Pressure enhanced dual-solid-surface ultra-rapid cooling improves post-thaw recovery in hepatocytes and precision cut liver slices

Abstract

Cell and tissue cryopreservation is essential in modern medicine for globally distributing biological materials; however, ice crystal formation hinders its efficacy. Non-equilibrium cryopreservation, which focuses on rapid cooling, shows promise in mitigating this issue and allows for reduced cryoprotective agent (CPA) concentrations, thereby minimizing cytotoxic effects. Increased pressure lowers both solution melting temperature (T_m) and solution homogeneous nucleation temperature (T_h), while simultaneously elevating solution glass transition temperature (T_g). These effects should improve vitrification outcomes by minimizing ice crystal formation. Here we designed, fabricated, and tested an apparatus capable of high-pressure dual-sided cooling for hermetically sealed tissue samples. Using this device, we demonstrated that dual-surface high-pressure solid-surface conductive cooling offers a more effective approach for increasing the cooling rate compared to convective methods like direct immersion in liquid nitrogen. The experiments were conducted on HepG2 cells and mouse precision-cut liver slices (PCLS, 300 μm thickness, 5 mm diameter) using dimethyl sulfoxide (DMSO) as the CPA at concentrations of 10 to 30% (v/v) and pressures ranging from 50 to 200 MPa. Optimal outcomes were achieved at 20% DMSO and 150 MPa, yielding the highest post-thaw tissue viability.

Introduction

The primary goal of cryopreservation is to enhance the post-thaw survival and functionality of biological specimens, including cells and tissues. This is achieved by halting biochemical reactions at extremely low temperatures, effectively stabilizing the specimens for prolonged preservation. This technique is crucial in various applications, from regenerative medicine to research, particularly in the context of liver cells and tissues like hepatocytes and precision cut liver slices (PCLS). These samples are invaluable for cell therapy, bioartificial liver systems, and pharmacotoxicology studies^1,2. The ability of the human liver to regenerate is a well-established feature in modern medicine³. In this context, the transplantation of hepatocytes stands as a primary area of interest within the field of liver cell therapy^4–6. Additionally, liver tissue transplantation is increasingly recognized as a promising therapeutic approach for tackling a range of liver pathologies and promoting liver regeneration. Traditional cryopreservation techniques, while moderately successful with cell suspensions, often face challenges with larger samples due to issues like tissue disruption from ice formation⁷. Optimizing cryopreservation is essential to address these challenges, improving the utility of cells and tissues in medical applications.

Cryoprotectant agents (CPAs), which include various chemical and biological materials, are fundamental in minimizing damage attributed to ice formation and excessive ion concentrations in cryopreservation processes⁸. However, high CPA concentrations can be harmful to cells and tissues, as they induce cellular osmotic and mechanical stresses. These stresses are caused by volume changes, including both shrinking and swelling, that occur during the addition and removal of CPA. These alterations can compromise the cellular structure, leading to potential damage or functionality loss⁹. Furthermore, the biophysical and biochemical interactions between CPAs and the cytoplasm may give rise to toxic effects. This relationship between the beneficial effects of CPAs in preventing ice damage and their potential harmful impacts on cells and tissues underlines the balance required in cryopreservation techniques.

From the physiochemical perspective, during the cooling process, as the temperature is reduced below the melting point of the solution (T_m) and reaches high sub-zero levels, water can exist in a metastable supercooled liquid state. However, as the temperature decreases further below T_m, reaching the homogeneous nucleation temperature (T_h), the likelihood of ice formation escalates dramatically. Despite this, it is still possible to pass through this region at ultra-rapid cooling rates, preventing ice crystallization in cases with high-CPA-concentration or resulting in only minimal ice formation with reduced CPA concentration. This process leads to an extremely low temperature, known as the glass transition temperature (T_g), which is around − 137 °C for pure water. In the absence of ice crystals, the liquid becomes a glassy, vitreous state in which biological samples can remain preserved and stable for extended periods. The minimum cooling rate, known as the critical cooling rate (CCR), to achieve vitrification of pure water is 3 × 10⁶ ℃/s ¹⁰, 1 × 10⁷ ℃/s ¹¹, or 1 × 10¹⁰ ℃/s ¹², indicating a range of values based on various studies¹³.

The relationship between ultra-rapid cooling protocols and the vitrification of biological samples has been explored in the literature using several different techniques. For instance, Risco et al. increased the cooling rate using a quartz capillary due to high thermal conductivity and high surface area to volume ratio while also reducing wall thickness and sample volume¹⁴. They used slush nitrogen as the cryogen and claimed to successfully vitrify a solution of 1.5 mol/L propane-1,2-diol (i.e. propylene glycol) and 0.3 mol/L sucrose in 1×PBS, considered to be a low-CPA-concentration solution for cryopreservation. Katkov et al. employed jet freezing with liquid nitrogen (LN₂) to vitrify an aqueous sample containing 15% (v/v) glycerol¹⁵. They reported successful vitrification, which they visually assessed as resulting in a transparent medium. Nevertheless, it is noteworthy that complete vitrification is not a necessity for successful cryopreservation or transparency if ice crystals are very small¹³. Cells appear to be resilient to the presence of minute ice crystals, those less than 1 μm in size, and can maintain their viability and functionality upon warming^13,16. This understanding allows for flexibility in cryopreservation techniques, potentially enabling effective preservation strategies that do not require full vitrification. According to research by Akiyama et al., a CCR of approximately 10⁴ ℃/s is recommended for successful CPA-free cell cryopreservation¹³. They achieved this by cryopreserving a 40 pL cell suspension using an inkjet cell printing technology. This rate is considered sufficient to traverse the thermodynamically unstable region between the homogeneous nucleation temperature (T_h) and the glass transition temperature (T_g).

Two primary methods have been employed to achieve rapid cooling rates in cryopreservation: liquid convection (moving a cryogen over the biological sample) and conduction (using solid-surface cooling)¹⁸. For some cryogens, the convection method encounters a significant limitation known as the Leidenfrost effect. When a biological sample comes into direct contact with a cryogen like LN₂, the temperature difference generates a vapor film between the biological sample and cryogen. Acting as an insulating barrier, this vapor film leads to reduced heat transfer coefficients and, consequently, a decrease in cooling rate within the biological sample¹⁵. This necessitates a higher CPA concentration to compensate for the decrease in cooling rate. For example, Glockner et al. identified an optimal concentration of 30% DMSO in Sack’s solution for the cryopreservation of rat PCLS¹⁹. Their methodology involved using cryovials plunged into LN₂ for freezing, followed by conventional warming procedures. They reported that the post-thaw slices retained 40–50% Glutathione (GSH) compared to fresh control groups. On the other hand, conduction methods (solid-surface cooling) avoid the limitations of the convection method by having the sample in direct contact with a cooled surface possessing high thermal diffusivity. This is commonly accomplished by using a pre-cooled solid surface, a technique referred to as metal block freezing or solid surface freezing^20–22. This method utilizes a metal block made from a high-thermal-conductivity material, such as copper, silver, or aluminum. The block is cooled using liquid nitrogen or helium and is kept within an insulating container to preserve its low temperature^23–25. Moreover, physically pressing the samples onto the metal block using a spring-loaded plunger is often employed to improve the cooling rate by enhancing contact between the biological sample and the metal block^26–28. However, since cooling occurs from only one side, the depth of cooling is restricted, potentially leading to insufficient cooling rates across the biological sample. Furthermore, this technique carries the risk of damaging the biological sample due to the force of impact on the cooled surface. In addition, these biological samples are exposed without any protective enclosure, which presents a challenge for use in clinical applications.

Another strategy to minimize the required CPA concentration is the use of high-pressure. The use of high pressure freezing to preserve cell and tissue ultrastructure is well established²⁹. Moreover, the relationship between T_g and T_m and pressure is also well understood: as Miyata et al. reported³⁰, T_g can be increased with pressure, and simultaneously, T_m can be reduced^31–33. Therefore, high-pressure reduces thermodynamically unstable and metastable regions with respect to ice formation in the phase diagram (Fig. 1). For instance, isochoric (constant volume) approaches can cause high-pressure inside the sample container due to the increasing ice volume during freezing^34–37. However, this technique of high-pressure freezing is typically not classified as a vitrification method because ice formation is a necessary part of the process.

Fig. 1.

Adapted phase diagram of cryoprotectant agent-water solution in the context of vitrification. The dashed line indicates that at high CPA concentrations, no ice formation by homogenous nucleation can occur, and the solution can remain supercooled until it reaches the glass transition temperature (T_g). Adapted with permission from Ref. ¹⁷, 2025, Elsevier.

Here we designed, built, and validated a new device that integrates two-sided solid-surface cooling with ultra-high pressures^38,39. While previous studies have explored the solid-surface method through just one-sided cooling of bare biological samples^40,41, our research took a different approach. We aimed to achieve a uniform and rapid cooling process on biological samples that were hermetically sealed. This method not only enhances the cooling efficiency but also ensures that the samples remain uncontaminated throughout the process. Additionally, applying high-pressure enhanced the cooling rate⁴², shortening thermodynamically unstable and metastable regions related to ice formation, which is the key to the nature of time-dependent ice formation (Fig. 1)^33,43. Furthermore, by increasing the contact area between surfaces, and thus, overcoming the thermal contact resistance⁴⁴, the heat transfer rate was maximized. To demonstrate the efficacy of this approach in suspended cells, this study employed the HepG2 cell line as a model system, since the HepG2 cell line serves as one of the most extensively utilized in vitro models for hepatocytes⁴⁵. For tissue studies, we employed mouse liver PCLS to provide an intermediary platform between monolayer cell culture and whole-organ experiments. The utilization of both HepG2 cells and mouse liver PCLS in our study allows for a comprehensive evaluation of cryopreservation techniques on cellular and tissue-specific levels. Further, this study focused on vitrification-based preservation using ultra-rapid cooling and a relatively low-CPA concentration. Our objective was to identify optimal conditions to maximize the post-thaw viability of liver cell and tissue.

Results

Biological compatibility with vitrification container

The compatibility of the biological samples with the aluminum sample container was assessed under standard laboratory conditions without applied pressure to control for possible effects of the container material, independent of the pressure. Suspended HepG2 cells were exposed to the aluminum container for 0, 5, and 10 min, representing the maximum handling durations prior to cooling and after warming, as the total time from CPA loading to immersion in LN₂ was approximately 7 min.

Figure 2 shows cell viability in terms of membrane integrity over time. The overall regression indicates that there is no statistically significant variation in cell viability over times relevant to container material during processing before and after cooling (p > 0.05).

Fig. 2.

Suspended HepG2 cell membrane integrity evaluated by PI exclusion after 0, 5, and 10 min to determine biological compatibility of the vitrification container. Data are mean ± s.d., n = 6 for each time point.

CPA cytotoxicity

DMSO-induced cytotoxicity assessment in HepG2 cells during 10 min incubation prior to cryopreservation was conducted. Since the present study emphasizes cooling techniques, a clinically straightforward one-step CPA addition protocol was employed for CPA exposure. Figure 3b reveals an ascending trend of cytotoxicity correlating with higher DMSO concentrations. Statistically significant cell membrane integrity reduction is evident at a DMSO concentration of 30% (P < 0.001). The difference between 10% and 20% DMSO was noted as marginally statistically significant (p = 0.04).

Fig. 3.

(a) Representative images of HepG2 cell membrane integrity post-treatment across different ultra-rapid cooling techniques without elevated pressure, and (b) comparative assessment of post-thaw HepG2 cell membrane integrity among various ultra-rapid cooling methods and CPA concentrations. Scale bar is 100 μm. Data are mean ± s.d., n = 7 for each group. p < 0.05.

Cell cryopreservation

Figure 3 demonstrates comparative analysis of post-thaw cell viability across different DMSO concentrations and ultra-rapid cooling techniques. The use of solid-surface cooling achieved through the device yielded higher cell viability compared to conventional methods, such as direct immersion in liquid nitrogen (LN₂). Optimal post-thaw viability was observed at 20% DMSO concentration across cooling techniques, illustrating the balance between cryoprotectant toxicity and ice crystallization at the specific cooling rate of each method. As evident from Fig. 3b, an increase in the CPA concentration correlates with a reduction in cell viability when normalized to the fresh control group. Conversely, the harmful impact of ice crystal formation on cell viability, compared to cell viability after CPA exposure becomes less pronounced as the CPA concentration increases.

PCLS pressure sensitivity

Figure 4 shows the effects of brief exposure (0.5 s) to varying high-pressure levels of 50, 100, 150, and 200 MPa on the PCLS viability, assessed using calcein-AM staining.

Fig. 4.

Assessment of the impact of short duration (0.5 s) high-pressure ranges without CPA on PCLS viability. The chart describes changes in tissue viability under various pressure magnitudes normalized to fresh PCLS as a control, ranging from 50 to 200 MPa. Data are mean ± s.d., n = 6 for each group. ***p < 0.001; NS, not significant.

It was observed that the application of high-pressure up to 150 MPa did not lead to a statistically significant reduction in tissue viability, thereby indicating that pressures within this range are tolerated by the tissue samples. However, at a pressure level of 200 MPa, there was a significant decrease of approximately 30% in tissue viability.

PCLS cryopreservation in different techniques

Figure 5 presents a comparative viability and histological analysis of post-thaw PCLS subjected to different vitrification methods through ultra-rapid cooling. In the context of this study, three distinct rapid cooling techniques were employed to assess their impact on the PCLS preservation. The convection method involves the direct immersion of the container including PCLS into LN₂. In contrast, in the solid-surface cooling technique (SS0), the container was positioned within the device, facilitating efficient heat transfer through enhanced thermal contact between the container and the aluminum thermal blocks. The third technique, known as the pressure-enhanced solid-surface method, also involves the use of the device but incorporates high-pressure. In light of our findings related to pressure induced toxicity, the Differential Scanning Calorimetry (DSC) technique⁴⁶, and the biological evaluation of CPA cytotoxicity, specific parameters were selected for this study. A high-pressure setting of 150 MPa (SS150) was chosen, and a DMSO concentration of 20% (v/v) was utilized, as this concentration was found to provide an effective balance between cryoprotection and minimization of cytotoxic effects. These preferred conditions aimed to perform as standardized parameters for comparing the efficiency and biological impact of the different high cooling techniques under investigation. It is evident that the pressure-enhanced solid-surface method yielded the highest viability when compared to other techniques. In Fig. 5a, confocal imaging technique utilizing Hoechst and propidium iodide (PI) staining were employed to evaluate cell membrane integrity across different cryopreservation methods. The pressure-enhanced solid-surface method (SS150) demonstrated higher cell membrane integrity when compared to other methods. Furthermore, tissue histology was evaluated through various methods, as detailed in Fig. 5b. Open tissue areas, which are indicative of ice crystal formation, were quantified as white spaces within the PCLS using the ImageJ software (Fig. 5d). These measurements were then normalized to the fresh control group. The open tissue area exhibited a significant increase when the convection method was employed for cryopreservation, contrast to the results obtained through conduction-based methods, both with and without the application of pressure.

Fig. 5.

Qualitative and quantitative analysis of post-thaw PCLS cryopreserved with different vitrification methods: a) confocal microscopy analysis of cell membrane integrity within PCLS using Hoechst and PI staining, b) histological evaluation of tissue integrity in PCLS, c) comparative analysis of viability in post-thaw PCLS, normalized to fresh PCLS as a control (n = 6). d) Open tissue areas in PCLS were quantified. These values were subtracted from and normalized to fresh PCLS as the control group. Five fields of view were observed (n = 3 mice/group). Scale bar is 50 μm. Data are mean ± s.d., *p < 0.05, **p < 0.01, ***p < 0.001.

Discussion

In low-CPA concentration or non-equilibrium vitrification, the cooling rate is the critical parameter that can significantly impact the overall success of the cryopreservation process. In such circumstances, the cooling rate dictates the formation and size of intracellular and extracellular ice crystals, which directly influence the viability and functionality of the cryopreserved cell or tissue upon thawing. Akiyama et al. suggest that a cooling rate of 10⁶ ℃/min could enable mammalian cell cryopreservation without the need for CPAs¹³. While their research provides valuable insights into the role of cooling rate in ultra-rapid cryopreservation, there are a number of limitations including an extremely small sample size (40 pL) and the unsealed cryopreserved sample, raising concerns about potential post-thaw contamination⁴⁷.

This concern about post-thaw contamination is elevated for clinical applications. One approach to mitigating this problem is to employ a hermetically sealed vitrification container. This usually involves a trade off in the characteristics that make samples easy to vitrify, including an increased volume and reduced heat transfer coefficients. In this study, we introduce a novel (but off the shelf) vitrification container to facilitate ultra-rapid cooling. This container features high thermal diffusivity (80 × 10^{− 6} m² s^{− 1}) ⁴⁸, an elevated surface-to-volume ratio compared to a sphere, and minimal wall thickness (0.2 mm). We previously studied the thermophysical characteristics of cryopreservation media under rapid cooling using a differential scanning calorimetry technique with this container^46,49. We also assessed the design of the container through physical and mathematical modeling⁴². This approach offers a scalable solution for cell suspensions compared to the inkjet cell printing method¹³, and makes it well suited for PCLS or other small tissue cryopreservation such as ovarian cortex pieces⁵⁰ or corneas⁵¹. Also, this study showed (cf. Figure 2) that the container is biocompatible, at least for hepatocytes and PCLS for time durations well longer than needed for freezing, and is consistent with findings from other studies⁵². For example, Pabon et al. developed a grid collector-container for cryopreserving human sperm that enabled the vitrification of 5–10 µL cell suspensions through direct contact with LN₂ ⁵³. They reported a post-thaw recovery rate of 70.9%, although the direct exposure to LN₂ raised concerns about potential contamination⁴⁷. Their reported post-thaw viability for human sperm exceeds our results for HepG2 cells (44.4%) using the convection method of plunging into LN₂, most likely due to their enhanced cryopreservation media (0.3 M trehalose-sucrose), smaller sample volume (5–10 µL), and consequently, higher cooling rate.

de Graaf et al. carried out a study on the cryopreservation of rat PCLS using cryovials, the most prevalent container in tissue cryopreservation, that were plunged into LN₂. In their study cryopreservation medium consisted of 18% DMSO, Williams Medium E, fetal calf serum, and University of Wisconsin medium⁵⁴. After thawing, the slices were cultured for an additional 4 h. Their findings indicated that cryopreserved slices retained 65% of the original potassium content and 60% of the number of intact cells when compared to fresh control groups incubated for 4 h at 37 °C. In comparison to their results, our findings on tissue viability (59%) using a cryopreservation container plunged into LN₂ (convection method) suggest that the use of complex media containing large protein molecules, along with an additional post-thaw incubation period, improved the viability of PCLS preserved at lower cooling rates in plastic cryovials. However, our results indicate higher tissue viability with the solid-surface (SS0) technique (70%) and even greater viability with high-pressure (SS150) cooling (80%).

A significant challenge in vitrification-based cryopreservation is the associated toxicity of the high-CPA concentration needed to prevent ice crystal formation²⁰. In the initial phase of our study, we examined the CPA toxicity on the biological sample. Given our focus on fast cooling techniques, we selected a one-step approach for CPA addition and removal during 10 min, a protocol that offers simplicity and is, therefore, preferred in research and clinical centers. As illustrated in Fig. 3, cell viability remains high at up to a 20% DMSO concentration. de Graaf et al. claimed that the viability of fresh rat liver slices was not influenced significantly by 18% DMSO^54,55, we found a significant difference at 20% DMSO with p = 0.04, suggesting that perhaps 18% DMSO is “statistically” tolerable for liver tissue but 20% is not for liver cell suspensions. However, a pronounced decline in cell viability is observed at 30% DMSO concentration, and is in agreement with de Graaf et al. ⁵⁴, indicating a critical threshold due to mechanical damage, possibly due to extreme cell volume excursions⁹ and cytotoxicity⁵⁶. Careful CPA equilibration⁵⁷ and cocktail design⁵⁸ may decrease viability loss in future studies. In our study, the CPA composition and loading conditions were selected based on commonly used biological sample cryopreservation protocols to maintain comparability with previous work. While this approach was suitable for demonstrating the effects of cooling rate and for validating the feasibility of the dual solid-surface high-pressure cooling technique, systematic optimization of CPA loading parameters will be important in future work including composition, temperature, duration, and concentration to further reduce cytotoxicity and enhance post-thaw viability. For example, dextran has often been used for high-pressure freezing for microscopy^59,60. Recently oocytes and embryos were cryopreserved using 15% DMSO + 15% ethylene glycol + 0.5 mol/L sucrose in a commercial high-pressure freezing device with reasonable success⁶¹.

Our data showed (Fig. 3) that increased DMSO concentrations are effective in mitigating the harmful effects of ice crystal formation on cell viability. Therefore, as cytotoxic effects of DMSO increase with concentration, the optimal concentration was identified as 20% DMSO, where cell viability was maximal. Hence, the damaging impacts of CPA concentration and ice formation on post-thaw viability need to be balanced in the context of cooling rate restrictions. These restrictions may arise from several factors including limited thermal diffusivity of material, vitrification container and biological sample thickness, low sample aspect ratio, and high sample volume. Moreover, Fig. 3 reveals that the solid-surface technique explored here (with a calculated cooling rate on the order of 10⁴ ℃/min⁴² results in higher post-thaw cell membrane integrity compared to the convection method (p < 0.01) with the cooling rate of 2 × 10³ ℃/min as reported by Sun⁶², while the critical cooling rate for a 20% DMSO solution in water has been reported to be on the order of 10⁵ ℃/min⁶³. For instance, de Kanter and Koster evaluated the impact of different cooling rates, ranging from 0.6 to 250 ℃/min, on the cryopreservation of rat PCLS⁶⁴ and found that a cooling rate of 250 ℃/min led to the highest post-thaw PCLS viability.

Convection-based cooling encounters the Leidenfrost effect⁶⁵, a phenomenon that hinders effective cooling by reducing heat transfer (cf. ^65–67). However, in solid-surface cooling methods, the sample avoids the Leidenfrost effect by interfacing with a surface characterized not only by high thermal conductivity but also by low thermal capacity and density, effectively resulting in high thermal diffusivity. In addition to empirical data, Amini and Benson conducted mathematical and physical simulations of the vitrification container, considering both convection and solid-surface (conduction) methods for cooling⁴². They demonstrated that the cooling rate achieved through the solid-surface method is about one order of magnitude higher than that of the convection method. This cooling method has been employed in cryo-electron microscopy, where it is commonly referred to as metal block freezing^24,26. In this method, the tissue is directly put in contact with a metal block with high thermal conductivity that is cooled by LN₂ or helium. This technique has an estimated cooling rate of 10⁶ °C/min, although this rate is only applicable to the surface of one side of the sample⁶⁸. However, in these existing protocols, the biological samples lack an enclosed container, thereby posing a significant obstacle for clinical applications. Additionally, the physical impact of the biological sample onto the cooled surface during this technique may result in damage to the sample¹⁸. Furthermore, because cooling occurs only from one side, the depth of vitrification is restricted, potentially leaving parts of the sample unvitrified. By implementing double-sided cooling, which applies cooling to the top and bottom of the sample, our solid-surface technique notably increased the cooling rate.

Material selection is critical for achieving optimal thermal performance in cryopreservation. The high thermal diffusivity of aluminum (80 × 10^{− 6} m²s^{− 1}) ⁴⁸ makes it preferable to commonly used materials like stainless steel (3.2 × 10^{− 6} m²s^{− 1}) ⁶⁹, or especially the plastics used in cryovials and straws (0.13 × 10^{− 6} m²s^{− 1}) ⁷⁰ for rapid cooling applications⁵². Thinner walls and an optimized surface area to volume ratio enable faster and more uniform cooling. For example, Day et al. studied the cryopreservation of rat PCLS with dimensions similar to those explored in our study, (8 mm in diameter and 250 μm thickness). For the cryopreservation process, they utilized aluminum plates (1.6 mm thickness) fitted with a gasket and plunged into LN₂ and subsequently rewarmed in a 37 ℃ water bath. The cryopreservation medium consisted of 10% DMSO in FBS. They observed cracking and fragmentation in the preserved slices. Additionally, they noted 70% lactate dehydrogenase (LDH) and alanine aminotransferase (ALT) levels compared to fresh groups. It appears that the inclusion of FBS, including large protein molecules, in the cryopreservation medium enabled them to reduce DMSO concentration while maintaining comparable viability to our solid-surface technique (SS0) at 70%. Nevertheless, we were able to enhance tissue viability using the high-pressure (SS150) method, with minimal cracking and fragmentation in the preserved PCLS. It is worth noting that FBS may not be suitable for all applications, especially those involving human samples. Their methodology did not effectively eliminate the Leidenfrost effect. Furthermore, the sealing process involving the use of a gasket is comparatively more complex and less reliable than the method presented in our study.

Tissue cryopreservation under ultra-rapid cooling conditions presents challenges. One of the primary limitations is tissue thickness, which can impact the efficiency of cooling, in contrast to single-cell suspensions, where ultra-rapid cooling can be more readily achieved (cf. ⁷¹). Thinner slices enable faster and more uniform heat transfer, facilitating higher cooling rates and thereby improving vitrification efficiency. In contrast, thicker samples exhibit a reduced cooling rate, especially at the tissue slice core, increasing the likelihood of partial ice formation. According to the literature^72,73, a maximum thickness of approximately 300 μm ensures complete and uniform diffusion of nutrients and oxygen to the inner cell layers, thereby maintaining tissue viability during culture. Therefore, a PCLS thickness of 300 μm was selected in this study, as it represents a practical and widely applicable standard for use in various experimental and biomedical applications. The total sample volume inside the cryopreservation container was kept constant for all experiments to ensure comparable cooling conditions.

The radial dimension of the tissue slice influences cooling uniformity and, consequently, preservation efficiency. Larger tissue dimensions can result in slower heat transfer toward the center of the slice. However, under the dual-solid-surface ultra-rapid cooling and high-pressure conditions used in this study, the radial gradients are minimal, a fact supported by our numerical simulations⁴².

The utilization of high-pressure as an adjunct to ultra-rapid cooling represents a sometimes used approach in tissue cryopreservation^74–76. High-pressure has the potential to modulate the thermophysical properties of the solution, effectively reducing the temperature at which ice crystallization occurs (T_m ,T_h) and enhancing glass transition temperature (T_g)⁷⁷. Amini and Benson’s findings indicate that the application of high-pressure can enhance the cooling rate through the solution⁴². These impacts mitigate the adverse effects of ice formation within the tissue.

The high-pressure influence on biological samples should not be overlooked. As observed in several studies, there can be a range of pressure magnitude and exposure duration that maintain cell viability without causing considerable disruption to cell functions^78,79. Moor and Heochli presented valuable insights into the high-pressure effects on the survival rate of Euglena gracilis cells⁸⁰. In their research, cells were subjected to pressures up to 200 MPa for varying exposure durations, up to a maximum of 4 s. It was observed that shorter durations of pressure exposure allowed for higher levels of pressure to be tolerated by the biological sample without compromising cell viability⁸¹. As indicated in Fig. 4, PCLS can tolerate pressures as high as 150 MPa for durations up to 0.5 s. The use of a 0.5 s pressure load was consistent with previous high-pressure studies, which demonstrated that short exposure durations permit the biological sample to tolerate pressures up to 200 MPa without significant viability loss⁸⁰. The comparable outcomes observed between 50 MPa and 150 MPa for PCLS suggest a threshold behavior of PCLS under high-pressure conditions, where pressures above a certain limit yield minimal incremental benefit. These findings align well with the results presented by Moor and Hoechli⁸⁰. It should be noted that in our study, pressure loading and cooling were conducted simultaneously.

When biological samples are cooled at higher pressures, the confined aqueous solutions may solidify into a vitreous state or develop ice II or III crystallites, which possess a higher density compared to Ice I. If the pressure is released while the biological sample remains below T_g, no alterations will take place⁸². Under these conditions, the applied pressure in this study could be safely released, allowing the pre-cooled aluminum thermal blocks, containing LN₂, to be detached and used to maintain the cryopreservation container at cryogenic temperatures during transfer and subsequent storage in LN₂.

However, during the warming process without pressure, Ice I formation is most likely to occur if the warming rate is insufficient⁸². In this study, the tissue slices were thawed within the sealed cryopreservation container by transferring it into a 37 °C water bath immediately after retrieval from LN₂ storage, under atmospheric (non-pressurized) conditions. Here, we reduced the likelihood of ice I formation during warming by employing a 20% DMSO concentration, a result supported by our previous work demonstrating that no measurable devitrification occurs in 20% DMSO solutions during thawing, even at lower warming rates, following ultra-rapid cooling⁴⁶.

According to Rios and Rabin⁸³, for tissue undergoing vitrification, thermal contraction occurs both above and below the glass transition temperature (T_g). In our study, the pressure reading wasn’t observed to significantly change during cooling, despite expected small contraction of the sample. This is because the pressure-supplying apparatus adjusted to maintain the applied pressure as the sample volume decreased slightly during cooling. Although vitrification inherently involves thermal contraction and the development of thermomechanical stress⁸³, the pressure-enhanced solid-surface cooling method (SS150) produced less visible physical damage compared to the other methods, as evidenced by the reduced open tissue areas in PCLS shown in Fig. 5. These findings, together with the viability results, suggest that although stress is not eliminated⁸³, the stresses generated during dual-solid-surface high-pressure cooling are likely lower than those associated with freezing.

According to Kauzmann⁸⁴, the specific volumes of vitrified materials are nearly identical to those of their corresponding liquids, suggesting that the glass transition is accompanied by only minimal volumetric change compared with crystallization. Under ultra-rapid cooling and high-pressure conditions, ice nucleation is strongly suppressed⁸², resulting either in a fully vitrified state or in the formation of extremely fine, non-harmful ice crystals¹³. These conditions substantially reduce volumetric expansion and the development of mechanical stress within the sample. The applied pressure suppresses ice nucleation and promotes uniform heat transfer across the sample, thereby minimizing temperature gradients that could otherwise induce internal stress.

Achieving a fully vitrified state in pure water and under a CPA-free condition is attainable at cooling rates on the order of 10⁷ ℃/min for a layer thickness of up to 0.1 μm ⁸². In cell and tissue samples, the presence of cytosol and extracellular matrix (ECM) contributes to a relatively high viscosity of the media⁸⁵. Moreover, it has been suggested that the compartmentalization of the solution within the tissue structure may increase the likelihood of achieving vitrification⁵⁵. These change the conditions required for vitrification as compared to pure water. In fact, it is possible to achieve complete or partial vitrification accompanied by minor ice crystal formation, known as non-harmful ice crystallization, at cooling rates on the order of 10⁵ °C/min for a sample layer thickness of 10 μm in the absence of CPA^13,82. This lowers the threshold for cooling rate required to avoid crystalline ice formation, making the process potentially more practical for larger and more complex biological samples. Achieving such cooling rates consistently across varying volumes of biological samples requires advances in vitrification container design and even the use of specialized cooling technology. For instance, the use of double-sided cooling under high-pressure described here seems to substantially lower the required cooling rate for effective vitrification, possibly reducing the necessary cooling rate to 10³-10⁴ °C/min for biological samples with a layer thickness of up to 600 μm. Moor stated that for a biological sample with a thickness of 600 μm, the high-pressure must be sustained for a minimum of 0.3 s with the cooling rate of 3 × 10⁴ ℃/min to achieve complete vitrification in the whole sample⁸². This represents a remarkable advantage, especially when dealing with thicker samples, such as tissues or organ slices.

In this study, the estimated cooling rates at the center of the cryopreservation container, the warmest point within the container, were approximately 1.2 × 10⁴ °C/min for the solid-surface cooling technique (SS0) and 3.8 × 10⁴ °C/min for the pressure-enhanced solid-surface method under 150 MPa (SS150), as determined by numerical simulations⁴². Therefore, it is reasonable to conclude that vitrification or near-complete vitrification likely occurred across most of the sample volume under SS150 conditions. Due to the high-pressure and hermetically sealed configuration of the cryopreservation apparatus, along with the ultra-rapid cooling rate and the limited thermal inertia of thermocouples, it was not feasible to directly measure temperature profiles inside the tissue slices.

Conclusion

In this study, we presented the novel application of double-sided solid-surface cooling to enhance the cooling depth, and the utilization of high-pressure to alter thermophysical properties during cooling. In our study, we designed, constructed, and evaluated an apparatus for small tissue cryopreservation. This involved testing that included viability assessments, cell membrane integrity evaluations, and histological examinations of the PCLS under various conditions. Figure 5c demonstrates that the pressure-enhanced solid-surface cooling technique yielded the most promising post-thaw tissue viability, when optimized at a pressure of 150 MPa and a DMSO concentration of 20%. Based on the findings through the DSC technique under ultra-rapid cooling conditions, as investigated by Amini and Benson⁴⁶, along with considerations for no pressure during the common warming procedures frequently used in research and clinical centers, it becomes essential to use 20% DMSO to inhibit devitrification. Further, we assessed structural damage to the PCLS during the cryopreservation by conducting H&E staining to evaluate tissue histology. As evidenced by Fig. 5b, the extent of open tissue area (represented as white space)⁸⁶ in the pressure-enhanced solid-surface technique signifies that the extracellular ice formation, particularly in the PCLS sinusoidal areas, was minimized due to the considerations employed during this technique.

Direct verification of the internal state of the sample (ice formation or vitrification) under simultaneous high-pressure and low-temperature conditions is technically challenging due to the sealed and ultra-rapid nature of the process. In this study, the internal physical state was inferred indirectly through post-thaw histological analysis and biological sample viability assessments, which revealed less open tissue areas, indicative of reduced ice crystal formation and damage, and correspondingly higher sample viability resulting from the minimized ice formation phenomenon. In addition, our previous studies using differential scanning calorimetry (DSC) under ultra-rapid cooling conditions⁴⁶ demonstrated reduced exothermic peaks associated with ice crystallization, further supporting the vitrified state of the samples. Although cryomicroscopy⁸⁷, Raman spectroscopy⁸⁸, and Synchrotron-based x-ray diffraction⁸⁹ could theoretically provide direct confirmation, such measurements are extremely difficult to implement within the current high-pressure sealed configuration.

Our study reveals the impact of cooling rate on cell and tissue cryopreservation when employing ultra-rapid cooling. We outlined the distinctions in cooling rates between convective and conductive heat transfer methods, attributing these differences to the Leidenfrost effect. Additionally, our study not only assessed the potential toxicity associated with high-pressure conditions but also found that high-pressure enhances the PCLS cryopreservation. This points the way for harnessing technological innovations at developing CPA-free and scaled-up biological sample cryopreservation protocols.

Materials and methods

Cell culture and treatments

The HepG2/C3A hepatocellular carcinoma cell line was obtained from the American Tissue Culture Collection (ATCC; CRL-10741). These cells were maintained in a controlled environment using Eagle’s Minimum Essential Medium (MEM), enhanced with components such as 1.5 g/L sodium bicarbonate, non-essential amino acids, L-glutamine, and sodium pyruvate (Sigma-Aldrich, Canada). Additionally, the medium was enriched with 10% Fetal Bovine Serum (FBS), 1% pen strep (P/S) and kept at a constant temperature of 37 °C within an atmosphere containing 5% CO₂. The growth medium was replaced at 48 h intervals. Once the cell population reached a confluency of 60–70%, subculturing was carried out using a 0.25% Trypsin-EDTA solution. The cells were then collected and resuspended in freshly prepared culture medium.

In the preparation of cell suspensions, the cells were used once they achieved a confluency of 60–70%. The suspension process involved treating the cells with a 0.25% Trypsin-EDTA solution, followed by centrifugation at 300 g for a duration of 5 min. Subsequently, the cells were suspended at a specific concentration of 1 × 10⁶ cells per milliliter in cryopreservation solutions.

Cryopreservation solutions

In order to assess the efficiency of various cooling techniques and methodologies applied to biological samples within this investigation, Eagle’s Minimum Essential Medium (MEM) was elected as the base medium. This choice was guided by MEM extensive use in cryopreservation^90,91. In addition, the composition of MEM, which encompasses essential and non-essential amino acids, vitamins, salts, and glucose⁹², but excludes large molecules like proteins, establishes a baseline for the examination of ultra-rapid cooling techniques and its effects on ice formation. For the preparation of cryopreservation solutions, dimethyl sulfoxide (DMSO; ThermoFisher Scientific, NJ, US), a commonly used CPA in cryopreservation, was combined with MEM at concentrations of 10%, 20%, and 30% (v/v).

Cell experimentation

Biological compatibility with vitrification container

To verify the biological compatibility of the cells with aluminum, the material used for the cryopreservation container in these experiments, biocompatibility tests were performed under standard laboratory conditions without applied pressure to specifically isolate the effect of the container material on cell viability. Tests were performed at time intervals of 0, 5, and 10 min, representing the maximum duration of cell handling prior to cooling and following warming. Accordingly, the maximum total time from the initiation of CPA loading to immersion in LN₂ was approximately 7 min. Since the objective of this experiment was to assess the compatibility of the aluminum cryopreservation container with the biological samples, independent of the effects of cooling rate enhancement associated with pressure application, the impact of cryo-storage duration on cell viability was not considered in this experimental design.

In this procedure, cells were suspended in culture medium at a concentration of 1 × 10⁶ cells/mL after being trypsinized. Prior to conducting the experiments, the containers were subjected to sterilization procedures, including autoclaving, cleaning with 99% isopropanol, and exposure to UV light for a minimum of 30 min. Following this preparation, 30 µL of the cell suspension was pipetted into a container (a 30 µL differential scanning calorimetry sample pan, approximately 5.5 mm x 2.5 mm (diameter x height); Shimadzu Co., Tokyo, Japan, 201-53090) which was hermetically sealed using a crimper (Shimadzu Co., Tokyo, Japan). This process was performed during intervals of 0, 5, and 10 min, representing the maximum time for cell preparation prior to cooling and after warming up. An analysis of cell membrane integrity was carried out using propidium iodide (PI) and Hoechst staining.

CPA cytotoxicity

To assess the potential cytotoxicity of CPA, an analysis was performed. Cells were suspended at a concentration of 1 × 10⁶ cells/mL within a microtube and then exposed in a single step to varying DMSO concentrations, including 10%, 20%, and 30% (v/v). This was done for a 1:1 dilution, resulting in final concentrations of 10%, 20% and 30%, over a period of 10 min at 4 °C. Following this exposure, the viability of the cells was evaluated and used to determine the appropriate DMSO concentrations for the cryopreservation process described below.

Cell cryopreservation

Cell suspensions, at a density of 1 × 10⁶ cells/mL, were prepared in cryopreservation media containing 10%, 20%, and 30% (v/v) DMSO mixed with MEM. These suspensions were pipetted into vitrification containers in 30 µL aliquots. After hermetically sealing the containers, they either were immersed in LN₂ or positioned in the device. This device was designed to freeze the biological samples using a cold surface technique, ensuring effective heat transfer by enhancing thermal contact between the container and the aluminum thermal blocks. Following this rapid freezing process, the containers were transferred into a LN₂ reservoir, where they remained for future use.

In the warming process, the containers were thawed by submerging them in a water bath maintained at 37 ℃. The containers were then opened with tweezers, contents were pipetted into a mini-centrifuge tube, the cell suspensions were centrifuged for 5 min at 300 g for the CPA removal, the supernatant was removed, and pellets were resuspended in fresh MEM for subsequent evaluation of cell viability (Fig. 6).

Fig. 6.

Schematic representation of the control and experimental procedure for cell and PCLS cryopreservation.

Tissue experiments

Animals

All research with animal tissues was conducted on tissues obtained from animals euthanized for other purposes under a “tissue share agreement”, and as such is exempt from animal use guidelines under Canadian Council on Animal Care policy. However, to be clear, all animal tissue use in this study was approved by the University of Saskatchewan’s animal ethics committee (UACC AREB Permit #026Exempt2021). C57BL/6J mice (12–14 weeks old, weighing about 30–35 g) were used in this study and all animals were housed at the Lab Animal Services Unit (LASU) at Health Sciences Department, University of Saskatchewan, Canada. Animals were housed at 22 °C, with 60% relative humidity, and a 12-hour light/dark cycle. All animal use in this study was approved by the University of Saskatchewan’s animal ethics committee under a tissue share agreement and exempt from animal use permits. The experiments conducted in this study were carried out in strict accordance with the guidelines of the Canadian Council on Animal Care, and all procedures adhered to the ARRIVE guidelines.

The standard method of euthanasia was inhalant isoflurane at 5%, followed by a secondary method of cervical dislocation.

Tissue Preparation as precision cut liver slices (PCLS)

Adult mice were anesthetized and euthanized and hepatectomy was performed where the liver was extracted and promptly immersed in a chilled, oxygen-rich excision buffer to minimize hypoxia. For the preparation of liver slices, the liver tissue was first punched by 5 mm diameter- punch biopsy (Integra-Mitex) to facilitate slicing. The tissue was transferred into a tube containing a 2% solution of low-melting-temperature agarose and subsequently cooled on ice for 5 min to allow the agarose to gel. This gelled agarose provided support for the liver tissue during the slicing process. A vibratome (Precisionary Instruments, Model VF-310-0Z) was employed to create the PCLS. This fully automated instrument ensured consistent slicing and was set at a speed of two with an oscillation at four specifically for the liver tissue. The PCLS thickness was 300 μm ⁷², and slices were placed in a chilled and oxygenated excision buffer, following a method described by Graaf et al. ⁵⁵.

Solutions

The excision buffers used in this procedure were formulated from Hank’s Balanced Salt Solution (1x HBSS), enhanced with 8.9 mM sodium bicarbonate. This was further supplemented with 10% Fetal Bovine Serum (FBS) and 5% penicillin streptomycin.

PCLS pressure sensitivity

An experiment was carried out to investigate the effect of different high-pressure levels on tissue viability. PCLS were placed within containers. The containers were then hermetically sealed. The sealed containers were positioned within the device. The device was employed to subject the PCLS to 50, 100, 150, and 200 MPa for maximum 0.5 Sect⁸². Following the application of the pressures, the containers were opened, and the tissue viability assessments were performed.

Tissue cryopreservation

PCLS were incubated in cryopreservation medium for 15 min at 4 °C which was optimized to comprise 20% DMSO, based on our previous DSC investigation of solution vitrification tendencies⁴⁹ and live cell toxicity assessments. Afterwards, they were placed into cryopreservation containers. After sealing the containers hermetically, they were subjected to one of three conditions: immersion in LN₂, using a cold surface technique to improve surface-to-surface contact areas (SS0), or application of an optimized high-pressure of 150 MPa using the solid-surface technique via the device (SS150). Containers were then transferred into an LN₂ reservoir, where they were kept for future use. Thawing was carried out in a water bath maintained at 37 ℃. The containers were opened, and the PCLS were washed with phosphate-buffered saline (PBS) several times for CPA removal. Finally, assessment of viability was made using the following metrics described in detail below. First, a 3D confocal microscopic examination of cell membrane integrity was conducted with Hoechst and PI staining to assess membrane and functional viability. Additionally, PCLS viability was assessed with calcein-AM staining, offering insights into the functionality of the tissue. Finally, histological examination using Hematoxylin & Eosin (H&E) staining was performed (Fig. 6).

The procedure for staining cell cytoplasm and nuclear elements employed three dyes: calcein-AM, propidium iodide (PI), and Hoechst 33,342. Calcein-AM is a membrane-permeable dye that turns into fluorescent calcein after intracellular esterases cleave the AM group within live, metabolizing cells. Propidium iodide (PI) emits red fluorescence when bound to double-stranded DNA, and it is utilized for staining the nuclei of dead cells since it cannot permeate living cell membranes. Hoechst 33,342 is a blue-fluorescent dye that binds to dsDNA. Hoechst is used to stain nuclei and is permeable to all cells, providing a counterstain for both live and dead cells. The staining buffer was formulated with calcein-AM (Catalog number: C1430), Hoechst 33,342 (Catalog number: H1399), and PI (Catalog number: P1304MP) in an incubation medium (All; ThermoFisher Scientific, Mississauga, ON, Canada).

Cell suspension assay

The post-thaw cell suspensions were incubated with Hoechst 33,342 (1 µg/mL) and PI (3 µM) staining buffer for 10 min at 37 °C. The stained cells were inspected under the fluorescence microscopy (Nikon, Eclipse80i, Tokyo, Japan). Hoechst 33,342 was excited at a wavelength of 350 nm and emission assessed at 460 nm, while PI was excited at 535 nm with emission assessed at 617 nm.

PCLS assay

Plate reader

The post-thaw PCLS viability was assessed using calcein-AM staining, following a modified version of the method described by Luckenbach et al. ⁹³. Following treatment, PCLS were subjected to incubation with calcein-AM (5 µM) for 45 min at 37 °C. Afterwards, the PCLS were washed with cold PBS. The PCLS were subsequently homogenized in ice-cold double-distilled water, which was supplemented with 50 mM Tris-HCl, pH 7.5 (10% w/v (g/mL))⁹⁴. The homogenate was subjected to centrifugation at 10,000 g for 20 min at 4 °C. Then, 100 µL of the resulting supernatant from each sample was pipetted into a 96-well plate. The intensity of calcein-AM fluorescence was quantified using a plate reader (Varioskan Flash), set at an excitation and emission wavelength of 485 and 530 nm, respectively. Each sample experienced duplicate testing. To evaluate the negative control for calcein-AM staining, PCLS were treated with ethanol for 30 min to induce cellular death. Next, these samples were stained and homogenized in the same manner as treated samples. To estimate the background fluorescence attributed to the calcein-AM dye, a number of wells were filled with calcein-AM but without any supernatant.

Laser scanning confocal microscopy

A staining buffer was prepared by incorporating PI (3 µmol/L) and Hoechst 33,342 (5 µg/mL) into the incubation medium. Following treatment, the tissue slices were placed in a 6-well plate partially filled with the prepared staining buffer. They were then incubated on an orbital shaker for 30 min at 37 °C. Subsequently, the tissues were washed twice with PBS. Imaging was conducted using a laser scanning confocal microscope (Zeiss LSM 880). Excitation and emission wavelengths were set at 350/460 nm for Hoechst 33,342 and 535/617 nm for PI, respectively.

Histological assessment

After treatments were applied to the PCLS, the specimens were placed in 10% Neutral Buffered Formalin. The embedded tissues were then kept in this state for a period ranging from 24 to 48 h. Subsequently, a microtome was employed to obtain precise sections of the tissue, each having a thickness of 5 μm. These sections were stained with Hematoxylin & Eosin (H&E). One observation made during this histological investigation was the effect of ice crystal formation in the tissue. The formation of these crystals can induce cracks, demonstrating as increased white spaces in the histology images⁸⁶. To collect quantitative insights from these observations, an analysis of the H&E-stained PCLS was conducted, focusing on the white space within the tissue area. By employing ImageJ software (ImagJ.nih.gov)⁸⁶, measurements were obtained, and the results were normalized to a fresh control sample. The categorization of samples was blinded until the completion of all analyses.

Pressure enhanced solid-surface ultra-rapid device

A device depicted in Fig. 7 was designed, fabricated, and evaluated to facilitate the cryopreservation of biological samples under high-pressure through ultra-rapid cooling while the biological specimens were frozen within a hermetically sealed environment.

Fig. 7.

Diagram of the apparatus and system for monitoring and regulating pressure.

The biological sample to be cryopreserved was enclosed within a vitrification container shown in Fig. 8. The vitrification container, constructed from a 0.2 mm thick sheet of aluminum, houses the biological sample, and is hermetically sealed with a crimper (Shimadzu Co., Tokyo, Japan). The device featured upper and lower aluminum thermal blocks due to the high thermal conductivity (k = 200 W/(m K)) (Fig. 9), moved and pressed by the hydraulic system.

Fig. 8.

Aluminum hermetic vitrification container.

Fig. 9.

Aluminum thermal block.

The hydraulic system was a designed network, where the upper and lower presses are linked to hydraulic lines. These lines, in turn, were connected to a splitter, acting as a connection for channeling hydraulic pressure. Subsequently, a line from the pressure limiter was directed to a hydraulic pump system. For pressure regulation, a load cell (FUTEK-LCM 350, USA) was installed on the hydraulic piston (ENERPAC-RC53, USA) and linked to a data acquisition unit (OM-DAQ-USB-2401, Omega, USA), which was in turn connected to a computer system that controlled the pressure limiter (SPX Hydraulic Technologies, No. 9625, USA).

The applied pressure was determined by dividing the load-cell force (FUTEK-LCM-350, USA) by the pressed surface area of the sample container (π × (5.5 mm / 2)²), as indicated in Fig. 8. The hermetically sealed container, filled with cryopreservation medium and biological samples, was placed between two aluminum thermal blocks and compressed by a hydraulic piston (ENERPAC RC-53, USA). This configuration produced a hydrostatic pressure within the confined volume. The absence of leakage after experiments confirmed that the applied force was converted into internal pressure.

The entire assembly was coordinated through a switch, granting control over the hydraulic pressure, the movement, and operation of the presses. The upper and lower presses are operated to move the corresponding thermal blocks towards each other, exerting a controlled pressure on the container for maximum 0.5 s⁸². The press exposure time applied was calculated based on the sample’s characteristics and pressure sensitivity of the biological material.

A holding platform was designed to support and suspend the container between the thermal blocks and was constructed to fit between the blocks without obstruction, allowing them to close entirely around the container. For cooling experiments, the thermal blocks were precooled using LN₂. During the press, the upper and lower thermal blocks approached each other and the container (and the biological sample within) simultaneously contacted both blocks, supported by the holding platform.

After freezing, the upper and lower pistons moved away from each other, and the upper and lower thermal blocks and the container were locked together. During the cooling phase, the container was maintained for a sufficient duration to ensure that the biological sample reached an equilibrium temperature with the thermal blocks, which had a surface temperature measured at around − 190 ℃. Then, the sample was removed and transferred to an LN₂ reservoir. The upper and lower thermal blocks were finally separated, leaving the container stored in the LN₂ reservoir for future use.

No frost accumulation was observed during operation. This was attributed to the short duration of the cooling procedure and the immediate transfer of the aluminum thermal blocks from the LN₂ source into the device, where they were automatically pressed together. The brief exposure to ambient air prevented moisture condensation on the cooling surfaces, ensuring stable and efficient heat transfer throughout the process.

Statistical analysis

Statistical analysis was conducted utilizing SPSS 25 (SPSS Inc., IBM). In each figure legend, the number of biological replicates is indicated. Normality for variables was assessed using the Shapiro–Wilk test. The homogeneity of variance was evaluated with the Levene’s test and ANOVA testing was employed. Comparisons were made using pairwise post hoc t-tests for individual contrasts, and the Tukey HSD test was applied for multiple comparisons with equal variance. Graphical representations of the data display the mean and standard deviation (s.d.). A significance level was selected at a 95% confidence interval, with P-values less than 0.05 to indicate statistical significance. All viability data were normalized based on the initial control.

Author contributions

M.A. designed and performed the experiments, analyzed experimental data, prepared figures, wrote and reviewed and edited the paper. J.B. conceived of the project and developed experimental design, coordinated sample collection, interpreted results, obtained funding, reviewed and edited the paper. All authors reviewed and approved the final draft.

Funding

Funding for this work was provided by the National Science and Engineering Research Council (RGPIN-2023-04007).

Data availability

All data relevant to this work are available from the corresponding author when a reasonable request is made.

Declarations

Competing interests

The authors declare no competing interests.

Human/Animal rights

The article does not contain any studies with human subjects performed by any of the authors. Studies with animal tissues were approved by the University of Saskatchewan Animal Care Committee (UACC; #026Exempt2021).