WO1998035021A1 - Preparation of spheroids and their use in medicin or diagnosis - Google Patents

Abstract

Spheroids can be cryopreserved by techniques which include step-wise cooling. The spheroids may be formed prior to cryopreservation by orbital rotation, which may be at two or more different spin speeds. Orbital rotation following cryopreservation may also be performed. Cryopreservation of spheroids allows hospitals and laboratories to be supplied with spheroids which can be thawed and used when desired, thereby avoiding the need for preparing spheroids in situ.

Description

PREPARATION OF SPHEROIDS AND THEIR USE IN MEDICIN OR DIAGNOSIS

The present invention relates to artificially produced aggregates of cells in the form of three dimensional structures which are known as spheroids.

During the 1950's Garber and Moscona discovered that single cell suspensions of fetal cells, taken at the right developmental stage from mammalian and avian sources, could spontaneously reaggregate on rotation to form three-dimensional 'spheroids', or 're-aggregate' cultures (Moscona, 1952; Moscona, 1961; Garber and Moscona, 1972). Whilst this may have remained just an interesting phenomenon, closer examination demonstrated that spheroids could reproduce the complex cellular organisation, development and maturation of the original tissue from which the single cells had been isolated. A common feature of monolayer cell cultures is their loss of phenotype the longer they remain in culture. However brain spheroids appear to restrict cellular division whilst enhancing biochemical and morphological differentiation (see Seeds et al., 1980) thereby more closely reflecting the in vivo development process compared with primary monolayer cultures. Spheroids are therefore suitable for long term culturing experiments and in the presence of Vitamin E can remain viable for up to two months (Halks-Miller et al., 1982).

Spheroidal cultures were initially grown from limb buds, liver and kidney tissue (Moscona, 1961) succeeded by whole brain and retinal spheroids (Garber and Moscona, 1972). In particular, brain reaggregate cultures have been studied extensively because of the lack of complex CNS cultures that can be maintained long term in vitro. In addition, the yield of tissue is sufficient for standard biochemical analysis of whole spheroids as well as subcellular fractions (Trapp and Richelson, 1980). Under certain circumstances the cells can migrate and organize themselves into aggregates which resemble the architecture of the brain area sampled. Similarly, spheroids can be produced which histotypically resemble a particular brain region so long as such discrete areas or nuclei have been used as the starting material (DeLong, 1970; Garber, 1977; Levitt et al., 1976; Tsai, 1976). This is useful for closer examination of selective CNS toxicity caused by specific agents for example, MPTP.

CNS spheroids comprise an integrated population of neurones, astrocytes and other glial cell types for example, oligodendrocytes. Spheroids can therefore be sampled at various developmental stages as the CNS cells develop from undifferentiated neuroepithelial cells to a population of mature neurones, astrocytes and oligodendrocytes, correlating with in situ CNS development (Seeds et al. , 1980). These cultures have been well characterised morphologically (DeLong, 1970; Trapp et al., 1979; Goldsmith and Berens, 1990), biochemically (Honegger and Richelson, 1976; Trapp and Richelson, 1980; Honegger and Schilter, 1992) and electrophysiologically (Stafstrom et al. , 1980). Various studies using electron microscopy have demonstrated the presence of all types of synapses within aggregating brain cell cultures (Kozak et al., 1977; Seeds and Hoffke, 1978; Seeds and Vatter, 1971; Stefanelli et al., 1977). Again, spheroids cultured from discrete brain regions exhibit the synaptic profile characteristic of that brain area. Aggregate cultures can also be enriched in either neurons or glial cells (Honegger and Werffeli, 1988; Atterwill, 1989) however the spheroid as a mixed cell culture enables the study of cellular interaction between neurones and glia.

Using various morphological techniques it has been shown that fetal cells, in a dissociated cell suspension, first form a group of randomly associated cells on rotation, which then migrate to produce an organised spheroid. Within the spheroid, astrocytes form a shell or 'glia limitans' around the outside of the structure with neurones and oligodendrocytes arranged within the interior. Smaller neurones, such as the granule cells are located innermost whilst larger neurones (including Purkinje cells) are found near the surface of the spheroid as in mature tissue in situ (Seeds et al. , 1980). An in depth developmental description of brain spheroids is provided by Goldsmith and Berens (1990).

During the 1970's brain spheroids were extensively characterised in term of expression of neurotransmitter systems. In the 1980's it was recognised that spheroid cultures offered great potential in neurotoxicology (see below). However spheroid cultures have also been used to investigate a range of different CNS conditions particularly with respect to CNS invasion by other cell types in order to understand HIV transmission (Pulliam et al. , 1991), multiple sclerosis (Loughlin et al., 1994) and cancer metastases (Lund-Johansen et al. , 1992) especially malignant glioblastoma invasion (Steinsvag et al., 1985; Bjerkvig et al. , 1986; deRidder et al. , 1987; Pulliam et al. , 1988). Neurotrophic viral infections have been studied in brain spheroids (Pulliam et al. , 1984) as well as myelination (Matthieu et al., 1979; Louglin et al., 1994) and the development of antioxidant enzyme systems (Aspberg and Tottmar, 1992) as well as pure developmental studies (Honegger and Schilter, 1992). More recently, brain spheroids have been used as a source of tissue for transplantation studies (Marienhagen et al., 1994). There are several good reviews on CNS spheroids highlighting their usefulness in neurobiology (Trapp and Richelson, 1980; Atterwill et al., 1989; Goldsmith and Berens, 1990; Honegger and Schilter, 1992). It has now been realised that brain spheroids appear to be equally well suited to pharmacological and toxicological research (Wehner et al., 1985 Atterwill et al., 1986; Honegger and Werffeli, 1988).

As well as brain spheroids Honegger & Schilter (1992) first noted that it was possible to culture hepatic (liver) spheroids from foetal tissue and then coculture them with adult rat brain spheroids. This is particularly useful for neurotoxicants and compounds requiring metabolic activation and avoids the inherent toxicity of liver microsomal S9 fractions to neural cultures. Their initial studies demonstrated that Phenobarbital- and 3-methylcholanthrene-exposed liver spheroids were able to produce neurotoxic metabolites of cyclophosphamide in vitro. Since then, interest in liver spheroids has arisen for hepatoxicological test models in their own right (Ueno et al, 1992; George et al, 1996). A method combining orbital rotation with substratum-coated plastic with PHEMA (2-hydroxyethyl methacrylate) allows liver spheroid culture from adult rat liver where the intraspheroidal hepatocytes retain many of the in vivo morphological characteristics (George et al, 1996).

Rat adult liver and foetal brain spheroids have now been co-maintained in the brain spheroid-based medium following separate initial cultivation in their optimal growth media in both our laboratory and that of Honegger & Schilter (1992). The two spheroidal subtypes co-exist for around a week without significant loss of structure or increase a basal toxicological indicators. It is anticipated that such methodology will further progress in vitro neurotoxicity testing strategies for compounds such as Ecstasy (MDMA) which require hepatic metabolic activation before neurotoxic effects.

In view of the foregoing comments it will be appreciated that spheroids are useful for many purposes. However their utility has been limited because they take a long time to prepare and typically only remain viable in culture for around 1-2 months.

According to the present invention there is provided a method comprising the steps of:

a) obtaining a plurality of cells derived from a tissue or an organ or from a part thereof; b) orbitally rotating the cells in a fluid at sufficient speed and for a sufficient time for spheroids to form; and c) cryopreserving a composition comprising the spheroids and a cryopreservant. Although the cryopreservation of a limited number of cell types (with varying degrees of success) is already known, there is no disclosure in the literature of the successful cryopreservation of spheroids prior to the present invention.

In order to function as useful in vitro models of tissues/organs, spheroids should substantially retain their general three-dimensional structures. Furthermore the cells of the spheroids should retain viability and should also retain any biological characteristics which may be desired for spheroids to function as models of in vivo systems. A skilled person would realise that extreme techniques such as cryopreservation can disrupt individual cells and can also disrupt intercellular interactions which are needed to maintain a spheroids three-dimensional structure.

The difficulty of developing high-viability cryopreservation procedures is clear when one considers the hostile environment to which cells and tissues are subject during the freezing process: typically the temperature can drop from -r-37°C to

-196°C, loss of over 95% of cell water can be incurred, the electrolyte concentration inside and outside the cells can increase by several orders of magnitude relative to isotonic conditions, concentrated organic solvents in the freezing media permeate the cells, ice crystals intercalate the tissue and mechanically deform cells, and ice may form inside cells, disrupting intracellular structures or between cells, disrupting cell-cell aggregation.

It is also important to note that cryopreservation cannot be equated with short term hypo hermic preservation at relatively high temperature since cryopreservation occurs under much more extreme conditions . Short term hypothermic preservation of porcine hepatocytes is discussed by Sakai et al in Cell Transplantation 5(4); 505-511 (1996), where it is indicated that porcine hepatocytes can be stored for a short period (up to 3 days) without significant loss in function. In any event it is clear that the techniques disclosed by Sakai et al are considered to be of limited application since it is admitted that the protocol described only provides low cell densities and that further improvement is required. Indeed the cell densities achievable by the techniques disclosed by Sakai et al are inadequate for bioartificial liver (BAL) applications.

It should be noted in particular that prior to the present invention a skilled person would not expect to be able to achieve cryopreservation of spheroids comprising neural cells (e.g. brain spheroids). Simple preservation techniques, such as refrigeration or tissue culture, have drawbacks including limited shelf-life, high cost, risk of contamination or genetic drift. Another technique is cryopreservation, an approach based on the principle that chemical, biological and physical processes are effectively 'suspended' at cryogenic temperatures.

While freezing of cells and tissue explants is not an uncommon procedure, one must be careful not to assume that current cryopreservation procedures can be applied with universal efficacy to engineered tissue material in the form of spheroid cultures. Many disclosed freezing protocols have resulted in low recovery rates, or altogether non-viable tissue.

With respect to brain spheroids, the necessity to preserve the complex three- dimensional architecture, cellular heterogeneity and cell-cell junctions (e.g. synapses and glial cell processes) to permit biochemical functionality is significant. Given the numerous different cell types comprising a brain spheroid (e.g. neurons, glial) and the knowledge that the cooling rate required for optimal survival varies by several orders of magnitude among different cell types, it has previously been considered impossible to simultaneously satisfy optimal cooling requirements for all cells in a tissue. Furthermore, freeze-thaw procedures must result in recovery of cell viability and tissue structure to permit neurophysiological, pharmacological and/or toxicological functionality and assessment thereof.

Spheroid cultures in particular present certain inherent difficulties with respect of developing a suitable cryopreservation protocol. Given their three-dimensional shape and size (typical range 100-1000 μm) it may be anticipated that surface cells may be exposed to toxic concentrations of cryoprotectant in order to attain the minimal necessary cryoprotectant concentration in the spheroid interior. Conversely, if care is taken not to harm the outer cells during cryopreservation, interior cells may have cryoprotectant concentrations too low to protect against freezing. The present invention therefore represents a significant advance over prior art, in that the cryopreservation protocols for spheroid cultures allow an even distribution of cryoprotectant across a spheroid to be maintained such that no region is significantly damaged by over- or under-exposure to the cryoprotectant.

Post-thaw assessment of morphology, histology and biochemical functionality show that cell-cell junctions and cellular heterogeneity are maintained following cryopreservation.

Various aspects of the present invention are set out in the claims provided herewith. Some of these will now be described in greater detail below:

Cells for use in producing cryopreserved spheroids can be derived from any suitable tissue source, including infected tissue (as will be discussed later). For spheroids comprising neuronal cells (e.g. brain spheroids), foetal tissue is preferred. For spheroids comprising other cells, tissue from both foetal and non- foetal (e.g. adult sources) can generally be used. Liver cells are particularly useful since they can be used to produce spheroids which have some of the characteristics of the liver and can therefore be used to model the in vivo metabolism of substances by the liver. This is useful, for example, in determining whether or not particular substances are likely to be toxic following metabolism by the liver. In one embodiment of the present invention spheroids derived from liver cells are co- cultured with spheroids comprising neurons (e.g. brain spheroids). This is useful in screening substances for possible neurotoxicity. Spheroids can in principle be produced from any desired tissue or organ from any animal by disrupting a sample of the tissue or organ, preferably to individual cells or to small groups of cells (e.g. by mechanical disruption via gentle trituration through a Pasteur pipette) and causing aggregation of the cells by orbital rotation. They can be used as in vitro models and are therefore advantageous alternatives to using live animals.

Preferred models are mammalian (e.g. human, non-human primate, porcine, rodent) or avian (e.g. chick) models.

The present invention allows the use of spheroids to become much more widespread because they can be prepared at one site in large numbers using the information provided herein and can then be cryopreserved and transported in the cryopreserved state over large distances and for long time periods to other sites (e.g. to hospitals or laboratories which can even be located overseas). They can then be stored, cultured with ease and used when desired, thereby avoiding the need to prepare spheroids in situ. These advantages are especially important in respect of human tissues.

The use of cryopreservation can allow spheroids to be preserved in a viable state for a long time until needed. It is anticipated that, like other cryopreserved cells and tissues, spheroids may be held in the cryopreserved state for a month or more without significant loss of functionality upon thaw. Indeed it is anticipated that they may be held for a year or more.

It will therefore be appreciated that the present invention represents a significant technical development likely to be of major significance to the pharmaceutical, chemical and agricultural industries. It is particularly useful in the healthcare sector (e.g. in transplantation for drug/chemical testing and for screening purposes). In one embodiment of the present invention the spheroids comprise lesions. Lesions can be provided (for example) by ibotenic acid or ethylcholine mustard aziridinium ion (ECMA) treatment of spheroids for cholinergic neurones (Pillar et al, 1993), MPTP or 6-hydroxydopamine (6-OHDA) for Dopaminergic or Dopaminergic/noradrenergic neurones respectively, Kainic acid, ecstasy (MDMA) or NMD A for excitotoxic lesions, iron (Fe) for free-radical/ oxidative stress- induced damage, 5,6-dihydroxy tryptamine for serotonergic neurones, in vitro ischaemic or anoxic manipulation to emulate stroke-induced damage, and aluminium or /3-amyloid (β-AP) for Alzheimer-like neuropathology.

They are useful for e.g. testing novel therapeutic drugs/molecules (NCE's) which could reverse or ameliorate such lesions for neurodegenerative conditions such as Alzheimer's Disease, Parkinson's Diseases, Stroke and lesions induced by pesticide, environmental/industrial chemical, or therapeutic (e.g. antibiotics) or recreational drug exposure, or through genetic predisposition (e.g. diabetic neuropathy). They are also useful for testing potentially useful therapeutic neurotrophic factors such as NGF, BDNF, GDNF etc since we have already shown e.g. that NGF treatment in vitro of rat brain spheroids can reverse ECMA- induced cholinergic damage (Atterwill & Meakin, 1990). The lesioned spheroid model could also be used to investigate the molecular and cellular neuropathological processes accompanying such lesions for comparison to the pathophysiological mechanisms in animal models and humans in neurodegenerative states.

In another embodiment of the present invention, the spheroids can contain 'foreign' genes. These can be introduced by standard molecular biological techniques for cell transfection, for example, electroporation. The appropriate reporter elements of promotor-reporter constructs are e.g. Chloramphenicol Acetyltransferase (CAT), Luciferase and Secreted Alkaline Phosphatase (SAP). These reporters can be used to detect activation of early response genes relevant to pharmacotoxicological applications (e.g. p53, bcl, c-fos, c-jun, NF-κB, etc). In another embodiment of the present invention spheroids are co-cultured with cells derived from blood vessels prior to cryopreservation (e.g. from capilliaries). This is useful in providing in vitro models of in vivo systems in which blood vessels are involved e.g. of the blood brain barrier (BBB).

In a further embodiment of the present invention, spheroids can be prepared from cells or tissues infected in vivo, ex vivo, or in vitro with microbiological organisms (e.g. viruses, bacteria, protozoa, etc.). Such 'infected' spheroids may be used to, for example, assess the efficacy of drugs on infective organisms, screening of novel anti-infective agents, follow the pathogenesis of infection, etc.

One model of the BBB involves the co-culture of EC (endothelial cells) with neural tissue cultures in the form of spheroids. These organotypic cultures comprise neural and glial elements which may provide a suitable matrix for EC attachment and growth hence producing an in vitro BBB. Whole rat brain spheroids have been shown to behave organotypically in toxicity studies (Atterwill et al, Br. J. Pharmacol. 83:89-102 (1984)). It has already been demonstrated that EC attach to reaggregates (Bystry et al, Hum. Expt. Tox. 2:171 (1996)).

The following description is of a method which has been used to prepare a BBB model (which can then be cryopreserved):

Brain spheroids were prepared by the method of Atterwill et al (supra). EC were prepared by the method of Abbott et al (J. Cell Sci. 103:23-27 (1992)). RBE4 cells were cultured by the method of Roux et al (J. Cell Physiol. 159:101-113

(1994)). Confluent cultures EC were stained at 5 days in vitro (DIV) and RBE4 were stained at 70% confluence with the fluorophore l,l'-Dioctadecyl-3,3,3',3'- tetramethylindodicarbocyanine perchlorate, (Dil), a non specific cell dye for two days. Dil was used to visualise EC and RBE4 cell attachment to the reaggregates. -lilt is important to note that the cryopreservation of spheroids is not only useful in respect of spheroids intended as animal models (e.g. of human or other mammalian systems). It is also useful in respect of spheroids which can themselves be used in treatment.

For example spheroids can be useful in providing bioartificial systems. One example of this is a bioartificial liver (BAL).

Porcine hepatocytes may be suitable for use in bioartificial liver (BAL) systems (Saki et al, 1996). Such systems can provide effective treatment of patients with acute liver failure or can provide life-prolonging systems for patients awaiting orthotropic liver transplantation. Porcine hepatocytes express high hepatic functions in vitro and resemble human hepatocytes in morphology and functions.

Recently, porcine hepatocytes have been reorganized into spheroids having a tissue-like structure in suspension culture, and they express higher hepatic functions in vitro as compared to conventional monolayers, as confirmed in the case of rat hepatocyte culture.

Spheroids can also be used to provide implants to be transplanted into and maintained in patients. They are therefore useful in restorative transplantation.

They are particularly useful in treating degenerative diseases or disorders since they can allow damaged cells or tissues or organs to be replaced. The degenerative diseases or disorders may be genetically-based or may be caused by environmental factors (e.g. by exposure to toxic drugs or other chemicals).

For example, neurodegenerative diseases (e.g. Parkinson's disease, Alzheimer's disease or stroke) may be treated. Neuronal degeneration caused by Ecstasy or by other drugs may also be treated. Liver disease or damage may also be treated (e.g. by using bioartificial livers as described above or by providing implants).

For the above medical uses it is desirable for a physician to have access to large numbers of cells reasonably quickly so that treatment of patients can be carried out. The present invention allows such access to be achievable since large numbers of spheroids can be stored in a cryopreserved state and can be thawed and prepared for use shortly before being required.

Cryopreservation methods of the present invention will now be discussed.

Various cryopreservants can be used, although DMSO is preferred.

Other cryopreservants include permeating cryoprotectants besides DMSO can however be used: e.g. glycerol, 1,2-propanediol, acetamide, ethylclycol and propylene glycol (Karlsson & Mehmet, 1996; Chao et al, 1994). Non-permeating cryopreservants may even be used: e.g. methy .cellulose poly vinyl pyrollidone, hydroxy ethyl starch and various sugar-based cryopreservants.

Desirably the cryopreservant is at a level of at least 5% v/v (e.g. from 10-20% v/v) with respect to a composition comprising the spheroids and the cryopreservant immediately prior to cryopreservation.

It is believed that the best results can be achieved if the spheroids are cooled in a step-wise manner. Therefore in preferred methods of the present invention spheroids are cooled and then maintained within a given temperature range before being cooled further. Without being bound by theory, it is possible that this procedure may give the cells constituting a spheroid sufficient time to acclimatize to cold temperatures and thereby to avoid cell death or damage which might otherwise occur. The spheroids can be cooled to a temperature of from 1 to 8°C (e.g. from 2 to 6°C and preferably of about 4°C) and held for a period before further cooling. This period may be a period of at least 10 mins, preferably a period of at least 30 mins and typically a period of 40-60 mins. Such cooling can be conveniently achieved by using a laboratory refrigerator.

The spheroids may then be cooled to a temperature of from 0 to -50 °C (e.g. from -10 to -30°C, preferably of from -15 to -25°C) and held for a period before further cooling. This can be conveniently achieved by using a laboratory freezer. This period may be a period of at least 30 mins and is preferably of at least 1 hour

(e.g. 1-6 hours).

The final step in the cooling procedure will usually be a rapid cooling to a temperature of below -100°C e.g. of below -190°C. This can be done using liquid nitrogen as a coolant, into which is placed a resilient container containing the spheroids and the cryopreservant.

A further preferred feature of methods of the present invention (which can be used in combination with the step-wise cooling described above) is that of forming the spheroids by a period of orbital rotation at a first speed followed by a period of orbital rotation at a second speed which is higher than the first speed. Preferably the orbital rotation occurs in a generally horizontal plane.

For example the first speed may be at least 50 rpm and the second speed may be at least 1 rpm higher than the first speed (e.g. from 1 to 10 rpm higher, preferably from 1 to 5 rpm higher).

Desirably the first speed is from 50 to 90 rpm (e.g. from 60 to 80 rpm). Most desirably it is up to 76 rpm, e.g. about 75 rpm. Desirably the second speed is from 60 to 90 rpm (e.g. from 65 to 85 rpm). Most desirably it is about 77 rpm. Typically orbital rotation at each of the first and the second speeds may be for a period of at least 24 hours. The total period of orbital rotation prior to cryopreservation would generally be from 2 to 20 days. (Such a period is also suitable if, less preferably, orbital rotation at only a single orbital rotation speed is used.)

Following cryopreservation the spheroids are thawed. This can be done by placing containers containing the spheroids in a water bath at 37 °C, typically for at least 2-3 mins. Once thawed, the cryopreservation medium can be removed e.g. by centrifugation. The spheroids may then be washed and further traces of the cryopreservation medium removed (e.g. by centrifugation).

It is desirable that orbital rotation is also performed after thawing of the spheroids. This can be done at one or more rotation speeds of at least 50 rpm (e.g. of from 50 to 90 rpm). If two or more different orbital rotation speeds are used then a period of rotation at a first speed will usually be followed by a period of rotation at a second speed which is higher than the first speed.

Preferably the spheroids are rotated (post-cryopreservation) at a rotation speed of from 50 to 70 rpm (e.g. of about 60 rpm). Typically this may be done for at least

6 hours (e.g. 12-48 hours). This may be followed by a period of rotation at a higher speed (e.g. of at least 70 rpm), preferably of about 75 rpm. This may typically be for a period of at least 7-10 days or longer.

The thawed spheroids can generally be maintained in cultures post-cryopreservation for a period of around 7-10 days (longest period so far tested). Maintenance in a viable state for longer periods than this is however possible.

A typical culture medium would include a balanced salt solution and glucose. For example, DMEM:F12 (3:1) plus 10 mg transferrin/100 ml (Sigma), 0.5 mg Insulin/100 ml, 30 nM L-triiodo-thyronine can be used (DMEM is Dulbecco's Minimal Essential Medium and F12 is Hams F12 Nutrient).

Overall, cryopreservation of avian embryo whole brain spheroids was optimal using the cryoprotectant DMSO (15%), as determined by comparison of fresh spheroids and those recovered after cryopreservation; assessment relied upon a comprehensive range of parameters (morphological, histological and biochemical). The post-thaw rotation speed was shown to have a significant effect on the 'recovery' of spheroid morphology. The presence of antioxidants, e.g. vitamin E, appeared to have a beneficial effect on maintaining certain enzyme levels, e.g.

AChE, in the post-thaw spheroid. In addition, spheroid diameter pre- cryopreservation was found to be important in that smaller spheroids grown in serum-free media cryopreserved better than larger spheroids grown in serum- supplemented media.

Once the spheroids have been thawed, cryopreservant has been removed and post- cryopreservation orbital rotation has been performed, the spheroids can be used for any desired purpose.

In addition to the aspects of the invention discussed above, the present invention provides an apparatus adapted to agitate material (e.g. cells, viruses or other biological material) present within a fluid i.e. to move said material within said fluid. The apparatus comprises a self-contained power source (e.g. one or more batteries) and is preferably rechargeable.

The apparatus may be adapted to impart any desired form of motion to said material within the fluid (e.g. random motion, motion, side-to-side motion, up-and- down motion, yawing, pitching, rolling, etc. However, if, as in a preferred embodiment of the present invention, the apparams is suitable for forming spheroids, then said motion will be orbital rotation. The apparatus of the present invention is advantageous in that it can be used in a sealed chamber (e.g. an incubator) without the need for connection to an external power supply - i.e. it can be used in stand-alone form. Sealed chambers are often used to reduce or minimise contamination of a sample. They may be provided under sterile or near-sterile conditions.

The apparatus may be provided with a container and means may be provided for releasably attaching the container to the apparatus. The container is adapted to hold one or more samples that are to be processed using the apparatus. The samples may be present in culture vessels. These may be well plates, e.g. 96-well plates.

The container may be adapted to hold a plurality of such plates in stacked form.

The apparatus and container may be provided together in a kit that optionally includes instructions for mounting the container to the apparatus. The kit may also optionally include one or more culture vessels, such as well plates, one or more rechargeable batteries, and an electrical connector.

The present invention and various techniques useful therein will now be discussed by way of example only with reference to the accompanying figures, wherein:

FIGURE 1 shows the glucose consumption (corrected for Protein) before and after cryopreservation in DMSO or in DMSO + Vitamin E for chick Brain spheroids grown in a serum-free medium.

FIGURE 2 shows the Protein content of spheroids before and after cryopreservation in DMSO or DMSO + Vitamin E for chick brain spheroids grown in a serum-free medium.

FIGURE 3 shows the Acetylcholinesterase (AChE) activity (corrected for protein) before and after cryopreservation in DMSO or in DMSO + Vitamin E for chick brain spheroids grown in a serum-free medium.

FIGURE 4 shows the lactate dehydrogenase (LDH) activity/content (corrected for protein) before and after cryopreservation in DMSO or DMSO + Vitamin E for chick brain spheroids grown in a serum-free medium.

FIGURE 5 shows a comparison of different DMSO concentrations (5-15 % DMSO) on LDH leaking into the culture medium (i.e. released from spheroids) after cryopreservation and at different times after cryopreservation for chick brain spheroids cultured in a serum-free medium.

FIGURE 6 shows the glutathione (GSH) level (corrected for protein) before and after cryopreservation in DMSO or DMSO + Vitamin E for chick brain spheroids grown in a serum-free culture medium.

FIGURE 7 shows the glucose consumption (corrected for protein) before and after cryopreservation in DMSO or in DMSO + Methylcellulose for chick brain spheroids grown in a serum-supplemented culture medium.

FIGURE 8 shows the protein content of chick brain spheroids grown in a serum-supplemented culture medium following cryopreservation in DMSO or in DMSO + Methylcellulose.

FIGURE 9 shows the Acetylcholinesterase (AChE) activity (corrected for protein) before and after cryopreservation in DMSO or in DMSO + Methylcellulose for chick brain spheroids grown in serum-supplemented culture medium. FIGURE 10 shows the lactate dehydrogenase (LDH) activity/content (corrected for protein) before and after cryopreservation in DMSO or in DMSO + Methylcellulose for chick brain spheroids grown in a serum- supplemented culture medium.

FIGURE 11 shows the glutathione (GSH) content (corrected for protein) before and after cryopreservation in DMSO or in DMSO + Methylcellulose for chick brain spheroids grown in a serum-supplemented culture medium.

FIGURE 12 gives a schematic representation of the development and use of cryopreserved brain and liver spheroids for use in toxicity screening and testing and/or neuroefficacy testing of new compounds. Cells for preparing the spheroids could be derived from a number of species and cocultured with other cell types (e.g. endothelial cells for the blood-brain barrier).

Alternatively, spheroids from liver and brain prepared independently could be cocultured to investigate metabolic activation of compound. The spheroids, or cells used to reaggregate to spheroids, could be also transfected with genomic reporter-promoter constructs for toxicity/efficacy biomarkers to facilitate the screening process and event marking

'downstream'. Following cryopreservation (independently, or together as shown in figure 12) the cells would be exposed to test compounds and then assessed either using the reporter gene markers, or other assays including cytotoxicological, morphological or neurotoxicologial/neurochemical markers.

FIGURE 13 outlines a proposed methodology for identifying relevant molecular toxicity biomarkers and incorporating reporter gene constructs for these markers into spheroids. Spheroids would be exposed to the test agent, mRNA isolated and a differential mRNA display produced by reverse transcripase PCR to enable identification and cloning of the response gene(s). The promoter element would then be identified and isolated. In Step 2, the dissociated cells from the desired tissue could be transfected with a reporter-promoter construct using a variety of technologies (e.g. electroporation, adenoviral vectors, etc.) and the spheroids prepared incorporating these constructs. Following toxin/compound exposure (as Fig. 12) the response would be detected optimally by designing a 96 well spheroid culture format (e.g. 1 spheroid/ well).

FIGURE 14. The plates in this figure show spheroids from chick brain before (at 7 DIV) and after cryopreservation following 24 hours in re- culture as described in experiment 1, using 15% DMSO as the cryopreservant. The images were similar in the presence of Vitamin E or Methylcellulose additives with the DMSO.

FIGURE 15. The plates in this figure show spheroids from chick brain before (at 7 DIV) and after cryopreservation following 7 days reculture as described in Experiment 1, using 15 % DMSO as the cryopreservant. These images were similar in the presence of Vitamin E or Methylcellulose additives with the DMSO.

FIGURE 16 shows in schematic form a liver and brain spheroid co-culture.

FIGURE 17 shows a rechargeable apparams that can be used in the preparation of spheroids. EXAMPLES

1. PREPARATION AND ANALYSIS OF SPHEROIDS

Examples A to E describe the general methodology which can be used to prepare and analyze fresh chick and rat brain spheroids. A method for the production of liver spheroids and the apparatus for liver-brain spheroid co-culture is discussed in Example F below. Example G describes a rechargeable apparams that can be used in the preparation of spheroids.

A) Preparation and Maintenance of Whole Rat Brain Spheroid Cultures

Brain tissue from 15-16 day old foemses produce successful rotation-mediated cultures (Garber and Moscona, 1972). At this age cells are actively proliferating and migrating, so that cells within the spheroid cultures are capable of biochemical and morphological differentiation as in normal brains (Honegger and Richelson, 1976; Seeds et al, 1980).

Pregnant female Wistar rats are sacrificed at 15-16 days of gestation (Atterwill, 1987). The uteri are removed to a sterile laminar flow cabinet and the dissection carried out under aseptic technique. The foetuses are removed from the uteri and whole brains including diencephalon and midbrain are removed by a single cut across the top of the head and the brain scooped out into ice-cold isotonic Hanks

D2 solution containing NaCl, 138 mM; KC1, 5.4 mM; Na₂HPO₄, 0.17 mM; KH₂PO₄, 0.22 mM; CaCl₂.2H₂O, 1.8 mM; MgCl₂.6H₂O, 0.8 mM; phenol red, glucose, 0.99 mg/ml; sucrose, 14.13 mg/ml; gentamycin, 10 mg/ml. The brains are then placed in a Nybolt gauze bag (205 mm) and washed three times in Hanks

Dl (as D2 without calcium or magnesium and containing 15.91 mg/ml sucrose).

Using a flattened glass rod the tissue is gently extruded through the filter. The suspension is then filtered by gravity through a 118 mm filter, producing a single cell suspension and removing contaminating meninges. The resultant suspension is centrifuged at 150 g for 5 min. The supernatant is discarded and the cells triturated with 10 ml fresh Dl . The cells are then centrifuged and triturated once more. After the third centrifugation the cells are triturated in 10 ml growth medium (Dulbecco's modified eagles medium(DMEM) supplemented with 10% fetal calf serum, 5 ml L-glutamine and 25 mg/ml gentamycin). The cells are counted and diluted to give a density of 1 x 10⁷ cells/ml. 3.5 x 10⁷ cells are added to 25 ml capacity Delong flasks. Spheroid cultures are grown in a 9% CO₂/humidified air incubator at 37°C on a gyrotatory shaker (New Brunswick) at 85 rpm. After two days in vitro (DIV) small spheroids have formed and these are transferred to 50 ml Delong flasks and 5 ml fresh growth medium added. Medium is changed on alternate days by removing half the old and replacing it with fresh medium. At 9 DIV spheroids are transferred into 6-well plates (Nunc) in a final volume of 2 ml, except for those spheroids which are to be assayed for lipid peroxidation. Treatments are performed on 12 DIV and repeated on 14 DIV at 50

% of the original dose. Spheroids are harvested and homogenised at 15 DIV for the TBA assay and at 16 DIV for all other assays. At this time growth media (1 ml) is removed from each well for LDH analysis. This is centrifuged at 13000g prior to freezing at -20 °C. The cultures are then transferred to 1.8 ml microcentrifuge tubes and washed three times in PBS at 37, 20 and 4°C respectively to avoid heat shock. The cultures are kept on ice prior to homogenising in homogenising buffer (NaH₂PO₄, 2 mM; EDTA, 0.5 mM; NaCl, 145 mM). The suspension is then aliquoted ready for analysis. The aliquots are stored at -70°C.

B) General Preparation and Maintenance of Chick Brain Spheroid Cultures

Chick brain spheroid cultures are prepared using a procedure modified significantly from that first described by Wylegyurech and Reinhardt (1991). Fertile eggs from White-leghorn hydrid hens are incubated for seven days at 37° C in a humidified egg incubator (Curfew). Embryos at approximately stages 27-29 (Hamburger and Hamilton, 1951) are removed into isolation buffer consisting of NaCl, 128.5 mM; KC1, 5.4 mM; glucose, 5.5 mM; sucrose, 51.8 mM; HEPES, 25 mM; BSA 0.1 %; pH 7.4 (Bruinink et al, 1987) at room temperamre and washed three times in isolation buffer. Whole brains without meninges are dissected out under a microscope and pooled. Isolated brains are rinsed three times in isolation buffer and then mechanically dissociated by gentle trituration through a Pasteur pipette. The resultant cell suspension is filtered through a 22 μm Nybolt membrane held in a filter holder, as described by Bruinink et al (1987). Cells are stored on ice, to reduce clumping during counting, and the cell viability assessed by either trypan blue dye exclusion or fluorescence diacetate hydrolysis (Freshney, 1992). Cells are diluted in culture medium (various, see later) to give a final concentration of 1.33 x 10⁶ viable cells/ml and plated out into 6 well pates (Nunc) with 3 ml per well. Cultures are incubated at 37 °C in a humidified incubator with 5 % CO₂ on a gyrotatory shaker (New Brunswick) at 80 rpm. Currently cell viability varies between 70-80% with each preparation, with approximately 1.5-2.5 x 10⁷ cells per chick brain. Typically, each culture well contains between 800-1100 spheroids by 14 DIV, and one well yields 1-1.4 mg of total protein. Spheroid development is monitored by measurement of diameter using a graticule under light microscopy.

Currently, two media are in routine use for growing 'fresh' spheroids: DMEM supplemented with 10% foetal bovine serum, and a serum-free formulation of DMEM/HAMS F12 (3: 1) supplemented with Bottensteins' and Sato's N3 neural cells supplement, to give a final concentration of the following: transferrin, human holo-form 50 mg/1; insulin, bovine 5 mg/1; progesterone, 20 nM; selenium, 30 nM, and putrescine, 100 μM (Bottenstein, 1985). Serum-free media is also supplemented with 30 μM L-triiodo-thryronine (LT3) after Atterwill et al (1984). Both media are supplemented with penicillin (200 IU/ml) - streptomycin (200 μg/ml) solution (Gibco) and L-glutamine (200 mM; Gibco). Two thirds of the culture media volume is replaced every two days with fresh LT3 added at each change. Both media variants have been used to prepare spheroids for cryopreservation (see Section (2)).

Chick brain spheroid culmres are harvested as described for whole rat brain spheroids and similar assays conducted on spheroid homogenates (see later). Expression of AChE, GFAP, GS, LDH, glucose consumption and total protein are followed over a period of culmre up to 30 DIV in several media formulations. Spheroids are harvested at various intervals after inoculation, washed in PBS (pH 7.4) and samples homogenised and frozen at -70°C. Monolayer culmres are used for initial 'dose-ranging' cytotoxicity measurements, and spheroids exposed to non-cytotoxic concentrations in order to assess neurotoxicity. Spheroid homogenates/supernatants are assayed for measures of cytotoxicity and neurotoxicity (neurone- and glial-specific markers) and spheroids processed for histology (paraffin and frozen sections).

C) General Assays - Rat Brain Spheroids

In order to evaluate CNS toxicity using the spheroid culmres, markers of specific and general toxicity are required. This can be achieved using both biochemical assays (Honegger and Richelson, 1976) and histological techniques (Trapp et al, 1979). Biochemical markers for both neuronal and glial cell function have been measured. Acetylcholinesterase (AChE), which is found in cholinergic neurones has been measured to estimate neuronal toxicity (Honegger and Richelson, 1976) and it is also used as a specific marker for organophosphate toxicity (Vargese et al, 1995). Astrocyte function is assessed by the measurement of the glial specific enzyme, glutamine synthetase (GS) (Martinez-Hernandez et al, 1977) and glial fibrillary acidic protein (GFAP) which is a marker of reactive gliosis (Eng, 1985). Oxidative stress is determined by measurement of lipid peroxidation (Aust and Buege, 1978). Release of lactate dehydrogenase (LDH), a cytosolic enzyme into the media is utilised as a marker of general cytotoxicity. All results are generally normalised against total protein.

Protein Assay Total protein is assessed according to the method of Bradford (1976).

Homogenates are diluted, 1:2, and 10 μl added per well of a microtitre plate. The Coomassie blue protein reagent (Bio-Rad) is diluted 1:5 with Millipore water and filtered through Whatman No. 1 filter paper. The diluted reagent is added to the homogenate (200 μlper well) and mixed well. The plate is allowed to stand for 5 min at room temperamre prior to recording the absorbance at 595 nm.

Acetylcholinesterase Assay (AChE)

AChE activity is measured according to the method of Ellman et al (1961). 100 μl of sample is placed in each well of a 96- well plate with 100 μl sodium phosphate buffer (0.1 M, pH 7.4,), 50 μl 5,5'-dithiobis-2-nitrobenzoate (DTNB)

(2 mg/ml in methanol,) and 25 μl acetylthiocholine iodide (2.5 mg/ml in buffer,). The resultant absorbance is monitored at 405 nm for 15 min, at 10 sec intervals, on a Biotek EL312 kinetic plate reader. Activity is calculated using a standard curve of pure acetylcholinesterase (AChE) from electric eel (Sigma). One unit of AChE can hydrolyse 1 mmole of acetylcholine to choline and acetate per minute at pH 8.0 and 37°C.

Glutamine Synthetase Assay (GS)

GS is assayed using the transf erase site of the enzyme, as reported by Thorndyke and Rief-Lehrer (1976).

Homogenate (80 μl) is transferred to a microcentrifuge tube and 330 μl reagent mix containing 0.4 M L-glutamine; 1.0 M hydroxylamine; 395.0 mM potassium arsenate; 823.0 mM sodium citrate; 82.3 mM manganese chloride; 4.0 mM adenosine diphosphate was added. The tubes are mixed and then incubated in a shaking water bath at 37°C for 15 mins. The reaction is terminated on ice by the addition of 400 μl FeCl₃ solution (1: 1: 1 mix of 7% FeCl₃ in 5 mM HC1: 15% TCA: 0.1 M HC1) . The mixtures are then centrifuged at 13000 g and absorbance at 500 nm measured. Results are compared against a standard of purified GS from sheep brain. One unit of GS is defined as that which will convert 1.0 mmole of

L-glutamate to L-glutamine in 15 min at pH 7.1 and 37°C.

Glial Fibrillary Acidic Protein Assay (GFAP)

GFAP is estimated using an adaptation of O'Callaghan's (1991) method. Immulon-2 plates are coated overnight at 4°C with mouse monoclonal GFAP antibody (1:1000). Excess antibody is removed by washing with PBS. All subsequent incubations are carried out at room temperamre on a rotatory shaker. Non-specific binding to the plastic is prevented by incubation with 5 % milk powder in PBS. This is washed off with PBS, and a 100 μl sample or standard added and left to incubate for two hours. Samples are centrifuged (13000 g) and diluted 1:2 before use. The plates are then washed with PBS-TWEEN (PBS and 0.05% TWEEN-20). Rabbit-derived GFAP antiserum, diluted 1:500 in BLOTTO (5% milk powder in PBS-TWEEN) is then added and left to incubate for 1 hr. After washing with PBS-TWEEN, anti-rabbit immunoglobulins conjugated to alkaline phosphatase are added, diluted 1:3000 in BLOTTO, for 1 hr. The wash step is repeated and finally 1 mg/ml p-nitrophenyl phosphate in 10% diethanolaminebuffer, the substrate for alkaline phosphatase, is added. This substrate is converted to p-nitrophenol, a yellow adduct which has an absorbance maxim at 405 nm, the absorbance being proportional to the concentration of GFAP in the sample.

Thiobarbituric Acid (TBA) Assay

Lipid peroxidation is assessed using the thiobarbituric acid (TBA) assay (Aust and

Buege, 1978). At 15 DIV spheroid culmres growing in Delong flasks are transferred to glass test tubes. The spheroids are washed three times in PBS and homogenised in 1 ml of homogenising buffer as before. 50 μl of the resulting suspension is removed and frozen (-20°C) for protein analysis. TBA reagent (2 ml) containing 26 mM TBA in 0.25 mM HC1 and 15% trichloroacetic acid is added to each sample homogenate, followed by incubation in a boiling water bath for 15 mins. The homogenates are then centrifuged at 150 g for 5 mins to remove any proteins and the absorbance of the resultant pink chromophore monitored at 532 nm. A standard curve of mM concentrations MDA (malondialdehyde bis (diethyl acetal)) against resultant absorbance is used to standardise results.

Lactate Dehydrogenase Assay (LDH)

Samples of homogenate and media are analysed for LDH, which is expressed as LDH released from spheroids compared to total cellular LDH according to the method of Korzeniewski and Callewaert (1983). Samples (100 μl per well) are added to 96-well plates along with 100 μl "LDH buffer" (0.2 M Tris buffer, pH 8.2; L(+) lactate, 54 mM; β NAD, 1.3 mM; phenazine methosulphate,

0.28 mM; p-iodonitrotetrazolium violet, 0.66 mM). The resultant absorbance at 490 nm is monitored for 15 min, at 10 sec intervals on a Biotek EL312 kinetic plate reader.

D) General Assays - Chick Brain Spheroids

Neuropathy Target Esterase (NTE)

The assay for NTE in chick brain spheroids is based on the methods of Johnson

(1977) and Correll and Enrich (1991) and measures the release of phenol, following the hydrolysis of phenyl valerate by NTE in the presence and absence of the specific NTE inhibitor, mipafox. Phenol release by hydrolysis is complexed with 4-aminoantipyrine to give an orange reaction product which is read spectrophotmetrically at wavelength 500 nm.

Chick brain spheroids are harvested as described previously, washed three times in phosphate buffered saline (PBS, pH 7.4) and resuspended in 0.5 ml of Tris/EDTA buffer (Tris, 50 mM; EDTA, 0.2 mM; pH 8). Spheroids are then homogenised in a motorised Potter homogeniser (10 strokes at 800 rpm), and total protein measured using the Biorad Coomasie blue assay with a gamma-globulin standard (Bradford, 1976), and then diluted to give a final concentration of 0.3 mg/ml protein in Tris/EDTA. Triplicate 50 μl aliquots of homogenate are plated into 96 well plates (Nunc) and incubated with either paraoxon (40 μM), paraoxon (40 μM) with mipafox (50 μM), or buffer (tissue blank). Plates are mixed using a Labsystems Multiscan RC plate reader for 30 sec and incubated for 20 min at 37 °C. Following incubation, 50 μl of a 1.06 mg/ml suspension of phenyl valerate in 0.03 % Triton X100 in water is added to wells containing mipafox and/or paraoxon and 50 μl of Triton XI 00 to tissue blanks. Plates are mixed again and incubated for a further 15 min at 37°C. The reaction is halted by the addition of 50 μl of 0.04% 4-amino-antipyrine/10%SDS in Tris buffer (0.5 M, pH 9.6). Liberated phenol is determined by the addition of 50 μl of an aqueous solution of

0.4% potassium ferricyanide. Plates are mixed for 1 min and incubated for 15 min at room temperamre to allow colour development and then read in a plate reader at 492 nm (Kayali et al, 1991). The concentration of phenol released is determined against a standard curve (0-100 μM phenol), run concurrently; appropriate tissue and reagent blanks are also included in the assay plate.

NTE activity in the sample homogenates of chick brain spheroids is expressed as μmoles of phenol released per minute per mg of protein, and is determined by subtracting the phenol concentration of tests containing mipafox and paraoxon from those containing paraoxon alone.

Glucose Consumption

Glucose in culture media is the major energy source for the spheroids and glucose consumption generally reflects the metabolic status and viability of the culmres. Glucose is measured in spheroid culture supernatant using a glucose kit (Sigma) based on an enzymic assay. The method is modified in our laboratories for use in a 96-well plate format. A 100 μl aliquot of media is taken at the end of a defined period of incubation (usually 24 hours) and diluted 1 :50 in distilled water. A 50 μl sample of diluted culture media or glucose standards are added to a 96 well plate and 250 μl of colour reagent added to each well and incubated at 37 °C for 30 min. Plates are read spectrophotometrically at wavelength 450 nm. In the chick brain spheroid culmres, single cells grow together to form three-dimensional spheroids and during this reorganization some cells locate in the inner part of the spheroid while others form the outer surface. The geographical isolation of the inner cells from the culture medium means that there will be a gradient of oxygen consumption across the spheroid (Breedel-Geissler et al, 1992), which would suggest that energy metabolism in the inner part is lower than that in the outer cells. Variations on this procedure for cryopreserved spheroid assessment is given in the next section.

E) General in vitro Histopathologv & Immunocvtochemistrv

Histological Preparation of Rat Brain Spheroid Cultures

13 DIV spheroids are transferred to a glass test tube and washed three times with phosphate buffered saline comprising (PBS) 137 mM NaCl; 2.7 mM KC1; 3.2 mM

Na₂HPO₄ and 1.3 mM NaH₂PO₄. Spheroids are fixed in 2 ml formal saline (10% formalin in 0.9% NaCl) for 30 min. The spheroids are then dehydrated by exposure to increasing concentrations of ethanol (70%, 90% and absolute). The absolute ethanol incubation is repeated to remove all traces of water and is then replaced with xylene, which enables wax to be added without emulsions forming.

The paraffin wax is kept molten at 60°C. The spheroids are washed in wax three times, then fresh wax is added, and transferred to a mould and allowed to set overnight. Sections of 7 μm are cut and fixed onto subbed slides. General Histological Preparation of Chick Brain Spheroids

Spheroids are harvested and transferred to 1.5 ml Eppendorf tubes, washed twice in PBS and resuspended in 1 ml phosphate buffered formalin (PBF) and left to fix at room temperamre overnight. Fixed spheroids are washed in PBS and resuspended in 0.5-1 ml molten agarose (1%). When the agarose has solidified, the plugs are removed from the tubes and cut into transverse sections, approximately 5 mm wide. These are fixed overnight in PBF and then processed in a Shandon tissue processor, dehydrated, cleared in xylene and embedded in paraffin wax. Sections are cut at 9 and 5 μm using a Shandon rotary microtome and attached to gelatin- or poly-D-lysine-coated slides. Sections are stained with either haematoxylin and eosin or immunohistochemically stained as described for rat brain spheroids.

Haematoxylin and Eosin Staining Slides are rehydrated by incubating with xylene, absolute alcohol, 90% and 70% alcohol, followed by PBS. Ehrlichs haematoxylin, which stains negatively charged chromatin within the nuclei, is added to each slide and left for 10 min. Excess haematoxylin is washed away with distilled water until the section turns blue. The slide is then exposed to the counterstain eosin, for 5 min. Excess stain is removed with 70% alcohol, followed by absolute alcohol and xylene prior to mounting in

DPX.

Immunostaining for GFAP and Neurofilament Protein

Slides are rehydrated as described previously for haematoxylin/eosin staining. H₂O₂ in methanol is added to the section to remove any residual peroxidase activity and 5% horse serum in PBS is added to prevent non-specific binding of antibodies. The section is then exposed to the primary antibody for 1 hr, at a dilution of 1 :400 of mouse monoclonal antibody for GFAP or 1:8 of the mouse antibody for neurofilament protein 160 (NFil). The antibodies are washed off with PBS. The binding of the primary antibody is identified using an ABC Ultrasensitive Anti-mouse Immunoglobin kit (Pierce). The slides are incubated for 30 min with a 1:400 dilution of biotinylated anti-mouse immunoglobulin. The slides are then incubated with a solution of avidin and biotinylated horseradish peroxidase for 30 min. The slides are again washed, and the substrate, 3,3' diaminobenzidine tetrachloride/urea added to form a brown precipitate. The slides are then dehydrated with 70%, 90%, absolute ethanol and xylene and embedded in DPX.

F) Production of Liver Spheroid Cultures and Co-Culture with Brain Spheroids

The following sections (A) and (B) describe the method for producing liver spheroids from rat liver material, and the co-culture technology for brain and liver spheroids. Modifications of this technology can be used for liver spheroids from other species prior to cryopreservations.

(A) Isolation of Rat Hepatocytes by CoIIagenase Perfusion

Preparation of rig

A standard 'hepatocyte isolation rig' is necessary. This basically consists of three flasks connected, via a peristaltic pump, to a platform with two cannulae.

Prior to liver collection, the flasks and tubing are rinsed with 70% IMS followed by sterile 'perfusion buffer'. The water bath is also switched on.

The rig is loaded with 500 ml 'chelating buffer' in flask 1, 250 ml 'perfusion buffer' in flask 2 and 100 ml 'collagenase buffer' in flask 3 (see appendix). All tubes are primed and the whole system finally primed with 'chelating buffer' .

Gas (95 % air, 5% CO₂) is gently bubbled through all three flasks. The liver is now ready for collection. Collection of the liver

A Male rat is killed by cervical dislocation. The abdomen is washed, and a long horizontal incision is made just below the rib cage. The liver is cut free from the connective tissue whilst being held over a beaker containing 'perfusion buffer' (see appendix).

Perfusion of the liver

The two largest lobes of the liver are taken and the cannulae inserted into the open, cut blood vessels. 'Chelating buffer' is allowed to flow (5 ml/min/cannula) and an almost immediate clearing of blood from the lobe should be seen. This is allowed to run for 15 mins. The tap is changed to allow the 'perfusion buffer' to flow, again for a further 15 mins. Nearing the end of this period the collagenase (see appendix) is dissolved in a little 'collagenase buffer' and returned to flask 3. The 'collagenase buffer' is allowed to perfuse for 30 mins, with recycling of the buffer by placing the waste mbe in flask 3. After this time the liver should appear spongy and digested, if not leave it for longer. The lobes are removed into sterile petri dishes.

The remainder of the procedure is performed in a laminar flow cabinet.

Dulbecco's Modified Essential Media (see appendix) is added to the lobes in separate dishes. The liver is teased apart with forceps and a spatula and the cells should flow into the media. Do not dissociate any badly digested parts of the liver. The cell suspension is poured through a 63 μm Nybolt filter into a centrifuge mbe and the petri dish washed. The cell suspension is centrifuged at

300 rpm for 3 mins. The supernatant is discarded and the pellet gently resuspended in 10 ml media. The centrifugation step is repeated and the pellet resuspended in media to give a final volume of 10 ml.

Hepatocyte viability is assessed by trypan blue exclusion. A 1:5 dilution of the hepatocyte suspension is usually necessary, followed by the addition of 200 μl trypan blue (0.4%) to 200 μl cell suspension.

Viable cells remain clear whilst dead cells stain blue. A viability of 80-85% is expected, below 70% should be discarded. There is usually a difference between the two lobes and the best viability suspension is used.

Cell count (cells/ml) = no. cells counted x dilution factor (10)x 10⁴.

From this cells can be plated out at the required density.

Appendix - Reagents

Perfusion buffer: 435 ml sterile 'millipore water' 50 ml Earles balanced salt solution xlO (without Ca ⁺,

Mg²⁺, phenol red) 15 ml sodium bicarbonate (7.5%) pH adjusted to 7.4, using sterile 1M HC1 (around 500 μl) Need to make 3x500 ml bottles for each perfusion.

Chelating buffer: 490 ml perfusion buffer

10 ml EGTA (25 mM, in 0.1 M NaOH, pH adjusted to 7.4 with 1M HC1)

Collagenase buffer: 100 ml perfusion buffer

200 μl calcium chloride (1M, in water) x mg collagenase A (Boehringer Mannheim)

where x = (0.12U/ml x 100)/specific activity (this is usually around 25-30 mg collagenase) Media: 425 ml sterile millipore water

50 ml Dulbecco's Modified Essential Media, xlO

25 ml sodium bicarbonate (7.5%)

2 ml foetal calf serum 5 ml L-glutamine (300 μg/ml)

.25 ml gentamycin (25 μg/ml)

pH adjusted with 1M NaOH until a deep cherry red colour

(B) Liver Spheroid Culture

Preparation of the plates

The plates need to be coated with poly(-2-hydroxyethylmethylacrylate) (p-HEMA) (Sigma) to prevent the cells adhering to the plastic.

A 2.5% w/v solution of p-HEMA is made in 95% absolute ethanol. This needs to be left to stir on a gently heated hot plate (50 °C) and will take around 3 hrs to dissolve.

6-well tissue culmre plates are coated by adding 2ml p-HEMA solution to each well and allowing them to air dry in a class II safety cabinet. Once dry (approx. 3hrs) these can be stored at 4°C.

Culmre procedure An isolated hepatocyte cell suspension is obtained by a collagenase perfusion method. Cells are plated at a density of between 2.5-5.0 x 10⁵ cells/ml (still under investigation), at 2 ml suspension per well of a 6-well p-HEMA coated plate.

Culmres are maintained in an incubator at 37 °C, 9% CO₂ and are gently rotated at between 85-90 rpm. Culture media 425 ml sterile millipore water

50 ml Dulbecco's Modified Essential Media, xlO

25 ml sodium bicarbonate (7.5%)

2 ml foetal calf serum 5 ml L-glutamine (300 μg/ml)

.25 ml gentamycin (25 μg/ml)

pH adjusted with 1M NaOH until a deep cherry red colour

This can be supplemented with insulin (50 ng/ml) and dexamethasone (5 ng/ml) for maintenance of hepatocyte function.

Sampling

The culmres are sampled for viability and functional assays. Two random wells are combined to give one sample. 500 μl of media is removed from each well and combined, microfuged and the supernatant frozen at -80°C for LDH and albumin assay. The spheroids are transferred from the well into a 1.5 ml eppendorf. The well is washed and the washings transferred. The spheroids are allowed to settle and the supernatant removed (checking for tissue loss under the microscope). The contents of one well are resuspended in 1 ml 'homogenising buffer' and this is transferred to the contents of the second well. The spheroids are allowed to settle again, the supernatant removed and finally resuspended in 1 ml homogenising buffer. The culture samples are sonicated, to disrupt the cells, with a probe for 10 sees and stored on ice. 200 μl aliquots of the samples are stored at -80°C until assay. Homogenising buffer - see rat brain spheroid culmre method.

G) Rechargeable Apparatus

A rechargeable apparams that can be used in the preparation of spheroids is within the scope of the present invention. It is shown in Figure 17 and is discussed below.

Figure 17 shows an apparams (1) that is adapted to allow orbital rotation of cells suspended in a fluid. Desirably this rotation occurs in a generally horizontal plane. However the apparams (1) may also be adapted to allow orbital rotation in other planes and may even be adapted to allow other forms of movement (e.g. side-to-side shaking, rolling, pitching, yawing, etc). This can be advantageous if the apparams (1) is to be used for other purposes than for forming spheroids.

The apparams (1) is provided with a control panel (6). This allows the rate of orbital rotation of cells to be maintained within a desired r.p.m. range (preferably 50-90 r.p.m.). An indicator (3) is also provided which allows r.p.m. values to be displayed.

A removable plate (5) is shown, behind which a rechargeable power source (e.g. one or more rechargeable batteries) is located. A port (4) is provided to allow a detachable electric cable to be attached to the apparatus (1) so that the power source can be recharged. When the apparams (1) is recharged the cable can be removed. The apparams (1) can therefore be used in stand-alone form (i.e. without the need for it to be connected to an external power source). This is advantageous in that the apparams (1) can be used in a sealed chamber that need not itself be provided with a power source. Such chambers are often used in order to minimise contamination of a sample. They may be maintained under sterile or near-sterile conditions.

A container (2) may be mounted to the apparams (1) via releasable securing means (not shown). The container (2) adapted to contain one or more culmre plates. A single 96 well culmre plate (8) is shown, but a plurality of such plates may be stacked in the container (2). The container (2) is mounted onto a platform of the apparams (2). The platform is operatively connected to a motor (not shown) to allow the platform to be moved in a desired manner. Controlled movement of the platform is used to impart movement to cells in suspension so that spheroids can be formed. Thus orbital rotation of cells can be imparted via orbital rotation of the platform.

The apparams includes a plurality of legs (7), which may be provided in a form so that their heights can be adjusted. This is advantageous if it is desired for the platform (not shown) to be generally horizontal since this can be achieved by adjusting the legs (7) accordingly.

2. CRYOPRESERVATION METHODOLOGY- SPECIFIC PREPARATION OF CRYOPRESERVED CHICK WHOLE BRAIN SPHEROIDS

The following describes how cryopreserved chick brain spheroids can be prepared taking as a starting point the general methodology described in the following examples.

2A. PROTOCOL FOR CHICKEN BRAIN SPHEROID INCUBATION AND CRYOPRESERVATION WITH SERUM FREE MEDIUM

A) Chick Brain Cell Preparation and Culture

Chicken eggs were incubated at 37°C in a water-saturated environment for 7 days. Foemses were removed from eggs and transferred into a dish containing isolation buffer (sterilised), and the foemses washed twice with isolation buffer. The whole brains of foemses were dissected out under stereo-microscope. The brain was washed three times with isolation buffer and dispersed into single cells by repeating drawing up into a pasteur pipette and then diluted with isolation buffer into 10ml cell suspension. The suspension was filtered by gravity through a filter made of Nybolt gauze (40 μm). The cells were counted and diluted with incubation medium to give a density of 1.33 x 106 cell/ml; 3ml diluted suspension was added to each well of 6-well plates. The incubation medium contains 300 ml Dulbeccos Modified Eagle's Medium (DMEM) x 1, 100 ml F12 (Ham), 20 mg transferrin, 2 mg insulin, 1.6 nmol progesterone, 40 μmol putrescine, 1.2 nmol selenium, 48 μg triiodothyronine and 5 ml penicillin and streptomycin. The plates were then placed on a shaker in 37°C 5% CO₂ incubator. The rotation speed of shaker is adjusted to 75 rpm for first 4 days and then to 77 rpm thereafter.

B) Cryopreservation

Seven day old spheroids in vitro (7DIV) in each well were transferred into a Cryovial. The vials were centrifuged at 800 rpm for 1 min. The medium is removed with a pipette and then 1 ml cryopreservative medium containing 15 % DMSO in medium either with or without 40 μM Vitamin E at 4°C is added. The vials are kept at this temperamre for 40-60 min and then transported to a -20°C freezer. After 2 hour freezing at -20°C, the vials are plunged into liquid nitrogen (-196°C).

C) Recovery

After 24 hour or longer storage in liquid nitrogen, the vials are taken out from liquid nitrogen and thawed at 37 °C in a water bath for 2-3 min. They are centrifuged at 800 rpm for 1 min and the cryopreservative medium removed.

Spheroids are washed once with 1 ml incubation medium and last step repeated. Then transferred to 6-well plate with 3 ml medium/vial/well. The plates are shaken at 60 rpm in an incubator at the standard incubation condition. The incubation is terminated at either 24 hours or 3 or 7 days after recovery from freezing for biochemical assays.

D) Assays

I) Glucose Consumption

Glucose assay is carried out by using Glucose Kit (Sigma, Catalog No. 510- A) which is based on Keston's enzymatic method (Keston, 1956). The method was modified in our laboratory to be suitable for 96-well Plate Reader assay which is quicker and more economic than the conventional method. Medium of 100 μl/well is taken at the end of a defined period of incubation (usually 24 hours) and diluted with distilled water (1:50). A blank control well must be used as the reference value for consumption calculation. Add 50 μl glucose standard serials or diluted medium to each well of 96-well Plate and then add 250 μl colour reagent solution (1 capsule of PGO enzyme dissolved in 100 ml water plus 1.6 ml o-dianisidine dihydrochloride). Each sample is run in duplicate. Incubate the plate at 37°C for 30 min to allow colour development. The concentration of glucose is determined by reading absorbance on a Plate Reader at 450 nm.

II) Protein Assay

Spheroids were homogenated in phosphate buffer (pH 7.4). Total protein was assayed according to the method of Bradford (1976). 10 μl homogenates were added per well of 96-well plate. The protein reagent (Bio-Rad) was diluted 1:5 with millipore water and filtered through Whatman No, 1 filter paper. The diluted reagent (200 μl per well) was added to the homogenate and mixed well. The plate was allowed to stand for 5 min at room temperamre prior to recording the absorbency at 595 nm.

HI) Acetylcholinesterase (AChE) Assay

AChE activity was measured according to the method of Ellman et al (1961). 100 μl of homogenate was placed in each well of a 96-well plate with 100 μl sodium phosphate buffer (0.1 M, pH7.4), 50 μl 5,5' -dithiobis-2-nitrobenzoate (DTNB) (2 mg/ml in methanol) and 25 μl acetylthiocholine iodide (2.5 mg/ml in sodium phosphate buffer). The resultant absorbency was monitored at 405 nm for 15 min, at 10 sec. intervals, on a Biotek EL312 kinetic plate reader. Activity was calculated using a standard curve of pure acetylcholinesterase from electric eel (Sigma). One unit of AChE can hydrolyse 1 μmole of acetylcholine to choline and acetate per minute at pH 8.0 and 37 °C.

TV) Lactate Dehydrogenase (LDH) Assay

Homogenate and medium samples were both analysed for LDH which was expressed as LDH released from spheroids compared to total cellular LDH according to the method of Korzeniewski and Callewaert (1983). Samples (100 μl per well) were added to 96-well plates along with 100 μl LDH buffer (0.2 M Tris buffer, pH8.2; L(+) lactate, 54 mM; 0-NAD, 1.3 mM; phenazine methosulphate, 0.28mM; piodonitrotetrazolium, 0.66 mM). The resultant absorbence at 490 nm was monitored for 15 min, at 10 sec. intervals on a kinetic plate reader.

V) Glutathione Assay

100 μl homogenate was diluted with equal volume of 13 % trichloroacetic acid (TCA). Vortex mixing and then centrifuged at 13,000 rpm for 3 min. 75 μl supernatant or standard was added to 2775 μl of phosphate/EDTA buffer (1 litre containing 13.6 g KH2PO4, 1.86 g EDTA, pH is adjusted to 8 with NaOH) and 150 μl of 0.1 % o-phthaldehyde (OPT) in methanol was added. The samples were mixed and kept at room temperamre for 30 min. They were read on a fluorimeter at 350 nm exitation and 420 nm emission.

2B. PROTOCOL FOR CHICKEN BRAIN SPHEROID INCUBATION AND

CRYOPRESERVATION WITH MEDIUM CONTAINING SERUM

I) Brain Cell Preparation

See Protocol 2A

II) Brain Spheroid Culture

The cells were counted and diluted with incubation medium to give a density of 1.33 x 10⁶ cell/ml. The incubation medium contains 10% foetal calf serum in the mixture of Dulbeccos Modified Eagle's Medium (DMEM) x 1 and F12 (Ham) (3: 1) and 1 ml penicillin and streptomycin per 100 ml medium. 3 ml diluted suspension was added to each well of 6-well plates. The plates were then placed on a shaker in 37°C 5 % CO₂ incubator. The rotation speed of the shaker is adjusted to 75 rpm for the first 4 days and then to 77 rpm thereafter. III) Cryopreservation

Seven day old spheroids in vitro (7DIV) in each well were transferred into a Cryovial. The vials were centrifuged at 800 rpm for 1 min. The medium is carefully removed with a pipette and 1 ml cryopreservative medium containing 15 % DMSO with or without 0.1 % methylcellulose (SIGMA) in

10% CFS medium at 4°C is added. The vials are kept at this temperature for 40-60 min and then transported to a -20 °C freezer. After 2 hour freezing at -20 °C, the vials are plunged into liquid nitrogen (-196°C).

VI) Recovery

After 24 hour or longer storage in liquid nitrogen, the vials are taken out from liquid nitrogen and thawed at 37 °C in a water bath for 2-3 min. Then they are centrifuged at 800 rpm for 1 min and the cryopreservative medium removed. Spheroids are washed once with 1 ml incubation medium and the last step repeated. Then spheroids are transferred to 6-well plate with 3 ml medium/vial/well. The plates are shaken at 60 rpm in an incubator for 24 hour and then adjusted to 75 rpm until the termination of the experiment. The incubation was terminated at either 24 hours or 3 or 7 days after recovery from freezing for biochemical assays.

3. RESULTS/DATA OBTAINED TO DATE:- CRYOPRESERVED CHICK BRAIN SPHEROIDS

The following describes two key smdies which are the culmination of a complete study series investigating optimal conditions for cryopreservation: -

A) EXPERIMENT ONE To evaluate DMSO at 15% (+/- Vitamin E) in a cryopreservation medium using spheroids prepared in the normal serum-free based incubation medium.

I) Procedures

7-day in vitro spheroids in 15% DMSO (with or without added Vitamin E (40 μm) were cryopreserved in liquid nitrogen by stepwise cooling (4°C for 40-60 min = 20 °C for 2 hr =» liquid nitrogen, -196°C). The spheroids were thawed in 37 °C water bath for 3 min after 2 days frozen and re-plated. Various assays (see below) were carried out at 24 hr, 3 days and 7 days after recovery.

II) Evaluation Parameters

Morphology (Light Phase Microscopy -(Plate 1) 24 hr glucose consumption

Protein content

AChE

LDH content of cells

LDH-leaking into culmre medium Glutathione (GSH)

GFAP

77/) Results

See Figs 1-6 and Table 1. The results depicted in Figs 1-6 (and Table 1) show the viability of chick brain spheroids grown initially in a serum-free based medium at -43- various timepoints after cryopreservation (as described in Methods) using either DMSO alone, or DMSO + Vitamin E. All parameters were corrected for protein content. Fig. 2 shows that the cryopreservation procedure reduced protein content to about 20% of original values indicate loss of a large number of the spheroids by the cryopreservation process. This was backed by a reduced lactate dehydrogenase (a soluble cytoplasmic marker) (LDH) spheroid content (Fig 4.). However, as Fig. 1 indicates, glucose consumption values for the surviving spheroids was good, being maintained at 'pre-values' at 24 hours after cryopreservation and reduced only by around 20% at 3 days and 7 days after. The presence of Vitamin E did not significantly improve this viability. These glucose consumption results were paralleled by the glutathione (GSH) data (Fig. 6) where no reductions in GSH at any timepoint were found after cryopreservation indicating normal/maintained cellular oxidative protectant levels. Similarly, the acetylcholinesterase (AChE) activity (as a neuronal marker) was maintained after cryopreservation with the presence of Vitamin E with DMSO slightly improving this parameter at 7 days after the cryopreservation procedure.

TV) Conclusion

1. So far 15% DMSO in the cryopreservation medium was the optimal cryopreservant concentration to date for chicken brain spheroids with values close to or approaching pre-cryopreservation controls for most parameters.

2. The orbital shaking speed has a significant influence on both spheroid growth or recovery from the cryopreservation state. The orbital shaking speed at 75 rpm for first 4 days and then 77 rpm before freezing achieved a better formation of spheroids. 60 rpm after recovery significantly reduced broken spheroids.

3. Although the absolute values of AChE and GSH were reduced, there was no significant reduction in values after freezing if their values were corrected for protein content. This may indicate that cholinergic neurons -44- may equally survive frozen as other neural cells except perhaps after longer time periods where at 7 days after freezing AChE was reduced. 4. Vitamin E provided added protective effect on cryopreserved spheroids in the case of AChE, which was greater with Vitamin E present. 5. Glucose consumption, protein content and GSH are relatively stable parameters in the evaluation of spheroid cryopreservation.

B_l EXPERIMENT TWO To evaluate the effect of using an incubation medium containing serum and the effects of Methylcellulose added to the cryopreservation medium on the viability of chick brain spheroids.

7) Procedures:- Brain spheroid culmre with 10% foetal calf-based serum medium

The cells were counted and diluted with incubation medium to give a density of 1.33 x 10⁶ cell/ml. The incubation medium contained 10% foetal calf serum in the mixmre of Dulbeccos Modified Eagle's Medium (DMEM) x 1 and F12 (Ham) (3:1) and 1 ml penicillin and streptomycin per 100 ml medium. 3 ml diluted suspension was added to each well of 6-well plates. The plates were then placed on a shaker in 37 °C 5% CO₂ incubator. The rotation speed of the shaker is adjusted to 75 rpm for 4 days and then to 77 rpm for 7 days.

Cryopreservation Seven day old spheroids in vitro (7DIV) in each well were transferred into a

Cryovial. The vials were centrifuged at 800 rpm for 1 min. The medium was carefully removed and then 1 ml cryopreservative medium added containing 15% DMSO and 0.1% methylcellulose (SIGMA) in 10% CFS medium at 4°C. The vials were kept at this temperamre for 40-60 min and then transported to a -20 °C freezer. After 2 hours freezing at -20° C, the vials were plunged into liquid nitrogen (-196°C).

Recovery

After 24 hours or longer storage in liquid nitrogen, the vials were taken out from liquid nitrogen and thawed at 37° C in a water bath for 2-3 min. Then they were centrifuged at 800 rpm for 1 min and the cryopreservative medium removed. They were washed once with 1 ml incubation medium and the last step repeated. They were then transferred to a 6-well plate with 3 ml medium/vial/ well. The plates were shaken at 60 rpm in incubator for 24 hours and then adjusted to 75 rpm until termination of the experiment. The incubation was terminated at either

24 hours or 3 after recovery from freezing for biochemical assays and evaluation parameters.

II) Evaluation Parameters As experiment one.

777) Results

See Figs 7-11 and Table 2. The results depicted in Figs 7-11 describe the viability of chick brain spheroids at various timepoints after cryopreservation following pre- cryopreservation culmre in the serum-based culmre medium and using either DMSO alone (15%) or DMSO + Methylcellulose (0.1%) as the cryopreservants. Furthermore, in this experiment the spheroids were rotated at 60 rpm for the initial 24 h following storage and then 75 rpm until experiment termination. All parameters were corrected for protein values.

Compared with the pre-cryopreservation values using this methodology, the glucose consumption parameter was approximately halved by the cryopreservation procedure and slightly improved by DMSO and methylcellulose at the 24 h post- cryopreservation timepoint. Although the absolute pre-cryopreservation value for this parameter was higher in this experiment than in Experiment 1 (presumably due to the improved maturity of spheroids due to serum addition), the reduction in viability as judged by this parameter was lower by the cryopreservation procedure possibly due to a more resricted access of cryopreservant into the spheroids due to their enhanced size in the serum-based culmre enviroment.

In parallel, Fig. 11 shows that glutathione (total GSH) content was not adversely markedly affected by the cryopreservation procedure at 24h and indicates a satisfactory cellular endogenous oxidant protectant level i.e. low oxidant stress level in the spheroids. At 3 days after the procedure a 50% pre-cryopreservation

GHS level was maintained. Protein content (Fig. 8) indicates that around 50% of the spheroids 'seeded' in the initial culmre were lost due to the cryopreservation procedure but of that 50% surviving, the other parameters indicate 50-75% viability overall by this culmre procedure. In line with this, the acetylcholinesterase activity (Fig. 9) was maintained at around 50% the pre- value showing that, in terms of a specific neuronal (as opposed to glial) marker, neuronal viability is also still fairly good.

The LDH activity, however, (i.e. cellular soluble enzyme content) of the cryopreserved spheroids was reduced to approximately 20% of original values in

'serum-grown' culmres by cryopreservation which, along with protein reductions, supports the notion that perhaps the larger spheroids grown in the presence of serum (compared with smaller diameter spheroids in a serum-free culmre - Figs. 1-6) do allow restricted access to the cryopreservant chemicals and thus final spheroid viability post-cryopreservation.

(Table II data relates to Figs. 7-11). IV) Conclusions

1. Chick brain spheroids grew faster and have a greater size/diameter in serum-based medium than in serum-free medium.

2. Rotation speed significantly influenced the size and homogenicity of these spheroids in the first 4 days of cell culmre.

3. The absolute values of all parameters significantly increased compared with spheroids cultured with serum-free medium, but post-cryopreservation viability was not as good as in the serum-free grown spheroids probably due to size/cry opreservant 'access' to spheroid mass.

GENERAL CONCLUSIONS

It is concluded that the cryopreservation procedure for chick brain spheroids using the technology given in experiment one and employing 15% DMSO is optimal for cryopreserving these cells given the available information to date. Spheroids grown in the serum-free based culmre medium lend themselves better to cryopreservation than those grown in a serum-supplemented medium and exhibit a good, intact morphological appearance by phase-contract light microscopy up to 7 days after the cryopreservation procedure (see Plate I). Vitamin E and Methylcellulose may have a beneficial/protective effect on these cryopreserved spheroids.

DEFINITIONS

1. For the purposes of this invention the term "spheroid" is taken to mean a three-dimensional strucmre which does not occur in namre and which consists of a reaggregate of cells of a tissue or of an organ.

A spheroid will preferably be generally spherical in shape. Desirably it will consist of at least 10³-10⁴ cells, more desirably of at least 10³ cells but this may vary according to species and rotation speed and tissue 'type' . Typically a spheroid will have a diameter of between 100 microns and 1000 microns. However it may be larger/ smaller than this. It may be made up of one or more different cell types. If a plurality of different cell types are present they may form different layers of the spheroid.

2. The term cryopreserving is taken to mean taking the spheroid(s) or cellular aggregate/re-aggregate strucmre down to a cryogenic temperamre (typically -196°C) where the biological sample is held in a state of 'suspended animation' in the presence of a cryopreservant (cryoprotectant), or several cryopreservant (cryoprotectant) chemicals in order to hold the material at this temperamre for prolonged periods (many months, and up to several years) maintaining significant cellular integrity and functionality during the cryopreservation process and upon subsequent retrieval after thawing and return to ambient and physiological temperatures (typically 37°C).

3. The term "tissue" is taken to mean an organised selection of cells having a common function. The term "organ" is taken to mean an organised collection of "tissues" having a common function.

The "tissue" or "organ" need not be completely intact to be used in the present invention since parts of whole tissues or organs (which may be obtained via biopsies) can be disrupted to individual cells/small groups of cells before being re-aggregated to form spheroids and then cryopreserved. REFERENCES

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Claims

Claims

1. A method comprising the steps of:

a) obtaining a plurality of cells derived from a tissue or an organ or from a part thereof, b) orbitally rotating the cells in a fluid at sufficient speed and for a sufficient time for spheroids to form, c) cryopreserving a composition comprising the spheroids and a cryopreservant.

2. A method according to claim 1 wherein the cryopreservant is a permeating cryopreservant (e.g. DMSO).

3. A method according to claim 1 or claim 2 wherein the cryopreservant is present at a level of at least 5% v/v (e.g. of from 10 to 20% v/v) with respect to the composition.

4. A method according to any preceding claim comprising stepwise cooling of the spheroids to a minimum temperamre which is no greater than -100°C (e.g. no greater than -190°C).

5. A method according to claim 4 wherein the stepwise cooling includes the step of cooling the spheroids to a temperamre of between 1 and 8°C and maintaining the spheroids within that temperamre range for a period of at least

10 mins before further cooling is undertaken.

6. A method according to claim 5 wherein the further cooling includes a step of cooling the spheroids to a temperature of between 0 and -50 °C (e.g. of between -10 and -30 °C) and maintaining the spheroids within that temperamre range for a period of at least 30 mins.

7. A method according to any preceding claim including the step of maintaining the spheroids in liquid nitrogen.

8. A method according to claim 7 wherein the spheroids are maintained in liquid nitrogen for a period of at least 12 hours (e.g. for at least 24 hours).

9. A method according to any preceding claim wherein the spheroids are formed by orbital rotation at a one or more speeds of at least 50 rpm (e.g. of from

50 to 90 rpm or from 60 to 80 rpm), e.g. for a total period of at least 48 hours.

10. A method according to claim 9 wherein the spheroids are formed by a period of orbital rotation at a first speed followed by a period of orbital rotation at a second speed which is higher than the first speed.

11. A method according to claim 9 or claim 10 wherein the spheroids are formed by orbital rotation at a speed of up to 76 rpm (e.g. for at least 24 hours) followed by orbital rotation at a higher speed (e.g. for at least 24 hours).

12. A method according to any preceding claim wherein prior to cryopreservation the spheroids or cells are incubated in the presence of one or more of the following: serum, methylcellulose, an antioxidant (e.g. Vitamin E) and an antibiotic.

13. A method according to any preceding claim wherein following cryopreservation the spheroids are thawed, the cryopreservant is removed and the spheroids are subjected to post-cryopreservation orbital rotation at one or more rotation speeds of at least 50 rpm (e.g. of from 50 to 90 rpm).

14. A method according to claim 13 wherein said post-cryopreservation orbital rotation includes a period of orbital rotation at 50 to 70 rpm (e.g. for a period of at least 6 hours).

15. A method according to claim 13 or claim 14 wherein said post- cryopreservation orbital rotation comprises a period of orbital rotation at a first speed followed by a period of orbital rotation at a second speed which is higher than the first speed.

16. A method according to any of claims 13 to 15 wherein said post- cryopreservation orbital rotation comprises a period of orbital rotation at 50 to 70 rpm (e.g. for a period of at least 6 hours) followed by a period of orbital rotation at a higher speed (e.g. of at least 70 rpm) suitably for a period of at least 6 hours.

17. A cryopreserved spheroid.

18. A cryopreserved spheroid according to claim 17 which comprises a plurality of layers of different cell types.

19. A cryopreserved spheroid according to claim 17 or claim 18 which comprises neuronal cells e.g. brain cells.

20. A cryopreserved spheroid according to any of claims 17 to 19 which comprises a layer of astrocytes and which also comprises a layer of neurons.

21. A cryopreserved spheroid according to claim 17 which comprises liver cells.

22. A composition comprising two different types of cryopreserved spheroids (e.g. cryopreserved spheroids which include neurons and cryopreserved spheroids which include liver cells).

23. A composition comprising a cyropreserved spheroid and cryopreserved cells derived from a blood vessel.

24. The use of a spheroid which has been cryopreserved (e.g. of a spheroid as described in any of claims 17 to 21) as an in vitro model.

25. The use of a spheroid which has been cryopreserved (e.g. of a spheroid as described in any of claims 17 to 21) in screening.

26. The use according to claim 25 in screening substances for pharmaceutical activity or for toxicity (e.g. neurotoxicity).

27. The use according to any of claims 24 to 26, wherein spheroids of at least two different types are used as a co-culture e.g. of spheroids which comprise neurons and spheroids which comprise liver cells.

28. The use according to any of claims 24 to 26 wherein a spheroid is co-cultured with cells derived from a blood vessel.

29. A spheroid which has been cryopreserved (e.g. as described in any of claims 17 to 21) for use in medicine.

30. A spheroid which has been cryopreserved (e.g. as described in any of claims 17 to 21) for use in transplantation or in treating a patient via a bioartificial system (e.g. a bioartificial liver).

31. A spheroid which has been cryopreserved (e.g. as described in any of claims 17 to 21) for use in treating a degenerative disease or disorder or in treating chemical or drug-induced damage.

32. The use of a spheroid which has been cryopreserved (e.g. as described in any of claims 17 to 21) in the preparation of a bioartificial system or of an implant, for treating a degenerative disease or disorder or in treating chemical or drug induced damage.

33. An apparams adapted to agitate non-fluid material within a fluid, said apparams comprising a self -contained power source.

34. An apparams according to claim 33, which is rechargeable.

35. An apparams according to claim 33 or 34, which is adapted to impart orbital rotation to said material.

36. An apparams according to claim 35, wherein said orbital rotation is at least 50 rpm (e.g. 50 to 90 rpm)

37. An apparatus according to any preceding claim, wherein the self-contained power source allows the apparams to operate continuously for at least 24 hours

38. An apparams according to any preceding claim, which is programmed to impart orbital rotation to said material for a plurality of hours at a first speed followed by orbital rotation for a plurality of hours at a second spin speed, said second spin speed being greater than said first spin speed.

39. A kit comprising an apparatus according to any of claims 33 to 38 and a container adapted to be releasably mounted to said apparams.

40. A kit according to claim 39 comprising one or more culmre vessels, said container being adapted to support a plurality of said culmre vessels when said apparams is in use.

41. The invention substantially as hereinbefore described.