**7. Cryopreservation**

Biopreservation has been characterized by a recent rapid growth since advances in cell therapy, stem‐cell research, personalized medicine, cell banking, etc. drive the need for optimized storage protocols. Nevertheless, this field still experiences significant issues with the current techniques including suboptimal survival, loss of poststorage cell function, addition of animal components in storage solutions, and activation of cellular stress pathways which can lead to changes in gene expression and protein denaturation [50]. The clinical application of autologous or allogeneic MSCs requires on demand access to a ready off‐the‐shelf amount of viable therapeutic doses of MSC and therefore necessitates fast availability to cryopreserved MSC stocks. The aim of cryopreservation is to preserve the therapeutic properties of those cells that maintain unaltered the characteristics of the freshly isolated samples, but the freezing and thawing procedures could determine an alteration of the cellular osmosis which can cause cell injury.

#### **7.1. Freezing**

The freezing rate is a fundamental factor for all biological systems in the determination of viability following cryopreserved storage. Several studies have shown that successful cryo‐ preservation of cells in suspension needs sufficiently high cooling rates to reach quickly low temperatures and avoid slow‐cooling injury, but low enough cooling rates to decrease the formation of intracellular ice and avoid rapid‐cooling injury [51]. The responses to cooling rates are cell‐type specific, as distinct cell types have different membrane permeability parameters. The intracellular dynamics during freezing or thawing could be influenced by many factors which influence cell viability after both of these procedures, affecting the therapeutic outcomes. Among these factors, the subtract desegregation stress before cryopre‐ servation of the cells attached to the plastic, the intracellular ice formation during freezing which can compromise the integrity of the cell membranes and, after thawing, the risk of impairing the membrane and altering other cellular functional characteristics can be listed [52]. Currently, there are two procedures to achieve the efficient cryopreservation of MSCs: conventional slow‐freezing and vitrification (rapid cooling). Both of these methods may lead to cell damage during loading/unloading of the cryoprotectant agents (CPAs), freezing, and thawing steps. The slow‐freezing procedure is the most commonly used cryopreservation technique in clinics and research laboratories today, because it allows the preparation of large amounts of vials at one time. Cryopreservation by vitrification has shown higher cell survival and it has been recognized as a promising strategy for long‐term cell banking. Nevertheless, the difficulty to generate a fast enough heating rate to minimize devitrification and recrystal‐ lization‐induced intracellular ice formation during rewarming is one of the major problems to be overcome. However, the high CPA concentration that is required to achieve vitrification results in osmotic dehydration to cells. For these reasons, new vitrification methods have emerged as alternative techniques, which have shown the ability to significantly reduce cryoinjury. This approach has been improved for the cryopreservation of organized tissues where even extracellular freezing causes several damages. In fact, in a recent study reported by Wang et al. [53], magnetic induction heating of superparamagnetic nanoparticles was successfully applied to enhance rewarming, with promising results of the vitrified human umbilical cord matrix MSC survival.

#### **7.2. Cryoprotectants**

adipose tissue into small portions, followed by enzymatic digestion with collagenase type I at 37°C and the subsequent centrifugation to separate the different cell fractions. The obtained supernatant was composed of mature adipocytes and the pellet fraction consisted of the stromal vascular fraction (SVF) components, which comprise a heterogeneous cell population, including circulating blood cells, fibroblasts, pericytes, and endothelial cells, as well as "pre‐ adipocytes" or adipocyte progenitors. Stem cells and progenitor cells represent about the 3% of all cell populations [42]. Stem cells derived from the adipose tissue (ASCs) represent a purified population of the adherent stem cells present in the adipose tissue, since all other cell types are removed or die with time. Currently, ASC recovery is quick and easy to perform from the subcutaneous adipose tissue, as it could be successfully collected via lipectomy or from the tail base in horses and from the inguinal region [43] or during ovariohisterectomies in dogs and cats. Stem cells derived from the adipose tissue have been increasingly used for cell therapy both in humans and animals [44], either as freshly isolated, SVF cells, or as cultivated ASCs [43]. ASCs proliferate rapidly with a high cellular activity, making them an ideal source to obtain MSCs [45]. The most important advantage of adipose‐derived stem cells is their

which gives 1–10% of stem cell yield [46]; in comparison, MSCs constitute only 0.001–0.01% of BM [15]. When autologous ASCs are used, the adipose tissue is collected 2 or 3 weeks before the treatment and the animal receives the cultivated cells, but long‐term cultivation of ASCs before therapeutic use is not recommended, since the cells may lose their progenitor charac‐ teristics [47]. The use of allogeneic ASCs has been also performed; since these cells have immunoregulatory properties [48], this approach would allow the use of species‐specific allogeneic cryopreserved cells, avoiding the need for collection of tissue from the patient [49].

Biopreservation has been characterized by a recent rapid growth since advances in cell therapy, stem‐cell research, personalized medicine, cell banking, etc. drive the need for optimized storage protocols. Nevertheless, this field still experiences significant issues with the current techniques including suboptimal survival, loss of poststorage cell function, addition of animal components in storage solutions, and activation of cellular stress pathways which can lead to changes in gene expression and protein denaturation [50]. The clinical application of autologous or allogeneic MSCs requires on demand access to a ready off‐the‐shelf amount of viable therapeutic doses of MSC and therefore necessitates fast availability to cryopreserved MSC stocks. The aim of cryopreservation is to preserve the therapeutic properties of those cells that maintain unaltered the characteristics of the freshly isolated samples, but the freezing and thawing procedures

The freezing rate is a fundamental factor for all biological systems in the determination of viability following cryopreserved storage. Several studies have shown that successful cryo‐ preservation of cells in suspension needs sufficiently high cooling rates to reach quickly low

could determine an alteration of the cellular osmosis which can cause cell injury.

SVF cells can be isolated,

abundance: from 1 g of adipose tissue an average of 0.5–2.0 × 106

**7. Cryopreservation**

42 Cryopreservation in Eukaryotes

**7.1. Freezing**

In addition to controlling the cooling rates, one of the major challenges to obtain an effective cryopreservation method is the selection of a suitable CPA, which minimizes the damaging effects of freezing. The most commonly employed CPA for cultured mammalian cells is dimethyl sulfoxide (DMSO) solution, because it is cheap and it has a relatively low cell toxicity. DMSO penetrates cell membrane, reduces intracellular ice formation, and prevents cell damage due to dehydration caused by extracellular ice formed during freezing; on the other hand, it can also decrease the survival rate [54, 55] or induce cell differentiation to neuronal‐ like cells when added to the cell culture medium [56]. The most common cryopreservation medium to store several types of stem cells has become a solution of 10% (v/v) DMSO and up to 90% (v/v) fetal bovine serum (FBS), despite showing disadvantages. To improve this procedure, MSCs have been cryopreserved using both DMSO and FBS free systems, compris‐ ing different polymers either alone or in combination with ethylene glycol, 1,2‐propylene glycol, trehalose, sucrose, and/or glucose. In contrast to DMSO that penetrates quickly into the cell, the high molecular weight polymers such as polyvinylpyrrolidone, polyethylene glycol, polyethylene oxide, or polyvinyl alcohol are nonpenetrating and seems to act extracellularly (at 10–40% concentrations), with the increasingly high viscosities at low temperature and avoiding that water molecules form ice crystals [52]. In a study reported by Renzi et al. [57], several cryopreservation solutions for MSCs isolated from equine, ovine, rodent bone marrow, and equine adipose tissue were compared: the best results regarding cell viability were obtained using a solution of fetal bovine serum added with 10% of DMSO. Conversely, in a previous study, Ock and Rho [58] reported that the survival and number of colonies formed by porcine MSCs were significantly decreased following short‐term storage (less than a month) into liquid nitrogen (−196°C) and the amount of this decrease was inversely proportional to the DMSO concentration. Those data strongly suggest the use of 5% DMSO instead of conventional 10% DMSO for the cryopreservation of porcine MSCs, for minimizing the CPA toxicity on cells. However, slow freezing with reduced concentration of CPAs has gained much interest in order to decrease the effect of the osmotic shock and chemical toxicity. Nevertheless, the commonly used CPAs are highly toxic at 37°C (body temperature) and could not be applied to patients. For this reason, multistep washing is required to completely remove the highly toxic, cell membrane‐permeable cryoprotectants from cryopreserved cells for clinical use, though this procedure is often associated with significant loss of precious cells (~10% during each washing step). Therefore, it is important to achieve cell cryopreservation with nontoxic CPAs. Recently, Rao et al. [59] demonstrate that nanoparticle‐mediated delivery of trehalose into mammalian cells has great potential for cryopreserving the human primary adipose derived stem cells (hADSCs) and possibly other types of stem cells to facilitate their ready availability for clinical use. In fact, successful results on cryopreservation of hADSCs using only trehalose as cryoprotectant has been achieved with high survival and undamaged function post cryopreservation.

#### **7.3. Thawing and viability assessment**

As well as cooling, optimizing the thawing method of frozen MSCs is also important. Furthermore, in clinical transplantation applications the post‐thaw viability assessment has shown to be of paramount importance. Several techniques have already been suggested for thawing frozen sample. A procedure of thaw and wash allows to remove DMSO and cell fragments, but may cause cell loss or cellular aggregation during centrifugation. Thaw, dilution, and wash procedure avoids the problem due to the centrifugation, allowing an osmolar equilibration, but the untoward effects of DMSO and cell debris infusion are not prevented. Currently, the standard method for thaw frozen MSCs, either from slow freezing or vitrification, is to warm them rapidly (>100°C/min) in a water bath at 37°C, until all ice crystals disappear. This method generally results in high post‐thaw recovery of viable cells without using high‐cost equipment, but it is safer to thaw cells using a dry warming procedure, due to the potential microbiological contaminations of the water bath [60]. Literature suggests that rapid thawing rates (>100°C/min) that can prevent damaging ice crystals during recrystallization are optimal choice and generally results in the best post‐ thaw recovery and viability of cells [61]. High post‐thaw viability of MSCs, comparable to those thawed with the standard method, were obtained by Thirumala et al. [62] with a thawing procedure in a controlled‐rate freezing/thawing chamber at 10°C/min. For evaluating the cryopreservation outcomes in terms of post‐thaw cell quality and quantity, the selection of the correct viability measurement is essential. The most commonly utilized test, owing to its easiness and quickness, is the Trypan blue dye exclusion assay; however, this method has the disadvantage that it generally overestimates the viable population. Several reports suggested that fluorescence dyes are more accurate and reliable indicators of cell viability [63].
