antibody was added to obtain 1� mouse anti-Ca19-9 (#NCL-L-CA19-9, Leica Microsystems, Germany) or 1� mouse anti-CD90 (#555593, BD Pharmingen, CA). After incubating for 25 min on ice at 75 rpm agitation, the cells were washed twice with MACS buffer. Magnetic labeling was performed using microbead-labelled goat or rat anti-mouse IgG1 antibody (#130-048-402 or #130-047-102, Miltenyi Biotec, Germany). Cells were re-suspended at 4휐 6 cells/mL and separated using the “possel” (for Ca19-9) or �pletes” (for CD90) program of an Automacs® (Miltenyi) cell separator. The total live cell yield from MACS was 60ଓ%.

Flow cytometry

Adherent cells were washed with PBS, trypsinized, triturated after adding FACS buffer (PBS⬐% FBS) and filtered through a 40 µm strainer. Cells kept on ice were then washed twice with FACS buffer, distributed to obtain 2.5휐 5 cells/sample, centrifuged and re-suspended in 50 µL. Then, 50 µL of primary antibody diluted in FACS buffer (same concentrations as for MACS) were added, followed by 25 min incubation at 75 rpm. The cells were washed twice, re-suspended in 50 µL and 50 µL of Alexa 647 labelled goat anti-mouse IgG (A21450, Invitrogen) were added to obtain a 1� dilution. After 15 min incubation in the dark at 75 rpm, 10 µL of propidium iodide at 100 µg/mL (Sigma, in PBS) was added. The cells were washed twice, re-suspended in 500 µL FACS buffer and analyzed on a BD FACSCalibur flow cytometer. Data analysis including compensation was performed with Flowjo 7.2.5 software (Tree Star, Ashland, OR).

Immunocytochemistry and Cellomics

Cell fixing and staining with BrdU was performed according to the BrdU kit instructions (Roche), except for diluting primary and secondary antibodies 1�. For samples that did not require BrdU labeling, the cells were fixed with Bouin's fixative for 15 min and stored in 70% ethanol. CK19 or Ki67 antigen retrieval was performed by microwaving 6 times for 5 s at 1000 W in 10 mM citrate (Sigma) at pH 6.0 and cells permeabilized by a 10 min incubation in 0.25% Triton X 100 (Sigma) dissolved in PBS. Unless otherwise mentioned, all subsequent incubations were at room temperature and 100 rpm agitation. All samples were then washed with PBS, incubated in blocking solution (Dakocyotmation, Glostrup, Denmark) for 15 min and then stained overnight at 4ଌ with primary antibodies in Antibody Diluent (Dako). Primary antibodies were 1� guinea pig anti-human insulin (Dako A0564), 1� rabbit anti-human amylase (Sigma A8273), 1� mouse anti-human CK19 (Dako M0888), 1� mouse anti-human vimentin (Dako M0725), 1� rabbit anti-human Ki67 (Santa Cruz Biotech sc-15402, Santa Cruz, CA) and/or 1� mouse anti-human Ki67 (BD Biosciences 556003). The next day, cells were washed with PBS and stained 1 h in the dark with secondary antibodies (Alexa 488 goat anti-guinea pig IgG, Alexa 488 or 568 goat anti-rabbit IgG and/or Alexa 568 goat anti-mouse IgG, all from Invitrogen) at 1� in Antibody Diluent. Samples were then washed, stained with 1 µg/mL DAPI for 15 min, and washed again. The plates were imaged on a Cellomics ArrayScan VTI. Slides were mounted with Vectashield medium (Vector Labs, Burlingame, CA) and imaged on a Zeiss Axioplan 2 microscope (Carl Zeiss, Oberkochen, Germany), using ImageJ analysis software (NIH).

Design of experiments and statistical analysis

Results represent the average values obtained from 3 to 5 pancreata ± standard error of the mean. Two-way comparisons were based on Student's t-tests with p-valuesπ.05 considered significant, with a paired t-test used in the case of the comparison between the CK19+ cell number and BrdU incorporation read-outs. For comparison between a basal response and responses normalized to the basal response, confidence intervals with α levels of 0.05 were used. The main and interaction effects of bFGF, EGF, HGF, KGF and VEGF were quantified by a two-level (low level 0 ng/mL high level 20 ng/mL) full factorial design with 8 centre points (10 ng/mL of all five factors). For each pancreas, these 40 conditions were repeated on three 96-well plates with different randomization on each plate. The results were analyzed with JMP 7.0 or 8.0 statistics software (SAS, Cary, NC). The growth factor concentrations Ci were transformed into scaled variables . The model was as follows: up to the fifth-order interaction effect , where βi values represent the fitted model parameters. The subscripts F, E, H, K, V represent the “i” factors bFGF, EGF, HGF, KGF and VEGF. The model was then reduced to exclude factors with p-valuesϠ.1.


Effect of EGF and FGF on the expansion properties of human umbilical cord mesenchymal cells

Mesenchymal stem cells have been increasingly introduced to have great potential in regenerative medicine, immunotherapy, and gene therapy due to their unique properties of self-renewal and differentiation into multiple cell lineages. Studies have shown that these properties may be limited and changed by senescence-associated growth arrest under different culture conditions. This study aimed to present the ability of some growth factors on human umbilical cord mesenchymal (hUCM) cells expansion and telomerase activity. To optimize hUCM cell growth, epidermal growth factor (EGF) and fibroblast growth factor (FGF) were utilized in culture media, and the ability of these growth factors on the expression of the telomerase reverse transcriptase (TERT) gene and cell cycle phases was investigated. TERT mRNA expression increased in the hUCM cells treated by EGF and FGF. So, the untreated hUCM cells expressed 30.49 ± 7.15% of TERT, while EGF-treated cells expressed 51.82 ± 12.96% and FGF-treated cells expressed 33.77 ± 11.55% of TERT. Exposure of hUCM cells to EGF or FGF also promoted the progression of cells from G1 to S phase of the cell cycle and induced them to decrease the number of cells entering the G2/M phase. Our study showed that EGF and, to a lesser extent, FGF amplify the proliferation and expansion of hUCM cells.

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Epidermal Growth Factor (EGF) Treatment on Multipotential Stromal Cells (MSCs). Possible Enhancement of Therapeutic Potential of MSC

Adult bone marrow multipotential stromal cells (MSCs) hold great promise in regenerative medicine and tissue engineering. However, due to their low numbers upon harvesting, MSCs need to be expanded in vitro without biasing future differentiation for optimal utility. In this concept paper, we focus on the potential use of epidermal growth factor (EGF), prototypal growth factor for enhancing the harvesting and/or differentiation of MSCs. Soluble EGF was shown to augment MSC proliferation while preserving early progenitors within MSC population, and thus did not induce differentiation. However, tethered form of EGF was shown to promote osteogenic differentiation. Soluble EGF was also shown to increase paracrine secretions including VEGF and HGF from MSC. Thus, soluble EGF can be used not only to expand MSC in vitro, but also to enhance paracrine secretion through drug-releasing MSC-encapsulated scaffolds in vivo. Tethered EGF can also be utilized to direct MSC towards osteogenic lineage both in vitro and in vivo.

1. Multipotential Stromal Cells/Mesenchymal Stem Cells (MSCs)

1.1. MSC Overviews

Adult bone marrow multipotential stromal cells / mesenchymal stem cells (MSCs) are multipotent cells with strong paracrine activities of various growth factors [1–7]. These cells were originally isolated as colony forming adherent fibroblast-like cells or colony forming unit fibroblastic cells (CFU-Fs) from bone marrow suspension [8], but it was subsequently realized that these cells carry multipotency capable of differentiating into multiple cell lineages including osteoblasts, chondrocytes, adipocytes, smooth muscle cells, skeletal and cardiac myocytes, endothelial cells, and neurons [3–6, 9, 10].

Initially MSC differentiation and direct incorporation into local tissues undergoing wound healing and tissue regenerations were regarded as a primary mechanism of MSC action however, the contribution of MSC differentiation and direct incorporation into regenerating tissues remains debated [11]. For example, some groups showed that MSCs were differentiated and incorporated as myocardiocytes or vascular cells (endothelial cell and vascular smooth muscle cells) in newly formed vessels in MSC-based cardioplasty models in rat (isogenic and allogenic MSC transplantation) and pig (allogenic transplantation) [11–14]. Human MSCs from adult bone marrow were engrafted and differentiated into cardiomyocytes within myocardium of SCID mice [15]. In contrast, another group showed that bone marrow-derived cells were not incorporated into newly formed blood vessels in hindlimb ischemia in mice allogenic bone marrow transplantation model [16]. Direct incorporation and differentiation of transplanted MSC into keratinocytes and vascular cells were also shown in mice dermal wound healing model (allogenic MSC transplantation) [17–19], whereas others showed that MSC differentiation of transplanted MSCs into keratinocytes and vascular cells was not observed in mice dermal wound healing model (allogenic MSC transplantation) and mice limb ischemia models (isogenic MSC transplantation) [20, 21]. Furthermore, even when there is an early incorporation noted into regenerating tissue, these cells are largely gone by one month[15]. The efficacy of engraftment of transplanted MSC was varied, suggesting the presence of other mechanisms of MSC-mediated promotion of tissue regeneration [7, 11].

One such mechanism is paracrine secretion of growth factors and cytokines. MSCs are known to have a strong paracrine capability of various growth factors and cytokines such as vascular endothelial growth factor (VEGF) or hepatocyte growth factor (HGF), which promote angiogenesis and wound healing [7, 22–24]. Indeed, conditioned medium of MSCs was also shown to promote angiogenesis or wound healing in animal models, suggesting the crucial role of MSC’s paracrine action in promotion of angiogenesis and wound healing [21, 23, 25].

1.2. In Vitro Expansion of MSCs

A major thrust is to use MSCs pharmacologically. Even if the physiological involvement of MSCs is debatable, studies have shown injected MSCs to home to wounded tissues [1, 2, 4, 5]. However, the availability of sufficient number of MSCs that retain their multipotency and paracrine activity is prerequisite for successful MSC-based therapeutics and tissue engineering. MSCs are present only in low frequency in the bone marrow (one in

bone marrow mononuclear cells, lower frequency in aged hosts) [5, 26], thus, MSCs harvested from the bone marrow for pharmacological uses need to be expanded in vitro. These cells are expandable in vitro [3, 27] and that is one of the desirable characteristics about MSCs.

Current in vitro expansion strategies generally rely on the use of fetal bovine serum, but this practice not only carries inherent disease risks [28] but also hampers standardization that is critical to establishing a broad clinical adoption. Another issue about current MSC expansion is the loss of differentiation, proliferative, and therapeutic potentials of MSCs through in vitro expansion process [29, 30]. Thus, there is a strong motivation to identify factors that might be used in serum-free formulations to expand MSC in vitro without losing differentiation capacity and to preserve self-renewal and therapeutic potentials of undifferentiated MSCs [31].

A recent report found that a combination of transforming growth factor-β (TGF-β), platelet-derived growth factor (PDGF), and basic fibroblast growth factors (bFGF) could replace serum component in cell culture medium to expand human MSCs ex vivo without compromising differentiation potentials, at least up to 5 passages [32]. Subsequently, serum and animal component-free MSC culture media (STEMPRO MSC SFM, from Invitrogen, Carlsbad, CA) (MesenCult-ACF Culture Kit, from STEMCELL Technologies, Vancouver, Canada) became available on the market. The manufacturers claim that the culture media exert superior MSC proliferation potentials while maintaining differentiation potentials and colony formation potentials, as supported by at least one study [33]. Those chemically-defined media should be safer and thus better for clinical settings, although the proprietary composition of these culture media may hinder acceptance in preclinical and clinical usage.

1.3. Heterogenous Populations within MSC Preparations

Although MSCs possess vast proliferative potential, have the capacity for self-renewal, and give rise to differentiated progenies, all MSC populations analyzed by clonal assays were shown to be heterogeneous, with individual cells capable of various differentiation potential and expansion capacity [34]. Thus, the International Society for Cellular Therapy proposed MSCs to be named as multipotential mesenchymal stromal cells, which can also be abbreviated as MSCs [35]. MSC populations in vitro are known to include early progenitors or rapidly self-renewing (RS)-cells as well as large slow replicating mature/senescent cells. It is early progenitors or RS-cells which retain strong multipotentiality for differentiation. In contrast, mature/senescent cells have only limited differentiation potentials, and these cells predominate in multiple passaged MSCs [27, 29, 36].

One of the prominent characteristics about MSCs is their ability to produce colonies after being seeded at low density [8]. Generation of a single-cell derived colony relies on the presence of early progenitors or RS cells in MSC preparations. In other words, assessment of the colony formation unit (CFU) can be used to gauge the proportion of colony forming early progenitors in MSC population [29, 37, 38]

The number of MSC within bone marrow mononuclear cells was shown to decrease with age [26]. Moreover, both MSCs from old donors and high passaged MSCs were shown to have decreased paracrine activity and reduced organ protective effects upon transplantation [30, 39]. These reports clearly suggest the importance of preserving early progenitors or RS cells in MSC preparation to maintain therapeutic potentials of MSCs.

1.4. MSC Expansion and Differentiation Potentials

To preserve early progenitors within MSC populations, self-renewal of these cells has to be maintained or even enhanced through in vitro MSC expansion process. Otherwise, early progenitors will be lost during in vitro expansion. Cell division is a central step of self-renewal and expansion of these cells. There are various growth factors and cytokines known to work as a mitogen, but the ideal growth factors for in vitro MSC expansion must reversibly suppress or at least not alter the subsequent differentiation process. In other words, these factors should not diminish differentiation potentials of MSCs. Growth factors or cytokines which promote differentiation of MSCs into certain lineages cannot be used for in vitro MSC expansion, as the differentiation process itself is antagonizing self-renewal of undifferentiated MSCs including early progenitors and the differentiation would compromise utilization of these cells.

Among those growth factors, we have focused on epidermal growth factor (EGF) as a candidate to utilize in vitro expansion of MSCs as EGF stimulates MSC proliferation without altering differentiation process and potentials [3].

2. EGF to Enhance Self-Renewal and Expansion of MSCs In Vitro

2.1. EGF and EGF Receptor

EGF was originally isolated from mouse salivary gland extract as a factor accelerating the corneal wound healing [40], but it was soon recognized that it is indeed a general growth factor exerting various actions including cell migration and proliferation on a wide variety of cells [41–43].

The EGF receptor (EGFR/ErbB-1 or human epidermal growth factor receptor 1(HER1)) is the prototypal growth factor receptor with intrinsic tyrosine kinase activity. It is widely expressed on many cell types, including epithelial and mesenchymal lineages [42]. Upon binding of at least five genetically distinct ligands (including EGF, transforming growth factor-

(TGF- ), and heparin-binding EGF (HB-EGF)), the intrinsic tyrosine kinase within EGFR/ErbB-1 is activated and phosphorylates the receptor itself (autophosphorylation) and numerous target downstream molecules. Intracellular signaling pathways downstream of EGFR/ErbB-1 include phosholipase C

(PLC ) and its downstream calcium- and protein kinase C (PKC)-mediated cascades, ras activation leading to various mitogen activated protein kinases (MAPK), other small GTPases such as rho and rac, multiple signal transducer and activator of transcription (STAT) isoforms, and heterotrimeric G proteins, phosphatidylinositol

-OH kinase (PI3K) and phospholipase D (PLD) [3, 42, 43].

Upon ligand binding and activation, EGFR/ErbB-1 undergoes internalization from the cell surface via the clathrin-coated endocytic system. Within the acidic late endosomal compartment, both EGF and EGF-bound EGFR/ErbB-1 undergo degradation as EGF is a nondissociative ligand for EGF, whereas TGF- -bound EGFR/ErbB-1 is recycled back to cell surface after dissociation of TGF- from EGFR /ErbB-1 [42]. There is a growing evidence suggesting that preferential and prolonged activation of EGFR/ErbB-1 from cell surface exerts a distinct activity from internalized EGFR/ErbB-1, as surface-tethered EGF promotes cell spreading and survival of MSCs, whereas soluble EGF does not (See discussion below) [44].

2.2. EGF Enhances MSC Proliferation

EGF is a prototypal mitogen for various types of cells. Human MSCs express EGFR/ErbB-1, and we and others demonstrated the mitogenic effect of EGF and HB-EGF on MSCs [3, 45]. Cell proliferation is an integral part of self-renewal and expansion of the cells, and thus these data supports our hypothesis that EGF can be used for in vitro MSC expansion, at least in short-term culture setting however, additive effects of EGF treatment on human MSC proliferation become less clear in the long-term culture (Figure 1), presumably due to down-regulation of EGFR/ErbB1, as discussed below.


(a)
(b)
(a)
(b) Effect of EGF on primary human MSC proliferation (a) and accumulative population doubling (PD)(b). (a) The cell number of primary human MSCs increases about 5.5-fold in the diluent (Ctrl) in culture medium supplemented with 17% FBS in 120 hours (5 days) time period. The addition of EGF (10 nM) gives an extra increase of cell counts to 8.5-fold. Total of 10000 cells was seeded per each well in 12-well plate and the cell count of each well was measured by Coulter Cell Counter Z2 (Beckman Coulter, Inc. Fullerton, CA). Shown are mean

s.e.m. of three experiments each performed in triplicate. The differences in proliferation were compared between growth factor and diluent (Ctrl) exposed (*

). (b) Accumulated PD of primary human MSCs in day 5 and 19 in the culture condition same as (a). After cell counting at day 5, equal number of cells (Total of 1000) was seeded per each well in 6-well plate and the cell count of each well was measured at day 19 by Coulter Cell Counter Z2 (Beckman Coulter, Inc. Fullerton, CA). Note that the PD is higher in EGF treated group (2.54 in Ctrl, 3.07 in EGF) for the initial 5 days (*

The second aspect of expansion is to maintain colony-forming units. In this aspect, EGF treatment is also successful. EGF leads to a statistically significant 25% increase in stainable colonies (Figure 2), suggesting that EGF treatment helps preserve early progenitors within human MSC populations.


(a)
(b)
(a)
(b) Effects of EGF treatment (10 nM) on primary human MSC colony formation. Five hundred cells were seeded in 10 cm dish within culture medium supplemented with 17% FBS and the number of formed colonies (Diameter

1.5 mm) was counted manually in day 14. (a) Representative image of MSC colonies stained with crystal violet. (b) Colony count of MSC. The number of colony is given per 1000 cells seeded initially (*

2.3. EGF and MSC Differentiation Potentials

Multidifferentiation potential is a key characteristic of MSCs, which attracts so much attention in the field of tissue engineering and regenerative medicine [4, 5]. Differentiation potential itself has to be preserved through the in vitro expansion of MSCs however, ongoing differentiation process itself should be suppressed or not induced at least as it antagonizes self-renewal and expansion of undifferentiated MSCs and limits further use of these cells.

Kratchmarova and her colleagues showed that EGF stimulation enhances osteogenic differentiation of human MSCs in the presence of chemical cues, whereas PDGF does not [46]. Through mass spectrometry-based proteomics approach, they identified PI3K as a molecular switch to turn off pro-osteogenic signal from PDGFR.

This report is contradictory to reports by our group and others. Our data show that EGF alone does not induce differentiation in the absence of chemical or other cues, and does not alter human MSC differentiation processes into osteogenic, adipogenic, and chondrogenic lineages by chemical cues in vitro [3]. This discrepant finding might be attributable to a different intracellular signaling, as PI3K-protein kinase B/akt pathway is activated in the downstream of EGFR/ErbB-1 in our report [3], whereas this pathway is not activated in their report [46]. The apparent reason for this discrepancy is unclear, but one prominent difference is EGF concentration 10 nM EGF was used in our report [3], whereas Kratchmarova and her colleagues used 83 nM of EGF in their report [46]. This speculation is supported by a recent report showing that 80 pM EGF inhibits osteogenic differentiation of human MSCs [47]. Krampera and his colleague also showed that HB-EGF (2.3 nM) inhibits osteogenic differentiation of MSC induced by chemical cues [45]. Human MSCs do not express ErbB-4, another receptor for HB-EGF both EGF and HB-EGF bind only EGFR/ErbB-1 on MSC, and downstream signaling cascade is similar [3, 45].

Do EGFR/ErbB1 agonists exert both positive and negative effects on osteogenic differentiation of MSC in a concentration-dependent manner? And if so, what is the underlying mechanism? This question is still unresolved, but we and our collaborators are utilizing immobilized tethered EGF surface in a manner that provides some important hints [44, 47]. Tethered EGF blocks EGFR/ErbB1 endocytic internalization and enhances osteogenic differentiation through providing sustained activation of downstream signaling through EGFR/ErbB1, whereas 80 pM of soluble EGF interferes with osteogenic differentiation through inducing receptor internalization and subsequent degradation [47], in agreement with Krampera’s data [45]. Thus, we hypothesize that weak and temporal signaling from low concentration of soluble EGF exerts anti-osteogenic effects, whereas strong sustained signaling from tethered EGF exerts pro-osteogenic signaling on MSC (Figure 3). Previous reports have shown that the activation of ERK/MAPK pathway, one of the major signaling pathways in the downstream of EGFR/ErbB-1 [3, 42], promotes MSC osteogenic differentiation [48–50]. In agreement with these reports, strong and sustained activation of ERK/MAPK pathway has been observed in MSCs cultured on tethered EGF surface [44, 47], and thus, ERK/MAPK pathway might be one of the main pro-osteogenic signaling pathways in the downstream of EGFR/ErbB-1.


Simplified model for the effects of EGFR/ErbB1 signaling on MSC osteogenic differentiation. Weak and temporal stimulation of EGFR/ErbB1 exerts anti-osteogenic effects, whereas strong and sustained stimulation of EGFR/ErbB1 exerts pro-osteogenic effects on MSC.

Other possible reasons for overall discrepancy about the effects of EGF on MSC differentiation include the heterogeneity of human MSC preparations including primary or immortalized. Kratchmarova and her colleagues used human MSCs immortalized by human telomerase reverse transcriptase (hTERT) [46], whereas Krampera and his colleague used primary human MSCs [45]. We and Griffith’s group also used AOC (Adipogenic, Osteogenic, Chondrogenic) clone of human MSCs immortalized by hTERT [3, 47, 51]. As immortalized MSCs were derived from single clone, it is possible that discrepant results between Kratchmarova’s group and our group might be due to clone selection bias. Heterogeneity exists even within primary preparation too. The expression levels of EGFR/ErbB-1 are highly variable in each clone of human MSC preparation, up to 77-fold difference among clones [52]. In this report, no significant correlation was observed between the levels EGFR/ErbB-1 and osteogenic differentiation capacity, although the levels of EGFR/ErbB-1 in average were higher in nonbone-forming colonies than bone-forming colonies. Patterns of protein tyrosine phosphorylation downstream of EGFR/ErbB-1 appeared heterogenous among colonies also. Thus, it is likely that heterogeneity does exist not only in EGFR/ErbB-1 expression levels, but also in the downstream signaling pathways from EGFR/ErbB-1 even within the same MSC preparation, which should also contribute to the overall discrepancy about the effects of EGF on MSC differentiation.

As we see above, reports about EGFR agonists and their effects on ongoing MSC differentiation still provide discrepant directives mainly due to different EGF conditions and external stimuli. Still, we need to emphasize that both our report and Krampera report agree that EGF or HB-EGF treatment does not diminish MSC differentiation potentials [3, 45]. There are no reports showing that EGF alone in the absence of osteogenic chemical cues promotes osteogenic differentiation of MSCs. In vitro MSC differentiation into adipogenic and chondrogenic lineages was not altered by soluble EGF [3]. Thus, soluble EGF can still be used to expand MSCs in vitro without inducing differentiation or sacrificing differentiation potentials.

3. EGF Treatment to Enhance Therapeutic Potentials of MSC

3.1. EGF Enhances Motility of MSC

Numerous cellular, hormonal, matrix and enzymatic activities are involved in wound repair and tissue regeneration processes. EGF is one of the pivotal growth factors present in the wound bed, accelerating wound repair along with other growth factors such as PDGF. Topical application of recombinant EGF was shown to accelerate epithelialization of wound healing process including diabetic foot ulcer [53, 54].

EGF is secreted from platelets and macrophages in wounded tissues [55]. HB-EGF is abundant in ECM [56]. Both EGF and HB-EGF stimulate proliferation and migration of fibroblasts and keratinocytes. Similarly, MSCs transplanted in the wounded tissues need to proliferate and repopulate themselves to promote wound healing and tissue regeneration processes in the MSC-based therapeutics. We and others showed that both EGF and HB-EGF elicit mitogenic and motogenic response of MSCs in vitro [3, 45, 52]. Thus, it is speculated that EGF-induced mitogenic and motogenic responses of MSCs play a role in regulating proliferation and repopulation of MSCs in the wounded tissues.

EGFR/ErbB1 ligands exist not only in soluble form, but also within multiple EGF-like repeats of extracellular matrix molecules such as tenascin and laminin in vivo [57]. We have previously shown that these EGF-like repeats within tenascin C bind to EGFR/ErbB1 and produce intracellular signaling promoting cell motility and adhesion, similar to tethered EGF [58, 59]. Tenacin C is produced by keratinocytes and fibroblasts during wound healing process, and thus it might serve as endogenous tethered EGF-like ligands and produce promigratory tracks for fibroblasts or implanted MSCs within the wound healing edges [57, 60].

Although motility enables MSC to reposition themselves in wounded tissues, it might make precise control of in vivo cell distribution difficult. One possible approach is to create concentration gradients or patterns of tethered EGF within MSC-embedded scaffolds. Motogenic activity of EGF is preserved in the tethered form, as human keratinocytes on tethered EGF gradients were reported to migrate in the direction of higher tethered EGF concentration [61]. Tethered form of EGF allows more precise control of EGF concentration and patterning within tissue microenvironments and one that lasts over a longer duration. This connecting of motogenic ligand to the space-forming matrix should be a strong tool in the field of tissue engineering.

3.2. EGF Enhances Paracrine Activities of MSC

MSC-based therapeutics heavily relies on the strong capability to secrete various growth factors and cytokines to promote angiogenesis, wound repair, and tissue regeneration [7, 21–25]. MSCs are required to be transplanted into the wounded tissues, which fail to heal otherwise. In vivo microenvironments of these nonhealing wounds are characterized by lack of oxygen and nutrients due to compromised blood flow and by exuberant proinflammatory mediators [62–64]. MSCs are needed to produce bioactive molecules even in those harsh environments to exert tissue regenerative effects. Indeed, the proinflammatory mediator tumor necrosis factor-alpha (TNF- ) or lipopolysacharide (LPS) was shown to enhance paracrine and autocrine functions of MSCs [65]. Also, TGF- , another EGFR/ErbB1 ligand, was shown to further increase VEGF secretion from MSCs already up-regulated by TNF- stimulation in a p42/44 MAPK dependent manner [66, 67].

Our in vitro data showed that EGF treatment of MSCs further promotes secretion of VEGF and HGF, but not bFGF (Figure 4), in agreement with a previous study [65]. Both VEGF and HGF play a pivotal role in MSC-mediated accelerated wound healing through inducing angiogenesis and improving oxygen supplies to the ischemic tissues [7, 21, 68–70].


(a)
(b)
(c)
(a)
(b)
(c) Effects of EGF treatment (10 nM) on paracrine activities of human primary MSCs. MSCs were cultured in serum-free culture medium with and without EGF (10 nM) for 24 hours. Concentrations of VEGF (a), HGF (b), and bFGF (c) within conditioned media were measured by ELISA and standardized to the total amount of cellular protein contents (*

Thus, it is likely that soluble EGFR/ErbB1 ligands (TGF- , EGF and HB-EGF) enhance paracrine and autocrine functions of MSCs not only in vitro, but also in vivo, even in the inflammatory microenvironments within nonhealing wounded tissues. It is also likely that tethered EGF enhances paracrine and autocrine functions of MSCs in vitro as well as in vivo through strong and sustained activation of p42/44 MAPK pathway in the downstream of EGFR/ErbB1 [44]. Both soluble and tethered EGFR/ErbB1 ligands are speculated to promote wound healing and tissue regeneration process through stimulating the secretion of angiogenic growth factors from transplanted MSCs in vivo. Further studies are warranted to elucidate the role of soluble EGFR/ErbB1 ligands and tethered EGF on the paracrine and autocrine effects of MSCs in vitro as well as in vivo.

3.3. Does EGF Enhance Therapeutic Potentials of MSC?

EGF stimulates cell proliferation and enhances self-renewal of MSCs, especially undifferentiated early progenitors within the MSC preparations in vitro. The presence of early progenitors is critical for MSC-based therapeutics, as MSCs from old donors and high passaged MSCs have decreased paracrine activity and reduced organ protective effects upon transplantation, presumably through loss of early progenitors [27, 29, 30, 36, 39]. EGF treatment also enhances cell motility, which is required for repopulation of MSCs within the wound bed. EGF treatment further increases paracrine secretion of VEGF and HGF, both of which enhance angiogenesis and promote wound healing and tissue regeneration also being stimulatory for the adherent cells resident within the wound bed. Taken together, it is reasonable to hypothesize that in vitro MSC treatment with EGFR/ErbB1 ligands enhances therapeutic potentials of MSCs. This could be tested by in vivo study.

It is also reasonable to hypothesize that EGFR/ErbB1 stimulation on MSC enhances therapeutic potentials of MSCs in vivo, presumably through augmenting paracrine activity and exerting both mitogenic and motogenic activities of MSCs. Biodegradable scaffolds are a promising approach to support cell delivery, guide proliferation and differentiation of the cells [71]. Drug delivery scaffolds or growth factor release scaffolds are also available, which enables the controlled release of growth factor [72]. Availability of EGF is less predictable in vivo setting however, EGF slow releasing scaffolds allow for better prediction of EGF concentration within microenvironments in vivo, thus, in vitro findings should be better translated to the in vivo settings and the role of EGF in MSC-based therapeutics could be better evaluated.

In addition to soluble EGFR/ErbB1 ligands, tethered EGF might bestow even stronger therapeutic potential on MSCs. First, tethered EGF exerts proliferative and cytoprotective effects on MSCs [44, 73]. Therapeutic effects of MSC largely depend on the number of injected MSC [19] however, low viability of postimplant MSCs limits the overall effectiveness of MSC-based therapeutics due to harsh microenvironments [15, 74]. Thus, improvement of postimplant MSC survival should increase the efficacy of MSC-based therapeutics. Second, tethered EGF provides pro-osteogenic cues for MSCs, thus it could be utilized in both in vitro and in vivo osteogenic differentiation of MSCs. This mechanism might play a significant role in vivo, as laminin 5, which contains EGF-like repeats, was shown to stimulate osteogenic differentiation of human MSCs through activation of ERK within bone tissue [75]. Thus, MSC-embedded scaffold with tethered EGF could be potentially applicable to current human studies such as osteogenic imperfecta [76–78].

The challenge of translating theoretical findings to bedsides always resides in moving from in vitro to in vivo studies and then into people. While we cannot foresee all the obstacles, the main challenge in this translation involves the inflammatory situation and immunological issues. In the case of the latter, this is moot if the MSCs are autologous, but allogenic MSCs are also useful, especially for aged patients, as MSC harvest and subsequent ex vivo expansion might be limited for those populations [26, 39]. The immunosuppressive nature of MSC makes allogenic transplantation feasible [26]. Nonspecific inflammation due to any foreign body is something not avoidable. Actually we propose that the tethered EGF confer resistance to death signals on MSCs [44]. Still the complex mixture of inflammatory cytokines and chemokines may alter the response to EGFR/ErbB1 ligands in unpredictable ways. Lastly, if the inflammatory response runs towards fibrosis, MSCs risk being walled-off from the site of injury. Another potential challenge of the translation into people is underlying disease conditions, as nutrient delivery, oxygen supplies, and removal of toxic metabolites are often severely compromised in these populations [62–64] in diabetes the hyperglycemia also impacts EGFR signaling pathways [79, 80]. These harsh microenvironments, with alter extracellular pH, will impinge on the MSC behavior in an unpredictable manner. That is why we are quickly moving to test these models in increasingly challenged animal models.

4. Epilogue

Previous studies suggest that EGF facilitates expansion of colony forming early progenitors in MSC population without inducing differentiation or compromising differentiation potentials. EGF treatment also promotes paracrine activity of MSC, at least the production of VEGF and HGF, both of which are pivotal for wound healing and tissue regeneration. EGF can be utilized to promote expansion and paracrine activities of MSCs in vitro, at least with short-term treatment with EGF. For in vivo settings, EGF can be incorporated in growth factor-releasing scaffolds encapsulating MSCs to augment MSC proliferation and paracrine action. Tethered form of EGF can also be incorporated in the scaffold to control osteogenic differentiation of MSCs or MSC distributions in vivo. The roles of EGFR/ErbB1 ligands and downstream signaling from EGFR/ErbB1 on MSC physiology are summarized in Figures 5 and 6. Overall, it should be reasonable to utilize EGF for MSC expansion in vitro, enhancement of MSC therapeutic potentials in vivo, and regulation of MSC differentiation both in vitro and in vivo.


Simplified diagram of EGFR/ErbB1 signaling pathways in MSC physiology. EGFR/ErbB1 ligands activate PLCγ pathway, p42/44 MAPK pathway, and PI3K/Akt pathways in MSCs [3]. PLCγ pathway plays a pivotal role in motogenic activity, whereas p42/44 MAPK pathway plays a key role in mitogenic activity and paracrine activities of certain factors such as VEGF [42, 43, 66, 67]. Sustained and strong activation of p42/44 MAPK pathway exerts cytoprotective and pro-osteogenic effects [44, 47], whereas PI3K/Akt pathway might exert anti-osteogenic effects [46].


The roles of soluble and tethered EGFR/ErbB1 ligands on MSC physiology. Soluble EGFR/ErbB1 ligands (EGF, HB-EGF, TGF-

Acknowledgments

Preparation of this article was supported by Grants from AHA Beginner-Grant-in-aid (09BGIA2050227) and start-up fund from Department of Pathology, OSU (K.T.) and from NIH (R01GM063569 and R01GM069668) (A.W.). We would like to thank Dr. J. Van Brooklyn (OSU, Columbus, OH) for his generous instrumental support.

References

  1. F. P. Barry and J. M. Murphy, “Mesenchymal stem cells: clinical applications and biological characterization,” International Journal of Biochemistry and Cell Biology, vol. 36, no. 4, pp. 568–584, 2004. View at: Publisher Site | Google Scholar
  2. D. G. Phinney and D. J. Prockop, “Concise review: mesenchymal stem/multipotent stromal cells: the state of transdifferentiation and modes of tissue repair—current views,” Stem Cells, vol. 25, no. 11, pp. 2896–2902, 2007. View at: Publisher Site | Google Scholar
  3. K. Tamama, V. H. Fan, L. G. Griffith, H. C. Blair, and A. Wells, “Epidermal growth factor as a candidate for ex vivo expansion of bone marrow-derived mesenchymal stem cells,” Stem Cells, vol. 24, no. 3, pp. 686–695, 2006. View at: Publisher Site | Google Scholar
  4. D. J. Prockop, “Marrow stromal cells as stem cells for nonhematopoietic tissues,” Science, vol. 276, no. 5309, pp. 71–74, 1997. View at: Publisher Site | Google Scholar
  5. M. F. Pittenger, A. M. Mackay, S. C. Beck et al., “Multilineage potential of adult human mesenchymal stem cells,” Science, vol. 284, no. 5411, pp. 143–147, 1999. View at: Publisher Site | Google Scholar
  6. M. Owen and A. J. Friedenstein, “Stromal stem cells: marrow-derived osteogenic precursors,” Ciba Foundation Symposium, vol. 136, pp. 42–60, 1988. View at: Google Scholar
  7. T. Kinnaird, E. Stabile, M. S. Burnett, and S. E. Epstein, “Bone marrow-derived cells for enhancing collateral development: mechanisms, animal data, and initial clinical experiences,” Circulation Research, vol. 95, no. 4, pp. 354–363, 2004. View at: Publisher Site | Google Scholar
  8. A. J. Friedenstein, J. F. Gorskaja, and N. N. Kulagina, “Fibroblast precursors in normal and irradiated mouse hematopoietic organs,” Experimental Hematology, vol. 4, no. 5, pp. 267–274, 1976. View at: Google Scholar
  9. J. Oswald, S. Boxberger, B. Jørgensen et al., “Mesenchymal stem cells can be differentiated into endothelial cells in vitro,” Stem Cells, vol. 22, no. 3, pp. 377–384, 2004. View at: Google Scholar
  10. K. Tamama, C. K. Sen, and A. Wells, “Differentiation of bone marrow mesenchymal stem cells into the smooth muscle lineage by blocking ERK/MAPK signaling pathway,” Stem Cells and Development, vol. 17, no. 5, pp. 897–908, 2008. View at: Publisher Site | Google Scholar
  11. M. F. Pittenger and B. J. Martin, “Mesenchymal stem cells and their potential as cardiac therapeutics,” Circulation Research, vol. 95, no. 1, pp. 9–20, 2004. View at: Publisher Site | Google Scholar
  12. S. Davani, A. Marandin, N. Mersin et al., “Mesenchymal progenitor cells differentiate into an endothelial phenotype, enhance vascular density, and improve heart function in a rat cellular cardiomyoplasty model,” Circulation, vol. 108, no. 10, supplement, pp. II253–II258, 2003. View at: Publisher Site | Google Scholar
  13. L. C. Amado, A. P. Saliaris, K. H. Schuleri et al., “Cardiac repair with intramyocardial injection of allogeneic mesenchymal stem cells after myocardial infarction,” Proceedings of the National Academy of Sciences of the United States of America, vol. 102, no. 32, pp. 11474–11479, 2005. View at: Publisher Site | Google Scholar
  14. W. Dai, S. L. Hale, B. J. Martin et al., “Allogeneic mesenchymal stem cell transplantation in postinfarcted rat myocardium: short- and long-term effects,” Circulation, vol. 112, no. 2, pp. 214–223, 2005. View at: Publisher Site | Google Scholar
  15. C. Toma, M. F. Pittenger, K. S. Cahill, B. J. Byrne, and P. D. Kessler, “Human mesenchymal stem cells differentiate to a cardiomyocyte phenotype in the adult murine heart,” Circulation, vol. 105, no. 1, pp. 93–98, 2002. View at: Publisher Site | Google Scholar
  16. T. Ziegelhoeffer, B. Fernandez, S. Kostin et al., “Bone marrow-derived cells do not incorporate into the adult growing vasculature,” Circulation Research, vol. 94, no. 2, pp. 230–238, 2004. View at: Publisher Site | Google Scholar
  17. Y. Wu, L. Chen, P. G. Scott, and E. E. Tredget, “Mesenchymal stem cells enhance wound healing through differentiation and angiogenesis,” Stem Cells, vol. 25, no. 10, pp. 2648–2659, 2007. View at: Publisher Site | Google Scholar
  18. M. Sasaki, R. Abe, Y. Fujita, S. Ando, D. Inokuma, and H. Shimizu, “Mesenchymal stem cells are recruited into wounded skin and contribute to wound repair by transdifferentiation into multiple skin cell type,” Journal of Immunology, vol. 180, no. 4, pp. 2581–2587, 2008. View at: Google Scholar
  19. V. Falanga, S. Iwamoto, M. Chartier et al., “Autologous bone marrow-derived cultured mesenchymal stem cells delivered in a fibrin spray accelerate healing in murine and human cutaneous wounds,” Tissue Engineering, vol. 13, no. 6, pp. 1299–1312, 2007. View at: Publisher Site | Google Scholar
  20. E. H. Javazon, S. G. Keswani, A. T. Badillo et al., “Enhanced epithelial gap closure and increased angiogenesis in wounds of diabetic mice treated with adult murine bone marrow stromal progenitor cells,” Wound Repair and Regeneration, vol. 15, no. 3, pp. 350–359, 2007. View at: Publisher Site | Google Scholar
  21. T. Kinnaird, E. Stabile, M. S. Burnett et al., “Local delivery of marrow-derived stromal cells augments collateral perfusion through paracrine mechanisms,” Circulation, vol. 109, no. 12, pp. 1543–1549, 2004. View at: Publisher Site | Google Scholar
  22. G. E. Kilroy, S. J. Foster, X. Wu et al., “Cytokine profile of human adipose-derived stem cells: expression of angiogenic, hematopoietic, and pro-inflammatory factors,” Journal of Cellular Physiology, vol. 212, no. 3, pp. 702–709, 2007. View at: Publisher Site | Google Scholar
  23. L. Chen, E. E. Tredget, P. Y. G. Wu, Y. Wu, and Y. Wu, “Paracrine factors of mesenchymal stem cells recruit macrophages and endothelial lineage cells and enhance wound healing,” PLoS ONE, vol. 3, no. 4, article e1886, 2008. View at: Publisher Site | Google Scholar
  24. T. Schinkothe, W. Bloch, and A. Schmidt, “In vitro secreting profile of human mesenchymal stem cells,” Stem Cells and Development, vol. 17, no. 1, pp. 199–205, 2008. View at: Publisher Site | Google Scholar
  25. M. Gnecchi, H. He, N. Noiseux et al., “Evidence supporting paracrine hypothesis for Akt-modified mesenchymal stem cell-mediated cardiac protection and functional improvement,” FASEB Journal, vol. 20, no. 6, pp. 661–669, 2006. View at: Publisher Site | Google Scholar
  26. A. I. Caplan, “Why are MSCs therapeutic? New data: new insight,” Journal of Pathology, vol. 217, no. 2, pp. 318–324, 2009. View at: Publisher Site | Google Scholar
  27. I. Sekiya, B. L. Larson, J. R. Smith, R. Pochampally, J.-G. Cui, and D. J. Prockop, “Expansion of human adult stem cells from bone marrow stroma: conditions that maximize the yields of early progenitors and evaluate their quality,” Stem Cells, vol. 20, no. 6, pp. 530–541, 2002. View at: Google Scholar
  28. N. Stute, K. Holtz, M. Bubenheim, C. Lange, F. Blake, and A. R. Zander, “Autologous serum for isolation and expansion of human mesenchymal stem cells for clinical use,” Experimental Hematology, vol. 32, no. 12, pp. 1212–1225, 2004. View at: Publisher Site | Google Scholar
  29. D. C. Colter, R. Class, C. M. DiGirolamo, and D. J. Prockop, “Rapid expansion of recycling stem cells in cultures of plastic-adherent cells from human bone marrow,” Proceedings of the National Academy of Sciences of the United States of America, vol. 97, no. 7, pp. 3213–3218, 2000. View at: Publisher Site | Google Scholar
  30. P. R. Crisostomo, M. Wang, G. M. Wairiuko et al., “High passage number of stem cells adversely affects stem cell activation and myocardial protection,” Shock, vol. 26, no. 6, pp. 575–580, 2006. View at: Publisher Site | Google Scholar
  31. P. A. Sotiropoulou, S. A. Perez, M. Salagianni, C. N. Baxevanis, and M. Papamichail, “Cell culture medium composition and translational adult bone marrow-derived stem cell research,” Stem Cells, vol. 24, no. 5, pp. 1409–1410, 2006. View at: Publisher Site | Google Scholar
  32. F. Ng, S. Boucher, S. Koh et al., “PDGF, TGF- ß , and FGF signaling is important for differentiation and growth of mesenchymal stem cells (MSCs): transcriptional profiling can identify markers and signaling pathways important in differentiation of MSCs into adipogenic, chondrogenic, and osteogenic lineages,” Blood, vol. 112, no. 2, pp. 295–307, 2008. View at: Publisher Site | Google Scholar
  33. H. Agata, N. Watanabe, Y. Ishii et al., “Feasibility and efficacy of bone tissue engineering using human bone marrow stromal cells cultivated in serum-free conditions,” Biochemical and Biophysical Research Communications, vol. 382, no. 2, pp. 353–358, 2009. View at: Publisher Site | Google Scholar
  34. E. H. Javazon, K. J. Beggs, and A. W. Flake, “Mesenchymal stem cells: paradoxes of passaging,” Experimental Hematology, vol. 32, no. 5, pp. 414–425, 2004. View at: Publisher Site | Google Scholar
  35. E. M. Horwitz, K. Le Blanc, M. Dominici et al., “Clarification of the nomenclature for MSC: the International Society for Cellular Therapy position statement,” Cytotherapy, vol. 7, no. 5, pp. 393–395, 2005. View at: Publisher Site | Google Scholar
  36. D. C. Colter, I. Sekiya, and D. J. Prockop, “Identification of a subpopulation of rapidly self-renewing and multipotential adult stem cells in colonies of human marrow stromal cells,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 14, pp. 7841–7845, 2001. View at: Publisher Site | Google Scholar
  37. J. R. Smith, R. Pochampally, A. Perry, S.-C. Hsu, and D. J. Prockop, “Isolation of a highly clonogenic and multipotential subfraction of adult stem cells from bone marrow stroma,” Stem Cells, vol. 22, no. 5, pp. 823–831, 2004. View at: Google Scholar
  38. R. Pochampally, “Colony forming unit assays for MSCs,” Methods in Molecular Biology, vol. 449, pp. 83–91, 2008. View at: Publisher Site | Google Scholar
  39. S. Jiang, H. Kh. Haider, R. P. H. Ahmed, N. M. Idris, A. Salim, and M. Ashraf, “Transcriptional profiling of young and old mesenchymal stem cells in response to oxygen deprivation and reparability of the infarcted myocardium,” Journal of Molecular and Cellular Cardiology, vol. 44, no. 3, pp. 582–596, 2008. View at: Publisher Site | Google Scholar
  40. S. Cohen and G. A. Elliott, “The stimulation of epidermal keratinization by a protein isolated from the submaxillary gland of the mouse,” Journal of Investigative Dermatology, vol. 40, pp. 1–5, 1963. View at: Google Scholar
  41. G. Carpenter and S. Cohen, “Epidermal growth factor,” Journal of Biological Chemistry, vol. 265, no. 14, pp. 7709–7712, 1990. View at: Google Scholar
  42. A. Wells, “EGF receptor,” International Journal of Biochemistry and Cell Biology, vol. 31, no. 6, pp. 637–643, 1999. View at: Publisher Site | Google Scholar
  43. A. Wells, M. F. Ware, F. D. Allen, and D. A. Lauffenburger, “Shaping up for shipping out: PLC γ signaling of morphology changes in EGF-stimulated fibroblast migration,” Cell Motility and the Cytoskeleton, vol. 44, no. 4, pp. 227–233, 1999. View at: Publisher Site | Google Scholar
  44. V. H. Fan, K. Tamama, A. Au et al., “Tethered epidermal growth factor provides a survival advantage to mesenchymal stem cells,” Stem Cells, vol. 25, no. 5, pp. 1241–1251, 2007. View at: Publisher Site | Google Scholar
  45. M. Krampera, A. Pasini, A. Rigo et al., “HB-EGF/HER-1 signaling in bone marrow mesenchymal stem cells: inducing cell expansion and reversibly preventing multilineage differentiation,” Blood, vol. 106, no. 1, pp. 59–66, 2005. View at: Publisher Site | Google Scholar
  46. I. Kratchmarova, B. Blagoev, M. Haack-Sorensen, M. Kassem, and M. Mann, “Mechanism of divergent growth factor effects in mesenchymal stem cell differentiation,” Science, vol. 308, no. 5727, pp. 1472–1477, 2005. View at: Publisher Site | Google Scholar
  47. M. O. Platt, A. J. Roman, A. Wells, D. A. Lauffenburger, and L. G. Griffith, “Sustained epidermal growth factor receptor levels and activation by tethered ligand binding enhances osteogenic differentiation of multi-potent marrow stromal cells,” Journal of Cellular Physiology, vol. 221, no. 2, pp. 306–317, 2009. View at: Publisher Site | Google Scholar
  48. R. K. Jaiswal, N. Jaiswal, S. P. Bruder, G. Mbalaviele, D. R. Marshak, and M. F. Pittenger, “Adult human mesenchymal stem cell differentiation to the osteogenic or adipogenic lineage is regulated by mitogen-activated protein kinase,” Journal of Biological Chemistry, vol. 275, no. 13, pp. 9645–9652, 2000. View at: Publisher Site | Google Scholar
  49. R. M. Salasznyk, R. F. Klees, M. K. Hughlock, and G. E. Plopper, “ERK signaling pathways regulate the osteogenic differentiation of human mesenchymal stem cells on collagen I and vitronectin,” Cell Communication and Adhesion, vol. 11, no. 5-6, pp. 137–153, 2004. View at: Publisher Site | Google Scholar
  50. D. F. Ward Jr., R. M. Salasznyk, R. F. Klees et al., “Mechanical strain enhances extracellular matrix-induced gene focusing and promotes osteogenic differentiation of human mesenchymal stem cells through an extracellular-related kinase-dependent pathway,” Stem Cells and Development, vol. 16, no. 3, pp. 467–479, 2007. View at: Publisher Site | Google Scholar
  51. T. Okamoto, T. Aoyama, T. Nakayama et al., “Clonal heterogeneity in differentiation potential of immortalized human mesenchymal stem cells,” Biochemical and Biophysical Research Communications, vol. 295, no. 2, pp. 354–361, 2002. View at: Publisher Site | Google Scholar
  52. K. Satomura, A. R. Derubeis, N. S. Fedarko et al., “Receptor tyrosine kinase expression in human bone marrow stromal cells,” Journal of Cellular Physiology, vol. 177, no. 3, pp. 426–438, 1998. View at: Publisher Site | Google Scholar
  53. G. L. Brown, L. B. Nanney, J. Griffen et al., “Enhancement of wound healing by topical treatment with epidermal growth factor,” The New England Journal of Medicine, vol. 321, no. 2, pp. 76–79, 1989. View at: Google Scholar
  54. J. P. Hong, H. D. Jung, and Y. W. Kim, “Recombinant human epidermal growth factor (EGF) to enhance healing for diabetic foot ulcers,” Annals of Plastic Surgery, vol. 56, no. 4, pp. 394–398, 2006. View at: Publisher Site | Google Scholar
  55. G. Schultz, D. S. Rotatori, and W. Clark, “EGF and TGF- α in wound healing and repair,” Journal of Cellular Biochemistry, vol. 45, no. 4, pp. 346–352, 1991. View at: Google Scholar
  56. M. Marikovsky, K. Breuing, P. Y. Liu et al., “Appearance of heparin-binding EGF-like growth factor in wound fluid as a response to injury,” Proceedings of the National Academy of Sciences of the United States of America, vol. 90, no. 9, pp. 3889–3893, 1993. View at: Google Scholar
  57. K. T. Tran, L. Griffith, and A. Wells, “Extracellular matrix signaling through growth factor receptors during wound healing,” Wound Repair and Regeneration, vol. 12, no. 3, pp. 262–268, 2004. View at: Publisher Site | Google Scholar
  58. A. K. V. Iyer, K. T. Tran, L. Griffith, and A. Wells, “Cell surface restriction of EGFR by a tenascin cytotactin-encoded EGF-like repeat is preferential for motility-related signaling,” Journal of Cellular Physiology, vol. 214, no. 2, pp. 504–512, 2008. View at: Publisher Site | Google Scholar
  59. C. S. Swindle, K. T. Tran, T. D. Johnson et al., “Epidermal growth factor (EGF)-like repeats of human tenascin-C as ligands for EGF receptor,” Journal of Cell Biology, vol. 154, no. 2, pp. 459–468, 2001. View at: Publisher Site | Google Scholar
  60. E. J. Mackie, W. Halfter, and D. Liverani, “Induction of tenascin in healing wounds,” Journal of Cell Biology, vol. 107, no. 6, part 2, pp. 2757–2767, 1988. View at: Google Scholar
  61. T. J. Stefonek and K. S. Masters, “Immobilized gradients of epidermal growth factor promote accelerated and directed keratinocyte migration,” Wound Repair and Regeneration, vol. 15, no. 6, pp. 847–855, 2007. View at: Publisher Site | Google Scholar
  62. G. C. Gurtner, S. Werner, Y. Barrandon, and M. T. Longaker, “Wound repair and regeneration,” Nature, vol. 453, no. 7193, pp. 314–321, 2008. View at: Publisher Site | Google Scholar
  63. G. Henry and W. L. Garner, “Inflammatory mediators in wound healing,” Surgical Clinics of North America, vol. 83, no. 3, pp. 483–507, 2003. View at: Publisher Site | Google Scholar
  64. C. K. Sen, “Wound healing essentials: let there be oxygen,” Wound Repair and Regeneration, vol. 17, no. 1, pp. 1–18, 2009. View at: Publisher Site | Google Scholar
  65. P. R. Crisostomo, Y. Wang, T. A. Markel, M. Wang, T. Lahm, and D. R. Meldrum, “Human mesenchymal stem cells stimulated by TNF- α , LPS, or hypoxia produce growth factors by an NF κ B- but not JNK-dependent mechanism,” American Journal of Physiology, vol. 294, no. 3, pp. C675–C682, 2008. View at: Publisher Site | Google Scholar
  66. Y. Wang, P. R. Crisostomo, M. Wang, T. A. Markel, N. M. Novotny, and D. R. Meldrum, “TGF- α increases human mesenchymal stem cell-secreted VEGF by MEK- and PI3-K- but not JNK- or ERK-dependent mechanisms,” American Journal of Physiology, vol. 295, no. 4, pp. R1115–R1123, 2008. View at: Publisher Site | Google Scholar
  67. Y. Wang, M. Wang, A. M. Abarbanell et al., “MEK mediates the novel cross talk between TNFR2 and TGF-EGFR in enhancing vascular endothelial growth factor (VEGF) secretion from human mesenchymal stem cells,” Surgery, vol. 146, no. 2, pp. 198–205, 2009. View at: Publisher Site | Google Scholar
  68. H.-F. Duan, C.-T. Wu, D.-L. Wu et al., “Treatment of myocardial ischemia with bone marrow-derived mesenchymal stem cells overexpressing hepatocyte growth factor,” Molecular Therapy, vol. 8, no. 3, pp. 467–474, 2003. View at: Publisher Site | Google Scholar
  69. T. Kinnaird, E. Stabile, M. S. Burnett et al., “Marrow-derived stromal cells express genes encoding a broad spectrum of arteriogenic cytokines and promote in vitro and in vivo arteriogenesis through paracrine mechanisms,” Circulation Research, vol. 94, no. 5, pp. 678–685, 2004. View at: Publisher Site | Google Scholar
  70. I. Rosová, M. Dao, B. Capoccia, D. Link, and J. A. Nolta, “Hypoxic preconditioning results in increased motility and improved therapeutic potential of human mesenchymal stem cells,” Stem Cells, vol. 26, no. 8, pp. 2173–2182, 2008. View at: Publisher Site | Google Scholar
  71. J. Guan, K. L. Fujimoto, M. S. Sacks, and W. R. Wagner, “Preparation and characterization of highly porous, biodegradable polyurethane scaffolds for soft tissue applications,” Biomaterials, vol. 26, no. 18, pp. 3961–3971, 2005. View at: Publisher Site | Google Scholar
  72. J. Guan, J. J. Stankus, and W. R. Wagner, “Biodegradable elastomeric scaffolds with basic fibroblast growth factor release,” Journal of Controlled Release, vol. 120, no. 1-2, pp. 70–78, 2007. View at: Publisher Site | Google Scholar
  73. M. O. Platt, C. L. Wilder, A. Wells, L. G. Griffith, and D. A. Lauffenburger, “Multipathway kinase signatures of multipotent stromal cells are predictive for osteogenic differentiation: tissue-specific stem cells,” Stem Cells, vol. 27, no. 11, pp. 2804–2814, 2009. View at: Google Scholar
  74. A. A. Mangi, N. Noiseux, D. Kong et al., “Mesenchymal stem cells modified with Akt prevent remodeling and restore performance of infarcted hearts,” Nature Medicine, vol. 9, no. 9, pp. 1195–1201, 2003. View at: Publisher Site | Google Scholar
  75. R. F. Klees, R. M. Salasznyk, K. Kingsley, W. A. Williams, A. Boskey, and G. E. Plopper, “Laminin-5 induces osteogenic gene expression in human mesenchymal stem cells through an ERK-dependent pathway,” Molecular Biology of the Cell, vol. 16, no. 2, pp. 881–890, 2005. View at: Publisher Site | Google Scholar
  76. E. M. Horwitz, P. L. Gordon, W. K. K. Koo et al., “Isolated allogeneic bone marrow-derived mesenchymal cells engraft and stimulate growth in children with osteogenesis imperfecta: implications for cell therapy of bone,” Proceedings of the National Academy of Sciences of the United States of America, vol. 99, no. 13, pp. 8932–8937, 2002. View at: Publisher Site | Google Scholar
  77. E. M. Horwitz, D. J. Prockop, L. A. Fitzpatrick et al., “Transplantability and therapeutic effects of bone marrow-derived mesenchymal cells in children with osteogenesis imperfecta,” Nature Medicine, vol. 5, no. 3, pp. 309–313, 1999. View at: Publisher Site | Google Scholar
  78. E. M. Horwitz, D. J. Prockop, P. L. Gordon et al., “Clinical responses to bone marrow transplantation in children with severe osteogenesis imperfecta,” Blood, vol. 97, no. 5, pp. 1227–1231, 2001. View at: Publisher Site | Google Scholar
  79. M. Portero-Otin, R. Pamplona, M. J. Bellmunt et al., “Advanced glycation end product precursors impair epidermal growth factor receptor signaling,” Diabetes, vol. 51, no. 5, pp. 1535–1542, 2002. View at: Google Scholar
  80. P. O. Prada, E. R. Ropelle, R. H. Mourao et al., “An EGFR tyrosine-kinase inhibitor (PD153035) improves glucose tolerance and insulin action in high-fat diet-fed mice,” Diabetes, vol. 58, no. 12, pp. 2910–2919, 2009. View at: Google Scholar

Copyright

Copyright © 2010 Kenichi Tamama et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


Generation of spinal motor neurons from human fetal brain-derived neural stem cells: Role of basic fibroblast growth factor

Neural stem cells (NSCs) have some specified properties but are generally uncommitted and so can change their fate after exposure to environmental cues. It is unclear to what extent this NSC plasticity can be modulated by extrinsic cues and what are the molecular mechanisms underlying neuronal fate determination. Basic fibroblast growth factor (bFGF) is a well-known mitogen for proliferating NSCs. However, its role in guiding stem cells for neuronal subtype specification is undefined. Here we report that in-vitro-expanded human fetal forebrain-derived NSCs can generate cholinergic neurons with spinal motor neuron properties when treated with bFGF within a specific time window. bFGF induces NSCs to express the motor neuron marker Hb9, which is blocked by specific FGF receptor inhibitors and bFGF neutralizing antibodies. This development of spinal motor neuron properties is independent of selective proliferation or survival and does not require high levels of MAPK activation. Thus our study indicates that bFGF can play an important role in modulating plasticity and neuronal fate of human NSCs and presumably has implications for exploring the full potential of brain NSCs for clinical applications, particularly in spinal motor neuron regeneration. © 2008 Wiley-Liss, Inc.


ACKNOWLEDGMENTS

This study was partly supported by a research grant from the National Natural Science Funding of China (81722028, 81802235, 81972150, 81572227, 81572237), Zhejiang Provincial Natural Science Foundation of China (LY17H060009, LR18H50001), Zhejiang Experimental Animal Science and Technology Project of China (2018C37112), and Wenzhou basic science research plan project (Y20180033). This study was also supported in part by the Australian Health and Medical Research Council (NHMRC, No 1107828, 1027932 to Jiake Xu). Granted by the Opening Project of Zhejiang Provincial Top Key Discipline of Clinical Medicine (No LKFJ017), Samuel Xu provided editorial support. The authors thank Jennifer Tickner and Nathan Pavlos for their comments on this manuscript.


Methods

MIAMI Cell Isolation

Whole bone marrow was obtained from the iliac crest of a 20 year old living male donor (Lonza Walkersville, Maryland MIAMI #3515), and were handled and processed following the guidelines for informed consent set by the University of Miami School of Medicine Committee on the Use of Human Subjects in Research. As previously described [3], isolated whole bone marrow cells were plated at a constant density of 1 × 10 5 cells/cm 2 in DMEM-low glucose media, containing 3% fetal bovine serum (FBS, Hyclone Waltham, MA, Lot#30039), 20 mM ascorbic acid (Fluka/Sigma St. Louis, MO, #49752), an essential fatty acid mixture (Sigma St. Louis, MO 12.9 nM arachidonic acid, (#A9673), 1.12 μM cholesterol (#C3045), 290 nM DL-alpha tocopherol-acetate (#T3376), 85.9 nM myristic acid (#M3128), 69.4 nM oleic acid (#01383), 76.5 nM palmitic acid (#P5585), 77.1 nM palmitoleic acid (P9417) and 68.9 nM stearic acid (#S4751) (modified from [21]) and antibiotics (100 U/mL penicillin, 0.1 mg/mL streptomycin) (Gibco Carlsbad, CA, #15140) on 10 ng/ml fibronectin (Sigma St. Louis, MO, #F2518) coated flasks (Nunclone Rochester, NY). Whole bone marrow cells, containing adherent and non-adherent cells, were incubated at 37°C under hypoxic conditions (3% O2, 5% CO2 and 92% N2). Seven days later, half of the culture medium was replaced. Fourteen days after the initial plating, the non-adherent cells were removed. Pooled colonies of adherent cells were rinsed with PBS and plated at low density for expansion (100 cells/cm 2 ) in 75 cm 2 fibronectin coated flasks.

MIAMI Cell Culture Conditions

MIAMI cells were grown in expansion media consisting of DMEM-low glucose (as described above) in low oxygen conditions (3% O2, 5% CO2 and 92% N2). Media was changed every 2-3 days and the cells were detached and pelleted using trypsin (Gibco Carlsbad, CA, #25300) upon reaching

60% confluency. Peleted cells were resuspended in media and plated in 10 ng/ml fibronectin (Sigma St Louis, MO, #F2518) coated flasks (Nunclon, Rochester, NY) at 100 cells/cm 2 . Prior to RNA isolation, adherent cells were rinsed 2× with PBS. MIAMI cells expanded for 3 passages were characterized using flow cytometry and were positive for MHC1, CD29, CD81, CD90 and 50% positive for CD63, and negative for MHC2, HLA-DR, CD49, CD109, CD54, CD56, CD36 (data not shown).

RS-1 Cell Culture Conditions

Human marrow stromal cells (hMSC, Donor#7081, 22yo male) were obtained from the laboratory of Dr. Darwin Prockop, Director, Texas A&M Health Science Center College of Medicine Institute for Regenerative Medicine. Bone marrow (BM) cells were isolated from human donors according to guidelines on the Use of Human Subjects in Research as described by all commercial vendors. The hMSC were cultured in Alpha-Minimum Essential medium (αMEM) with L-glutamine, but with no ribonucleosides or deoxyribonucleosides (Invitrogen/Gibco Carlsbad, CA, #12561-056), supplemented with 16.5% FBS (Hyclone Waltham, MA, #31752), 2 mM GlutaMAX (#35050) and antibiotics (Gibco Carlsbad, CA, #15140). To enrich for RS-1 cells, hMSC(P1) were plated at 37°C under normoxic conditions (21% O2, 5% CO2 and 74% N2) onto 10 ng/ml fibronectin (Sigma St. Louis, MO, #F2518) coated flasks (Nunclon Rochester, NY) overnight. The cells were detached using trypsin (Gibco Carlsbad, CA #25300) and seeded at 50 cells/cm 2 . RS-1 enriched hMSCs were detached at 30-40% confluency and re-plated at low density (50 cell/cm2) [7]. RS-1 cells were harvested for RNA isolation at each passage. MSC derived RS-1 cells, passage 2, were positive for CD29, CD90, CD105 and CD73 as determined by Tulane University Center for Gene Therapy (hMSCs #7801). RS-1 cells derived in our facilities, passage 3, were positive for MHC1, CD81, CD90, CD29 (20%), CD63 (45%) and negative for MHC2, HLA-DR, CD49, CD109, CD54, CD56, CD36, as determined using flow cytometry analysis (data not shown).

MSC Cell Culture Conditions

Human mesenchymal stem cells (MSC) derived from the iliac crest were purchased from Lonza (Walkersville, Maryland (PT-2501: 21yo female)). Bone marrow (BM) cells were isolated from human donors according to guidelines on the Use of Human Subjects in Research as described by all commercial vendors. The MSC were plated at 6,000 cells/cm 2 in DMEM-high glucose media (Gibco Carlsbad, CA, #31053) supplemented with 15% FBS (Hyclone Waltham, MA, #30039), ascorbic acid, antibiotics and essential fatty acids (as described above), and expanded at 21% O2, 5% CO2 and 92% N2. The entire culture media was changed every 3-4 days and the cells were detached and replated every 7 days [16]. MSC purchased from Lonza were positive for CD105, CD166, CD29, CD44, and negative for CD14, CD34 and CD45, as determined by flow cytometry (Lonza Wlkersville, Maryland (Document # TS-PT-212-8 06/09).

Neural Pre-treatment

Epidermal growth factor (EGF) and basic fibroblast growth factor (bFGF) treatment of MIAMI cells was performed using 20 ng/mL each of EGF (#AF-100-15) and bFGF (Peprotech Rocky Hill, NJ, #AF-100-18B) alone or in combination. The pre-treated cells were detached using trypsin and replated after day 5, followed by a second 5 day pretreatment period. Pre-treated cells were grown in expansion media under expansion conditions (3% O2, 5% CO2 and 92% N2). Media was changed every 2-3 days and the cells were split using trypsin (Gibco Carlsbad, CA, #25300) upon reaching

Endothelial Differentiation

For endothelial differentiation, MIAMI cells were plated at 20,000 cells/cm 2 in 6 well plates (Nunclone Rochester, NY) in DMEM-low glucose media, containing 100 μM Ascorbic Acid, antibiotics, essential fatty acids, angiogenic growth factor cocktail [Sigma St. Louis, MO: 10 ng/ml bFGF, 10 ng/ml EGF, 10 ng/ml IGF R&D Systems, Inc. Minneapolis, MN: 100 ng/ml VEGF], 100 nM Hydrocortisone, in atmosphere of 21% O2, 5% CO2 and incubated at 37°C for 21 days, with media changes every 5 days. Cells were harvested at day 10 and 21 and evaluated by RT-qPCR for the endothelial marker CD 31.

Total RNA Sample Preparation and cDNA Synthesis

MIAMI cells were detached (Trypsin) and centrifuged to form a cell pellet. RNA was isolated using the RNAqueous ® -4PCR kit (Ambion Austin, TX, #AM1914) according to manufacturer's directions. Total RNA was quantified on the Nanodrop ND-1000 Spectrophotometer (Nanodrop Wilmington, DE). Reverse transcription of 2 μg total RNA to cDNA was done with random hexamer primers using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems Foster City, CA, #4368814). The cDNA was diluted 1:20 (Nuclease-Free Water: Gibco#10977-015) to a final cDNA concentration of 5 ng/μl, aliqoted, and stored at -20°C until next use. Only RNA with a 260/280 ratio between 1.9-2.0 was used for PCR analysis.

Quantitative real-time RT-PCR (RT-qPCR)

Quantitative real-time PCR (RT-qPCR) was done using 10 μl of 1:20 diluted cDNA (50 ng) on the Mx3005P Multiplex Quantitative PCR System (Stratagene#401513) using RT-qPCR SYBR GREEN Reagents (Brilliant ® II SYBR ® Green QPCR Master Mix, Agilent Technologies) with ROX reference dye. Forward and reverse primer pairs were reconstituted in Nuclease Free Water (Gibco#10977-015). A 2 μM stock solution containing both forward and reverse primer pairs was mixed and stored at -20°C. A final concentration of 160 nM forward and reverse primer pairs was used for each RT-qPCR reaction. The cycling conditions were as follows: an initial 95°C for 10 minutes, followed by 40 cycles of 95°C for 30 sec, 58°C for 30 sec, 72°C for 15 sec. MxPro-Mx3005P v4.10 software was used to determine the CP for each amplification reaction. Results were exported to Microsoft Excel for analysis.

Analysis of RT-qPCR data

All of the corresponding RT-qPCR data was analyzed using the ΔΔCP method [13] and normalized against one negative control, and two reference genes (housekeeping genes).

The crossing point (CP) is defined as the point at which the fluorescence rises appreciably above the background fluorescence. The 'Fit Point Method' was used by the Mx3005P software to determine the CP for each reaction. The control sample was set to a value of "1" in all cases and error bars in the respective figures are displayed as standard deviation. The number of independent experiments is designated as "N" with 2-3 individual data points collected per experiment.

Normalization Genes

Eight genes were tested for normalization [beta-actin (ACTB, NM_001101), beta-2-microglobulin (B2M, NM_004048), eukaryotic translational elongation factor 1 alpha (EF1α, NM_001402), glyceraldehyde-3-phosphate dehydrogenase (GAPDH, NM_002046), Hypoxanthine phosphoribosyltransferase 1 (HPRT1, NM_000194), ribosomal protein L13a (RPL13a, NM_01242), tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein, zeta polypeptide variant 1 & 2 (YWHAZ, NM_003406 & NM_145690), and ubiquitin C (UBC, NM_021009)]. A list of primer pair sequences used are in Table# 1.

Determination of Primer Pair Efficiency

The determination of each genes' primer pair efficiency (E) for RT-qPCR was calculated using this equation: E = 10^(-1/m) [22]. The slope (m) was calculated by plotting the cycle number crossing point (CP) calculated during the exponential phase of the amplification plot (PxPro-Mx3005P v4.10 software) against the total cDNA concentration. Concentrations of cDNA ranged from 50-1 ng per reaction. The percent efficiency (%E) was also calculated: %E = (E-1)*100. N = 4 (2-3 data points per experiment) (Additional file # 1).

Construction of species-specific primer pairs

In order to create species-specific primer pairs that detect only human mRNA sequences or only rat mRNA sequences within a human-rat cDNA library, the corresponding human and rat mRNA sequences must have a unique region of at least 60 bp or more. Using the human and corresponding rat FASTA mRNA sequences for EF1α, RPL13a and YWHAZ, we used Blast-n http://blast.ncbi.nlm.nih.gov/Blast.cgi to compare the sequences. EF1α had 99% sequence coverage (100% identity) between the human and rat mRNA sequences. RPL13a had 57% sequence coverage (87% identity) and YWHAZ transcript variants 1 and 2 had 92% - 63% sequence coverage (100% identity). Therefore, RPL13a and YWHAZ both were candidate human species-specific normalization genes while EF1α did not have a region containing a unique sequence (≥60 bp) in order to create primer pairs. Human species-specific primer pairs were constructed for the 2 normalization genes RPL13a and YWHAZ and for 3 target genes stanniocalcin-1 (STC-1), tumor necrosis factor, alpha-induced protein 6 (TSG6), and latent transforming growth factor binding protein 2 (LTBP2). NCBI Primer-BLAST http://www.ncbi.nlm.nih.gov/tools/primer-blast/ was used for primer pair sequence construction [23] using the species-unique mRNA sequences (FASTA format). Gradient PCR was used to determine optimum annealing temperature. All human and rat specific primer pairs were validated with RT-qPCR using cDNA from human MIAMI cells H3515(3) or rat hippocampal organotypic cultures either separately or in combination. All primer pairs produced 1 species-specific amplicon, with minimum off-target amplification. This was determined by the melting curve of each amplification reaction (Additional File # 2) and agarose gel electrophoresis (data not shown). Approximately 3-5 primer pairs were tested per human or rat species-specific normalization or target gene. All RT-qPCR results were normalized against a negative control, and the 2 normalization genes hRPL13a and hYWHAZ (human), or rRPL13a (rat). Using this same method rat specific primer pairs were also constructed for RPL13a, IGF1, IGFBP3, and IGFBP5.

Model of ex vivo global cerebral ischemia for cross-species RT-qPCR analysis

All animal experiments were performed according to approved guidelines established by the University of Miami IACUC. The rat hippocampal organotypic slice preparation has been described in detail [19, 24]. Briefly, 400 μm brain slices were obtained from rat pups of either sex between postnatal days 9 and 10. Slices were cultured for two weeks in a medium consisting of 25% heat inactivated horse serum, 50% minimal essential medium, and 25% Hank's balanced salt solution, 5.5 mg/mL D-glucose and 1 mmol/L glutamine. For ischemia we used an established model consisting of combined oxygen and glucose deprivation ([19, 24]) during 40 mins. For OGD, oxygen is replaced with nitrogen and glucose with sucrose. MIAMI cells were pre-treated with bFGF and EGF (7 days: 50 ng/ml) prior to injection in the CA1 region of the hippocampus (7,500 cells/μl per injection (3 injections)). One hour after OGD induction and 24 hours after OGD total RNA was isolated from rat hippocampal organotypic slice cultures (described in [25]) with or without injected MIAMI. As described previously, 2 μg of total RNA was used for cDNA synthesis. RT-qPCR analysis was done using 5 μl of undiluted cDNA. Human and rat specific primer pairs are designated by (h) and (r) respectively (Table# 2: human specific primer pairs are designated by (*)).


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