Semen Collection Technique
2-1 Semen Collection and Evaluation
2-1-1 Introduction
Semen collection and evaluation is fundamental for the development of viable artificial insemination techniques, which are at present primarily used to selectively breed domestic animals at present (Foote, 2002). The evaluation of semen involves measuring a number of parameters, such as semen volume and pH, as well as sperm concentration, motility, and viability, each of which are important for increasing successful artificial insemination attempts (Foote, 2002). Previous studies on sea turtles have reported the use of electro-ejaculation to collet semen (green turtles: Platz et al., 1980; Wood et al., 1982; olive ridley and hawksbill turtles: Tanasanti et al., 2009).
However, the ejaculated semen of just one hawksbill has been previously examined (Tanasanti et al., 2009); hence, insufficient data are available to date on this topic for this species.
In animals where the reproductive period is limited (i.e., seasonal), males are known to exhibit seasonal spermatogenesis, which is associated with changes in testosterone levels. Physiological studies of sea turtles have also reported this trend (Jessop et al., 2004; Licht et al., 1985; Rostal, 2005; Valente et al., 2011; Wibbels et al., 1990). Furthermore, Wibbels et al. (1990) and Rostal (2005) reported that the
spermatogenic process of adult male turtles may be deduced by the change in testosterone levels and the observation of seminiferous tubules collected from testis.
In this chapter, the author evaluated the semen of two captive hawksbill turtles by the analysis of semen characteristic and serum testosterone over a 15-month (i.e., at least one year). Preliminary information on semen collection by electro-ejaculation and seminal characteristics will be obtained to assist in the future establishment of an artificial insemination technique for hawksbill turtles.
2-1-2 Materials and Methods
In 1998 and 2000, two adult male hawksbill turtles were captured in the set-nets of fishing boats operating at sea, and were kept in captivity at the Ocean Expo Park, Motobu-tyo, Okinawa Prefecture, Japan in and indoor holding tank under Okinawa Prefecture permits (based on the Fishery Act in Japan). The holding tank (5 m × 5 m × 1 m) was an open water system, had 4 equal divisions, with the turtles being kept separately in 2 of the divisions. The water temperature of the tank was measured daily, and ranged between 20 °C and 30 °C across a 12-month period. This water temperature was approximately similar to that experienced by hawksbills under natural conditions, at least around Okinawa Island. The light condition of the tank was maintained at 12 h light/12 h dark. The turtles were fed at 24-h intervals, with a diet that included fish and squid in quantities equivalent to 1.0–2.0% of their body weight (BW). The straight carapace length (SCL) and BW of the 2 turtles were as follows: Turtle no. 1, SCL 79.5 cm and BW 54.5 kg; Turtle no. 2, SCL 83.0 cm and BW 71.0 kg.
Semen was collected at 1–2-month intervals between January 2008 and March 2009 (Turtle no. 1: 9 attempts; Turtle no. 2: 10 attempts). The turtles were removed from the tank, and kept stationary with a soft cloth belt on a retention stand (diameter:
500 mm, height: 500 mm), which was assembled from four car tires (Fig. 2-1-1A). The tail was allowed to hang unsupported beyond the edge of the stand. An electro-transformer for domestic animals (Lane Manufacturing Inc, Lane Pulsator IV-Auto AdjustTM, USA) was used, which produces an electric stimulus ranging from 3.0 V to 19.5 V (Fig. 2-1-1B). The electronic stimulus probe, which was assembled from an acrylic bar, was 700 mm in length and 8 mm in diameter, and had 2 stainless steel electrodes (diameter: 1 mmLength: 20 mm) on either side of the tip covered acrylic resin, which was made by us (Fig. 2-1-1B and C). After measuring the distance from cloacal entry point to the urogenital papilla by endoscopy (Olympus, Japan), the probe, which was washed (lubricated) by normal saline, was inserted 250–350 mm into the cloaca (from which the penis also extends) by leading the probe dorsally around the urogenital papilla, which was then electro-stimulated. Over a 5-min period, an electronic stimulus was given for 5 sec, followed by a 5-sec rest (1 cycle). A total of 3 cycles (1 set) was performed, using 3.0 V in the first cycle, 10.0 V in the second cycle, and 19.0 V in the third cycle, and 3 sets were performed in maximum. The urine of turtles was excreted from the cloaca during the early stages of electronic stimulus, and was collected into a vial by suctioning it up with a pipette. When semen was observed on the probe, and/or flowed along the dorsal sulcus to the tip of penis, it was counted as an ejaculation. At this point, I collected all ejaculated semen into a vial by suctioning it
up with a pipette within 10-min. The defecation of 2 turtles were not observed during semen collection. After semen collection, the cloaca and penis were washed with normal saline. After a 30-min rest, the turtles were returned to the holding tank. The health of both turtles was then assessed by veterinarians and aquarium staff of Okinawa Churashima Foundation, to check that no trauma had been incurred from the electronic stimulus. This study was performed based on the ethical guideline for animal exhibition and research of JAZA (Japanese Association of Zoos and Aquariums) and was permitted by the board of directors of the Okinawa Churashima Foundation.
After semen collection, semen volume (mL) was measured using a 15-mL high-clarity polypropylene conical tube (Becton, Dickinson and Company, USA).
Sperm concentration (×106/mL) was measured using a hematocytometer, Thoma (Sunlead Glass Corp., Japan). Sperm viability (%) was calculated by Eosin-nigrosin staining, which differentiates between live and dead sperm. The measurement of sperm concentration and staining has been described in previous studies (Bjorndahl et al., 2003). The pH of semen and urine was measured using a compact pH meter (Horiba Corp., Japan). The sperm motility (%) of normal semen and semen mixed with urine (mixed semen) was measured as described previously (Platz et al., 1978; Tsutsui, 2002).
The total number of surviving sperm (TNSS) was calculated for each ejaculation based on the results of semen volume, sperm concentration and viability, using the following equation:
T N S S = × Sperm viability (%) 100
Semen volume (ml) Sperm concentration (×106/ml)
×
Blood samples were collected from the 2 turtles immediately before semen collection. Ten milliliters of blood was sampled from the jugular vein on the left or right side of the neck using a 70-mm 20-gauge needle (TERUMO Inc., Japan) and a 10-mL syringe (TERUMO Inc.). Blood samples were stored in heparin vacutainers, after which plasma was collected by centrifugation (speed: 3000rpm, time: 20 min). The serum concentrations of testosterone were determined by modified double antibody enzyme immunoassay (EIA) as described previously (Prakash et al., 1987). Serum samples were extracted with a diethylether before assay. The EIA used antibodies of sheep anti testosterone-3-CMO-BSA (GDN250, Colorado State University, Fort Collins, CO, USA, Taya et al., 1985) and horseradish peroxidases conjugated testosterone -3-CMO. Serial dilutions of serum sample from male captive hawksbill turtle resulted in dose-response curve that was parallel to the standard curve generated with testosterone.
Assumptions for normality were tested using the Shapiro-Wilks test. We used a Wilcoxon signed-ranks test to test for a significant difference in median sperm motility between normal and mixed semen. Statistical significance was assumed as P < 0.05.
2-1-3 Results
Ejaculate containing sperm was obtained from 4 out of 9 attempts for Turtle no. 1 and from 10 out of 10 attempts for Turtle no. 2. Turtle behavior during the 14 combined ejaculations, included the curling of the rear flippers (Fig. 2-1-2A), the elongation and erection of the penis (Fig. 2-1-2B and C), the formation of the urethral fissure on the penis (Fig. 2-1-2D), and the anteroposterior shaking of the neck, which was composed
of approximately 5-shakes per second. In particular, the anteroposterior shaking of the neck was frequently observed just before ejaculation. However, when ejaculation was not observed (in the 5 attempts for Turtle no. 1), the elongation and erection of the penis and the anteroposterior shaking of the neck were not observed.
All samples containing semen had a high viscosity. The median of semen volume in the 14 successful attempts was 0.5 mL (inter-quatile range: 0.2–1.5 mL), the sperm concentration was 325 × 106/mL (100–645 × 106/mL), the semen pH was 7.4 (7.1–7.7), the urine pH was 5.9 (5.9–6.2), and the TNSS was 180.5 × 106 (12.6–750.5 × 106) (Table.
2-1-1). There was a significant difference in the median sperm motility between normal and mixed semen for Turtle no. 2 (from 2% to 54%; Wilcoxon signed-ranks test, P <
0.05; Fig. 2-1-3). Although, small sample size prevented us from statistical analysis, a similar pattern was also observed in Turtle no. 1 (from 2.5% to 39.5%; Fig. 2-1-3).
There was a clear change in the mean water temperature in each month over a 12-month period, with the temperature ranging from 20.5 °C to 29.3 °C. The highest temperature was recorded between July and October, and the lowest temperatures being recorded between January and March (from winter to early spring, < 22°C, Fig. 2-1-4A).
When we divided semen collection attempts between low (<22 °C) and high (>22 °C) water temperatures, the TNSS median for Turtle no. 1 was 1035 × 106 (inter-quatile range: 733.5–1336.5 × 106, n = 2) and 20.9 × 106 (11.8–30.0 × 106, n = 2), respectively (Fig. 2-1-4B). Correspondingly, the motility of mixed semen was 69.5% (59.3–79.8%, n
= 2) and 23.5% (20.3–26.8%, n = 2) for low and high temperatures, respectively (Fig.
2-1-4C). The serum testosterone concentration was 49.0 ng/mL (43.3–53.4 ng/mL, n =
5) and 11.5 ng/mL (10.2–33.1 ng/mL, n = 4) for low and high temperatures, respectively (Fig. 2-1-4D). In contrast, the TNSS median for Turtle no. 2 were 795 × 106 (616.9–931.0 × 106, n = 5) and 3.8 × 106 (1.9–39.0 × 106, n = 5) for low and high temperatures, respectively (Fig. 2-1-4B). In addition, the motility of mixed semen was 79.5% (79.5–96.0%, n = 5) and 31.5% (13.0–41.5%, n = 5) for low and high temperatures, respectively (Fig. 2-1-4C). The serum testosterone concentration was 75.3 ng/mL (73.7–83.7 ng/mL, n = 5) and 31.5 ng/mL (22.5–31.7 ng/mL, n = 5) for low and high temperatures, respectively (Fig. 2-1-4D).
Discussion
This study presents novel information about the semen evaluation of captive male hawksbill sea turtles, based on an extended 15-month study using the electro-ejaculation technique. The author collected information that is useful for developing optimal semen collection techniques for future artificial insemination programs of this and other endangered sea turtle species.
Wood et al. (1982) first studied the use of electro-ejaculation to obtain semen from green sea turtles, followed by Tanasanti et al. (2009) for olive ridley and hawksbill turtles. The former study reported that ejaculate containing semen was obtained from 74.3% of the 74 attempts from green turtles (n = 28), while the latter study reported a success rate of 75.0% out of 8 attempts from olive ridley turtles (n = 3) and 33.3% out of 3 attempts from a single hawksbill turtle. Except for the single hawksbill turtle (Tanasanti et al., 2009), the current study obtained similar results to the previous
observations, with a 73.7% success rate out of 19 semen collection attempts from two individuals. Therefore, the author considered this method to be a viable tool for semen collection from hawksbill turtles.
In the semen collection study of green turtles by Wood et al. (1982), the rear flippers were observed to curl in response to the electronic stimulus. This behavior was also observed in the current study (Fig. 2-1-3A). The author suggests that the curling of the rear flippers is a diagnostic behavior of electoro-ejaculation. Furthermore, the author frequently observed the anteroposterior shaking of the neck just before ejaculation.
Previous studies have not reported the behavior of turtles during ejaculation; however, the observed shaking of the neck in the current study was very similar to the behavior frequently observed in mating captive male hawksbill turtles (Kawazu, unpubl. data).
Therefore, the author suggests that the anteroposterior shaking of the neck is a diagnostic behavior with ejaculation in male hawksbill turtles. During direct observations of mating, it is not possible to observe the moment of ejaculation, due to the penis being inserted into the female cloaca. Hence, the author suggests that the anteroposterior shaking of the neck presents a viable indicator for ejaculation during mating.
A previous study reported that semen pH was 5.5 for a single hawksbill turtle (Tanasanti et al., 2009). The semen pH for hawksbills in the present study was noticeably higher compared to the previously reported value (median: 7.4, Table. 2-1-1).
Previous studies that have collected semen from numerous animals have suggested that contamination with urine causes the lower semen pH values (Griggers et al., 2001;
Miyake, 2006). This hypothesis was supported by our results; the pH of the collected urine (median pH: 5.9) was lower compared to that of the semen, and was similar to the previously reported values. The pH of semen from domestic mammals and fishes is 6.5–7.7 (Miyake, 2006) and 7.5–8.5 (Alavi and Cosson, 2005), respectively, which is approximately similar compared to that obtained in the current study (pH inter-quartile range: 7.1–7.7, Table. 2-1-1). Therefore, the author suggested that the semen samples collected in the current study were less (or not at all) contaminated with urine compared to the previous studies. This pH difference might also explain the variability in sperm motility between the current study and the previous studies. In the previous study of the single hawksbill (Tanasanti et al., 2009), sperm motility was 60%, whereas the median of sperm motility in the current study rose from 2.5% to 39.5% and from 2% to 50% for Turtles no. 1 and 2, respectively, when normal semen was mixed with urine (Fig. 2-1-3).
This increase in motility in the presence of urine is extremely interesting, with this study being the first to report this phenomenon.
The sperm of marine and freshwater fish are mainly activated by osmolality that is created between the seminal fluid and surrounding medium (seawater and freshwater), which serve as osmotic and ionic signals (Cosson, 2012). When sea turtles mate, the male first mounts the female, then the male curls its tail under the female carapace to come to contact with the female cloaca, and the erected penis is inserted into the female cloaca prior to ejaculation (Miller, 1997). Thus, it is unlikely that marine water enters the cloaca of female sea turtles during mating, and the semen of male turtles is not surrounded by water at any point in this process, as is observed in fish (Cosson, 2012).
Therefore, sperm motility after ejaculation might be induced by the presence of urine in the cloaca of the male or female, and by the natural secretions in the female cloaca.
Sperm motility of several mammals tends to be compromised by highly anisosmotic solutions, such as urine; however, pH is probably not associated with the deleterious effects of urine on semen quality, since both substances have a similar pH (Santos et al., 2011). However, the pH value differed between semen and urine in the current study (Table. 2-1-1). This difference might indicate that the sperm motility of hawksbill turtles is regulated by changes in pH. A similar case in which semen is activated under low pH conditions has been reported for sea urchin (Christen et al., 1982). However, optimum sperm motility in rainbow trout occurs at pH 9.0 (Alavi and Cosson, 2005), which is the opposite of our results, in which the sperm of hawksbills was activated under low pH conditions (Fig. 2-1-3). Gist et al. (2000) reported that the sperm motility of painted turtles had little effect on spermatozoa at pH range of 5.9–8.4.
The semen collected in the current study was highly viscous, which would be alleviated when mixed with urine. Thus, sperm might also be activated through the relaxation of semen viscosity. Further study on the factors that activate sperm, as conducted on fish (Cosson, 2012), would help to clarify the presence of such relationships.
The serum or plasma testosterone levels of other sea turtles are known to change during the course of the year (termed a seasonal testosterone cycle), in which testosterone peaks during winter and early spring (Jessop et al., 2004; Licht et al., 1985;
Rostal, 2005; Valente et al., 2011; Wibbels et al., 1990). On the basis of the histological observations of the testis, Wibbels et al. (1990) and Rostal (2005) reported that
spermatogenesis is induced by increased testosterone levels in loggerhead and olive ridley turtles. In the current study, the testosterone concentration was high when the water temperature was low, 20–22 °C (between winter and early spring) (Fig. 2-1-4A and D). During this period, we collected semen samples showing relatively high TNSS and sperm motility from the 2 turtles (Fig. 2-1-4B and C). In male sea turtles, spermatogenesis is completed prior to the mating period (Licht et al., 1985; Wibbels et al., 1990; Rostal et al., 1998), with good quality semen (January–March, between winter and early spring) also being collected just before the mating season in the current study, because captive hawksbill turtle mates during April–May (spring) (Kobayashi et al, 2006; 2010a; Shimizu et al., 2005). Therefore, my results are consistent with the spermatogenic cycle of sea turtles recorded in previous studies that were based on histological and physiological analyses.
The sperm of male sea turtles is stored in the epididymis for 2–3 months after spermatogenesis (Hamann et al., 2003). However, in the current study, semen quantity and quality were low from April onward (Fig. 2-1-4B and C), which probably reflects the mating season (Kobayashi et al, 2006). In other words, this observation might be influenced by the frequency of semen collection. For instance, in the current study, semen was collected at 1–2 month intervals. This result indicates that the intervals between semen collections should be carefully selected; for instance, the author made one to two attempts per year spring onwards in the current study. Further study is needed to try to collect semen from non-ejaculated hawksbills (at least over a 1-year period) from spring onwards to determine the optimum time for semen collection.
Table. 2-1-1. Evaluation of semen collected by electro-ejaculation from 2 captive hawksbill turtles (n = 14 semen samples combined). The total number of surviving sperm (TNSS) was calculated for each ejaculation based on the results of semen volume, sperm concentration and viability.
Median Inter-quatile
range Median Inter-quatile
range Median Inter-quatile range
Semen volume (mL) 0.6 0.4-0.9 0.5 0.2-1.5 0.5 0.2-1.5
Sperm concentraion
(×106/mL) 450 247.5-1100 325 100-645 325 100-645
Semen pH 7.5 7.3-7.6 7.4 7.1-7.7 7.4 7.1-7.7
Urine pH 5.9 5.9-5.9 6 5.9-6.2 5.9 5.9-6.2
TNSS (×106) 235.5 29.9-733.5 180.5 12.6-750.5 180.5 12.6-750.5
Semen evaluations
No.1 turtle (n=4) No.2 turtle (n=10) All turtles (n=14)
Fig. 2-1-1. Photograph of a restrained hawksbill turtle and the instrument used for electrical stimulation. A: Hawksbill turtle on a retention stand, which is composed of 4 stacked car tires. B: Electro-transformer and electronic stimulus probe. C: Tip of the electronic stimulus probe. Black arrows indicate the 2 stainless steel electrodes. Photo by Isao Kawazu.
Fig. 2-1-2. Response of hawksbill turtles during electro-ejaculation. A: curling of the rear flippers. B: elongation of the penis. C: erection of the penis. D: formation of the urethral fissure. White arrow indicates semen. Photo by Isao Kawazu.
Fig. 2-1-3. Sperm motility before and after the semen was mixed with urine in 2 captive hawksbill turtles. Whiskers indicate range, box represents inter-quartile range (percentile: 25–75%) with median.
20 22 24 26 28 30
0 600 1200 1800
0 20 40 60 80
J F M A M J J A S O N D J F M 100
Water temperature(℃)TNSS(×106)Serum testosterone concentration(ng / mL)
Month
Sperm motility(%)
0 20 40 60 80 100
A
B
C
D
Fig. 2-1-4. Seasonal change in water temperature, TNSS, sperm motility, and serum testosterone concentration. Water temperature indicates the monthly means in holding tanks, TNSS indicates the total number of surviving sperm, and sperm motility indicates the sperm motility of semen mixed with urine. Black and light gray bars indicate Turtle no. 1 and 2, respectively.
2-2 Optimal Period of Semen Collection by Electro-ejaculation
2-2-1 Introduction
The second capter revealed that electro-ejaculation was viable for the semen collection of hawksbill turtles. However, the quality of the semen collected in the current study was poor from April onward (Fig. 2-1-4B and C), which probably reflects the mating season, based on previous success captive breeding programs (May−June;
Kobayashi et al., 2006; 2010a; Shimizu et al., 2005). For the development of an artificial insemination program for hawksbills, the author believes that semen must inject to female during mating season.
The result that the quality of semen was poor during mating season might be influenced by the frequency of semen collection. For instance, in the Chapter 2-1, semen was collected at 1–2 month intervals; hence, good quality semen that formed at low temperatures might have been ejaculated before March. I belive that further study is needed to try to collect semen from non-ejaculated hawksbills (at least over a 1-year period) from spring onwards to determine the optimum time for semen collection.
Again, two male turtles, which are indentical to Chapter 2-1, are also selected in this chapter, which are isolated from approximately 1 to 2 years (389−728 days) and are collected semen by electro-ejaculation. Subsequently, the author compared between total number of surviving sperm (TNSS) of this and second capter (inter-semen collection intervals: 1−2-month).
2-2-2 Materials and Methods
Two male turtles (Turtle no.1 and 2), their husbandury, semen collection and evaluation were indentical to those of Chapter 2-1; hence these informations were cut here (see Materials and Methods of Chapter 2-1).
Two male turtles were kept separately after their semen was collected in the Chapter 2-1. Semen were collected from 2 males at July of 2010, 2011, and 2013 (728 days), their inter-semen collection intervals were 485, 389, and 728 days, respectively.
The seminal characteristics of each ejaculate were measured, including semen volume, sperm concentration, and viability, the TNSS was caluculated based on there characteristics.
Mann-Whitney U-test was used to test for a significant difference in median of TNSS between this (inter-semen collection intervals: 1−2-year) and Chapter 2-1 (1−2-month). Spearman’s rank correlation coefficient was calculated for the correlation between inter-semen collection interval and total number of suviving sperm, including the results of this and Chapter 2-1.
2-2-3 Results
The mean meddian of TNSS in this chapter (1941×106, n = 6; Table 2-2-1) were significantly higher than that in chapter 2 (180.5×106, n = 20; Table 2-2-1;
Mann-Whitney U-test, P < 0.01). In particular, the TNSS obtained from Turtle no. 1 and 2 at July of 2013 were 7769.1×106 and 2453.0×106, respectively, which indicate the maximum values of both turtles so far (Table 2-2-1). Positive correlation between
inter-semen collection intervals and total number of suviving sperm at July, including the results of this and Chapter 2-1, was significant (Spearman’s rank correlation coefficient = 0.71、n = 8、P < 0.05; Fig. 2-2-1).
2-2-4 Discussion
Mating season is May–June, based on previous studies of mating success in captivity (Kobayashi et al., 2006; Shimizu et al., 2005). Despite semen were collected at July (after mating season), many surviving sperms were collected. Hence, this reslt indicates the turtles kept separately long-term (1−2-year) were collected many surviving semen during mating season. Therefore, the author suggests that the quality of semen would be affected by the frequency of semen collection. This fact is supported by positive collection interval and total number of suviving sperm at July, as shown in Fig.
2-2-1. Hence, the results indicate that the turtle kept in the sepatare tank (i.e. non-mating and ejaculation) is possible to be collected good semen mating season onwards.
Sea turtles have a seasonal spermatogenic cycle; many sperms are formed at early spring (Rostal, 2005; Wibbels et al., 1990; Chapter 2, 2-1), which are stored in ductus epididymis of turtles (Owens, 1980). This spermatogenesis is completed prior to mating season (Wibbels et al., 1990). Overall, these results indicate that sperms are stored between early spring (spermatogenesis) and mating season (summer). To date, however, this hypothesis of hawksbill turtles has not proved. This demonstration will exceed our knowledge about reproductive cycle of male sea turtles.
Seasonal spermatogenesis of sea turtles is associated with changes in testosterone
levels (Rostal, 2005; Wibbels et al., 1990; Chapter 2, 2-1). The teststrone when many surviving sperms were collected in this chapter (i.e., July) is evaluated minimum values baced on the result of Chapter 2, 2-1 as shown in Fig. 2-1-4. This minimum testosterone le el indicates that sperms are formed at July no longer. Therefore, the result that many surviving sperms were collected at July (Table 3-1) suggests that sperms would be stored in ductus epididymis, at least between May (early spring; maximum spermatnensis) and July (summer). This is strongly consistent with the hypothesis that sperms is stored from spermatogenesis to mating season.
The collection of viable semen is fundamental for successful artificial insemination.
The semen collection technique used here (Chapter 2, 2-1 and this chapter) contributes significant information about the utility of artificial insemination for endangered sea turtles. For the development of an artificial insemination program for hawksbills, we must also consider certain problems with respect to females, such as reproductive biology on vitellogenesis (i.e. age and body of the onset of vitellogenesis, vitellogenic cycle) and estrus. These reproductive information would be usual for the selection technique of females that is a good reproductive state and for the optimal period of insemination (i.e. semen injection), which contribute to improve the fertilized success.
Table 3-1-1 Seasonal change in the TNSS (total number of surviving sperm). The TNSS was calculated for each ejaculation based on the results of semen volume, sperm concentration, and sperm viability. *The data presented from January 2008 to March 2009 were extracted from Chapter 2-1.
2010 2011 2013
Jan. Mar. May. Jul. Aug. Oct. Nov. Jan. Feb. Mar. Jul. Jul. Jul.
Turtle no. 1 0 1638.0 39.0 0 0 2.7 0 432.0 0 - 114.7 1359.8 7769.1
Turtle no. 2 795.0 931.0 39.0 1.9 3.7 0.2 318.0 950.4 616.9 42.9 1466.3 997.2 2453.0 TNSS(×106)
Year Month
*2008 *2009
Ｉnter-semen collection interval (day) Total number of surviving sperm(×106)
0 2000 4000 6000 8000
0 200 400 600 800
Fig. 2-2-1 Relationship between the TNSS total number of surviving sperm from semen collected at July and inter-semen collection. The TNSS was calculated for each ejaculation based on the results of semen volume, sperm concentration, and sperm viability. Open and solid circles indicate Turtle no. 1 and 2, respectively. The data that inter-semen collection is 59 days were extracted from Chapter 2-1. The regression line is shown by Y = 7.0X - 1090.2 (Spearman’s rank correlation coefficient = 0.71, n = 8, P
< 0.05).