A novel, simplified method to prepare and preserve freeze-dried mouse sperm in plastic microtubes

Abstract

Although freeze-drying sperm can save space, reduce maintenance costs, and facilitate the transportation of genetic samples, the current method requires breakable, custom-made, and expensive glass ampoules. In the present study, we developed a simple and economical method for collecting freeze-dried (FD) sperm using commercially available plastic microtubes. Mouse epididymal sperm suspensions were placed in 1.5 ml polypropylene tubes, frozen in liquid nitrogen, and dried in an acrylic freeze-drying chamber, after which they were closed under a vacuum. The drying duration did not differ between the microtube and glass ampoule methods (control); however, the sperm recovery rate was higher using the microtube method, and the physical damage to the sperm after rehydration was also reduced. Intracytoplasmic sperm injection (ICSI) using FD sperm stored in microtubes at −30°C yielded healthy offspring without reducing the success rate, even after 9 months of storage. Air infiltration into all microtubes stored at room temperature (RT) within 2 weeks of storage caused a drastic decrease in the fertilization rate of FD sperm; underwater storage did not prevent air infiltration. RT storage of FD sperm in microtubes for 1 week resulted in healthy offspring after ICSI (5–18%), but the addition of silica gel or CaCl₂ did not improve the success rate. Our novel microtube method is currently the simplest and most effective method for treating FD sperm, contributing to the development of alternative low-cost approaches for preserving and transporting genetic resources.

The preservation of mammalian genetic resources is of great importance. It is applied in diverse scientific fields, including human medicine, farm animal production, laboratory animal record keeping, and wildlife conservation [1]. Preserving genetic resources in oocytes and embryos is unrealistic because of the difficulty in obtaining sufficient numbers of oocytes and/or embryos from each individual, even when they are in good health. In contrast, a large number of spermatozoa can be obtained from a single male. Thus, the genetic banking of spermatozoa provides an efficient and cost-effective approach to preserving genetic resources [2, 3].

Spermatozoa are almost exclusively preserved via cryopreservation [4, 5]. Long-term frozen sperm storage is inconvenient because of the need for a steady supply of liquid nitrogen (LN₂), considerable storage space, and the inherent risk of cross-contamination [6, 7]. The maintenance of frozen stock is also energy-dependent and requires continuous monitoring and an uninterrupted power supply, which makes biobanks susceptible to natural disasters and power outages. These characteristics make such biobanks unrealistic and often unobtainable in developing countries and small businesses where LN₂ or the power supply is unreliable or impractical. LN₂ is also dangerous because of its exceptionally low temperature and risk of oxygen deficiency. In addition, the transportation of cryopreserved sperm requires the use of an LN₂ container or dry shipper that conforms to international standards, which is associated with additional disadvantages in terms of labor and costs [8].

Freeze-drying is an alternative method for sperm preservation that can overcome the disadvantages of cryopreservation. The freeze-drying process causes spermatozoa to lose their motility and viability but preserves their DNA and activation potential; therefore, freeze-dried (FD) spermatozoa injected into oocytes via intracytoplasmic sperm injection (ICSI) can still successfully form viable embryos [9-11]. The live birth of viable offspring obtained from FD sperm has been reported in mice [9], rabbits [12], rats [13, 14], and horses [15]. In addition, sperm from livestock [16, 17], rare animal species [18], and humans [19] can be FD. We have also succeeded in FD preservation of spermatids [20] and somatic cells [21]. FD sperms can be stored and transported for short periods at room temperature (RT) without using LN₂ or dry ice as cooling agents [9, 22, 23]. Furthermore, the nuclei of FD mouse sperm are highly resistant to extreme conditions, including frequent temperature changes, extremely high and low temperatures [24], and space radiation, as evidenced by the long-term storage of FD mouse sperm at the International Space Station [25, 26]. Unfortunately, FD sperm results in a lower live birth success rate than fresh sperm when using ICSI; however, this drawback is minimized by improved drying methods [27] and the addition of trehalose as a freeze-drying protectant [28].

The current freeze-drying methods require breakable glass ampoules. These ampoules must be able to resist low temperatures and vacuum treatments, which considerably increase their costs. Additionally, glass ampoules require a heat-sealing process that carries the risk of explosion, thus putting the experimenters in potential danger. Glass ampoule specifications (e.g., volume and inner diameter) also vary, making it difficult to share results between laboratories. Attempts to place dehumidifying agents and/or oxygen absorbents in glass ampoules to improve the live birth success rate have been unsuccessful because of the small opening of the ampoules and the resulting inability to insert protective agents [22].

Plastic microtubes, commonly referred to as microcentrifuge polypropylene tubes, can provide a potential solution to these drawbacks when used to store FD sperm. Microtubes are easy to handle and commercially available globally in standardized sizes. They have a wide entrance that enables the insertion of various protective agents and can be closed by simply pushing down the cap, eliminating the need for heat sealing. Despite these potential benefits, no study has evaluated the possibility of using microtubes for FD sperm. Here, we aimed to develop a safer and more cost-effective method for preparing and preserving mouse FD sperm than glass ampoule using commercially available 1.5 ml microtubes. This method makes it possible to safely manage a large number of mouse sperms using easily accessible, low-cost materials without requiring any special skills.

Materials and Methods

Animals

Female and male Institute of Cancer Research (ICR) mice aged 8–10 weeks were obtained from Japan SLC, Inc. (Shizuoka, Japan). Sperm were collected from the epididymides removed from male mice after sacrifice via cervical dislocation. The females were mated with vasectomized ICR males whose sterility had been previously demonstrated and used as surrogates to receive embryos fertilized with FD sperm. The mice were euthanized via CO₂ inhalation or cervical dislocation either on the day of the experiment or upon the completion of all experiments. All animal experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Committee of Laboratory Animal Experimentation of the University of Yamanashi, which followed the ARRIVE guidelines (A4-10).

Media

Human tubal fluid (HTF) medium [29] was used for the capacitation and freezing of dry spermatozoa. CZB [30] and HEPES-CZB [31] media were used for oocyte/embryo incubation (in 5% CO₂ in air at 37°C) and manipulation, respectively. Unless otherwise stated, all materials were purchased from Sigma Aldrich (St Louis, MO, USA).

Preparation of FD sperm

We preserved FD sperm using the standard glass ampoule method [24, 27] and 1.5 ml microtubes (polypropylene, colorless) (Eppendorf Inc. Hamburg, Germany; microtube-FD method; Fig. 1A). To prepare the samples, semen was suspended in 1 ml HTF medium and incubated for 30 min at 37°C in 5% CO₂ in air. After sperm preculture, 50 μl aliquots of the sperm suspension were dispensed into glass ampoules or microtubes. In the glass ampoule method, the ampoules were flash-frozen in LN₂ and freeze-dried using an FDU-2200 freeze-dryer (EYELA, Tokyo, Japan), as previously described [22]. In the microtube method, the microtubes were put in a microtube rack and frozen in nitrogen vapor above the surface of LN₂ for 10 min, then dried in a vacuum dryer at −50°C (FreeZone2.5, Labconco, MO, USA). The pressure was maintained at 0.100 mBar throughout the drying process. After drying, the microtubes were sealed manually by closing their caps. The sealing procedure was performed inside the acrylic chamber of a freeze drier to obtain vacuumed microtubes. The sperm-filled microtubes were removed from the chamber and analyzed using a Tesla coil leak detector (Tesla lamp checker HF-20; Shinko Electric & Instrumentation, Osaka, Japan). Only tubes determined to be under high vacuum were selected for further use (Fig. 2A). All samples were stored at −30°C or room temperature (RT; 15 to 25°C) until further use.

Fig. 1.

Schematic presentation of the freeze-drying protocol for sperm using the microtube method. (A) Sperm freeze-drying procedure. (B) Preservation of Freeze-dried sperm using moisture absorbents via the microtube method.

Fig. 2.

Sample drying time and retrieval rates of freeze-dried sperm. (A) Freeze-dried (FD) sperm preserved in glass ampoules (left) or microtubes (right). (B) Sample drying time. The horizontal axis indicates the drying time and the vertical axis shows the weight ratio of a container filled with 50 μl of frozen sperm suspension at various times during the FD process compared to the initial weight before vacuum drying (n = 10 replicates/samples per each group). (C) Retrieval rates and head-tail separation rates of FD sperm using the conventional ampoule method and the microtube method (n = 5 replicates/samples per each group). The asterisks indicate significant differences between groups (P < 0.05).

Assessment of drying duration

We compared the drying rates of frozen spermatozoa using three different methods: the standard method using glass ampoules and a manifold freeze-dryer, the microtube method using 1.5 ml plastic microtubes and an acrylic FD chamber, or a method using glass ampoules and an acrylic FD chamber (Fig. 2B). Microtubes containing frozen sperm were dried for 3, 6, 9, 12, or 24 h before ICSI to determine the effects of drying time on the quality of FD sperm. The weight of the microtube with 50 μl of a fresh sperm suspension was measured before and after the freezing treatment and measured every 3 h during the drying process up to 24 h. For the drying duration experiment, the samples were rehydrated immediately after the completion of each drying duration and then used for ICSI or stored at −80°C for later use.

Effect of storage duration and temperature on fertilization

We examined the effect of storage duration by storing FD sperm at −30°C for 3 days, 1–2 months, and 9 months and performing ICSI immediately after rehydration. We also studied the effect of storage temperature on the fertilization ability of microtube-FD spermatozoa by storing additional sperm samples at 4°C or RT for 1 or 2 weeks, respectively.

Counting of spermatozoa and measurement of head-tail separation rate

Glass ampoules or microtubes containing 50 μl frozen sperm were attached to an FD apparatus (standard method) or put in an acrylic freeze-drying chamber (microtube-FD sperm method), respectively, and dried for 6–8 h, which is the routine drying duration for FD sperm used in previous research [24]. Once drying was complete, the samples were rehydrated with distilled water, then 5 μl of the sperm suspension was transferred to a microtube containing 45 μl ultrapure water to obtain a dilution of 1:10. The samples were then immediately analyzed to determine the retrieval rate of sperm and the mechanical damage rate, as judged by the number of sperm without a tail (Figs. 2C and 2D). Spermatozoa were counted using a hemocytometer (Burker-Turk line, ERMA, Tokyo, Japan) at a magnification of 100 ×, with 80 of 400 squares counted for each assessment. The same counting chamber and cover glass were used in all experiments. The retrieval rate was measured by dividing the total number of sperm after rehydration by the total number of fresh sperm before FD. The head-tail separation rate was measured by dividing the number of sperm without a tail by the total number of sperm.

Detection of trapped air in microtubes using a Tesla coil leak detector

Microtubes containing air were identified using a Tesla coil leak detector (Shinko Electric & Instrumentation) according to the manufacturer’s instructions. Briefly, microtubes were exposed to a Tesla detector in the dark. Microtubes containing only a small amount of residual air become bright because of the ionization of the low-pressure gas inside them when the Tesla coil tip is brought close to the microtube; these microtubes were considered positive and vacuumed. When a significant amount of air is trapped inside a microtube, it cannot be ionized. These microtubes remained dark and were considered Tesla-negative and non-vacuumed. The vacuumed microtubes were stored for 2, 3, or 4 weeks in water or in a desk drawer at RT, and the amount of air trapped in the microtubes was measured.

Measurement of air trapped in microtubes stored at RT

Each microtube was opened in water to measure the amount of air trapped within it. The bubbles of air released from the microtube were captured using a 1 ml syringe and the volume was measured. Unless otherwise stated, RT storage refers to storing specimens in a drawer. Some specimens stored in water at RT were placed in water-filled bags, from which the air was expelled until no air bubbles were visible to the naked eye.

Absorption of residual moisture in the microtubes

We examined whether the quality of microtube-FD sperm could be improved by adding silica gel or calcium chloride (CaCl₂) to the microtubes to absorb moisture. A simple device was designed to prevent the desiccant from coming into direct contact with FD sperm. Briefly, aluminum foil was folded into a cone and embedded with a paper wipe to cover its inner wall and form a two-layer, cone-shaped container (Fig. 1B). Silica gel or CaCl₂ was added to the container and inserted into the microtube immediately before completing the FD process. The microtubes were subjected to vacuum for at least two hours and then sealed. For the control experiment, FD sperm were prepared as previously described using microtubes without any additional devices or desiccants. The microtubes were stored at RT for 1 or 2 weeks, the fertilization rate was determined, and two-cell embryos were transferred into recipient females to determine whether the increase in the rate of fertilized oocytes and two-cell embryos after moisture absorbent treatment was associated with an increased rate of offspring production.

Oocyte preparation

Female mice were superovulated via injection of 5 IU equine chorionic gonadotropin, followed by 5 IU human chorionic gonadotropin for 48 h. Fourteen to sixteen hours after hCG injection, cumulus-oocyte complexes (COCs) were collected from the oviducts of female mice and transferred to a Falcon dish containing HEPES-CZB medium. To disperse the cumulus, COCs were transferred to a 50 μl droplet of HEPES-CZB medium containing 0.1% bovine testicular hyaluronidase for 3 min. Cumulus-free oocytes were washed twice and added to a 20 μl droplet of CZB for culture.

ICSI and embryo transfer

ICSI was performed as previously described [31]. Immediately before ICSI, the microtube caps were opened, 50 μl sterile distilled water was added, and the cell suspension was mixed using a pipette. For the microinjection of spermatozoa, 1-2 μl of the spermatozoa suspension was transferred directly to a droplet of 10% polyvinylpyrrolidone in the injection chamber. The sperm suspension was replaced every 30 min during ICSI [24, 27]. Several piezopulses were applied to separate the sperm head from the tail, and the head was injected into the oocyte. The oocytes that survived ICSI were incubated in CZB medium at 37°C in 5% CO₂ in air. Pronucleus formation was verified at 6 h after ICSI. Embryos at the two-cell stage were transferred to day-0.5 pseudo-pregnant ICR female mice that were mated with a vasectomized male the night before the transfer [32]. Five to 15 embryos were transferred to each oviduct. On day 18.5, offspring were delivered via cesarean section and allowed to mature. The remaining unused embryos were cultured for up to four days to evaluate their potential to develop into blastocysts.

Statistical analysis

Data calculated as percentages were subjected to arcsine transformation for each replicate before performing one-way ANOVA. The Tukey-Kramer test was used for multiple comparisons. Fertilization and in vitro development rates were analyzed using nonparametric Mann-Whitney U tests. Offspring rates were analyzed using Fisher’s exact probability test (Prism, GraphPad Software, USA). P values less than 0.05 were considered to indicate statistical significance.

Results

Drying rates of spermatozoa in microtubes

Standard glass ampoules reached a stable state after 3 h of drying (Fig. 2B). The acrylic FD chamber exhibited a stable weight after 3 h, regardless of whether microtubes or glass ampoules were used.

Retrieval rate and mechanical damage of spermatozoa after freeze-drying using microtubes

When FD sperm were stored in glass ampoules, the retrieval rate of spermatozoa was lower than the initial concentration and head-tail separation rate, suggesting that physical damage to the spermatozoa increased significantly (P < 0.05). In contrast, when FD sperm were stored in microtubes, the retrieval rate and head-tail separation rate of spermatozoa improved compared to glass ampoules, although the differences were not significant (Fig. 2C).

Optimal drying duration for the microtube-FD sperm method

We sought to optimize the drying duration for the microtube-FD sperm method by assessing whether the drying duration influenced the developmental capabilities of the sperm after ICSI. Differences in drying duration did not result in significantly different rates of development to the blastocyst stage; however, more desirable results were obtained when sperm were dried for 6, 9, and 12 h (56%, 60%, and 58%, respectively; Table 1). Based on these results, we determined that 6–12 h was the ideal drying duration for the easy preparation of microtube-FD sperm.

Table 1. Effect of different drying times on the in vitro fertilization and developmental rates of freeze-dried mouse sperm using the microtube method.

Drying duration (h)	No. of oocytes used	No. of oocytes surviving after ICSI *	No. (%) of fertilized embryos	No. (%) of embryos developed to
				2-cell	4–8-cell	Morula	Blastocyst
3	83	55	50 (91)	47 (94)	42 (84)	36 (72)	25 (50)
6	181	116	93 (80)	87 (94)	75 (81)	64 (69)	52 (56)
9	169	99	85 (86)	84 (99)	75 (88)	67 (79)	51 (60)
12	118	82	76 (93)	71 (93)	64 (84)	55 (72)	44 (58)
24	144	96	83 (86)	76 (92)	50 (60)	44 (53)	43 (52)

* Intracytoplasmic Sperm Injection. There were no significant differences in the fertilization or blastocyst rates between the groups.

Integrity of FD sperm stored for long-term in microtubes at –30°C, 4°C, or RT

High fertilization rates of 92%, 99%, and 85% were obtained for FD sperm stored in microtubes for 3 days, 1–2 months, and 9 months, respectively. The two-cell embryos obtained from each group had comparable rates of two-cell stage embryos and births, even when microtube-FD sperm were stored at −30°C for 9 months (29%; Table 2). The reproductive potential of randomly selected offspring derived from microtube-FD sperm stored for 1–2 months was examined after they had grown to adulthood to determine whether they had normal reproductive potential. All examined offspring (male, n = 4; female, n = 9) demonstrated normal reproductive potential and were delivered to the next generation 3 to 4 months later. Thus, FD spermatozoa prepared in microtubes and stored at −30°C for 9 months had adequate fertilization ability to produce offspring at practical efficiencies for preserving genetic strains.

Table 2. Full-term development of mouse embryos derived from freeze-dried sperm preserved at –30°C for up to 9 months using the microtube method.

Storage period	No. of used oocytes	No. of oocytes surviving after ICSI	No. (%) of fertilized zygotes	No. (%) of 2-cell embryos	No. of transferred embryos (No. of recipients)	No. (%) [min–max] of offspring *	Mean body weight (g) ± SD
3 days	74	50	46 (92) ab	45 (98)	45 (4)	5 (11) [1–22]	2.07 ± 0.11
1–2 months	120	68	67 (99) a	61 (91)	61 (5)	15 (25) [0–54]	1.82 ± 0.15
9 months	167	92	78 (85) b	74 (95)	68 (6)	20 (29) [15–80]	1.83 ± 0.22

* Offspring rate: number of offspring/transferred embryos. Offspring rates were evaluated using Fisher’s exact test. Significant differences are indicated by superscript letters (^a,b P < 0.05).

The mean fertilization rates of FD sperm stored for 1 week at 4°C or RT were 92% and 93%, respectively (Fig. 3A). Most fertilized oocytes developed to the two-cell stage, with 51% and 39% of the zygotes developing into morulae and blastocysts, respectively. When FD sperm were stored for 2 weeks, the fertilization rates decreased significantly for sperm stored at RT compared to those stored at 4°C, with only 19% of the zygotes developing into morulae and blastocysts from the FD sperm stored at RT (Fig. 3B).

Fig. 3.

Developmental potential of embryos fertilized with FD sperm preserved at RT in microtubes. In vitro development of embryos derived from freeze-dried (FD) sperm preserved at 4°C or room temperature (RT) for 1 week (4°C, n = 193; RT, n = 177) (A) or 2 weeks (4°C, n = 78; RT, n = 92) (B). Mor/blast: morulae/blastocyst. Asterisks indicate statistically significant differences in developmental rates between the groups (P < 0.05). (C) Relative proportions of air in the Tesla-positive vacuumed microtubes immediately after measurement (n = 9). (D) Microtube-FD sperms were preserved in water. (E) Measurement of air trapped in microtubes containing FD sperm underwater and in a drawer at RT (n = 5 replicates per group). Data are expressed as the mean ± standard deviation (SD). W/o vacu: Without vacuum. Significant differences are indicated by superscript letters (P < 0.05). Note: Only Tesla-positive microtubes were used for storage. (F) In vitro development of embryos derived from microtube-FD sperm with or without moisture absorbents stored for 2 weeks at RT (control, n = 46; silica, n = 59; CaCl₂, n = 65). Asterisks indicate statistically significant differences compared to the standard microtube method without absorbent agents (P < 0.05). (G) Full-term development of offspring derived from microtube-FD sperm preserved at RT for 1 week.

Exploration of the cause of impaired development in microtube-FD sperm stored under RT

The amount of air inside the microtubes was quantified after RT storage. It was found that 33% of the vacuumed tubes contained less than 20 µl of air, and 44% of vacuumed tubes contained approximately 50 µl of air prior to storage (Fig. 3C). After 2 weeks of storage at RT, the amount of air in the microtubes increased to 420 μl when stored under water and 560 µl when stored in a desk drawer (Figs. 3D and 3E). In both instances, negative results were confirmed using the Tesla-detector. A weight increase of both samples was also observed after storage.

Effect of moisture absorbents on the full-term development of RT-preserved microtube-FD sperm

When microtube-FD sperm were stored for 2 weeks at RT, the control spermatozoa did not dissolve upon rehydration, and compact aggregates were observed. Thus, only a few spermatozoa were collected and used for ICSI, and the mean two-cell development rate was low. However, spermatozoa with normal morphology were easily retrieved when silica gel or CaCl₂ was inserted into the microtubes. The two-cell development rates were significantly increased in FD sperm stored with CaCl₂ compared to those in the control group. When the embryos were cultured for 4 days after ICSI, only a few developed into blastocysts, regardless of the moisture absorbent used (Fig. 3F).

The mean birth rate of the control sperm stored for 3 days at RT was 18%; it further decreased to 5% when stored for 1 week (Table 3, Fig. 3G). Contrary to our prediction, a lower offspring birth rate (1% or 2%) was observed in both groups. When FD sperm was stored for 2 weeks at RT, no offspring were obtained, regardless of the moisture absorbent used.

Table 3. Full-term development of mouse embryos derived from freeze-dried sperm preserved at RT for 1 week without an absorbent agent (control) or with silica gel or calcium chloride.

Moisture absorbent	Preservation period	No. of used oocytes	No. of oocytes surviving after ICSI	No. (%) of fertilized zygotes	No. (%) of 2-cell embryos	No. of transferred embryos (No. of recipients)	No. (%) of [min–max] offspring *	Mean body weight (g) ± SD
Control	3 days	86	44	41 (93)	34 (83)	33 (3)	6 (18) a [17–20]	1.93 ± 0.22
Control	1 week	292	203	184 (91)	168 (91)	168 (9)	8 (5) b [0–14]	1.99 ± 0.15
Silica gel		318	232	216 (93)	198 (92)	198 (11)	1 (1) bc [0–5]	2.02
CaCl2		313	205	198 (97)	187 (94)	187 (12)	3 (2) c [0–11]	2.15 ± 0.06
Control	2 weeks	93	50	33 (66)	33 (100)	33 (3)	0	-
Silica gel		80	47	47 (100)	38 (81)	38 (3)	0	-
CaCl2		161	90	84 (93)	62 (74)	62 (5)	0	-

* Offspring rate: number of offspring/transferred embryos. There were no significant differences in fertilization or two-cell fertilization rates. Significant differences in offspring rates determined using Fisher’s exact test are indicated by different superscript letters (^a,b,c P < 0.05).

Discussion

Glass ampoules have been predominantly used for FD sperm since healthy offspring were obtained from FD mouse spermatozoa approximately a quarter-century ago [9]. Despite the drawbacks of this method (e.g., high cost and risks) and the urgent need to overcome these issues, new methodologies and substitutes for glass ampoules have rarely been reported. In a previous study, we successfully preserved FD sperm in a thin plastic sheet, which was cheaper, more resistant to glass breakage, and generally less bulky than the glass ampoule method [23, 33]. However, that method has many challenges that need to be addressed, such as its RT storage limitation of less than 3 days. In addition, that method requires the removal of air from the gaps between the plastic sheets; thus, inserting additives without negating the space-saving features of that method was difficult. In the present study, we succeeded in preserving FD sperm in commercially available microtubes with an extended storage duration of 7 days at RT, which indicates great potential for international delivery. This new method is a simpler, more effective, and secure alternative to current plastic sheet-based methods for preserving genetic resources.

Although the weight of FD sperm decreased to a stable state after 3 h of vacuum treatment, a higher blastocyst rate was observed in samples dried for up to 24 h. This result indicated that vacuum treatment and dehydration within the microtube did not attenuate the developmental ability of embryos fertilized with FD sperm. The dry state is essential for the long-term storage of FD sperm at RT, but we were unable to determine the effect of drying duration on the sperm. Similarly, precise analytical methods have not yet been used to quantify residual humidity in FD sperm. In this study, air accumulated in all microtubes stored in a desk drawer at RT. Less air accumulated in the microtubes when they were stored in water; however, it was not possible to completely stop air accumulation. This may be due to the ability of gases (e.g., O₂ and water vapor) to penetrate the materials that form the microtube, an intrinsic characteristic of propylene [34]. Submergence in water may slow the rate of gas penetration; however, it does not provide an effective barrier. Wrapping microtubes with gas-impermeable metal films may help maintain a vacuum state and preserve sperm quality.

The degradation of sperm chromosomes is related to membrane damage [35,36,37]; the membranes of sperm heads have been shown to be severely damaged after freeze-drying [9]. It is known that the separation of the sperm tail from the head is commonly found in FD sperm [38]; thus, the separation rate of the sperm tail from the head would be a clear indicator of the extent of damage caused by freeze-drying treatment. Despite this finding, embryos fertilized with microtube-FD sperm that came into contact with air in the microtubes after vacuum treatment had poorer developmental potential to the blastocyst stage than embryos fertilized with FD sperm from the vacuumed glass ampoules. This finding indicates that the penetration of air into the microtubes reduced the quality of FD sperm when stored for a short period at RT.

The developmental rates of FD sperm to the two-cell stage significantly increased when moisture-absorbent agents were included in the microtubes; however, this effect was limited, and development to the blastocyst stage decreased slightly (Fig. 3F). No offspring were obtained from embryos fertilized with sperm stored in these conditions (Table 3). It is very likely that both FD sperm and moisture-absorbing agents exhibit hygroscopic behavior in the system, thus competing randomly to absorb any moisture that enters the microtube. If FD sperm absorbed moisture first, the moisture-absorbing agents would have reacted by absorbing water from the FD sperm. This results in FD sperm repeatedly undergoing hydration and dehydration, which increases cell damage with each alteration in moisture content. Another possible explanation for this phenomenon is that the moisture-absorbing agents dehydrate the sperm long-term, thus absorbing the water bound to the FD sperm. Because sperm need to maintain a fraction of bound water covalently bonded to its various biological macromolecules (e.g., proteins, lipids, and nucleic acids) to develop successfully [39, 40], removing these water molecules would be detrimental to development. This is particularly true for DNA, which requires water molecules to support its characteristic double-helical form and the proper decoding of its instructions [41].

Lyophilized proteins and DNA are prone to moisture-induced, solid-state aggregation during storage [42, 43]. During this process, these molecules undergo structural changes in the dry state because of their hydrogen-bonded crosslinks, which are dependent on their hydration level and controlled by the relative humidity of their environment. Furthermore, the denaturation of proteins and DNA is accompanied by a loss of aqueous solubility. In the present study, FD sperm stored at RT for 2 weeks formed insoluble aggregates upon rehydration. Adding moisture-absorbing agents to the microtubes seemed to prevent this aggregation; however, the possibility that aggregation continued to occur at the molecular level cannot be ruled out.

Plastics have a lower air resistance than other materials (e.g., glass or metals), which was the main cause of the limited storage duration of FD sperm at RT in this study. We showed that this drawback could be mitigated using a regular freezer by successfully producing offspring from FD sperm stored for 9 months at −30°C at a rather high birth rate, similar to that of sperm stored in glass ampoules. In addition, we also found that the microtubes could remain Tesla-positive longer when stored at −30°C than when stored at RT. Thus, microtubes are a practical and efficient substitute for glass ampoules when used to store FD sperm at −30°C.

In conclusion, we developed a protocol for freeze-drying sperm using conventional microtubes. Microtubes are easy to handle and provide a simpler and more effective method for obtaining FD mouse sperm compared to the currently used protocols. However, additional experiments are needed to develop more suitable storage conditions for FD sperm, especially because we observed a decline in the developmental ability of FD sperm when stored at RT. Nevertheless, our study contributes to the development of alternative low-cost approaches for preserving and transporting genetic resources.

Although freeze-drying can overcome several disadvantages of cryopreservation, it has many disadvantages. For example, freeze-drying equipment is required before the preservation of spermatozoa, and the production of offspring could incur a certain cost because various microscopic manipulators, as well as the training of technicians, are required. Therefore, it would be best to choose the most convenient method or a combination of both, depending on the purpose of use, mouse strains used, and the environment of the places (countries) where they are used.

Conflict of interests

The authors declare no conflicts of interest.