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Abstract

Development of effective cryopreservation protocols will be essential to realizing the potential for clinical application of neural stem and progenitor cells. Current cryopreservation protocols have been largely employed in research, which does not require as stringent consideration of viability and sterility. Therefore, these protocols involve the use of serum and protein additives, which can potentially introduce contaminants, and slow cooling with DMSO/glycerol-based cryopreservation solutions, which impairs cell survival. We investigated whether serum- and protein-free vitrification is effective for functional cryopreservation of neurosphere cultures of neural stem or progenitor cells. To protect the samples from introduction of other contaminants during handling and cryostorage, an original “straw-in-straw” method (250 μl sterile straw placed in 500 μl straw) for direct immersion into liquid nitrogen and storing the samples was also introduced. The protocol employed brief step-wise exposure to vitrification solution composed of ethylene glycol (EG) and sucrose (40% v/v EG, 0.6 M sucrose) and removal of vitrification solution at room temperature. Evaluation of the effects of vitrification revealed that there were no differences between control and vitrified neural stem or progenitor cells in expression of the neural stem or progenitor cell markers, proliferation, or multipotent differentiation. This sterile method for the xeno-free cryopreservation of murine neurospheres without animal or human proteins may have the potential to serve as a starting point for the development of cryopreservation protocols for human neural stem and progenitor cells for clinical use.

Keywords

Vitrification Cryopreservation Neural progenitor cells Neural stem cells Neurospheres

Introduction

The potential clinical implications of neural stem and progenitor cell (NSPCs) therapies in the repair of nervous system injuries and disorders are immense. It is likely that safe and efficient cryopreservation of the NSPCs will be a prerequisite for quality assurance in the storage and distribution of the cells for clinical use (26). Effective storage to ensure reproducible supply of cells would benefit both future clinical applications and current basic research into the functions and properties of NSPCs. While stem cells self-renew, NSPCs cannot be maintained indefinitely by continuous passaging in culture. For example, when cultured long term, telomere shortening in NSPCs leads to senescence and associated dysfunctions in proliferation and neurogenesis (4, 37). While telomere shortening can be avoided by immortalization with overexpression of human telomerase reverse transcriptase (hTERT) (1), long-term culture of human neural progenitor cells immortalized in this manner can result in accumulation of karyotypic abnormalities (36). Such chromosomal abnormalities can in turn lead to abnormal proliferation and transformation of progenitor or stem cell characteristics, although this is not necessarily an inevitable consequence of all immortalization protocols (8, 21, 27). This could influence experimental results and their interpretation and, on clinical application, may be associated with increased risk of tumorigenesis. Development of efficient cryopreservation techniques is therefore essential.

It is anticipated that sterile and xeno-free culture and storage protocols will be a requirement for the ultimate realization of the clinical application of NSPCs. Sterile and reproducible cryostorage protocols would also facilitate basic research. As has been discussed in the fields of mesenchymal and embryonic stem cell research [for review see (13, 23, 24)], the use of animal derivatives and other organic components increases the risk of contamination and batch-to-batch variability. In view of the requirement for sterility, we demonstrated that cryopreservation of biological material by vitrification can be achieved using only nonbiological additives (17, 19, 22, 32, 38). We have also developed a protocol using a sealed “straw-in-straw” configuration (250 μl sterile straw placed in 500 μl straw), which enables maintenance of sterility during cryopreservation (18, 19). The protocol is both cost- and time-effective as it does not need expensive slow-cooling apparatus and requires only direct immersion into liquid nitrogen for cooling.

Approaches to cryopreservation can be classified as “freezing” and “vitrification.” Cryoprotectants are used in both approaches. In the case of freezing, the aim is to reduce damage to cells by the formation of extracellular ice crystals rather than intracellular ice. In contrast, vitrification aims to avoid ice crystal formation on both cooling and warming by achieving glass-like solidification [for review see (17)]. Having established the first clinical application of vitrification in the cryopreservation of oocytes for fertility treatment (15), we showed that vitrification protocols are preferable to slow-cooling freezing protocols for the preservation of oocytes and embryos (17). Slow-cooling freezing of human embryonic stem cell colonies in suspension is reportedly plagued by poor viability and abnormal differentiation upon recovery (12). Most studies have concluded that slow-cooling freezing protocols are suboptimal for cryopreservation of embryonic stem cells [for review see (7)]. A rapid cooling procedure for cryopreservation of hematopoietic progenitor cells from human cord blood in solutions employing serum was described in an earlier report (20). There have been interesting reports on cryopreservation of human and other primate embryonic stem cells avoiding conventional slow-cooling method by directly plunging into liquid nitrogen (5, 28, 29, 31, 39). Although these studies were a step forward, all of them employed serum or proteins of human or animal origin. Additionally, various concerns over the achievement of the glass-like vitreous state of samples need to be addressed.

We first sought to select the most promising concept to adopt as a basis for the development of protocols for the cryopreservation of NSPCs. On explorative investigation of freezing strategies, we found that freezing of cells is detrimental to cell viability and the structural integrity of neurospheres compared to cooling in solutions containing solute concentrations supporting vitrification (32). To establish the efficacy of a method for the cryopreservation of stem cells, it is essential to demonstrate that the cells preserve expression of stem cell markers and the capacity for multipotent differentiation. Preservation of the capacity for multipotent differentiation is important because any application in replacement therapy would require that stem cells differentiate into functional cell types for integration into neural networks to replace the particular neural cell types that are damaged or lost. In the present study, we aim to establish whether our protein- and serum-free vitrification protocol has the potential for effective stem cell cryopreservation for clinical and research applications. Using NSPCs we evaluate whether expression of neural stem cells markers, proliferative capacity, and the potential for multipotent neural differentiation are maintained following vitrification.

Materials and Methods

Culture of NSPCs

Pregnant C57BL/6J mice were anesthetized and embryonic day 14 (E14) fetuses were surgically extracted. The hippocampi were dissected out and digested in 0.25% trypsin-EDTA solution (Gibco, CA, USA) for 30 min with trituration every 15 min.

Twice the volume of PBS was added to stop the digestion and the tissues were sieved with a 70-μm cell strainer. The samples were centrifuged to pellet the cells and the cells were then plated with neurosphere medium. The neurosphere medium consisted of DMEM/F12 (Gibco) supplemented with N2 supplement (Gibco), 20 ng/ml EGF (Gibco), and 10 ng/ml bFGF (Gibco). After 2–3 weeks in culture visible neurospheres were present and the culture was passaged. For passage, neurospheres were allowed to settle in a microcentrifuge tube and then mechanically dissociated using a 200-μl pipette before replating in fresh neurosphere medium. All animal experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of the National University of Singapore and were conducted in accordance with the Guide for the Care and Use of Laboratory Animals, National Institutes of Health, USA.

Vitrification of Neurospheres

The protocol for vitrification of NSPCs is outlined in Figure 1. All cryoprotectant solutions were prepared in DMEM/F12. After three passages, neurospheres were either taken through vitrification and warming or kept as untreated controls. All experiments were performed at least in triplicate on batches of neurospheres isolated from tissue derived from different pregnant females. The procedure comprised a three-step exposure to cryoprotectants, where neurospheres were sequentially exposed to solutions containing increasing concentrations of cryoprotectans as detailed in Figure 1. In each of the first two steps, the neurospheres were allowed to sink to the bottom of the microcentrifuge tube before the removal of the cryoprotectant solution and its subsequent substitution with cryoprotectant of increased concentration. While exposed to the final vitrification solution (40% v/v EG, 0.6 M sucrose, resulting in 65% w/v total solute concentration), neurospheres were packed in a “straw-in-straw” configuration as previously described (18). Briefly, neurospheres were aspirated into a 250-μl sterile, plastic straw. This 250-μl straw was then placed within a 500-μl straw. The 500-μl straw was then sealed with an impulse heat sealer. The entire procedure was performed at room temperature (23 ± 2°C), with a total duration of 9–10.5 min. Finally, the straws were immersed in liquid nitrogen. The durations of the various steps in the procedure are schematically presented in Figure 1.

Figure 1.

Schematic representation of the vitrification–warming and dilution procedures for the cryopreservation of NSPCs. Total solute concentrations refer to EG solutions, except for the solution indicated by an asterisk (^*), which consists of EG + sucrose (40% v/v EG and 0.6 M sucrose).

Warming of Neurospheres and Dilution of Cryoprotectant

The straws containing neurospheres were warmed by immersion in a water bath at 38 ± 0.5°C for 30 s with continuous shaking. Next the neurospheres were unpacked and expelled into 1 M sucrose in DMEM/F12 in a 15-ml centrifuge tube. The sucrose concentration was reduced stepwise first to 0.7 M by dilution with 0.25 M sucrose in DMEM/F12 and subsequently by 0.175 M on stepwise dilution with DMEM/F12. The durations of the warming and dilution steps are schematically presented in Figure 1. The whole dilution procedure took ~15 min. All treatments were performed at room temperature (23 ± 2°C). The neurospheres were then allowed to sink. The excess solution was then removed and the neurospheres were washed in neurosphere culture medium before being placed in the incubator. After 30 min they were transferred to fresh culture media for routine culture until further analysis.

Assay for Neural Stem Cell Markers

To determine whether the vitrification affected the neural stem or progenitor cell state, dissociated NSPCs were plated on poly-L-ornithine/fibronectin-coated slides in a monolayer and maintained for 3 days in neurosphere medium. The progenitor or stem cells were then fixed by treatment with 4% paraformaldehyde for 20 min. Immunostaining was conducted sequentially using anti-Sox2 (AB5603, Chemicon, Tenecula, CA, USA) and anti-nestin (MAB353, Chemicon) antibodies for identification of neural stem or progenitor cell markers. Fluorescence-conjugated Alexa Fluor secondary antibodies (Invitrogen, CA, USA) were used to visualize the primary antibodies and the coverslips were counterstained with DAPI in Prolong Antifade Gold mounting medium (Invitrogen). The immunostained cells were then imaged by sequential scanning with a confocal microscope (LSM 510, Carl Zeiss Microimaging GmbH, Germany).

Cell Proliferation Assay

In order to assay the rate of cell proliferation of the NSPCs, the control neurospheres were washed thoroughly using culture medium, prior to dissociation and staining, mimicking the dilution process of the vitrified group for procedure consistency. The neurospheres were dissociated using 0.25% trypsin-EDTA solution and the cells were replated on poly-L-ornithine/laminin-coated coverslips. The cells were allowed to adhere to the coverslips and 5-bromo-2′-deoxyuridine (BrdU; Sigma-Aldrich, MO, USA) was then added to the culture medium to a final concentration of 10 μM and incubated with the dissociated NSPCs for 24 h. The cells were then fixed with 4% paraformaldehyde for 20 min. Immunostaining was conducted to detect the BrdU incorporation using an anti-BrdU antibody (1:100, BRD.3, Neomarkers, Lab Vision Corporation, CA, USA) and detected using Alexa Fluor 555 conjugate secondary antibody (Invitrogen). The cells were counterstained with DAPI to reveal all nuclei. The cells were imaged by sequential scanning with a confocal microcope (Fluoview 1000, Olympus, Japan). An investigator blind to the treatment conditions counted the number of BrdU-immunoreactive cells and DAPI-positive nuclei and the number of BrdU-positive cells was expressed as a percentage of the total cell count.

Assay for Multipotent Differentiation

To assay for multipotent differentiation into neurons, astrocytes, and oligodendrocytes, dissociated NSPCs were plated on poly-L-ornithine/laminin-coated coverslips. Differentiation was induced with 0.5% fetal calf serum (Hyclone Laboratories Inc., UT, USA) in neurosphere medium without EGF and bFGF. After 1 day, the medium was replaced with DMEM/F12 with N2 supplement and the cells were allowed to differentiate for a further 6 days. The differentiated cells were then fixed by treatment with 4% paraformaldehyde for 20 min. Immunostaining was conducted sequentially using an anti-MAP2a+b antibody (MAB378, Chemicon) for identification of neurons, an anti-glial fibrillary acidic protein (GFAP) antibody (Z0334, Dako, Denmark) for identification of astrocytes and an anti-CNPase antibody (MAB326R, Chemicon) for identification of oligodendrocytes. Fluorescence-conjugated Alexa Fluor secondary antibodies (Invitrogen) were used to visualize the primary antibodies and the coverslips were counterstained with DAPI. The differentiated cells were then imaged by sequential scanning with a confocal microscope (Fluoview 1000, Olympus). The numbers of neurones, oligodendrocytes and astrocytes were counted by an investigator blind to the treatment and expressed as a percentage of the total cell count.

Statistical Analysis

Values obtained for vitrified samples were compared to those of the controls by Student's t-test (SPSS version 12, SPSS Inc., Chicago, IL, USA). A value of p < 0.05 was considered statistically significant. All data are reported as mean ± SEM.

Results

Effects of Vitrification on Expression of Stem or Progenitor Cell Markers

To investigate whether the process of vitrification had an effect on the stem or progenitor cell state of NSPCs, expression of the neural stem or progenitor cell markers, nestin and Sox2, was assayed. The cells were plated on poly-L-ornithine and fibronectin-coated coverslips and allow to attached and recover for 3 days. The vitrified NSPCs were found to express nestin and Sox2 (Fig. 2). To investigate whether the NSPCs would lose their stem/progenitor cell state upon further passaging, the neurosphere cultures were passaged three times and the assay was conducted again. After three passages, the NSPCs were still found to express nestin and Sox2. This indicates that the neural stem or progenitor cell state was maintained for at least three passages after vitrification (Fig. 2B).

Figure 2.

Vitrified NSPCs maintain expression of progenitor or stem cell markers. Vitrified NSPCs were plated as a monolayer and immunostained using anti-nestin (green) and anti-Sox2 (red) antibodies. NSPCs cultured (A) after vitrification and (B) three passages after vitrification. All cells were double-stained for nestin and Sox2. Representative examples of double-stained cells at higher magnification are inset. Scale bar: 50 μm.

Effect of Vitrification on the Rate of Proliferation

The proliferation rate of the vitrified NSPCs was compared with the untreated group by a BrdU incorporation assay. The proliferation rate over 24 h was calculated as the percentage of BrdU-immunoreactive cells in the total DAPI-stained cell count. Vitrification did not significantly alter the number of cells labeled with BrdU compared to the untreated control (Fig. 3B compared with Fig. 3A). Quantification of the BrdU-positive cells did not reveal any significant change in the rate of proliferation in vitrified NSPCs compared to the untreated control (Fig. 3C).

Figure 3.

BrdU cell proliferation assay of NSPCs after vitrification. NSPCs dissociated from neurospheres (A) untreated, and (B) after recovery from a complete vitrification–warming cycle. The dissociated cells were plated on poly-L-ornithine and laminin-coated coverslips and assayed for proliferation by BrdU incorporation assay. The cells with BrdU incorporation were identified using an anti-BrdU antibody (red) and the coverslips were counterstained with DAPI (blue). Representative examples of BrdU-positive cells at higher magnification are inset. Scale bars: 80 μm. (C) The number of cells undergoing division within 24 h was counted and expressed as a percentage of the total DAPI-positive cell count. The data are mean ± SEM of five replicates. There were no significant differences.

Multipotent Differentiation After Vitrification

The neurons, astrocytes, and oligodendrocytes were identified by immunostaining with cell type-specific markers (Fig. 4). Both untreated and vitrified NSPCs differentiated into cells expressing either GFAP or MAP2a+b (Fig. 4A and B, respectively) and GFAP or CNPase (Fig. 4C and D, respectively). The number of each of the three cell types was quantified as a percentage of the total cell number counted by DAPI counter-staining. Both untreated and vitrified NSPCs were able to differentiate into 7–11% neurons, 80–86% astrocytes, and <1% oligodendrocytes (Fig. 4E). There were no significant differences in the differentiation in untreated and vitrified cells.

Figure 4.

Multipotent differentiation of neurospheres. (A) and (C) untreated control; (B) and (D) NSPCs after recovery from a complete vitrification-warming cycle. The neurospheres were differentiated with 0.5% fetal calf serum and double immunostained for (A) and (B) astrocytes and neurons using anti-GFAP (green) and anti-MAP2a+b (red) antibodies, respectively, and (C) and (D) astrocytes and oligodendrocytes using anti-GFAP (green) and anti-CNPase (red) antibodies, respectively. Nuclei were counterstained with DAPI (blue). Scale bars: 20 μm. (E) Percentage of the neurons, astrocytes and oligodendrocytes differentiated from untreated and vitrified NSPCs expressed as a percentage of the total DAPI-positive cell number. The data are mean ± SEM of three replicates. There were no significant differences between the untreated and vitrified groups.

Discussion

The present study aimed to investigate whether cryopreservation of stem cells without the use of proteins and serum was feasible. Specifically, we have established a vitrification protocol without any protein or serum additives for the sterile cryopreservation of functional NSPCs. It is essential that NSPCs maintain their multipotency after cryopreservation for the benefit of basic research and for future clinical application. Therefore, after the vitrification–warming cycle, neurospheres were assayed for their ability to differentiate into neurons, astrocytes, and oligodendrocytes. All three neural cell types were found in differentiated NSPCs after vitrification–warming cycle. There was no significant difference in the proportion of the three cell types between the untreated and vitrified NSPCs. Thus, these results indicate that the vitrification process does not affect the multipotent differentiation of NSPCs

Expression of nestin and Sox2, two commonly used neural stem or progenitor cell markers, was studied to ensure that the state of the NSPCs was not affected by vitrification. Both the markers were detected in NSPCs recovered from vitrification and also three passages after recovery from vitrification. This shows that the vitrification procedures did not alter the expression of these important stem cell markers. In our study, there was no evidence for stimulation of spontaneous differentiation of NSPCs following recovery from vitrification, which had been mentioned as a problem following cryopreservation of embryonic stem cells in a protocol involving the use of dimethylsulfoxide (DMSO) (12). This problem could be attributed to the role of DMSO in promoting differentiation, mechanical and osmotic stresses, and other chemical and physical factors (2, 9). Maintenance of progenitor or stem cell properties following vitrification is likely to be important in clinical application as expansion of the NSPCs will probably be required to reach the desired cell number for cell replacement therapy.

Careful selection of the constituents of the cryopreservation solutions is important to avoid toxic effects. To ensure low toxicity, we carefully assessed a number of vitrification solutions and conditions for vitrification (22). The vitrification solution evaluated here used EG and sucrose as cryoprotectants. EG is regarded as a promising cryoprotectant due to its high permeability and relatively low molecular weight (62 Da) (35). Sucrose is an easily soluble, cheap cryoprotectant additive with little or no toxicity and, in contrast to EG, is a nonpenetrating cryoprotectant. Our previous experience in cryopreservation by vitrification had shown that the use of mixtures of penetrating and nonpenetrating cryoprotectants in certain proportions is a feasible approach for cryopreservation of human oocytes (17), encapsulated hepatocytes (19, 38), and self-assembled aggregates (22). Therefore, we employed this methodology in vitrifying NSPCs. In our protocol, the cells were only exposed briefly to the penetrating cryoprotectant; after that the EG was removed completely through the dilution process. Hence, we believe that the use of EG as one of the cryoprotectants in our vitrification protocol did not cause toxic effects in the model we used. This cryopreservation protocol was cautiously designed so that no penetrating cryoprotectant will remain in cells or will be transferred into patient or culture medium with cells. The vitrification solution evaluated in the current study was invented by us and lead to the first live human birth following vitrification of human oocytes (15). In this regards, it can be consider likely to be safe for the prospect of cryopreservation of human stem cells for clinical applications. We had found that vitrification, even without any components of human or animal origin, allows preservation of neurospheres with high cell viability. In contrast, freezing, even incorporating fetal calf serum, led to significant lost of viability (32). A comprehensive study on the long-term impact of “freezing” on NSPCs also showed a decline in cell viability (26).

Recent studies that moved away from the slow-cooling “freezing” concept for the cryopreservation of embryonic stem cells showed no evidence of altered pluripotency or karyotype abnormalities (28, 29, 31, 39). We have also conducted studies to investigate the effects of vitrification on the chromosomal status of human oocytes (15) and mammalian neurospheres (32) and did not find evidence of chromosomal abnormality following vitrification.

Currently, NSPCs are typically cryopreserved by freezing. Freezing is usually achieved by either rapid freezing or slow-cooling protocols. Protocols that involve freezing, which inevitably results in the formation of ice crystals, may damage sensitive cells. Moreover, slow cooling has also been shown to have persistent adverse effects on embryonic stem cells (14). DMSO with or without fetal calf serum has been commonly used as a cryoprotectant (3, 6, 26). Although not evident in a recent study by Milosevic and colleagues (26), the use of DMSO as a cryoprotectant can have adverse effects on differentiation (10). The inclusion of fetal calf serum in some freezing protocols is also undesirable, as it may induce differentiation of the NSPCs. If cryopreservation protocols are to serve as the foundation for the development of protocols for clinical cell transplantation, it is also important that they should be free from foreign proteins and sera so as to avoid any possible risk of contamination (23). Xeno-free cryopreservation protocols have sometimes replaced animal proteins with human proteins (29), but ideally even human proteins would be eliminated to prevent all possibility of contamination. Our vitrification protocol does not require the use of any protein. A broader implication of our current result is that we have shown that cell cryopreservation without serum and proteins can be effective when vitrification is employed. Therefore, we propose vitrification as a preferred method for cryopreservation of NSPCs.

A major drawback of existing cryopreservation protocols has been that it is difficult to ensure sterility. Not only can pathogens survive cryopreservation but, as experience in the field of assisted reproduction has shown, cross-contamination can even occur during cryostorage (25, 30, 34). We have invented a “straw-in-straw” method using readily available 250- and 500-μl straws, which we arranged into a “straw-in-straw” configuration (18), as a simple and cost-effective strategy to prevent contamination. This approach means that samples can be cooled and warmed by direct immersion in liquid nitrogen and a water bath, respectively. Previously, we have pioneered the “straw-in-straw” technique in the cryopreservation of embryos (15, 16, 18), cell–matrix systems (19, 38), and hepatocyte spheroids (22). Together with our previous work, the present data suggest that this strategy can be incorporated in an effective cryopreservation protocol for NSPCs.

Here we have discussed a cost-efficient and simple protocol for the cryopreservation of neurospheres by vitrification. While it is uncertain whether neurospheres will be the preferred culture system for potential future clinical application of NSPCs (11), the ability to preserve the unsupported three-dimensional structure of neurospheres by vitrification suggests that this approach could be considered as a method to the preservation of NSPCs in three-dimensional microfabricated structures or scaffolds (e.g., nerve conduits for spinal cord repair) (33). Importantly, we have shown in the present report that this protocol did not affect the proliferation or differentiation of NSPCs. With the complete avoidance of products of human or animal origin, we hope that this protocol can serve as starting point for development of protein- and serum-free vitrification protocols for the cryopreservation of human stem cells, especially human neural stem cells that may eventually be used in clinical settings.

Footnotes

Acknowledgments

The authors gratefully acknowledge the support of Biomedical Research Council, Singapore to Dr. Lilia L. Kuleshova (BMRC Grant No.: 04/1/21/19/317) and the National University of Singapore Young Investigator Award to Dr. Gavin S. Dawe. We thank Ms. Siew Ping Han for her excellent technical and administrative support.

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Effective Cryopreservation of Neural Stem or Progenitor Cells without Serum or Proteins by Vitrification

Abstract

Keywords

Introduction

Materials and Methods

Culture of NSPCs

Vitrification of Neurospheres

Warming of Neurospheres and Dilution of Cryoprotectant

Assay for Neural Stem Cell Markers

Cell Proliferation Assay

Assay for Multipotent Differentiation

Statistical Analysis

Results

Effects of Vitrification on Expression of Stem or Progenitor Cell Markers

Effect of Vitrification on the Rate of Proliferation

Multipotent Differentiation After Vitrification

Discussion

Footnotes

Acknowledgments

References