PLURIPOTENT STEM CELLS

Field of the invention

The present invention relates to methods of cryopreserving pluripotent stem cells which result in improved cell viability, survival and maintenance of the stem cell phenotype upon recovery.

Background of the invention

Pluripotent stem cells are a potential source of cells for cell therapy and regenerative medicine. However, working with pluripotent stem cells is complicated because the cells are particularly sensitive to their environment. Thus, for example, culture of pluripotent stem cells typically requires complex culture conditions, often involving the use of feeder cells to supply as yet incompletely identified support factors. It may also require the addition of supplements derived from animal serum, which thus renders the cells unsuitable for administration to human patients.

Similarly, storage of pluripotent stem cells by standard cryopreservation techniques is characterised by low recovery and a slow growth rate, as well as a high incidence of short-term spontaneous differentiation such that the original stem cell phenotype is lost. These problems are particularly apparent when working with pluripotent stem cells, particularly human pluripotent stem cells such as human embryonic stem cells (hESCs) or human induced pluirpotent stem cells (hiPSCs).

Standard suspension cryopreservation of hESCs or hiSCs using cryoprotective agents (CPAs) and slow-cooling has yielded particularly low recoveries. Stem cell markers such as Oct4 have expression levels as low as 10% of the recovered cell population indicating that the stem cell phenotype is poorly maintained. Post-thaw growth rates can be so slow and unpredictable that recovery of original stocks takes weeks to months. Some additional problems with suspension cryopreservation include the sharp, sudden exposure to high CPA concentrations (which are often toxic) and the need to use expensive apoptosis inhibitors, such as inhibitors of the Rho- associated protein kinases (ROCKs).

Thus, there is a need for methods of cryopreserving pluripotent stem cells which do not suffer from the problems associated with existing methods. Summary of the invention

The present inventors have surprisingly demonstrated that by encapsulating pluripotent stem cells in a three-dimensional extracellular matrix, the problems with existing methods of cryopreserving pluripotent stem cells can be overcome. In particular, the inventors have shown that encapsulation in accordance with the method of the invention increases post-thaw survival rate and inhibits spontaneous differentiation.

Thus, the present invention provides a method of cryopreserving a pluripotent stem cell comprising:

- encapsulating said cell in a three-dimensional extracellular matrix;

freezing said cell; and

thereby cryopreserving said cell.

A cryoprotective agent may be added before freezing the cell. The pluripotent stem cell is preferably a human pluripotent stem cell.

The present invention also provides a method of recovering a pluripotent stem cell cryopreserved according to the method of the invention, wherein the method comprises thawing the cell cryopreserved according to the method of the invention and thereby recovering said cell. Preferably, the recovery method does not comprise the addition of an inhibitor of apoptosis. For example, the recovery method does not comprise the addition of an inhibitor of a Rho-associated protein kinase (ROCK).

The recovery method of the invention may further comprise suspending said cell in a culture medium and/or removing the extracellular matrix. The culture medium in which the cell is suspended preferably does not comprise an inhibitor of apoptosis. Most preferably, the culture medium does not comprise an inhibitor of a ROCK.

Description of the Figures

Figure 1 : Comparison of Transmission FTIR spectra for unmodified alginate, RGDS-alginate and mixture of alginate and RGDS.

Figure 2: (Images) Analysis of cytoskeletal remodeling by actin-staining of mouse ES cells. Cell nucleus was counter-stained with DAPI. (A) Cell seeded on a gelatin surface; (B) cell seeded on RGDS-modified alginate surface; (C) cell encapsulated in RGDS-modified alginate and (D) cell encapsulated in unmodified alginate. Figure 3 : A comparison of post-thaw cell survival rate using trypan blue assay for suspension and 3D cryopreservation (unmodified and modified alginates) based on three independent trials (n= 3). All 3D experiments were performed with a 1.2 wt % alginate concentration, 1.2 mm mean bead diameter, l°C/min cooling rate, 10 % vol. DMSO concentration, a 30-minute loading time and at a le6 cells/mL seeding density. Suspension cryopreservation was performed with the same parameters but without encapsulation in alginate. Asterisk indicates that 3D cryopreservation with RGDS-alginate is better than using unmodified alginate or in suspension (* p < 0.05).

Figure 4: Cell health tracked for 6 hours after thawing. Cells were

cryopreserved in RGDS-alginate, unmodified alginate or in suspension recovered and cultured on 2D gelatinized surfaces. Error bars represent the standard errors (n=3). All 3D experiments were performed with a 1.2 wt % alginate, 1.2 mm mean bead diameter, l°C/min cooling rate, 10 % vol. DMSO concentration, a 30-minute loading time and at a le6 cells/mL seeding density. Suspension cryopreservation was performed with the same parameters but without encapsulation in alginate. Asterisk indicates that 3D cryopreservation with RGDS-alginate is better than using unmodified alginate or in suspension (*, p < 0.05) using a single factor ANOVA and a Tukey-Kramer post-hoc test.

Figure 5: Analysis of cells by flow cytometry of octamer-4 (Oct-4), Stage Specific Antigen 1 (SSEA-1) and Alkaline Phosphatase (ALP) for cells cryopreserved in suspension (filled, grey), in unmodified alginate beads (filled, black) and RGDS- alginate (grey line, unfilled) 24 hours after thawing (n=3). Asterisk indicates that 3D cryopreservation with RGDS-alginate is better than cryopreservation in suspension (* p < 0.05) while hash indicates that 3D cryopreservation with RGDS-alginate is better than cryopreservation in unmodified alginate.

Figure 6: mouse ES cells were differentiated into three different lineages after cryopreservation in 3D RGDS-alginates. These are (A) endoderm-derived endomucin- positive cells (B) adipocytes stained with Oil Red O and (C) neurons stained with anti-beta-III-tubulin antibody. Cells are representative of the endodermal, mesodermal and ectodermal lineages respectively.

Figure 7: Influence of RGDS ligand spacing on cell survival rate. Asterisk indicates that 12.5 mg RGDS/g alginate was significantly better than all lower RGDS concentrations when tested in a homogeneous and in a heterogeneous concentration (* p < 0.05). Figure 8: Cell survival rate against CPA loading. Asterisk indicates that cell survival rate for cells encapsulated in RGDS-alginate, after a 30 minute loading time is significantly higher than all other loading times (* p < 0.05).

Figure 9: Influence of cooling rate on survival in 3D and in suspension cryopreservation. In all three loading times, and for both suspension and 3D cryopreservation, a l°C/min cooling rate gave the highest cell survival rate but was only statistically significant for the 20 and 30 minute loading times (* p < 0.05).

Figure 10: Comparison of cell survival rate for 3D-, Su+ and Su- cell populations (via a trypan blue assay and assessed within 4 minutes of staining) for hESCs cryopreserved in RGDS-alginate spheres without RI supplementation (3D-), in suspension ("Su") with RI supplementation (Su+) and without RI supplementation (Su-). Asterisk indicates that cell survival rate for 3D- cells is significantly higher than for Su+ and Su- cells (* p < 0.05) using a paired t-test. Error bars represent the standard errors of at least three independent trials. {Experimental conditions: mean bead diameter 1.2 mm in 1.2 wt% RGDS-alginate beads, 10 % vol. DMSO, l°C/min cooling rate, le6 cells/mL, 30 minute loading time, 0.01 mM RI supplementation (Su+ cells) and 4°C loading temperature}

Figure 11 : Influence of freezing rate on cell survival rates (determined via a trypan blue assay and assessed within 4 minutes of staining) for 3D- cells. Asterisk indicates that cell survival rate for cells cooled at l°C/min is significantly higher than faster cooling rates (* p < 0.05) based on a single factor ANOVA and a Tukey- Kramer post-hoc analysis. Error bars represent the standard errors of at least three independent trials. {Experimental conditions: mean bead diameter 1.2 mm in 1.2 wt% RGDS-alginate beads, 10 % vol. DMSO, l°C/min cooling rate, le6 cells/mL, 4°C loading temperature and -80°C storage temperature}

Figure 12: Cell growth after cryopreservation compared between cells cryopreserved in 3D/RGDS-alginate beads without RI (3D-), cells cryopreserved in suspension with 0.01 mM RI supplementation post-thaw (SU+), following suspension cryopreservation without RI supplementation (SU-), after trpsinization from culture (CLTR). {Experimental conditions: 1.2 wt % alginate (3D-), mean bead diameter 1.2 mm (3D-), 10 % vol. DMSO, l°C/min cooling rate, le6 cells/mL, 4°C loading temperature and -80°C storage temperature }

Figure 13: Alamar Blue reduction rate for cells cryopreserved in 3D or in suspension (SU) with (+) or without (-) RI supplementation after thawing, after trpsinization from culture (CLTR); Assuming a linear first-order reduction rate, 3D- cells have a higher gradient (0.035 h ^"1 ) than the Su+ cells (0.02 h ^_1 ). Error bars represent the standard errors of at least three independent trials. {Experimental conditions: mean bead diameter 1.2 mm in 1.2 wt% RGDS-alginate beads, 10 % vol. DMSO, l°C/min cooling rate, le6 cells/mL, 4°C loading temperature, 0.01 mM ROCK inhibitor supplementations (Su+) and -80°C storage temperature}

Figure 14: Analysis of apoptosis rates in cells cryopreserved in 3D/RGDS- alginate beads without RI (3D-), cells cryopreserved in suspension with RI supplementation post-thaw (SU+), following suspension cryopreservation without RI supplementation (SU-), after trpsinization from culture (CLTR). {Experimental conditions: 1.2 wt % alginate (3D-), mean bead diameter 1.2 mm, 10 % vol. DMSO, - l°C/min cooling rate, le6 cells/mL, 0.01 mM ROCK inhibitor supplementations (Su+) and -80°C storage temperature}

Figure 15: Measurement of DNA damage via the comet assay:

cryopreservation in RGDS-alginates with no RI supplementation (3D-), following suspension cryopreservation with RI supplementation (Su+), following suspension cryopreservation without RI supplementation (Su-), after trpsinization from culture (Culture) and after shortwave UV treatment for 1.5 hrs from a low pressure mercury vapor discharge tube (UV). {Experimental conditions: 1.2 wt % alginate (3D-), mean bead diameter 1.2 mm, 10 % vol. DMSO, -l°C/min cooling rate, le6 cells/mL, 0.01 mM ROCK inhibitor supplementations (Su+ and culture) and -80°C storage temperature }

Figure 16: Stem cell characteristics from cryopreserved and cultured (cltr) cells as measured by flow cytometry for Oct-4 , Nanog, ALP, SSEA-4. 3D- cells were cryopreserved in RGDS-alginate beads at passage 30, thawed, allowed to recover then assayed at a minimum of 10,000 events at le6 cells/ml. Su+ and Cltr cells were supplemented with RI while 3D- and Su- cells were not. The loading time was fixed at 30 minutes & cell densities at le6 cells/ml. Asterisk indicates that the expression rate for cells cryopreserved in 3D alginate-RGDS beads is significantly higher than for cells cryopreserved in suspension with RI supplementation post-thaw (* p < 0.05) based on a paired t-test. Error bars represent the standard errors of at least three independent trials. {Experimental conditions: loading temperature: 4°C, l°C/min cooling rate, le6 cells/mL} Figure 17: Expression levels for cell markers for endoderm-derived (Sox 17+), mesoderm-derived (FABP4+) and ectoderm-derived (β-ΙΠ Tubulin+) cells from RGDS -Alginates without RI supplementation (3D-) , in suspension with RI supplementation (Su+), in suspension without RI supplementation (Su-) and in culture after RI supplementation (Cltr). Cells were cryopreserved in RGDS-alginate beads or in suspension at passage 31, thawed, allowed to recover then assayed at a minimum of 10,000 events at le6 cells/ml. The loading time was fixed at 30 minutes & cell densities at le6 cells/ml. Asterisk indicates that expression rate according to the adipogenic lineage for cells cryopreserved in 3D alginate-RGDS beads without RI supplementation is significantly higher than for cells cryopreserved in suspension with RI supplementation post-thaw (* p < 0.05) based on a paired t-test. Error bars represent the standard errors of at least three independent trials. {Experimental conditions: loading temperature: 4°C, l°C/min cooling rate, le6 cells/mL} Detailed description of the Invention

It is to be understood that different applications of the disclosed products and methods may be tailored to the specific needs in the art. It is also to be understood that the terminology used herein is for the purpose of describing particular embodiments of the invention only, and is not intended to be limiting.

In addition as used in this specification and the appended claims, the singular forms "a", "an", and "the" include plural referents unless the content clearly dictates otherwise. Thus, for example, reference to "a cell" includes "cells", reference to "a ligand" includes two or more such ligands, reference to "a protein" includes two or more such proteins, and the like.

All publications, patents and patent applications cited herein, whether supra or infra, are hereby incorporated by reference in their entirety.

Extracellular matrix, ligand and cell surface protein

The present inventors have shown that encapsulation in a three-dimensional extracellular matrix improves cell viability, survival and maintenance of the stem cell phenotype upon recovery from cryopreservation. Without wishing to be bound by any particular hypothesis, the inventors believe that cell attachment to the matrix acts as a transitory anti-apoptotic signal for the cells. This signal also stabilises the stem cell phenotype. Accordingly, the surface of the encapsulated stem cell is preferably attached to the extracellular matrix

For use in the encapsulation of stem cells, the three-dimensional extracellular matrix must comprise a substance which is biocompatible, mechanically stable, chemically stable, and which allows for the production of microcapsules of an appropriate size. The substance should typically allow for attachment of the stem cell surface to the substance, following encapsulation. The substance is typically a polymer. Suitable polymers include an alginate, a collagen, a gelatine, a chitosan, a carrageenan, a cellulose (e.g. carboxymethyl cellulose) and an agarose. Thus, the matrix of the invention may comprise at least one of these polymers. The matrix preferably comprises an alginate. Alginate is a block co-polymer of guluronic and manuronic acids. It is an attractive biomaterial because of its low immunogenicity, easy sol-gel transition, abundance, low-cost and amenable chemistry. The amenability of alginate allows the inclusion of peptide repeats to facilitate cell attachment.

The extracellular matrix may additionally comprise at least one ligand which is capable of binding to a receptor molecule on the surface of the stem cell, thereby attaching the surface of the stem cell to the extracellular matrix. The extracellular matrix may be chemically linked to the ligand. For example, the matrix may be covalently coupled to the ligand.

The binding of the ligand to the receptor molecule on the surface of the stem cell is preferably of high affinity. By high affinity binding it is meant that the ligand binds to the receptor with a Kd (dissociation constant) of less than ΙΟμΜ, 9μΜ, 8μΜ, 7μΜ, 6μΜ, 5μΜ, 4μΜ, 3μΜ, 2μΜ, or ΙμΜ. Methods for assessing binding affinity are known to the person skilled in the art, for example surface plasmon resonance assays, competition binding assays and the like.

The binding of the ligand to the receptor molecule on the surface of the stem cell is preferably specific. By specific binding it is meant that the ligand binds to the receptor molecule with no significant cross-reactivity to any other unrelated stem cell surface receptor molecule. For example, a ligand for an integrin will show no significant cross-reactivity with any non-integrin receptor present on the surface of a stem cell. The ligand may show no significant cross-reactivity to any other receptor molecule present on the surface of a stem cell. Cross-reactivity may be assessed by any suitable method. Suitable methods are known to the person skilled in the art, for example competition binding assays and the like. The ligand may comprise or consist of any suitable molecule which is capable of specific binding to a receptor molecule on the surface of a stem cell. The receptor molecule may be any molecule present on the surface of the stem cell. The receptor molecule may typically be a carbohydrate or a protein. The receptor molecule is preferably a protein. The protein is preferably a cell adhesion molecule (CAM). CAMs are typically classified as (1) integrins, (2)immunoglobulins, (3) cadherins or (4) selectins. Most preferably, the protein may be an integrin and the ligand may be a molecule capable of specific binding to said integrin. Preferably, the integrin is an alpha-v integrin. Particularly preferred integrins include alpha-v beta-3 and alpha-v beta-5 integrin.

The ligand may be a peptide comprising or consisting of the sequence arginine-glycine-aspartic acid (RGD) or arginine-glycine-aspartic acid-serine

(RGDS). Such peptides may be linear or cyclic. Examples include cyclo(Arg-Gly- Asp-D-Phe-[Nme]Val, commonly referred to as cilengitide or EMD-121974.

Alternatively, a non-peptide mimetic for the sequence RGD or RGDS may be used as a ligand. Suitable mimetics include the non-peptide RGD mimetic S36578-2 (sodium salt of 5-(s)-7-{[4-pyridin-2-ylamino-butyrl amino]-methyl-6.9-dihydro-5H- benzocyclohepten-5-yl} acetic acid).

The ligand may comprise a galactose moiety. The ligand may be a fibronectin, a vitronectin, an osteopontin, a collagen, a thrombospondin, a fibrinogen, a laminin or von Willebrand factor, or a part of any thereof capable of specific binding to the receptor molecule present on the cell surface.

In a preferred embodiment of the invention, the ligand is RGDS. The RGDS sequence is an important recognition site for cell adhesion. Many cell-adhesive proteins present in the extracellular matrix (ECM) and in blood contain RGDS. These proteins include fibronectins, vitronectins, osteopontins, collagens, thrombospondins, fibrinogens, laminins, and von Willebrand factor. RGDS domains on these ECM molecules bind to transmembrane domains such as alpha- V beta-3 integrins which play significant roles e.g. cell survival during embryonic development. On a macro- scale, integrin-RGDS interactions also affect tissue permeability. Such interactions, when applied to, e.g. cryoprotective agent (CPA) loading may help improve cell response to changes in CPA concentration.

In a particularly preferred embodiment of the invention, the extracellular matrix comprises an alginate covalently coupled to a peptide consisting of the sequence RGDS. In another preferred embodiment, the extracellular matrix comprises a chitosan, preferably a galactosylated chitosan.

Encapsulation of cells in the extracellular matrix may be achieved by any suitable method. Such methods are well known to a person skilled in the art. See for example, Nigam et al; Biotechnology Techniques Vol 2 No 4 271-276 (1988) and Nir Kampf; Polymers for Advanced Technologies, 13, 896-905 (2002). Examples include: dropping methods, wherein a suspension of cells in a polymeric solution is added drop-wise into an appropriate cross-linking solution or passed through an appropriate cross-linking aerosol; method using moulds, wherein a suspension of cells in a polymeric solution is injected into a mould and allowed to set; and coacervation methods, wherein the cell suspension and the polymeric solution are kept separate, and are prepared in such a way as to form a droplet with two separate phases, which is then added to an appropriate cross-linking solution to form a liquid cell suspension surrounded by a thin film. By way of illustration, suitable methods are described in more detail below, using a matrix comprising alginate or chitosan as a representative example.

Encapsulation of cells in a matrix comprising alginate may typically be achieved by a dropping method. This comprises resuspending the cells in a first solution of a monovalent cation salt of alginate, e.g. sodium alginate, at an appropriate concentration, typically around 1.2 wt%. Droplets of the alginate solution containing the cells are then typically generated in air. Droplet generation may be carried out using any droplet generation device. For example, droplets may be generated by gravitational flow from a tube or needle into air, where the effects of surface tension result in the formation of spherical droplets of the suspension. Alternatively, electrostatic droplet generators may be used whereby an electrostatic differential is created between an alginate solution and the collection solution, such that the alginate is drawn through a tube or needle in small droplets. Other devices are equally applicable, including the use of a spinning disc droplet generator or laminar air flow extrusion device. Once generated, the droplets are then collected in a separate solution comprising divalent metal cations, e.g., Ca ²⁺ , Sr ²⁺ or Ba ²⁺ . Generally, CaCl ₂ BaCl ₂ and SrCl ₂ solutions are preferred. The divalent metal cation solution may be supplemented with a buffer, such as a citrate buffer, and/or a culture medium such as Dulbecco's Modified Eagle Medium (DMEM). The droplets are gelled by the interactions between the alginate and the divalent cations in the collection solution, encapsulating the cells suspended within the gelled alginate droplets. The size of the resulting encapsulated compartments is determined by the size of the droplets.

Encapsulation of cells in a matrix of alginate/Poly (L-lysine) may typically use the same basic approach as described above. The exterior surface of alginate capsules prepared by the method described above is modified by treating with a poly (L- lysine) solution. The treated capsules are washed to remove uncomplexed poly (L- lysine).

Encapsulation in a matrix comprising a cross-linked polymer such as collagen may be achieved by following the same method as for alginate, but with the additional step of coating the gelled alginate droplets with the cross-linked polymer. The alginate droplet may then optionally be dissolved by, for example, chelating the divalent cation that was used in the initialling gelling step.

A similar technique may be applied to achieve encapsulation of cells in a matrix comprising agarose by using a mixture of alginate and agarose in place of the alginate solution described above.

Other encapsulation techniques include emulsification methods, where an emulsion of alginate (or other polymer) is added directly to an appropritate cross- linking solution leading to bead formation.

Chitosan forms an ionotropic gel similar to the one formed by alginate.

However chitosan is a cationic polymer and requires negatively charged muhivalent ions (eg. polyphosphate ions) to form the gel structure. Typically, therefore, the cells are prepared in a solution containing chitosan, which is then dropped into a second solution comprising the multivalent ions, typically a sodium-tri-poly-phosphate solution. A chitosan polyphosphate membrane is instantly formed enclosing the droplet. Capsules are removed from the chitosan solution and typically further treated in multivalent ion solution after washing away excess chitosan using acetic acid.

Once encapsulation is complete, the encapsulated cell is typically transferred to a suitable culture medium prior to carrying out the subsequent steps of the method. Any appropriate medium may be used. Examples include Dulbecco's Modified Eagle Medium (DMEM) and RPMI- 1640 medium. Other standard culture medium components may be added.

Freezing The present invention provides a method of cryopreserving a stem cell comprising:

encapsulating said cell in a three-dimensional extracellular matrix; freezing said cell; and

thereby cryopreserving said cell.

By "cryopreseving" it will be understood that the method of the invention involves storage of the stem cell at a temperature below its freezing point. Thus, freezing of the cell involves reducing the temperature of the cell such that it is no higher than 0°C, preferably no higher than - 4°C, -10°C, -15°C, -18°C, -20°C, -30°C, -40°C, -50°C, -60°C, -70°C, -75°C, -78°C, -80°C, or -100°C . The temperature of the cell will typically not be reduced to below the boiling point of liquid nitrogen, -196 °C.

Freezing in accordance with the method of the invention may be carried out using any suitable technique. Such techniques are well known to a person skilled in the art. Suitable techniques include slow freezing techniques in which vials or other containers of cells are frozen over a number of minutes or hours. Such techniques may reduce the temperature of a cell by 1°C per 5 seconds, per 20 seconds, per 30 seconds, per 40 seconds, per minute, per 2 minutes, per 3 minutes, per 5 minutes, per 10 minutes, or per 20 minutes. Cooling at 1°C per minute or slower is preferred.

Controlled cooling may be achieved for example by use of an electronically controlled freezing device in which the temperature is reduced slowly in a controlled way, or more crudely by placing the cells in a lidded freezing box (for example an isopropanol jacketed freezing box), which is placed into a -20°C or -80°C freezer so that the freezing temperatures of the freezer penetrate the freezing box slowly and it takes a number of minutes or hours (for example 30 minutes or 2 or 3 or 4 hours) for the cell to freeze.

Techniques involving fast freezing may also be used. Such techniques may employ liquid nitrogen or frozen carbon dioxide, and/or special cooling liquids and vitrification techniques.

The method of the invention may comprise the addition of a cryoprotective agent (CPA) before freezing the cell. CPAs are agents which protect against injury to the cell caused by freezing. Examples of CPAs include DMSO (dimethyl sulfoxide), ethylene glycol, glycerol, erythritol, dimethylformamide, 2-Methyl-2,4-pentanediol (MPD), propylene glycol, dextrans, polyethylene glycol (PEG), polyethylene oxide (PEO), sucrose, trehalose and glucose. The CPA may be biocompatible and / or nontoxic to cells. The CPA preferably does not induce differentiation of stem cells.

If a CPA is added, it is typically present before freezing the cell at a concentration of less than 2.5% by volume (%vol), less than 5%vol, less than 7.5 %vol, less than 10%vol or less than 15 %vol.

Recovery

The methods of the invention also comprise recovering a stem cell cryopreserved according to the invention, wherein the method comprises thawing the frozen cell obtained according to a method of the invention and thereby recovering said cell. The recovery method preferably does not comprise the addition of an inhibitor of apoptosis. Inhibitors of apoptosis, and particularly inhibitors of Rho- associated protein kinases (ROCKs), can be expensive and may be disadvantageous for and/or detrimental to the use of the cells after recovery.

Inhibitors of apoptosis are well known to the person skilled in the art and include, for example, caspase inhibitors and inhibitors of ROCKs. An example of a ROCK inhibitor is Y-27632. The recovery method of the invention preferably does not comprise the addition of a ROCK inhibitor.

Thawing of the cell may be carried out using any suitable method. Typically, the cell is rapidly warmed to 37°C, for example by immersion in a water bath, until fully thawed. Culture medium may be added. Typically this is done dropwise to slowly dilute any CPA which may be present. The culture medium preferably does not comprise an inhibitor of apoptosis. Most preferably, the culture medium does not comprise an inhibitor of a ROCK. The cell will then typically be allowed to rest at room temperature for approximately 10 minutes prior to any further treatment.

The recovery method of the invention may comprise suspending said cell in a culture medium. The culture medium in which the cell is suspended preferably does not comprise an inhibitor of apoptosis. Most preferably, the culture medium does not comprise an inhibitor of a ROCK.

The recovery method of the invention may comprise removing or not removing the extracellular matrix. Removal of the extracellular matrix may be achieved by any suitable method. Such methods are well known to the person skilled in the art. For example, where the extracellular matrix comprises alginate, removal may be achieved by the addition of a solution comprising sodium citrate to dissolve the alginate. The cells may then be centrifuged and resuspended in culture medium. The culture medium in which the cell is resuspended preferably does not comprise an inhibitor of apoptosis. Most preferably, the culture medium does not comprise an inhibitor of a ROCK.

The viability (cell survival rate) of the recovered cells is preferably >50%,

>60%, >70%, most preferably >80%. Viability is typically assessed within 1 hour after thawing, and may be assessed by any suitable method. For example, a trypan blue assay may be used.

The apoptosis rate of the recovered cells is preferably < 5%, <2.5% and most preferably < 1%. Apoptosis rate is typically assessed within 2 hours or 4 hours after thawing. Apoptosis may be assessed by any suitable method. Such methods are well known to a person skilled in the art. For example, methods involving staining with Annexin V and propidium iodide are well known.

DNA damage in the recovered cells may be assessed by the comet assay (single cell gel electrophoresis assay), and if so the recovered cells will typically display a mean extent tail moment of <25%, preferably <20%.

The recovered cells will typically display no statistically significant reduction in expression of stem cell surface markers relative to cells which have not undergone cryopreservation by the methods of the invention. Suitable surface markers for human embryonic stem cells include Oct-4, Nanog, ALP and SSEA-4. Similar markers may be used for mouse embryonic stem cells. Methods for assessing cell surface markers are well known to a person skilled in the art.

The recovered cells should show no statistically significant reduction in the ability to differentiate into other cell types, relative to cells which have not undergone cryopreservation by the methods of the invention. For example, the recovered cells should show no statistically significant reduction in the ability to differentiate into endoderm-derived cells such as liver cells, mesoderm-derived cells such as adipogenic cells or ectoderm-derived cells such as neurons. Differentiation may be triggered and assessed by any suitable method, for example as set out in the Examples. Methods for differentiating pluripotent stem cells are well known to a person skilled in the art.

Cell types

The pluripotent stem cells for use in accordance with the invention are typically animal cells, preferably mammalian cells. The cells may be from livestock animals (for example, cows, pigs or chickens), pets/companion animals (for example, cats, dogs, horses), sporting animals (for example racehorses), research animals (for example, mice, rats, rabbits, monkeys, apes), or primates (especially including humans). Most preferred are human or other primate cells, mouse, rat or bovine cells.

The pluripotent stem cells may be embryonic stem cells (ESCs). In particular they may be mouse or human embryonic stem cells (mESCs or hESCs). Human embryonic stem cells (hESCs) are pluripotent, self-renewing cells isolated directly from preimplantation human embryos that recapitulate organogenesis in vitro. As used herein, the term "human embryonic stem cells (hESCs)" is intended to include undifferentiated stem cells originally derived from the inner cell mass of developing blastocysts, and specifically pluripotent, undifferentiated human stem cells. hESCs useful in the practice of the methods of this invention are derived from

preimplantation blastocysts as described by Thomson et al., in U.S. Patent No.

6,200,806. Multiple hESC cell lines are also currently available in US and UK stem cell banks. Suitable hESC lines include, but are not limited to, SA01, SA02, ES01, ES02, ES03, ES04, ES05, ES06, BG01, BG02, BG03, HUES1, HUES2, HUES3, HUES4, HUES5, HUES6, HUES7, HUES8, HUES9, HUES10, HUES11, HUES 12, HUES 13, HUES14, HUES 15, HUES 16, HUES17,TE03, TE04, TE06, UCOl, UC06, WA07, WA09, WA13 and WA14, all of which are available from WISC Bank via WiCell Research Institute. Techniques for culturing ESCs are well known to a person skilled in the art.

The stem cells may be induced pluripotent stem cells (iPSCs). In particular they may be mouse or human induced pluripotent stem cells (miPSCs or hiPSCs).

Methods producing iPSCs are known in the art. A method for inducing pluripotency of differentiated cells, such as somatic cells, was first disclosed by Yamanaka (WO

2007/069666). In this method, somatic cells are reprogrammed using three main nuclear reprogramming factors, namely an Oct family gene, a Klf family gene and a Sox family gene (preferably Sox2). The factors are preferably Oct3/4, Klf4 and Sox2. A fourth reprogramming factor, namely the product of a Myc family gene (preferably c-Myc), may also be used. Numerous different methods have since been disclosed for inducing pluripotency in somatic cells. Such methods are reviewed in Hanna et al, Cell. 2010 Nov 12; 143(4):508-25; and Stadtfeld & Hochedlinger, Genes Dev. 2010 Oct 15;24(20):2239- 63. A preferred method is described in Carpenter, L. et al Blood In Press (2011). Human iPSCs typically display the characteristic morphology of human embryonic stem cells (hESCs), express the pluripotency-associated markers SSEA-4 and TRA1-60, the transcription factors Oct-4 and Nanog and differentiate in vitro into cell types derived from each of the three embryonic germ layers. The iPSCs may be an established cell line or may be produced from somatic cells taken from a subject.

Human iPSCs may be derived from any human somatic cell. Suitable cells include, but are not limited to, keratinocytes, dermal fibroblasts or leukocytes derived from peripheral blood. The iPSCs are preferably derived from dermal fibroblasts.

Techniques for culturing iPSCs are well known to a person skilled in the art.

Ethical and legal constraints on sources of cell

The Inventor and Applicant are fully aware of potential ethical and legal issues surrounding the use of stem cells and surrounding use of materials derived from embryos and foetuses. Those issues are particularly acute when the material is derived from humans. The Applicant reserves the right to disclaim any subject matter excluded from patentability by statute on moral grounds or in the interest of public policy (ordre public). In particular, the Applicant reserves the right to restrict any of the claims of this application (or parts of claims or products recited in the claims) to non-human products or to the use of non-human cells or cell-derived products, to disclaim the use of embryo derived embryonic stem (ES) cells (either primary or cell lines) and to disclaim commercial and/or industrial uses of embryos.

Kits

The present invention also provides a kit comprising the components of an extracellular matrix suitable for encapsulating a stem cell, as described above. The kit preferably comprises alginate. The kit optionally provides one or more culture media, and / or instructions for carrying out a method of the invention. Instructions may also recite features of methods disclosed in the examples of this patent application. Examples

The invention is exemplified by the non-limiting examples presented below: EXAMPLE 1

Use of alginate encapsulation for improved cryopreservation of mouse

embryonic stem cells The following experiments demonstrate that encapsulation in alginate improves mouse stem cell responses during CPA loading and post-thaw cell survival rate, whilst maintaining their sternness, even when using a simple constant-rate cooling protocol.

Materials and methods

Unless otherwise stated, all materials were obtained from Sigma- Aldrich, UK.

Mouse Embryonic Stem Cell (mESC) culture: The ESF 183 mESC cell line was derived from mouse strain 129 S2/Sv by the Gardener Lab (University of Oxford, UK) using techniques previously described (Brook FA, Gardner RL (1997) Proc Natl Acad Sci U S A 94, 5709-5712.). Mitomycin-treated primary mouse embryonic fibroblasts (feeder cells) were isolated from mouse foetus and initially cultured in Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% foetal calf serum (FCS), 1% (v/v) glutamine, 1% (v/v) beta-mercaptoethanol and 1% (v/v) penicillin/streptomycin. The mESCs were then seeded onto the feeder layer in mESC culture medium consisting of DMEM supplemented with 15% (v/v) FCS

supplemented with 1% (v/v) nonessential amino acids, 1% (v/v) β-mercaptoethanol (β-ΜΕ), 1% (v/v) penicillin/streptomycin and mouse leukaemia inhibitory factor (LIF) (1000 units/ml, Millipore, UK). The mESCs were incubated at 5% vol. C02 in air and 37 °C.

For feeder- free culture, mESCs were initially grown on feeders to 60%> confluence and then passaged. Cells were washed in Ca2+/Mg2+ free phosphate buffered saline (PBS) and dissociated using accutase at 37 °C for 5 minutes. Basal Medium (Millipore, UK) was added to the culture flasks and the cells were pipetted up and down to obtain a single cell suspension. The cells were centrifuged at 148 g for 5 minutes and resuspended in Clonal Grade Medium (CGM) (Millipore, UK) before seeding at 5E4 cells/cm ² on a 0.1 % gelatin coated surface. Mouse ESCs grown in feeder-free cultures were used for cryopreservation experiments.

Modification of alginate with RGDS and characterization: Alginate

modification was carried out using a "solution modification" method (Genes NG, Rowley JA, Mooney DJ, Bonassar LJ (2004) Arch Biochem Biophys 422, 161 - 167; Rowley JA, Madlambayan G, Mooney DJ (1999) Biomaterials 20, 45-53.)· Briefly, medium viscosity alginate (2000 cP at 2 wt. % polymer at 25 ⁰ C ) from brown algae was dissolved overnight in a buffer made of 50 mM 2-(N-morpholino) ethanesulfonic acid (MES) in 0.15M NaCl (pH 6.5) at a concentration of 1 g alginate/ lOOmL buffer. RGDS was covalently coupled to the alginate using carbodiimide chemistry with stabilization against hydrolysis using highly water soluble N-hydroxysulfosuccimide (sulfo-NHS). The sulfo-NHS to l-ethyl-(dimethylaminopropyl) carbodiimide (EDC) ratio was at 1 :2 while the peptide to EDC ratio was at 1 :4. The sulfo-NHS, EDC and peptide (in that order) were added in quick succession (after the dissolution of each reagent) to alginate while stirring at room temperature. The reaction was quenched with hydroxylamine hydrochloride (HA-HC1) after 20 hours at 18: 1000 (HA-HC1: alginate weight) ratio. The modified alginate was dialyzed against decreasing concentrations of NaCl solution from 15 g/L to Og/L Milli-Q water and then freeze- dried.

Two techniques were used to adjust ligand spacing. First, modified alginates were mixed with unmodified alginate at different volume ratios to give final RGDS concentrations of 12.5, 5, 2 and 0.8 mg RGDS/g alginate. Second, ligand spacing was adjusted by using different peptide concentrations during synthesis, specifically 5, 12.5 and 31.25 mg RGDS/g alginate. Ratios for stabilizers and linkers were unchanged.

Alginate was characterized by using T-FTIR and ninhydrin assay. FTIR analysis was performed on a Tensor 37 FTIR spectrophotometer (Bruker, Ettlingen, Germany) set at 32 scans and 4 cm ^"1 resolution. Alginate samples were analyzed in solid pellets after freeze-drying, grinding with potassium bromide (at 0.5 wt. %) and being formed into a pellet at 95,000 N using a hydraulic press (Apex Construction, UK) for 1 minute using a 13 mm diameter KBr pellet press (Specac, UK).

The ninhydrin assay was used to quantify the amount of RGDS in the alginate as previously described (Ito Y, Kajihara M, Imanishi Y (1991) J Biomed Mater Res A 25, 1325-1337.). Briefly, the freeze-dried alginate was reconstituted in distilled water to a preset 1.34 mg/ml peptide concentration (assuming 100% efficient coupling and no loss of peptide during dialysis). 100 uL Alginate sample was immersed in 400 uL of 5M HC1 and autoclaved for lh at 120 °C. The resulting acidic solution was neutralized by adding 400 uL of 5M NaOH. Then 900 uL of the ninhydrin solution was added to the hydrolyzed alginate sample. The ninhydrin solution was prepared as follows: 0.027 g of ninhydrin and 0.004 g of hydrindantin were added to 1ml 2- methoxyethanol and then diluted in 3ml potassium acetate buffer (pH 5.2). The alginate-ninhydrin solution was then incubated in a 100°C water bath for 3 minutes and allowed to cool for 8 minutes. Further dilutions may be performed with isopropanol where necessary to ensure readings are within calibration limits. All samples were filtered through a 0.22 μιη pore syringe filter prior to analysis using a UV-Vis spectrophotometer (model: Varian Cary 50®, Yarton, UK). Absorbances were measured at 570 nm at 25 °C. Unmodified alginate was used as blank. Solutions of free peptide in distilled water were used as standards for quantification.

Encapsulation: The mESCs were suspended in 1.2 wt. % alginate solution in

0.9 wt. % NaCl at a density of le6 cells/ml. The cell suspension was transferred into a syringe (BD Bioscience, UK) and ejected, drop by drop, through a 30 gauge needle (BD Bioscience, UK) into a beaker of 102 mM CaCl ₂ solution. The alginate beads thus formed were allowed to cross-link for 10 min before the excess CaCl ₂ was removed by decantation. Excess CaCl ₂ was removed by washing the alginate beads twice with 0.9 wt. % NaCl. The alginate beads were then resuspended in culture medium.

Actin Staining: The mESCs were washed with pre-warmed PBS, fixed in 3.7% formaldehyde and extracted in 0.1 % triton X-100 in PBS at -20°C for 3 minutes (encapsulated cells were allowed to proceed for 12 minutes). Cells were then pre- incubated with 1 % BSA in PBS for 20 minutes prior to staining with the Texas Red Phalloidin (Invitrogen, UK) staining solution at room temperature for 30 minutes. Finally, cells were washed twice with PBS, air-dried and stained with Slow Fade Gold Reagent with DAPI TM (Invitrogen, UK). Cells were imaged using an inverted microscope (Model: Ti-E® inverted microscope coupled to a Ds-Qil camera system, Nikon Instruments Europe B.V., The Netherlands).

Flow Cytometry (FC): The mESCs were cryopreserved under three different conditions and analyzed 24 hours after thawing. The three cryopreservation conditions were— in suspension, encapsulated in unmodified alginate beads and in RGDS- modified alginate beads. Cells in each of the three groups were stained with antibodies to octamer 4 (Oct-4), stage specific embryonic antigen 1 (SSEA-1) and Alkaline Phosphatase (ALP), which were conjugated to FITC (Invitrogen, USA), Cyanine-3 (Millipore, USA) and phycoerythrin (RnD Systems, USA) respectively. After staining, at least 10,000 cells were analyzed on a flow cytometer (model: BD FACSCalibur® BD Biosciences, USA) and the results were processed using WinMDI software (Scripps Institute, USA).

Cryopreservation and thawing: The cryopreservation process was optimized to improve the post-thaw cell survival rate. DMSO was used as a cryopreservation agent (CPA). The CPA was prepared by diluting stock solution (20% (v/v) DMSO, 20% (v/v) FBS, 60%) (v/v) DMEM) with mESC culture medium at a volume ratio of 1 : 1. Cells in suspension and cells encapsulated in alginate (at le6 cells/mL) were loaded with the CPA for 20, 30 and 45 minutes at 4°C. All samples were gently shaken intermittently during CPA loading by hand. All samples were slow cooled at the rate of 1, 5 or 10°C/min using a controlled rate freezer (Model: Kryo 560-16, Planer Pic, UK) until -80°C. For recovery from cryogenic storage, cells in suspension and cells in alginate beads were rapidly warmed by immersion in a 37°C water bath until fully thawed. Then mESC culture medium was added dropwise (reducing CPA

concentration to below 1.8 % vol.) and the samples were allowed to stand for 10 minutes at room temperature. Before analysis for post-thaw cell survival rate, cells in alginate beads were resuspended in 0.055M sodium citrate in 0.9 wt% NaCl to dissolve the alginate. The cells were then centrifuged at 148 g for 5 minutes then resuspended in mESC culture medium. Supernatant was checked for the presence of cells before discarding. Additionally, cell concentration after resuspension was reconciled to concentration during encapsulation (1 million cells/ mL).

Cell viability after thawing was evaluated by using trypan blue exclusion assay. Cells were resuspended in culture medium then an equal volume of sterile filtered 0.4%> wt. /vol. trypan blue was added to the suspension. Viability was assessed on a hemacytometer (Neubauer, UK) under a light microscope within 4 minutes of preparation. Viability was expressed as the number of live cells over the total cell number and expressed as a percentage.

Alamar blue assay was employed to monitor cell viability over 6 hour period post-thaw. Ie5 cells were maintained in 100 uL of medium supplemented with 10% Alamar blue in the incubator. Cells in alginate beads were recovered after alginate dissolution and plated on 2D gelatinized surfaces. Absorbance values were measured at 1 hour, 4 hours and 6 hours using a TECAN Genios® spectrophotometer (TECAN, UK) at 570 nm with reference readings at 600 nm. Reduction percentages were derived as previously described (Larson EM, Doughman DJ, Gregerson DS, Obritsch WF (1997) Invest Ophthalmol Vis Sci 38, 1929-1933). Cell differentiation: Unless otherwise specified all induction media and supplements were purchased from Millipore, UK. Mouse ES cells were differentiated to test for pluripotency. The three lineages were: endomucin-positive cells (endoderm- derived), adipocytes (mesoderm derived) and neurons (ectoderm-derived). Mouse ES cells were first developed into embryoid bodies (EBs) by passaging cells into an ultra low attachment dish (Corning, UK) using mESC medium without the addition of mLIF. After three days, the EBs were dissociated and plated onto respective substrates with the relevant media formulation for the desired lineage: (1) to generate endothelial cells, 2 mM VEGF-165 in basal medium and gelatin substrata were used. (2) For adipogenesis, 500 nM of 3-isobutyl- 1 -methylxanthine (IBMX), 850 nM insulin and, ΙμΜ dexamethasone, 100 μΜ indomethacin in basal medium were used. Basal medium, supplemented with 10 ug/ml insulin, was used for maintenance in between inductions. Cells were grown on a 0.1 wt % gelatin (in PBS) surface. (3) For neurogenesis, 0.1 μΜ retinoic acid in a basal medium were used for induction while the basal medium with no supplementation was used for maintenance. Cells were grown on a laminin (10 mg/mL) / poly-L-ornithine (50 ug/mL) -coated surface. For all three experiments, a total of five cycles of induction (3 days each) and

maintenance (1 day each) were performed prior to staining for respective lineages. Negative controls maintained in a basal medium and in mESC medium (without induction-maintenance cycles) were run in parallel. Immunocytochemical

characterization was determined for the differentiated cells after completion of differentiation. For endoderm-derived cells, the cells were washed and blocked. For intracellular staining of β-ΙΙΙ tubulin, the cells were fixed and permeabilized. Primary antibodies goat anti-endomucin antibody, neuron-specific mouse anti- β-ΙΙΙ tubulin antibody (both from RnD systems, UK) were added to the cells, incubated for three hours then washed. Secondary antibodies, Northern Lights 493 nm conjugated donkey anti-goat IgG and Northern Lights 493 nm (NL 493) conjugated donkey anti-mouse IgG (RnD systems, UK) were used to stain endoderm-derived cells and ectoderm derived cells by incubation for 60 min at room temperature then washed. For Oil Red O staining, cells were washed before a 0.36 wt. % oil Red O stain (solvent was 40% MilliQ water and 60 % isopropanol) was applied to the cells for 15 minutes. Cells were washed with PBS before observation in bright field. The slides with fluorescent stains were mounted with the SlowFade kit containing DAPI (Invitrogen GIBCO, UK). All slides were imaged using an inverted microscope (Model: Ti-E® inverted microscope coupled to a Ds-Qil camera system, Nikon Instruments Europe B.V., The Netherlands).

Statistical analysis: Each experiment was performed in triplicate and repeated three times. Statistical significance between groups was determined by analysis of variance (ANOVA) test with a multiple comparison test using the Tukey-Kramer method. Data was presented as mean ± standard error and a p-value of less than 0.05 was considered statistically significant.

Results and discussion

To confirm the presence of RGDS in the modified alginate, absorption spectra for modified and unmodified alginates were compared. RGDS-alginate has distinct peaks at 1280 and 960 cm ^"1 that were previously attributed to aspartic acid (in an RGDS peptide coupled to a polyurethane-polyethylene glycol backbone) (Lin YS, Wang SS, Chung TW, Wang YH, Chiou SH, Hsu JJ, Chou NK, Hsieh TH, Chu SH (2001) Artif Organs 25, 617-621); unique peaks close to these sites were identified (figure 1). Comparing RGDS-alginate to a positive control (mixture of alginate and RGDS) confirms these peaks are contributions from the peptide.

The concentration of RGDS incorporated into the scaffold was quantified by measuring the absorbance of RGDS-alginate samples that had been assayed using the ninhydrin reaction. Previous work has shown that amino acid derivatives maintain a peak absorbance at 570 nm. Hence, all measurements were performed at this wavelength. For the pre-dialysis sample, the concentration of RGDS was 1.38 mg/ml which was comparable to the estimated value (1.34 mg/ml) at only 3% difference. However, the post-dialysis sample had an RGDS concentration of 1.21 mg/ml at 10% difference. These differences may be attributable to inefficient coupling of peptide during solution modification or loss of peptide during dialysis. Nonetheless, the ninhydrin assay only serves to estimate the amount of RGDS peptide in the alginate sample. It has been previously reported that for peptide concentrations ranging from 1-10 mM, percent errors of about 16-25 % can be encountered.

Biological assays for cytoskeletal reorganization demonstrate the effect of

RGDS on cell attachment in the alginate matrix. Figure 2A and B show mESC grown on either gelatin surface (A) or RGDS-alginate surface (B). The cells show evidence of spreading, polarity and organization of actin into filaments. Figure 2C shows mESC encapsulated within RGDS-modified calcium alginate (bead radius: 0.6 mm). The cells show some evidence of cell spreading and actin remodelling, which is similar to cells in A and B. Figure 2D shows an mESC encapsulated in unmodified alginate. In contrast to the first three images, Figure 2D shows a strong actin staining but no evidence of remodelling and no polarity. The circular shape of the image is similar to the cell shape observed for encapsulated cells in bright field immediately after encapsulation. The evidence of mESCs response to RGDS-coupled alginate is consistent with reports confirming the expression of νβ3 integrins in mESC (Sasaki T, Timpl R (2001) FEBS Lett 509, 181-185). RGDS domains have been identified on ECM molecules within the blastocyst which explains the ability for mESCs to recognize RGD-containing peptides. Integrins identified on mESCs include the νβ3 integrins which play significant roles e.g. cell survival during embryonic development (Sasaki T, Timpl R (2001) FEBS Lett 509, 181-185). Alpha-V integrins are receptors of vitronectin, fibronectin and laminin which are all constituents of the basement membrane in blastocysts and possess the RGD repeat. They play a role in embryo implantation and development. Genetically, this mechanism of spreading is connected to the regulation of transcriptional activity of osteopontin by two other genes also associated with sternness i.e. oct-4 and sox-2. Osteopontin contains the RGDS motif and regulates cell adhesion and migration. Mammalian cells can attach, demonstrate attachment and spreading on RGD-containing surfaces within 20 minutes, unaffected by DMSO. The concentration of cross-linker used (CaCl ₂ ) can improve cell adhesion rates at concentrations as low as 25 mmole/g alginate (Genes NG, Rowley JA, Mooney DJ, Bonassar LJ (2004) Arch Biochem Biophys 422, 161-167.); low temperature binding (4°C) of RGD-containing peptides to alpha- V-beta-3 integrins has been observed to achieve saturation faster (30 minutes) than saturation at 37°C (2 hours).

Alginate was tested as a potential substrate for cryopreserving mESCs. A comparison of post-thaw viability of mESCs cryopreserved in modified, unmodified alginate and suspension by trypan blue exclusion assay confirms that attached cells ("modified alginate") had significantly higher post-thaw viability than those in unmodified alginate and suspension (Figure 3).

Alamar Blue assay was used to monitor the metabolic rate of cells after thawing as shown in Figure 4. Cells cryopreserved in RGDS-alginates had a significantly higher metabolic rate than both cells cryopreserved in suspension and encapsulated in unmodified alginate. This, together with the results from trypan blue assay, showed that cells cryopreserved in RGDS-alginate retained higher viability than cells cryopreserved in suspension or encapsulated in unmodified alginate. This is consistent with reports that encapsulation of cells in gelled high molecular weight polymers (HMWP) reduces membrane damage and the breakage of cell- substrate interactions. Physically, these HMWP inhibit the growth of extracellular ice by depressing the melting and homogeneous nucleation temperatures of their solutions. Furthermore, alginate cross-linkage over a 10 minute duration is diffusion- limited and only 55% of the maximum gel- volume is achieved based on experimentally- determined constants for gelation kinetics; the alginate capsule is therefore, a sol-gel aqueous system (hydrogel) retaining many of the properties observed in the polymer solutions before cross-linkage. The added benefit of cell attachment may be attributed to increased hydraulic conductivity allowing for cell dehydration and prevention of intracellular ice formation. Cell attachment does not immobilize cells, rather, it provides traction for cell movement and improves osmotic cell response to the changing chemical environment during cryopreservation (inventors observed sustained cell survival for cells in RGDS-alginate capsules as temperatures are progressively reduced at l°C/min on a graded freezing procedure, data not shown) resulting in increased post-thaw cell survival. When both DMSO and extracellular HMWP are used, the cryoprotective strengths of the two components are reported to be increased.

In addition to testing post-thaw survival rates, the post-thaw sternness of mESCs was examined. Flow cytometry data is shown in Figure 5 to compare the expression of some stem cell specific markers for cells cryopreserved in RGDS- alginate beads, unmodified alginate beads and in suspension. The data indicated that higher percentage of cells cryopreserved in RGDS-modified alginate beads expressed stem cell markers of ALP and Oct-4.

The sternness of the mESCs cryopreserved in RGDS-modified alginate were tested for their pluripotency by differentiating them along the three lineages and then staining for specific markers (neuronal and endoderm-derived lineages) or for inclusions (lipids for adipocytes). The dominant marker for epithelial cells lining major internal organs e.g. the ileum is endomucin. The dominant and exclusive marker for neurons is β-ΙΙΙ-tubulin. Tubulin is a component of microtubules which have a structural, transport and cell reproductive function. Figure 6 showed that after cryopreservation, these mESCs were still capable of differentiating into all three lineages.

Having observed a high cell survival rate and sternness of cells when RGDS- alginates were used as encapsulation material, it was necessary to see whether this factor can be optimized. To alter the ligand spacing, RGDS concentration was increased to 2.5 times the initial value (31.25 mg RGDS/ g of alginate) and to 0.4 times the initial concentration (5 mg RGDS/ g alginate) during chemical modification. The resulting RGDS-modified alginates would have "homogenous local RGDS concentration." The initial RGDS concentration (12.5 mg peptide/g alginate) is 25% higher than previous published values (Genes NG, Rowley JA, Mooney DJ, Bonassar LJ (2004) Arch Biochem Biophys 422, 161-167.) and is sufficient for cell attachment. The incorporated peptide varies linearly and directly in proportion to the input peptide for the peptide concentrations. Figure 7 (black line) shows the impact of ligand concentration on post-thaw cell survival. The results show that increasing the ligand concentration yielded no significant change in the post-thaw cell survival rate.

Conversely, decreasing the RGDS concentration reduced the cell survival rate to levels that were similar to that of unmodified alginate. Secondly, RGDS-alginate was mixed with unmodified alginate creating a "heterogeneous local RGDS

concentration". This set of experiments is shown in Figure 7 (grey line on secondary ordinate). Decreasing the alginate concentration shows statistically significant differences between post-thaw cell survival rates at 12.5 mg peptide/g alginate and lower peptide concentrations. Previous work for 2D adhesion studies using bovine chondrocytes has shown that optimal cell attachment occurs at around 10 mg peptide per gram of alginate (Genes NG, Rowley JA, Mooney DJ, Bonassar LJ (2004) Arch Biochem Biophys 422, 161-167.) which is close to what we observed for maximum post-thaw cell survival rates for mESCs. The results are consistent with observations that ligand spacing below the identified optimum yields unstable focal adhesions which are typical of non-specific cell-substrate interactions (e.g. in unmodified alginate) and are likely to contribute to cell death. Integrins may help cells sense the mechanical properties of the synthetic ECM which, if adequate, may improve cell function. The attachment of cells has been shown to increase the water permeability parameters of cells while slowing loss of water permeability with decrease in temperature so that cells dehydrate sooner, for longer and take up CPA faster during CPA loading -activities that correlate well with high cell survival rates. The loading time was also optimized while fixing the 12.5 mg RGDS /g alginate concentration. Figure 8 shows that attachment improved cell survival across all the loading times attempted. The 30 minute loading time was the best loading time when compared against each loading time used for cells encapsulated in RGDS- alginates. The improvement in cell survival across all loading times was attributed to the improved response of cells to CPA loading. The benefit of cell attachment has been demonstrated previously in 2D cryopreservation systems where surface-bound cells have been reported to endure and respond to the cryopreservation process better than cells in suspension .

Higher cooling rates were investigated as potential ways to reduce process time. Figure 9 shows a comparison of different cooling rates for chosen loading times. For the 20 and 30 minute loading times, a cooling rate of l°C/min yields the highest cell survival rates followed by 10°C/min while 5°C/min gave the lowest cell survival. This holds true for both suspension and 3D cryopreservation data. Furthermore, when loading time was extended to 45 minutes, there was no statistically significant difference in the cell survival rate between the cooling rate of 10°C/min and l°C/min. In general, cooling rates of 5 and 10°C/min are not typically used in cryopreservation by slow-cooling; these results confirmed that they are not as good as a l°C/min cooling rate.

Conclusions

In this investigation, it is demonstrated that mESCs cryopreserved using a linear programmed cooling rate of l°C/min in RGDS-modified alginate beads maintain a higher expression of stem cell markers (including Oct-4) within 24 hours after thawing, a higher cell survival rate and better cell health compared with cells cryopreserved in suspension or encapsulated in unmodified alginate using the same cooling rate. The current results indicate that encapsulation in RGDS-functionalised alginate provides better protection for cells during cryopreservation. EXAMPLE 2

Use of alginate encapsulation for improved cryopreservation of human embryonic stem cells The following experiments demonstrate that encapsulation in alginate improves human embryonic stem cell survival rate and cell health after thawing, while maintaining their sternness, even when using a simple constant-rate cooling protocol and in the absence of a ROCK inhibitor.

Materials and methods

Unless otherwise stated, all materials were obtained from Sigma- Aldrich, UK.

Culture of human embryonic stem cells (hESCs): As previously described (X. Xu et al; Biotechnol. Prog. 26 (2010) 781-788. doi:10.1002/btpr.358 ER), the human embryonic stem cell line, HUES2 (Howard Hughes Medical Institute, Department of Molecular and Cellular Biology, Harvard University, USA), approved by the UK Stem Cell Bank Steering Committee, was cultured using two methods: feeder- dependent culture and feeder- independent culture (only the cells from feeder- free culture were used for cryopreservation experiments). For the feeder-dependent culture, hESCs were cultured on a feeder layer of mitomycin C-inactivated mouse embryo fibroblast (MEF) on a 0.1% gelatin-coated plate, in hESC culture medium containing knockout (KO) Dulbecco's modified Eagle's medium, supplemented with 10% KO-Serum Replacement, 1% nonessential amino acids, 2 mM Glutamax-I (all from Invitrogen GIBCO, UK), 0.055 mM β-mercaptoethanol (GIBCO, UK), and 10 ng/mL basic fibroblast growth factor (bFGF) (R&D systems, UK) at 37°C, under 5% C0 ₂ , and 21 ) 0 ₂ in air. Culture medium was changed daily. For feeder- independent culture, the hESC cells were cultured on matrigel-coated plates diluted 1 : 100, in MTeSR™ culture medium (Stem Cell Technologies, France), at 37°C, under 5% C0 ₂ and 21% 0 ₂ . Culture medium was completely changed daily. For passaging, hESC colonies were detached by TrypLE™ Express (Invitrogen, UK) at 37°C for 5-7 min after 5-7 days of culture, followed by gentle flushing with pipette several times to detach hESC. Undifferentiated dissociated hESC were transferred to fresh MEFs plates or matrigel-coated plates which were prepared in advance, in the presence of 10 μΜ ROCK inhibitor Y-27632 during the first day of culture. The split ratio was 1 :5- 1 : 10.

Modification of alginate with RGDS and characterization: Alginate modification was carried out using "solution modification" method (J.A. Rowley et al, Biomaterials 20 (1999) 45-53). Briefly, medium viscosity alginate (2000 cP at 2 wt. % polymer at 25°C ) from brown algae was dissolved overnight in a buffer made of 50 mM 2-(N-morpholino) ethanesulfonic acid (MES) in 0.15M NaCl (pH 6.5) at a concentration of 1 g alginate/ lOOmL buffer. RGDS was covalently coupled to the alginate using carbodiimide chemistry with stabilization against hydrolysis using highly water soluble N-hydroxysulfosuccimide (sulfo-NHS). The sulfo-NHS to 1- ethyl-(dimethylaminopropyl) carbodiimide (EDC) ratio was at 1 :2 while the peptide to EDC ratio was at 1 :4. The sulfo-NHS, EDC and peptide (in that order) were added in quick succession (after the dissolution of each reagent) to alginate while stirring at room temperature. The reaction was quenched with hydroxylamine hydrochloride (HA-HC1) after 20 hours at 18 : 1000 (HA-HC1: alginate weight) ratio. The modified alginate was dialyzed against decreasing concentrations of NaCl solution from 15 g/L to Og/L Milli-Q water and then freeze-dried. Successful coupling of the peptide to the alginate was confirmed via Transmission-FTIR and ninhydrin assay.

Cell Encapsulation; Cells were centrifuged then resuspended in 1.2 wt % alginate (in 0.9 wt % NaCl) at a density of le6 cells/ml. The cell suspension was transferred into a syringe (BD, UK) and ejected, drop by drop, through a 30 gauge needle (BD, UK) into a beaker of 102 mM CaCl ₂ solution giving an average bead radius of 0.6 mm (standard error of 0.1 mm). The CaCl ₂ solution was supplemented with Dulbecco's Modified Eagle Medium (DMEM) to 547 mOsm. The alginate cell suspension was dropped from a 3 cm height. Alginate beads were allowed to crosslink for 10 min before the excess CaCl ₂ was removed by washing the alginate beads twice with 0.9 wt. % NaCl. The alginate beads were then resuspended in culture medium. For encapsulation in RGDS-coupled alginate, mouse ESCs were

encapsulated as above substituting RGDS-alginate for unmodified alginate. Beads were thereafter loaded with CPA and cryopreserved.

Cryopreservation and thawing: The cryopreservation process was optimized to improve the post-thaw cell survival rate. DMSO was used as a cryopreservation agent (CPA). The CPA was prepared by diluting stock solution (20% (v/v) DMSO, 20% (v/v) Foetal Bovine Serum (FBS), 60% (v/v) DMEM) with mTeSR culture medium at a volume ratio of 1 : 1. Cells encapsulated in alginate (at 1 e6 cells/mL) were loaded with the CPA for 20, 30 and 45 minutes at 4°C. All samples were gently shaken intermittently during CPA loading by hand then slow cooled at the rate of 1 or 5°C/min using a controlled rate freezer (Model: Kryo 560-16, Planer Pic, UK) until - 80°C. For recovery from cryogenic storage, cells were rapidly warmed by immersion in a 37°C water bath until fully thawed. Then mTESR ^® culture medium was added dropwise (reducing CPA concentration to below 1.8 % vol.) and the samples were allowed to stand for 10 minutes at room temperature. Before analysis for post-thaw cell survival rate, cells in alginate beads were resuspended in 0.055M sodium citrate in 0.9 wt% NaCl to dissolve the alginate. Cells were then centrifuged at 148 g for 5 minutes then resuspended in culture medium. Supernatant was checked for the presence of cells before discarding. Additionally, cell concentration after resuspension was reconciled to concentration during encapsulation (1 million cells/ mL). Cells in suspension were cryopreserved and recovered similarly without performing alginate dissolution, all resuspension culture medium was supplemented with Rho-associated Kinase (ROCK) Inhibitor (RI) at 0.01 mM except for controls without RI

supplementation ("Su-").

Trypan Blue Assay: Cell viability after thawing was evaluated by using trypan blue exclusion assay. Cells were resuspended in culture medium then an equal volume of sterile filtered 0.4% wt. /vol. trypan blue was added to the suspension. Viability was assessed on a hemacytometer (Neubauer, UK) under a light microscope within 4 minutes of preparation. Viability was expressed as the number of live cells over the total cell number and expressed as a percentage.

Colony Counts: Culture flasks were prepared for scoring by tracing 400 mm ² unit grid onto the bottom outer surface of a T75 culture flask (Corning, UK). After thawing, cells were seeded on gelatin-coated plates at 0.5e5 cells/cm ² density

(suspension cryopreserved cells were RI supplemented while cells recovered from RGDS-alginate beads were not) and monitored over 3 days. On the first and third day, cells colonies were counted; cells detached, dissociated and counted on a

hemacytometer (Neubauer, UK) before further culture. To eliminate dead cells, cell viability was assessed using a trypan blue assay (viable cells were > 99% in all counts). Counting criteria was 40 cells/ colony or more.

Alamar blue assay: Alamar blue assay was employed to monitor cell viability over 6 hour period post-thaw. Ie5 cells were maintained in 100 uL of medium supplemented with 10% Alamar blue (Invitrogen, UK) in the incubator. Cells in alginate beads were recovered after alginate dissolution and plated on 2D gelatinized surfaces without RI supplementation. Absorbance values were measured at 2 hours, 4 hours and 6 hours using a TEC AN Genios® spectrophotometer (TEC AN, UK) at 570 nm with reference readings at 600 nm. Reduction percentages were derived as previously described (E.M. Larson et al, Invest. Ophthalmol. Vis. Sci. 38 (1997) 1929-1933).

Apoptosis Assay: cells were thawed and resuspended in media containing RI (Su+), passaged from culture (Cltr) or recovered from RGDS-alginate beads and cultured in media without RI supplementation (3D-) for a fixed duration (2, 4 or 24 hours); controls were suspension cryopreserved cells without RI supplementation. At the appointed times, cells were fixed. All sample conditions were stained using FITC- conjugated Annexin V and propidium iodide from kit as supplied (Trevigen, UK), incubated at room temperature in the dark for 15 minutes, resuspended in a binding buffer and analyzed by flow cytometry at a minimum of 10000 events per condition.

Comet Assay: DNA damage was tested using the comet assay (also known as single cell gel electrophoresis assay) kit (Cambridge Bioscience, UK). Cells were recovered and resuspended at a density of le5 cells/mL in ice cold PBS (no Mg ²⁺ and Ca ²⁺ ) then le4 cells were mixed with pre-warmed 1 wt % agarose at 37 °C at a 1 : 10 volume ratio and applied on a 3-well comet assay slide (pre-warmed at 37 °C) at 75 μΐ _^ per single well. The sample was gelled for 15 minutes at 4 °C before immersion in a pre-chilled (on ice) lysis solution (provided in kit) containing EDTA solution (supplied with kit) 10 % vol. DMSO and 14.6 wt % NaCl in MilliQ water . After 30 minutes in the dark at room temperature, lysis buffer was aspirated and replaced with pre-chilled alkaline solution (50 mL per slide) composed of 1.2 wt % NaOH pellets, EDTA solution (provided in kit) in MilliQ water; sample was left for another 30 minutes in the dark at 4 °C. Slides were then rinsed with MilliQ water and placed in a GIBCO BRL Model H5 horizontal gel electrophoresis apparatus. Slides were covered with 300 mM NaOH, 1 mM EDTA, in MilliQ water, pH > 13. Slides were left in the apparatus at 1 V/cm uniform electric field (assuming no edge effects) and 300 niA for 30 minutes, removed, rinsed with MilliQ water and incubated for 15 minutes in the dark with Vista Green Dye (provided in kit). After staining, slides were rinsed in distilled water, placed in a humidified air-tight container and viewed immediately on an upright Nikon Eclipse 80i microscope connected to an ORCA-AG CCD camera. The extent tail moment was calculated as a product of tail DNA percent (percent ratio of tail DNA intensity over cell DNA intensity) and tail moment length. At least 50 cells were assayed per sample condition. Controls were ran after a 90 minute exposure to short-wave UV from a low pressure mercury vapour discharge tube. Cell differentiation: Human embryonic stem cells were differentiated to endoderm-derived Soxl7+ cells, adipogenic and neuronal cells. Human ES cells were first developed into embryoid bodies (EBs) by plating recovered 3D- cells into an ultra low attachment dish (Corning, UK) using embryoid formation medium

(Millipore, UK) after alginate dissolution without RI supplementation. EBs were dissociated into single cells using TrypLE™ Express (Invitrogen, UK) at 37°C for 5 minutes before plating at 3e6 cells per 60 mm plate. After three days, the EBs were dissociated and plated onto respective substrates with the relevant media formulation for the desired lineage. For derivation of endoderm-derived cells and adipogenesis, cells were seeded on 0.1 wt % gelatin at a 0.5e5 cells/cm ² cell density; for

neurogenesis, cells were plated onto (10 mg/mL) laminin (Millipore, UK) / (50 μg/mL) poly-L-ornithine-coated surface at 0.5e5 cells/cm ² . For endoderm

differentiation, dissociated cells were cultured on EB formation medium until post- confluence then conditioned in EB formation medium supplemented with human Fibroblast Growth Factor (FGF) basic for 4 hours before being cultured in Endoderm Differentiation Base Media (EDBM) supplemented with FGF basic, human Activin and human Wnt-3a (RnD Systems, UK) for 1 day. The medium was changed twice on the second day to EDBM supplemented with FGF basic and Activin A (differentiation base medium (DBM) II). On the third day, the cells were cultured with DBM II twice a day. The three day differentiation cycle was repeated for three weeks at which point cells were observed to be ready for tests by flow cytometry. For adipogenesis, dissociated cells from EBs were conditioned by culturing in EB formation medium supplemented with adipogenic supplement (RnD Systems, UK) for 4 hours before being cultured in adipogenic base media with adipogenic supplement, changed daily, for 21 days at which point cells were observed to be ready for testing by flow cytometry. For neurogenesis, dissociated cells from EBs were conditioned by culturing in EB formation medium supplemented with neural inducer (Millipore, UK) for 4 hours before being cultured in embryoid body formation medium supplemented with neural inducer, changed daily, for 21 days at which point cells were observed to be ready for testing by flow cytometry.

Suspension cryopreserved cells and cells that were previously in culture were differentiated in a similar fashion but were not required to be recovered from constructs via alginate dissolution. However, cells were first developed into embryoid bodies (EBs) by plating thawed suspension-cryopreserved cells into an ultra low attachment dish (Corning, UK) using embryoid formation medium (Millipore, UK) supplemented with RI before commencing differentiation process. Controls were run without RI supplementation (Su-).

Flow Cytometry: Cells at passage 30 were stained with antibodies to octamer 4 (Oct-4), Nanog, stage specific embryonic antigen 4 (SSEA-4) and Alkaline

Phosphatase (ALP) which were conjugated to phycoerythrin (RnD Systems, USA). After staining, at least 10,000 cells were analyzed on a flow cytometer (model: BD FACSCalibur® BD Biosciences, USA) and the results were processed using WinMDI software (Scripps Institute, USA).

Following differentiation of hESC at passage 31 (cells trypsinized from culture using TrypLE Express (Invitrogen, UK) ("culture" or "cltr"), (suspension cryopreserved cells with ROCK inhibitor (RI) supplementation post-thaw [Su+], suspension cryopreserved cells without ROCK inhibitor (RI) supplementation post- thaw [Su-] and RGDS-cryopreserved cells without RI supplementation post-thaw [3D-)]), cells were stained using Sox 17 (endoderm differentiation), Fatty Acid Binding Protein 4 (FABP4, adipogenesis) and β-ΙΙΙ Tubulin (ectodermal

differentiation) antibodies (Ab). The cells were trypsinized using TrypLE™ Express (Invitrogen, UK) and resuspended in PBS then fixed, permeabilized and blocked at room temperature before overnight incubation in primary antibodies (RnD Systems, UK) at room temperature in the dark. Cells were washed before incubation in secondary antibody conjugated to Northern Light (NL) 637 for FABP4 staining (RnD Systems, UK) and Alexa Fluor 568 (Invitrogen, UK) for the β-ΙΙΙ Tubulin antibody for 60 minutes in the dark; Sox 17 samples were stained using NL557-conjugated Sox 17 antibody for 3 hours. All samples were washed, resuspended in cold PBS and placed on ice awaiting measurement on a flow cytometer. At least 10,000 cells were analyzed on a flow cytometer (model: BD FACSCalibur® BD Biosciences, USA) and the results were processed using WinMDI software (Scripps Institute, USA).

Statistical analysis: Each experiment was performed in triplicate and repeated three times. Statistical significance for single paired comparisons was determined using a paired t-test assuming unequal variances. Statistical significance between groups was determined by analysis of variance (ANOVA) test with a multiple comparison test using the Tukey- Kramer method. Data was presented as mean ± standard error and a p-value of less than 0.05 was considered statistically significant. Results and discussion

The extent of cryoinjury experienced by cells during cryopreservation was investigated. In figure 10, cells cryopreserved in RGDS-alginate had a lower incidence of cryoinjury as indicated by the higher percentage of cells excluding trypan blue compared to cells cryopreserved in suspension (Su+ and Su-). This observation is in accordance with reports that encapsulation of cells in gelled high molecular weight polymers (HMWP) reduces damage to membrane and cell-substrate structures.

Physically, these HMWP may inhibit the growth of extracellular ice by depressing the equilibrium melting and homogeneous nucleation temperatures of their solutions. The added benefit of cell attachment may be due to the increased water permeability allowing for faster cell dehydration and prevention of intracellular ice formation during slow-cooling. Cell attachment facilitates the transmission of tractive force for cell movement and improves the osmotic cell response to the changing chemical environment during CPA loading (inventors observed progressively high cell survival rate (CSR) for cells in RGDS-alginate capsules as temperatures are progressively reduced at 1° C/min on a graded freezing procedure, data not shown) resulting in higher post-thaw cell survival. When both DMSO and extracellular HMWP are used, the cryoprotective strengths of the two components are reported to be synergistically increased. These results show that the addition of RI does not affect the degree of cryoinjury since the Su+ and Su- cells were no different from one another.

The cooling rate was also varied for the best performing method (3D-) as was the loading time as per figure 11 indicating that the l°C/min cooling rate gives significantly higher post-thaw cell survival rate (CSR) than a higher rate (5°C/min). Generally, cooling rates of 5 C/min or more are not typically used in cryopreservation by slow-cooling; these results confirmed that a slower cooling rate at 1 °C/min yields higher CSR.

The trypan blue assays were corroborated with colony counts from cells seeded after cryopreservation. From figure 12, 3D- cells have higher number of colonies three days after thawing compared to Su+ cells consistent with results from vital staining; the 3D- cells also have a higher growth rate than the Su+ cells. The positive control (CLTR) has the highest cell growth while the negative control (Su-) has the lowest growth rate. The post-thaw cell health was also monitored at 2, 4 and 6 hours after thawing and showed that the metabolic rate of 3D- cells at the 4 ^th and 6 ^th hours is significantly higher than the Su-cells as indicated by its higher Alamar blue reduction rate in figure 13 consistent with vital staining (upstream) and colony counting (downstream). The Su+ cells have the lowest metabolic rate consistent with reports of RI supplementation inhibiting some energy-dependent activities such as cell motility and centrosome positioning. The Su- cells have a higher metabolic rate than the Su+ cells which may indicate a high biochemical activity some of which may be associated with the execution of apoptotic signals. The CLTR positive control has the highest metabolic rate consistent with the absence of negative effects of cryopreservation.

Beyond looking at the cell metabolic rate, the apoptosis rate for the cells in 3D- and Su+ were investigated showing that the 3D- cells have lower apoptosis rates for the 2 ^nd and 4 ^th hours post-thaw as per figure 14. There is no difference between the two cell populations 24 hours post-thaw where the benefits of 3D cryopreservation appear to be overwhelmed by cell doubling and acclimation to the post-thaw incubation conditions - this can be gleaned from the similarity in the apoptotic rate observed in the CLTR group and in the Su+ and 3D- groups. The Su- negative control shows a consistently higher apoptosis rate of all the groups tested. The apoptotic rate observed at 2 hours for the Su+ cells is corroborated from previous studies. The decline in apoptosis rates observed in cells supplemented with RI shows that apoptosis is reversible - this has previously been confirmed.

The extent of DNA damage was also investigated as per figure 15 showing that the Su+ cells have a slightly higher level of DNA damage sustained during cryopreservation compared to 3D- cells (at an 82% level of confidence based on student's t-test i.e. a 0.82 probability of sample means between Su+ and 3D- specimens being different on a random draw). For both 3D- and Su+ cells, the levels of DNA damage are significantly lower than those sustained during exposure to UV light and for Su- cells. The culture control has some degree of DNA fragmentation comparable to that of the Su+ and 3D- cells. Previous research has indicated that hESCs have far better DNA repair mechanisms than differentiated human cells so that even when higher DNA damage is sustained than that observed in the 3D- and Su+ cell populations, hESC can still recover the status quo.

Although a given cell population may have a strong stem cell marker expression during culture, the population may lose this feature after cryopreservation (I.I. Katkov et al, Cryobiology 53(2006) 194-205. oi: 10.1016/j.cryobiol.2006.05.005); hence tests for sternness were run after thawing. Figure 16 shows that ALP, Nanog, Oct4 and SSEA4 expression is better conserved in 3D- group than in the Su+ group confirming reports of a decrease in stem cell marker expression after suspension cryopreservation; cells that were continuously cultured still show a strong expression of stem cell markers (Cltr) while those suspension cryopreserved without RI supplementation show a low expression of stem cell markers (Su-) which may indicate that some of the surviving cell population from earlier experiments may have already lost their stem cell phenotype; RI supplementation or the provision of a substratum during cryopreservation appears to diminish this loss as shown by the 3D- and Su+ cells.

The decreased expression of stem cell markers may indicate a loss of pluripotency (S. Koestenbauer et al, Am J Reprod Immunol 55 (2006) 169-180). From figure 17, although both Su+ and 3D- cells have the ability to differentiate, 3D- cells show a significantly higher success rate in adipogenesis. This may be attributed to the greater complexity of adipogenesis relative to endoderm and ectoderm differentiation implying a greater demand on cell pluripotency than has been observed for other primary germ layers. The mesoderm is a developmental feature that distinguishes diploblastic animals from triploblastic animals with underlying processes of more recent evolutionary origin and embryogenic complexity than the ectoderm and endoderm. Immunocytochemical results qualitatively confirmed the successful differentiation of both stem cell populations (data not shown). The positive control (Cltr) has a high success rate during differentiation given the cells were not exposed to the negative effects of cryopreservation while the Su- cells show a significantly lower success rate during differentiation consistent with its low expression of stem cell markers that means the survival cell population may not be as stem as the cells before cryopreservation.

Conclusions

Cryopreservation of hESCs in calcium RGDS-alginate capsules without RI supplementation conserves sternness, improves adipogenesis (and thus pluripotency), diminishes cryoinjury and diminishes apoptosis in comparison to cryopreservation of hESC in suspension. These findings indicate that attachment mediated mechanisms may be used to successfully store and repopulate hESC stocks without RI supplementation. Cryopreservation in RGDS-alginate capsules is likely to be beneficial for all cell types that abundantly express RGDS-recognising integrins e.g. alpha- V beta-3 integrins, subject to suitable optimization of process parameters such as ligand density, distribution, cell recognition kinetics and cryopreservation parameters.