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Active peristaltic movements and fluid production of the mouse oviduct: their roles in fluid and sperm transport and fertilization

Active peristaltic movements and fluid production of the mouse oviduct: their roles in fluid and... Abstract To study how the oviduct behaves in relation to fluid secretion and sperm transport, ovary–oviduct–uterus complexes of the mouse were installed in a fluid-circulating chamber without disturbing the blood circulation or parasympathetic innervation. Injection of a bolus of Indian ink into the lower isthmus revealed very active adovarian peristalsis of the isthmus, which was most prominent during the periovulatory period. Oviduct fluid, secreted by the entire length of the isthmus, was rapidly transported to the ampulla and ovarian bursa before draining into the peritoneal cavity. The upper isthmus, in particular the isthmic–ampullary junction, was responsible for this adovarian fluid flow. Peristalsis of the oviduct, undisturbed flow of oviduct fluid from the isthmus to the peritoneal cavity, and the spermatozoon's own motility all contribute to efficient sperm ascent and to fertilization within the oviduct. Therefore, chemotaxis, rheotaxis, and thermotaxis of spermatozoa toward oocyte–cumulus complexes in the ampulla are all unlikely mechanisms for explaining sperm–oocyte contact and successful fertilization, given the rapid adovarian flow of oviduct fluid in this species. Introduction The mammalian oviduct is a long, thin tube linking the ovary to the uterus. Cumulus–oocyte complexes released from the ovary are picked up by the oviduct's fimbria and transported to the ampulla of the oviduct where they meet spermatozoa. Spermatozoa are normally deposited in the lower segment of the female tract, either the vagina or uterus. They pass through the uterotubal junction (UTJ) and the oviduct's isthmus before reaching the ampulla. Although sperm transport in the female tract has been studied extensively (for reviews, see [1–6]), there are still many subjects not fully understood and controversial. For example, how do spermatozoa ascend the oviduct from its lower segment to the ampulla? Some investigators maintain that molecules such as progesterone secreted by oocyte–cumulus complexes diffuse down to the isthmus and “direct” spermatozoa chemotactically toward the oocytes in the ampulla (e.g. [7–9]). Others claim that the ampulla is slightly warmer than the isthmus (e.g. [10]) and this thermal gradient directs spermatozoa to the ampulla (e.g. [11]). Still others have claimed that oviduct fluid drains down to the uterus and that spermatozoa swim rheotactically against this gradient toward the ampulla [12]. One must be aware that the oviduct is by no means a static structure. It displays a very active adovarian contractile movement as reported in the rabbit [13] and the golden hamster [14, 15]. A bolus (or boluses) of stained oil or India ink injected in the oviduct's isthmus is quickly transported to the ampulla by active peristaltic movements. Whereas Blandau and Gaddum-Rosse [13] observed this in the oviduct separated from animal's body, Battalia and Yanagimachi [14] examined it in the oviduct in situ. Nakaso [16] was one of the first to observe active peristaltic contractions of oviducts in a live animal (the rabbit). That study used a transparent celluloid (nitrocellulose-based plastic) window installed in the rabbit's abdomen. Injection of a bolus of staining solution in oviduct's lumen made observations of oviduct's contractility easier. Muro et al. [17] observed a steady migration of mouse spermatozoa through the UTJ after pulling the entire ovary–oviduct–uterus complex out of the body through a flank incision. Ishikawa et al. [18] isolated oviducts of mated female mice and examined the movement of spermatozoa within the oviducts, which had been immobilized by prior injection of padrin (3-diphenylmethylen-1,1-dieyl-2-methylpyrrolidine bromide) into the animals. They found that the oviduct's contractile movements are important for sperm transport from the low to the mid-isthmus, and that sperm motility by itself is necessary for sperm migration from the isthmus to the ampulla. Certainly, we are able to obtain much important information about sperm–oviduct interactions by using oviducts isolated from an animal's body, but we must be aware that what happens in isolated organs does not necessarily happen in situ. Here, we adopted the technique of Battalia and Yanagimachi [14], which allowed us to examine oviducts still attached to the animal's body without disturbing the blood circulation or parasympathetic innervation. The use of transgenic mice producing doubly fluorescent spermatozoa made our examination of sperm transport within the oviduct easier. Retrospective examinations of video recordings enabled analyses of the oviduct's contractility and fluid flow within it. Materials and methods Animals ICR and B6D2F1 strain mice were obtained from the Charles River Co. (Kanagawa, Japan). Transgenic (TG) male mice [19] were obtained from RIKEN through the National Bio-Resource Project of the Ministry of Education, Culture, Sports, Science and Technology, Japan. Spermatozoa of these TG male mice express green fluorescent protein in their acrosomes and red fluorescent protein in the midpiece mitochondria. All mice were maintained in air-conditioned rooms (23 ± 2°C, 45 ± 5% humidity) with light between 5:00 and 19:00 h. Under these lighting conditions, females are expected to mate between 20:00 and 01:00 h, ovulate around 03:00 h [20], and most oocytes are fertilized between 06:00 and 09:00 h [21]. For some experiments, female mice were maintained in a room with reversed lighting conditions (light between 15:00 and 05:00 h) for 2 weeks or more. This made females mate between 06:00 and 11:00 h and ovulate around 13:00 h, which allowed us to perform operations during ordinary working hours. For some experiments (e.g. comparison of transport of acrosome-intact and acrosome-reacted spermatozoa in the oviduct), female mice were induced to ovulate by intraperitoneal injection of equine chorionic gonadotropin (eCG; 2.5–5.0 IU) irrespective of the stage of the estrous cycle followed by an injection of human chorionic gonadotropin (hCG; 2.5–5.0 IU) 48 h later. Ovulation is known to take place around 12 h after hCG injection. All experiments conformed to the Guidelines for Animal Experiments of Asahikawa Medical University. Chemicals and media All chemicals were purchased from Nacalai Tesque Inc. (Kyoto, Japan) unless otherwise stated. The medium used for perfusion of an exposed ovary–oviduct–uterus complex was Dulbecco's Ca2+- and Mg2+-containing phosphate-buffered saline (PBS+) supplemented with 0.1% bovine serum albumin (BSA; Sigma-Aldrich, St. Louis, MO) (PBS-BSA, pH 7.3). The medium used for the examination of oocytes and spermatozoa as well as insemination in vivo and in vitro was bicarbonate-buffered TYH medium [22], which consisted of 119.37 mM NaCl, 4.79 mM KCl, 1.71 mM CaCl2, 1.19 mM KH2PO4, 1.19 mM MgSO4, 25.07 mM NaHCO3, 5.56 mM glucose, 1.00 mM sodium pyruvate, 4 mg/ml BSA (AlbuMax; GibcoBRL, Auckland, New Zealand), 50 μg/ml streptomycin sulfate, and 75 μg/ml penicillin G potassium, pH 7.4 when equilibrated with 5% (v/v) CO2 in air. Observations of oviduct movements and oviduct fluid flow in situ The setup we used for examination of oviducts in situ was essentially the same as that used by Battalia and Yanagimachi [14] (Figure 1). Briefly, a female mouse was anesthetized with isoflurane vapor, and the ovary/oviduct and part of the uterus were pulled out through a flank incision. They were then bathed with running warm PBS-BSA. Temperature of the PBS-BSA and the complete coverage of exposed organs with PBS-BSA were of critical importance for keeping the oviduct actively motile. At 30°C or below, oviducts became completely immotile. Complete coverage of organs with PBS-BSA was also important, or peristaltic motion of the oviduct stopped or slowed down noticeably. Preliminary experiments showed that oocytes could be fertilized normally in oviducts during 4 h of anesthesia of the animal. Figure 1. View largeDownload slide Diagrams of the apparatus used for observing an ovary–oviduct–upper uterus complex. (A) Entire view. Warmed (37°C) PBS is pumped through the perfusion collar attached to the dorsal surface of an anesthetized female mouse with the reproductive tract exposed via a flank incision. (B) Temperature of the circulating medium is monitored using a temperature sensor. (C) An oviduct undergoing active peristalsis can be seen under the dissecting microscope. Fertilization can proceed normally in the oviduct under such conditions. Figure 1. View largeDownload slide Diagrams of the apparatus used for observing an ovary–oviduct–upper uterus complex. (A) Entire view. Warmed (37°C) PBS is pumped through the perfusion collar attached to the dorsal surface of an anesthetized female mouse with the reproductive tract exposed via a flank incision. (B) Temperature of the circulating medium is monitored using a temperature sensor. (C) An oviduct undergoing active peristalsis can be seen under the dissecting microscope. Fertilization can proceed normally in the oviduct under such conditions. To visualize the fluid flow within the oviduct lumen and to ease our ability to observe the oviduct's contractions, approximately 0.1 μl of India ink was injected into the lower part of the oviduct isthmus through a fine injection pipette (∼20 μm in diameter). The ink had been dialyzed overnight against PBS+ and diluted with PBS-BSA before use. In one series of experiments, muscular contraction of the oviduct was inhibited by circulating PBS-BSA containing 20 μM nicardipine (Merck KGaA, Darmstadt, Germany) through the perfusion collar. Oviducts arrested peristalsis by 5–10 min after this treatment. India ink was then injected into the lower isthmus. Movements of oviducts and the behavior of ink within the oviducts were recorded with a digital movie camera. The time needed for ink to reach the ampulla was determined by retrospective examination of the movies. Measurement of the volume of fluid secreted by the oviduct To determine how much fluid the oviduct secretes during the estrous period, we ligated one of the two oviducts at two places using sterile 11–0 nylon threads (Bear Medic Corp, Ibaraki, Japan): one at the upper ampulla near ovarian bursa and the other at the UTJ. The oviduct of the other side was left untouched and served as the control. One hour later, the mouse was euthanized and the ligated oviduct (now distended) was isolated along two loops of the threads. The oviduct surface was carefully wiped clean using absorbent tissue paper, removing any blood and body fluid. After weighing the oviduct, it was put on a filter paper, cut into seven to eight pieces, and compressed under another filter paper to remove all the fluid. The oviduct was then reweighed. We assumed that the relative density of the oviduct fluid would be about the same as that of distilled water (1 mg = 1 μl) and estimated the volume of fluid that had accumulated within the isolated region of the oviduct for 1 h. The contralateral control oviduct was ligated at two places—the upper ampulla and the UTJ—separated from the animal body and weighed before and after removal of fluid to estimate the volume of fluid in this region of the oviduct. In another experiment, oviducts were ligated at four points: (1) the UTJ, (2) the lower- and mid-isthmus junction, (3) the mid- and upper isthmus junction, and (4) upper isthmic–ampullary junction (IAJ). The oviducts were examined 30 min or 2 h later to determine which parts were swollen from fluid accumulation. Observation of sperm ascent in the oviduct A postovulatory female mouse (14–15 h after hCG injection) was anesthetized and its ovary–oviduct–uterus complex was installed in the perfusion collar. After the oviduct was completely submerged in running PBS-BSA, a small amount (0.1 μl) of sperm suspension was injected into the lower isthmus. The spermatozoa were obtained from the cauda epididymidis of the TG male mice described above. The time and number of spermatozoa reaching the ampulla were determined. Spermatozoa injected into the isthmus were acrosome-intact (fresh cauda epididymal spermatozoa suspended for 5–10 min in TYH medium), 100% acrosome-reacted (AcR), or 100% dead. The 100% AcR spermatozoa were obtained by preincubating cauda epididymal spermatozoa for 2 h in TYH medium at 37°C under 5% CO2 in air before treatment with 20 μM Ca2+-ionophore A23187 [23]. Spermatozoa immobilized by the ionophore were centrifuged (550 g, 5 min) and pelleted spermatozoa were overlaid with fresh TYH medium to allow them to swim up into the medium. Virtually 100% of swim-up spermatozoa were AcR and vigorously motile. We also prepared 100% dead spermatozoa by freezing the suspension of cauda epididymal spermatozoa in liquid nitrogen for a few minutes without cryoprotection. Insemination was done by depositing approximately 0.1 μl of sperm suspension (∼1–5 × 103/μl) into the lower isthmus. Various times after insemination, oviducts were isolated, put on a glass slide, and compressed slightly under a coverslip before examination of spermatozoa in the lumen using a fluorescence microscope with a highly sensitive digital camera (α7s, Sony). We could see spermatozoa in oviducts under fluorescence microscopy after oviducts were separated from the animal's body. In some experiments, TG spermatozoa were suspended in TYH medium containing a small amount of India ink before injection into the isthmus. This enabled us to examine the movement of spermatozoa and fluid (ink) within the oviduct simultaneously. Assessment of fertilization In some experiments, sperm fertilizing capacity within oviducts was assessed. A drop of TYH medium containing 100% AcR spermatozoa was injected into the lower isthmus. At 4 h later, oocytes were collected from the ampulla (or the uterus in some experiments) and examined for evidence of fertilization. An oocyte with two pronuclei and two polar bodies (6–8 h after sperm deposition in the isthmus) was considered fertilized. Statistical analysis All percentile data were transformed into arcsine values for statistical analysis to normalize them. When data were compared between two groups (distributions of spermatozoa in the oviduct after sperm injection into the lower isthmus), one-way analysis of variance (ANOVA) and Welch's t-test were used. When the data were compared among more than two groups (fertilization rate), one-way ANOVA and Tukey–Kramer methods were used. All statistical analyses were performed with R software (version 3.2.3; https://www.r-project.org). Results Structure of the mouse ovary–oviduct–uterus complex The mouse oviduct is a tortuous tube. Its upper end opens to the ovarian bursa and the lower end connects to the uterus via the UTJ. Figure 2 shows photographs of the oviduct (A and B), and semi-diagrammatic (C) and highly diagrammatic (D) illustrations of the ovary, the oviduct and the upper region of the uterus. The oviduct comprises (a) the infundibulum, (b) the ampulla, (c) the isthmus, and (d) the UTJ. Both the ovary and the infundibulum of the oviduct are encapsulated by the ovarian bursa, a membranous sac with a small hole that opens into the peritoneal cavity (Figure 3D) [24]. As described below, oviduct fluid drains into the ovarian bursa and then passes through the bursa hole into the peritoneal cavity. Figure 2. View largeDownload slide (A) A mouse oviduct, slightly stretched, as seen through an ordinary dissecting microscope. (B) An oviduct extensively stretched, flushed with PBS containing FITC-conjugated WGA and observed using fluorescence microscopy. (C , D) Schemata of the anatomy of the ovary, oviduct, and uterus. The site of ink and sperm injection is shown in diagram D. IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. Figure 2. View largeDownload slide (A) A mouse oviduct, slightly stretched, as seen through an ordinary dissecting microscope. (B) An oviduct extensively stretched, flushed with PBS containing FITC-conjugated WGA and observed using fluorescence microscopy. (C , D) Schemata of the anatomy of the ovary, oviduct, and uterus. The site of ink and sperm injection is shown in diagram D. IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. Figure 3. View largeDownload slide (A) Diagrams showing the behavior of ink injected into the lower isthmus of an estrous female. A bolus of ink in the lower isthmus ascends the oviduct in a to-and-fro fashion, is fragmented, and drains into the peritoneal cavity via the bursa hole. IAJ, isthmic-ampullary junction; IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. (B) Oviduct and uterus in situ showing the site of Ink injection; this image was captured from Supplemental Movie 1 showing the behavior of injected ink in the oviduct. (C, D) Diagrams of the structural relationships among the uterus, oviduct, ovary, and ovarian bursa. The ovary is completely encapsulated by the bursa membrane. Fluid within the bursa drains into the peritoneal cavity via the bursa hole. Note the presence of the bursa hole (arrow) in the mesosalpinx covering the top of the uterus. Figure 3. View largeDownload slide (A) Diagrams showing the behavior of ink injected into the lower isthmus of an estrous female. A bolus of ink in the lower isthmus ascends the oviduct in a to-and-fro fashion, is fragmented, and drains into the peritoneal cavity via the bursa hole. IAJ, isthmic-ampullary junction; IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. (B) Oviduct and uterus in situ showing the site of Ink injection; this image was captured from Supplemental Movie 1 showing the behavior of injected ink in the oviduct. (C, D) Diagrams of the structural relationships among the uterus, oviduct, ovary, and ovarian bursa. The ovary is completely encapsulated by the bursa membrane. Fluid within the bursa drains into the peritoneal cavity via the bursa hole. Note the presence of the bursa hole (arrow) in the mesosalpinx covering the top of the uterus. The thin-walled ampulla is where oocytes and spermatozoa meet during normal fertilization in vivo. Numerous motile cilia line the ampulla, infundibulum, and the inner aspect of the fimbria. The transitional region between the ampulla and the isthmus is the IAJ. The isthmus, except for its uppermost region, has numerous transverse ridges and pockets on its inner wall. In mice and many other animals, the isthmus serves as the sperm reservoir prior to fertilization in vivo [3, 25, 26, 27]. The UTJ is a thick-walled “straight” tube with longitudinal mucosal folds, connecting the isthmus to the uterus. Active contractions of the oviduct and rapid transport of luminal fluid The oviduct of estrous females displayed a very active contraction–relaxation movement. This movement could be seen clearly when a bolus of India ink (in PBS-BSA) was injected into the lower isthmus (Figure 3A and B; see Supplemental Movie S1 in real time). The ink bolus was quickly divided into smaller boluses by the repeated contraction–relaxation (segmenting) movements of the isthmus wall. These moved up (i.e. adovarian) and down. Because the upward movement was greater than the downward movement, all boluses moved steadily toward the oviduct's ampulla, some faster than others. This back and forth movement of each ink bolus continued until it reached the upper isthmus. In this region, the ink did not go back (i.e. toward the uterus) because of the unidirectional (adovarian) contractions of the upper one third of the upper isthmus. The most prominent adovarian motion of the oviduct was seen in the IAJ (Supplemental Movie S2 in real time). Ink quickly entering the ampulla hit the cumulus mass, stayed for a while but gradually moved to the end of the upper ampulla and to the ovarian cavity, and finally drained into the peritoneal cavity via the bursa hole (Figure 3C and D; Supplemental Movie S3). To our surprise, the ink put into the isthmus was transported to the ampulla at any time during the estrous cycle and even during early pregnancy (Figure 4; Supplemental Movie S4 comprising nine movies). However, the fastest ink transport to the ampulla occurred during the periovulatory period. We found that during estrous cycle (days 1–4) most of the ink injected into the lower isthmus was transported to the ampulla, whereas during early pregnancy (days II–IV) most of the ink remained in the lower and mid isthmus for a long time. Interestingly, nicardipine (20 μM) in PBS-BSA running through the perfusion collar did not prevent ink from ascending to the ampulla, even though this drug stopped the oviduct's muscular contractions completely. Apparently, the ink was able to ascend the quiescent oviduct along with the fluid secreted from the isthmus (see below). Figure 4. View largeDownload slide Time (in second) the India ink bolus took to travel from the lower isthmus to the ampulla. Each dot represents a single experiment. Note that the fastest ink (fluid) transport is seen during the peri-ovulatory period. Figure 4. View largeDownload slide Time (in second) the India ink bolus took to travel from the lower isthmus to the ampulla. Each dot represents a single experiment. Note that the fastest ink (fluid) transport is seen during the peri-ovulatory period. The entire length of the oviduct's isthmus secretes fluid continuously When both the IAJ and the UTJ were ligated and examined 1.5 h later, the entire length of the isthmus was seen to be swollen extensively. When the isthmus was ligated at four places as shown in Figure 5, all segments, in particular the upper isthmus with a thinner wall, were swollen extensively, indicating that the entire length of isthmus secretes fluid actively. Apparently, under normal conditions the oviduct fluid drains into the ovarian capsule, then into the peritoneal cavity via the bursa hole. We estimated that the isthmus of estrous females contains approximately 0.9 μl of fluid at any time and secretes about 2.2 μl of fluid per hour (averages of three determinations for each). Even when oviducts were immobilized by nicardipine, they kept secreting the same volume of the fluid. Ink injected into the lower isthmus of such oviducts ascended to the ampulla within 2–4 min, sometimes in more than 5 min. Figure 5. View largeDownload slide Isthmus immediately after 0 min and 30–120 min after four ligations of the isthmic region of the oviduct. Note that all segments of the isthmus expand with fluid accumulation. Figure 5. View largeDownload slide Isthmus immediately after 0 min and 30–120 min after four ligations of the isthmic region of the oviduct. Note that all segments of the isthmus expand with fluid accumulation. Both the oviduct's contractility and the upward flow of oviduct fluid are important for efficient sperm ascent and fertilization To investigate whether the oviduct's contractility is important for sperm transport and fertilization, we first immobilized oviducts using nicardipine, and then deposited about 100 live AcR TG spermatozoa in the lower isthmus of each oviduct. Oviducts bathed with PBS-BSA free of nicardipine served as controls. The results (Figure 6A) show that in the control (motile) oviducts, both live and dead spermatozoa, in particular live AcR spermatozoa, were rapidly transported from the lower isthmus to the ampulla. When live, “fresh” spermatozoa (which were mostly uncapacitated and acrosome-intact) were injected, more than 90% attached to epithelium of the lower isthmus and did not ascend to the ampulla. Within immobilized oviduct, on the other hand, none or only few spermatozoa ascended to the ampulla. When 100% AcR live spermatozoa were put in the isthmuses of intact oviducts and the eggs were examined 4 h later for evidence of fertilization, 94% of eggs were fertilized, whereas only 20% were fertilized in immobilized oviducts (Figure 6B), indicating the importance of oviduct's contractility for efficient fertilization in vivo. It should be noted that both immobilized and motile (control) isthmuses produced the same amount of oviduct fluid (∼2.3 μl/h). Figure 6. View largeDownload slide (A) Sperm ascent in normal (motile) and immobilized oviducts. About 100 TG spermatozoa, live or dead, were put in the lower isthmus. Five and twenty minutes later, the number of spermatozoa in the ampulla was determined. Note that inhibition of oviduct contractility by nicardipine results in a drastic reduction of sperm ascent to the ampulla. (B) Immobilization of oviduct by nicardipin results in a drastic reduction in fertilization rate. Mean ± SD. (C) Prevention of oviduct fluid flow into ovarian bursa/peritoneal cavity by ligation of the upper ampulla results in fertilization failure. Mean ± SD. Figure 6. View largeDownload slide (A) Sperm ascent in normal (motile) and immobilized oviducts. About 100 TG spermatozoa, live or dead, were put in the lower isthmus. Five and twenty minutes later, the number of spermatozoa in the ampulla was determined. Note that inhibition of oviduct contractility by nicardipine results in a drastic reduction of sperm ascent to the ampulla. (B) Immobilization of oviduct by nicardipin results in a drastic reduction in fertilization rate. Mean ± SD. (C) Prevention of oviduct fluid flow into ovarian bursa/peritoneal cavity by ligation of the upper ampulla results in fertilization failure. Mean ± SD. What will happen when oviduct's fluid flow from the isthmus to the peritoneal cavity is blocked? Will oocytes be fertilized normally? Figure 6C summarizes the results of experiments in which the upper ampulla was ligated to prevent the ascent of oviduct fluid to the ovarian bursa. When 100% AcR spermatozoa were put in the lower isthmus 15 min after ligation of the upper ampulla, about half of the oocytes were fertilized by 4 h. The oocytes were found either in the oviduct (3/5 animals) or both the oviduct and uterus (2/5 animals). In contrast, none of the oocytes was fertilized when spermatozoa were put into the lower isthmus 1 h after ligation of the upper ampulla. We found the oviducts very much distended by 1 h after ligation of the upper ampulla. India ink put into the lower isthmus along with spermatozoa did not ascend to the ampulla. Instead, both ink and spermatozoa descended to the uterus. Surprisingly, some oocytes also descended from the ampulla to the uterus. In one instance, we saw the ampulla rupture. In control (nonligated) oviducts, all oocytes were in the ampulla and all or most of them were fertilized by the time of examination. Discussion Role of the upward flow of oviduct fluid for fertilization in vivo As reported here, the entire region of the mouse oviduct secretes fluid very actively. The fluid is transported from the isthmus to the ampulla because of the predominantly one-way (upward) pumping action of the upper one-third of the isthmus, in particular the IAJ. The fluid eventually drains into the peritoneal cavity via the bursa hole. Because the distance between the lower isthmus and the ampulla is about 12 mm in the mouse and ink travels this distance in about 30 s in a to-and-fro manner, the fluid in the oviduct must move faster than about 400 μm/s. Although such upward flow of the oviduct fluid was seen during any time of the estrous cycle, it was most prominent during the periovulatory period. Such upward flow of the oviduct fluid must be important for fertilization in vivo. Prevention of upward flow of oviduct fluid by ligation of the upper ampulla after ovulation results in not only swelling of the ampulla, but also flashing of oocytes down to the uterus without fertilization. The swollen ampulla may even rupture. Role of the spermatozoon's own movement for fertilization in vivo Chang and Suarez [28] maintained that the spermatozoon's movement alone is enough for sperm transport from the isthmus to the ampulla to fertilize oocytes. This was based on experiments in which isolated oviducts of mated female mice were maintained in a medium containing the muscle-immobilizing drug nicardipine. Ishikawa et al. [18], who did similar experiments concentrating on the contractile role of the myosalpinx, considered that the spermatozoon's own motility is necessary for ascent from the mid-isthmus to the ampulla. Here, we found that (i) adovarian peristaltic movements of the isthmus; (ii) secretion and adovarian flow of oviduct fluid; and (iii) the spermatozoon's own active movement all contribute to efficient sperm ascent from the isthmus to the ampulla. A scheme of fertilization in vivo Based on information obtained from previous and this study, we propose that the following events likely happen in the female tract during natural fertilization in the mouse and common laboratory rodents. During and/or after coitus, millions of spermatozoa are deposited in the uterus, but only a small percentage passes through the UTJ. Those in the UTJ do not exhibit high amplitude tail oscillations, yet they are able to advance steadily to the lower isthmus of the oviduct [17]. Some spermatozoa reaching the isthmus may die there. They are quickly transported to the upper oviduct before draining into the peritoneal cavity (they are the so-called “vanguard” spermatozoa but play no part in fertilization) [29, 30]. Spermatozoa reaching the lower isthmus have not completed capacitation [17, 31]. They are held there, with heads firmly attached to the isthmic epithelium until they are capacitated and hyperactivated [5, 32]. It is unlikely that all spermatozoa are capacitated synchronously. Some must be capacitated and AcR faster than others [21]. Some might never be capacitated and simply die in the isthmus. Capacitated ones with lower binding ability to isthmic epithelium than uncapacitated ones [32], lift off the epithelium assisted by their vigorous (hyperactivated) tail movement [26, 33]. After swimming free for a while, they reattach to the isthmic wall. Adovarian oviduct peristalsis and fluid flow help to direct spermatozoa toward the ampulla. It is known that there are fewer spermatozoa around the oocytes in the ampulla during the progression of fertilization until almost all oocytes are fertilized [17, 34]. This does not necessarily mean that only a few spermatozoa reach the ampulla. It is more likely that both live and dead spermatozoa keep ascending the oviduct before, during, and even after fertilization. When a cumulus–oocyte complex is in the ampulla, live spermatozoa are trapped by the cumulus and enter the oocytes [21, 35]. Even after all the oocytes have been fertilized, spermatozoa that did not participate in fertilization drain out of animal's body or are phagocytized by epithelial cells [36, 37] or keep ascending the oviduct and drain into the peritoneal cavity (see Supplemental figure). On chemotaxis, rheotaxis, and thermotaxis of spermatozoa during fertilization in vivo How spermatozoa reach oocytes in the oviduct's ampulla has been the subject of much debate. Many investigators speculated that factors (e.g. progesterone) secreted from the oocyte and surrounding cumulus cells guide spermatozoa chemotactically toward the egg (e.g. [7, 9, 38]). Even though such chemoattractants might help orient spermatozoa toward oocytes in the immediate vicinity, it is difficult to conceive that the attractant could diffuse far down to the isthmus to guide spermatozoa. The oviduct's predominant fluid flow is upward (adovarian) rather than downward, as we report here. It has been known for many years that the presence of ovulation products (oocytes, cumulus oophorus, and follicular fluid) within the ampulla enhances sperm ascent from the isthmus to the ampulla (e.g. [39, 40]), but this can be explained by the sperm-trapping action of the cumulus oophorus rather than by sperm chemotaxis. Some other investigators have postulated that oviduct fluid flows down from the oviduct to the uterus and that spermatozoa swim rheotactically (upward) from the uterus to the ampulla of the oviduct [12]. Even though spermatozoa may swim rheotactically when observed using a microscope [41], it is most unlikely that spermatozoa within the lower isthmus of oviduct move toward the ampulla rheotactically, because the oviduct fluid is pumped up from the isthmus to the ampulla by adovarian peristalsis of the upper isthmus, in particular the IAJ. Other investigators have suggested that the temperature at the oviduct ampulla is slightly higher (1–2°C) than at the lower isthmus and uterus, and this temperature difference directs spermatozoa toward the ampulla [42]. Here, we bathed the mouse uterus–oviduct–ovary complex in running PBS with a constant temperature of 37°C. Because the temperature of the uterus, oviduct, and ovary is expected to be the same and fertilization proceeded normally under this experimental condition, it is most unlikely that any higher temperature of the ampulla is of critical importance for successful fertilization in vivo. Biological importance of the oviduct and problems to be studied further Even though the oviduct can be bypassed in assisted reproductive technologies such as in vitro fertilization and intracytoplasmic sperm injection, it is the oviduct where oocytes are normally fertilized and start to develop. Any errors in interactions between spermatozoa and oocytes and the development of embryos within the oviduct could have lasting detrimental effects on the well-being of offspring. The oviduct is not a simple tube; it provides an ideal environment for the spermatozoon's preparation for fertilization, fertilization itself, and the preimplantation development of zygotes. Several oviduct-related topics to be investigated further include (1) the origin and fate of oviduct fluid; (2) the effect of inhibition of oviduct fluid secretion; (3) the site and mechanism of sperm capitation, hyperactivation, and acrosome reaction within the oviduct, which are surprisingly still not yet fully understood [5, 43–45]; (4) the fate of surplus spermatozoa in the uterus, oviduct, and peritoneal cavity; (5) hormonal control of oviduct fluid secretion, oviduct peristalsis, and sperm transport in the oviduct; (6) confirmation of the proposed role of cell surface progesterone receptors of ciliary cells in the ampulla [46]; (7) the nature and function of “pacemaker cells” (interstitial cells of Cajal or telocytes) in the oviduct [47]; and (8) role of oviduct's peristalsis and fluid secretion in the transport of fertilized eggs and preimplantation embryos through the oviduct. These all undoubtedly play crucial roles in the coordinated movement of the oviduct before and during fertilization as well as in the development of preimplantation embryos. In this study, we have emphasized the importance of oviduct fluid flow from the lower to the higher segments of the oviduct for sperm ascent and fertilization. During the course of this study, we found that India ink injected into the ovarian bursa does not descend to the oviduct's ampulla (unpublished data), whereas cumulus-enclosed and cumulus-free oocytes as well as live and dead spermatozoa, if not all, are transported from the ovarian bursa down to the ampulla, apparently propelled by ciliary movement of the epithelia of both the fimbria and infundibulum. That mouse spermatozoa deposited into the ovarian bursa can fertilize oocytes in the ampulla [48] can be explained by ciliary movement of the epithelia of both fimbria and infundibulum. Fertilization of oocytes in the oviduct following intraperitoneal injection of spermatozoa is well known in various animals [3]. The differences among species with regard to the movement of spermatozoa from the isthmus to the ampulla as well as the role of the oviduct's pacemaker cells (interstitial cells of Cajal or telocytes; [47, 49, 50]) in sperm and egg transport must be the subject of future investigations. Supplementary data Supplemental Figure. An ICR female at D4 was caged with a TG male in the evening. Next morning, the formation of a copulation plug was confirmed. The mated female was sacrificed and the peritoneal cavity was washed thoroughly with 10 mL PBS (+) with 1% Triton-X and 0.1% BSA, which was thereafter centrifuged. The presence of the spermatozoa was examined under the microscope. Seven spermatozoa, of which six were separated heads only, were recovered from the peritoneal cavity after coitus. Supplemental Movie S1. A real-time movie showing ascendance of India ink injected in the lower isthmus. At 23:00 h of day 4 of estrous cycle. Supplemental Movie S2. A real-time movie of the isthmic–ampullary junction showing unidirectional (upward) pumping contractions. At 5:00 h of day 1 of estrous cycle. Supplemental Movie S3. A movie of India ink draining from the ovarian bursa into the peritoneal cavity via the bursa orifice. Supplemental Movie S4. Nine movies of oviducts displaying peristaltic movement at various times of estrus cycle (day 1–4 of estrous cycle) and early pregnancy (day 2–4 of pregnancy). Acknowledgment We thank Dr Hiroyuki Tateno for providing invaluable information and advice. We are grateful to Dr. Susan S. Suarez (Cornell University) and Dr. James M. Cummins (Murdoch University) for their invaluable advice. 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Interstitial Cajal-like cells (ICLC) as steroid hormone sensors in human myometrium: immunocytochemical approach . J Cell Mol Med 2006 ; 10 : 789 – 795 . Google Scholar Crossref Search ADS PubMed WorldCat © The Author(s) 2019. Published by Oxford University Press on behalf of Society for the Study of Reproduction. This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Biology of Reproduction Oxford University Press

Active peristaltic movements and fluid production of the mouse oviduct: their roles in fluid and sperm transport and fertilization

Hino,, Toshiaki; Yanagimachi,, Ryuzo
Biology of Reproduction , Volume 101 (1) – Jul 1, 2019
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Oxford University Press
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© The Author(s) 2019. Published by Oxford University Press on behalf of Society for the Study of Reproduction.
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0006-3363
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1529-7268
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10.1093/biolre/ioz061
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Abstract

Abstract To study how the oviduct behaves in relation to fluid secretion and sperm transport, ovary–oviduct–uterus complexes of the mouse were installed in a fluid-circulating chamber without disturbing the blood circulation or parasympathetic innervation. Injection of a bolus of Indian ink into the lower isthmus revealed very active adovarian peristalsis of the isthmus, which was most prominent during the periovulatory period. Oviduct fluid, secreted by the entire length of the isthmus, was rapidly transported to the ampulla and ovarian bursa before draining into the peritoneal cavity. The upper isthmus, in particular the isthmic–ampullary junction, was responsible for this adovarian fluid flow. Peristalsis of the oviduct, undisturbed flow of oviduct fluid from the isthmus to the peritoneal cavity, and the spermatozoon's own motility all contribute to efficient sperm ascent and to fertilization within the oviduct. Therefore, chemotaxis, rheotaxis, and thermotaxis of spermatozoa toward oocyte–cumulus complexes in the ampulla are all unlikely mechanisms for explaining sperm–oocyte contact and successful fertilization, given the rapid adovarian flow of oviduct fluid in this species. Introduction The mammalian oviduct is a long, thin tube linking the ovary to the uterus. Cumulus–oocyte complexes released from the ovary are picked up by the oviduct's fimbria and transported to the ampulla of the oviduct where they meet spermatozoa. Spermatozoa are normally deposited in the lower segment of the female tract, either the vagina or uterus. They pass through the uterotubal junction (UTJ) and the oviduct's isthmus before reaching the ampulla. Although sperm transport in the female tract has been studied extensively (for reviews, see [1–6]), there are still many subjects not fully understood and controversial. For example, how do spermatozoa ascend the oviduct from its lower segment to the ampulla? Some investigators maintain that molecules such as progesterone secreted by oocyte–cumulus complexes diffuse down to the isthmus and “direct” spermatozoa chemotactically toward the oocytes in the ampulla (e.g. [7–9]). Others claim that the ampulla is slightly warmer than the isthmus (e.g. [10]) and this thermal gradient directs spermatozoa to the ampulla (e.g. [11]). Still others have claimed that oviduct fluid drains down to the uterus and that spermatozoa swim rheotactically against this gradient toward the ampulla [12]. One must be aware that the oviduct is by no means a static structure. It displays a very active adovarian contractile movement as reported in the rabbit [13] and the golden hamster [14, 15]. A bolus (or boluses) of stained oil or India ink injected in the oviduct's isthmus is quickly transported to the ampulla by active peristaltic movements. Whereas Blandau and Gaddum-Rosse [13] observed this in the oviduct separated from animal's body, Battalia and Yanagimachi [14] examined it in the oviduct in situ. Nakaso [16] was one of the first to observe active peristaltic contractions of oviducts in a live animal (the rabbit). That study used a transparent celluloid (nitrocellulose-based plastic) window installed in the rabbit's abdomen. Injection of a bolus of staining solution in oviduct's lumen made observations of oviduct's contractility easier. Muro et al. [17] observed a steady migration of mouse spermatozoa through the UTJ after pulling the entire ovary–oviduct–uterus complex out of the body through a flank incision. Ishikawa et al. [18] isolated oviducts of mated female mice and examined the movement of spermatozoa within the oviducts, which had been immobilized by prior injection of padrin (3-diphenylmethylen-1,1-dieyl-2-methylpyrrolidine bromide) into the animals. They found that the oviduct's contractile movements are important for sperm transport from the low to the mid-isthmus, and that sperm motility by itself is necessary for sperm migration from the isthmus to the ampulla. Certainly, we are able to obtain much important information about sperm–oviduct interactions by using oviducts isolated from an animal's body, but we must be aware that what happens in isolated organs does not necessarily happen in situ. Here, we adopted the technique of Battalia and Yanagimachi [14], which allowed us to examine oviducts still attached to the animal's body without disturbing the blood circulation or parasympathetic innervation. The use of transgenic mice producing doubly fluorescent spermatozoa made our examination of sperm transport within the oviduct easier. Retrospective examinations of video recordings enabled analyses of the oviduct's contractility and fluid flow within it. Materials and methods Animals ICR and B6D2F1 strain mice were obtained from the Charles River Co. (Kanagawa, Japan). Transgenic (TG) male mice [19] were obtained from RIKEN through the National Bio-Resource Project of the Ministry of Education, Culture, Sports, Science and Technology, Japan. Spermatozoa of these TG male mice express green fluorescent protein in their acrosomes and red fluorescent protein in the midpiece mitochondria. All mice were maintained in air-conditioned rooms (23 ± 2°C, 45 ± 5% humidity) with light between 5:00 and 19:00 h. Under these lighting conditions, females are expected to mate between 20:00 and 01:00 h, ovulate around 03:00 h [20], and most oocytes are fertilized between 06:00 and 09:00 h [21]. For some experiments, female mice were maintained in a room with reversed lighting conditions (light between 15:00 and 05:00 h) for 2 weeks or more. This made females mate between 06:00 and 11:00 h and ovulate around 13:00 h, which allowed us to perform operations during ordinary working hours. For some experiments (e.g. comparison of transport of acrosome-intact and acrosome-reacted spermatozoa in the oviduct), female mice were induced to ovulate by intraperitoneal injection of equine chorionic gonadotropin (eCG; 2.5–5.0 IU) irrespective of the stage of the estrous cycle followed by an injection of human chorionic gonadotropin (hCG; 2.5–5.0 IU) 48 h later. Ovulation is known to take place around 12 h after hCG injection. All experiments conformed to the Guidelines for Animal Experiments of Asahikawa Medical University. Chemicals and media All chemicals were purchased from Nacalai Tesque Inc. (Kyoto, Japan) unless otherwise stated. The medium used for perfusion of an exposed ovary–oviduct–uterus complex was Dulbecco's Ca2+- and Mg2+-containing phosphate-buffered saline (PBS+) supplemented with 0.1% bovine serum albumin (BSA; Sigma-Aldrich, St. Louis, MO) (PBS-BSA, pH 7.3). The medium used for the examination of oocytes and spermatozoa as well as insemination in vivo and in vitro was bicarbonate-buffered TYH medium [22], which consisted of 119.37 mM NaCl, 4.79 mM KCl, 1.71 mM CaCl2, 1.19 mM KH2PO4, 1.19 mM MgSO4, 25.07 mM NaHCO3, 5.56 mM glucose, 1.00 mM sodium pyruvate, 4 mg/ml BSA (AlbuMax; GibcoBRL, Auckland, New Zealand), 50 μg/ml streptomycin sulfate, and 75 μg/ml penicillin G potassium, pH 7.4 when equilibrated with 5% (v/v) CO2 in air. Observations of oviduct movements and oviduct fluid flow in situ The setup we used for examination of oviducts in situ was essentially the same as that used by Battalia and Yanagimachi [14] (Figure 1). Briefly, a female mouse was anesthetized with isoflurane vapor, and the ovary/oviduct and part of the uterus were pulled out through a flank incision. They were then bathed with running warm PBS-BSA. Temperature of the PBS-BSA and the complete coverage of exposed organs with PBS-BSA were of critical importance for keeping the oviduct actively motile. At 30°C or below, oviducts became completely immotile. Complete coverage of organs with PBS-BSA was also important, or peristaltic motion of the oviduct stopped or slowed down noticeably. Preliminary experiments showed that oocytes could be fertilized normally in oviducts during 4 h of anesthesia of the animal. Figure 1. View largeDownload slide Diagrams of the apparatus used for observing an ovary–oviduct–upper uterus complex. (A) Entire view. Warmed (37°C) PBS is pumped through the perfusion collar attached to the dorsal surface of an anesthetized female mouse with the reproductive tract exposed via a flank incision. (B) Temperature of the circulating medium is monitored using a temperature sensor. (C) An oviduct undergoing active peristalsis can be seen under the dissecting microscope. Fertilization can proceed normally in the oviduct under such conditions. Figure 1. View largeDownload slide Diagrams of the apparatus used for observing an ovary–oviduct–upper uterus complex. (A) Entire view. Warmed (37°C) PBS is pumped through the perfusion collar attached to the dorsal surface of an anesthetized female mouse with the reproductive tract exposed via a flank incision. (B) Temperature of the circulating medium is monitored using a temperature sensor. (C) An oviduct undergoing active peristalsis can be seen under the dissecting microscope. Fertilization can proceed normally in the oviduct under such conditions. To visualize the fluid flow within the oviduct lumen and to ease our ability to observe the oviduct's contractions, approximately 0.1 μl of India ink was injected into the lower part of the oviduct isthmus through a fine injection pipette (∼20 μm in diameter). The ink had been dialyzed overnight against PBS+ and diluted with PBS-BSA before use. In one series of experiments, muscular contraction of the oviduct was inhibited by circulating PBS-BSA containing 20 μM nicardipine (Merck KGaA, Darmstadt, Germany) through the perfusion collar. Oviducts arrested peristalsis by 5–10 min after this treatment. India ink was then injected into the lower isthmus. Movements of oviducts and the behavior of ink within the oviducts were recorded with a digital movie camera. The time needed for ink to reach the ampulla was determined by retrospective examination of the movies. Measurement of the volume of fluid secreted by the oviduct To determine how much fluid the oviduct secretes during the estrous period, we ligated one of the two oviducts at two places using sterile 11–0 nylon threads (Bear Medic Corp, Ibaraki, Japan): one at the upper ampulla near ovarian bursa and the other at the UTJ. The oviduct of the other side was left untouched and served as the control. One hour later, the mouse was euthanized and the ligated oviduct (now distended) was isolated along two loops of the threads. The oviduct surface was carefully wiped clean using absorbent tissue paper, removing any blood and body fluid. After weighing the oviduct, it was put on a filter paper, cut into seven to eight pieces, and compressed under another filter paper to remove all the fluid. The oviduct was then reweighed. We assumed that the relative density of the oviduct fluid would be about the same as that of distilled water (1 mg = 1 μl) and estimated the volume of fluid that had accumulated within the isolated region of the oviduct for 1 h. The contralateral control oviduct was ligated at two places—the upper ampulla and the UTJ—separated from the animal body and weighed before and after removal of fluid to estimate the volume of fluid in this region of the oviduct. In another experiment, oviducts were ligated at four points: (1) the UTJ, (2) the lower- and mid-isthmus junction, (3) the mid- and upper isthmus junction, and (4) upper isthmic–ampullary junction (IAJ). The oviducts were examined 30 min or 2 h later to determine which parts were swollen from fluid accumulation. Observation of sperm ascent in the oviduct A postovulatory female mouse (14–15 h after hCG injection) was anesthetized and its ovary–oviduct–uterus complex was installed in the perfusion collar. After the oviduct was completely submerged in running PBS-BSA, a small amount (0.1 μl) of sperm suspension was injected into the lower isthmus. The spermatozoa were obtained from the cauda epididymidis of the TG male mice described above. The time and number of spermatozoa reaching the ampulla were determined. Spermatozoa injected into the isthmus were acrosome-intact (fresh cauda epididymal spermatozoa suspended for 5–10 min in TYH medium), 100% acrosome-reacted (AcR), or 100% dead. The 100% AcR spermatozoa were obtained by preincubating cauda epididymal spermatozoa for 2 h in TYH medium at 37°C under 5% CO2 in air before treatment with 20 μM Ca2+-ionophore A23187 [23]. Spermatozoa immobilized by the ionophore were centrifuged (550 g, 5 min) and pelleted spermatozoa were overlaid with fresh TYH medium to allow them to swim up into the medium. Virtually 100% of swim-up spermatozoa were AcR and vigorously motile. We also prepared 100% dead spermatozoa by freezing the suspension of cauda epididymal spermatozoa in liquid nitrogen for a few minutes without cryoprotection. Insemination was done by depositing approximately 0.1 μl of sperm suspension (∼1–5 × 103/μl) into the lower isthmus. Various times after insemination, oviducts were isolated, put on a glass slide, and compressed slightly under a coverslip before examination of spermatozoa in the lumen using a fluorescence microscope with a highly sensitive digital camera (α7s, Sony). We could see spermatozoa in oviducts under fluorescence microscopy after oviducts were separated from the animal's body. In some experiments, TG spermatozoa were suspended in TYH medium containing a small amount of India ink before injection into the isthmus. This enabled us to examine the movement of spermatozoa and fluid (ink) within the oviduct simultaneously. Assessment of fertilization In some experiments, sperm fertilizing capacity within oviducts was assessed. A drop of TYH medium containing 100% AcR spermatozoa was injected into the lower isthmus. At 4 h later, oocytes were collected from the ampulla (or the uterus in some experiments) and examined for evidence of fertilization. An oocyte with two pronuclei and two polar bodies (6–8 h after sperm deposition in the isthmus) was considered fertilized. Statistical analysis All percentile data were transformed into arcsine values for statistical analysis to normalize them. When data were compared between two groups (distributions of spermatozoa in the oviduct after sperm injection into the lower isthmus), one-way analysis of variance (ANOVA) and Welch's t-test were used. When the data were compared among more than two groups (fertilization rate), one-way ANOVA and Tukey–Kramer methods were used. All statistical analyses were performed with R software (version 3.2.3; https://www.r-project.org). Results Structure of the mouse ovary–oviduct–uterus complex The mouse oviduct is a tortuous tube. Its upper end opens to the ovarian bursa and the lower end connects to the uterus via the UTJ. Figure 2 shows photographs of the oviduct (A and B), and semi-diagrammatic (C) and highly diagrammatic (D) illustrations of the ovary, the oviduct and the upper region of the uterus. The oviduct comprises (a) the infundibulum, (b) the ampulla, (c) the isthmus, and (d) the UTJ. Both the ovary and the infundibulum of the oviduct are encapsulated by the ovarian bursa, a membranous sac with a small hole that opens into the peritoneal cavity (Figure 3D) [24]. As described below, oviduct fluid drains into the ovarian bursa and then passes through the bursa hole into the peritoneal cavity. Figure 2. View largeDownload slide (A) A mouse oviduct, slightly stretched, as seen through an ordinary dissecting microscope. (B) An oviduct extensively stretched, flushed with PBS containing FITC-conjugated WGA and observed using fluorescence microscopy. (C , D) Schemata of the anatomy of the ovary, oviduct, and uterus. The site of ink and sperm injection is shown in diagram D. IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. Figure 2. View largeDownload slide (A) A mouse oviduct, slightly stretched, as seen through an ordinary dissecting microscope. (B) An oviduct extensively stretched, flushed with PBS containing FITC-conjugated WGA and observed using fluorescence microscopy. (C , D) Schemata of the anatomy of the ovary, oviduct, and uterus. The site of ink and sperm injection is shown in diagram D. IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. Figure 3. View largeDownload slide (A) Diagrams showing the behavior of ink injected into the lower isthmus of an estrous female. A bolus of ink in the lower isthmus ascends the oviduct in a to-and-fro fashion, is fragmented, and drains into the peritoneal cavity via the bursa hole. IAJ, isthmic-ampullary junction; IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. (B) Oviduct and uterus in situ showing the site of Ink injection; this image was captured from Supplemental Movie 1 showing the behavior of injected ink in the oviduct. (C, D) Diagrams of the structural relationships among the uterus, oviduct, ovary, and ovarian bursa. The ovary is completely encapsulated by the bursa membrane. Fluid within the bursa drains into the peritoneal cavity via the bursa hole. Note the presence of the bursa hole (arrow) in the mesosalpinx covering the top of the uterus. Figure 3. View largeDownload slide (A) Diagrams showing the behavior of ink injected into the lower isthmus of an estrous female. A bolus of ink in the lower isthmus ascends the oviduct in a to-and-fro fashion, is fragmented, and drains into the peritoneal cavity via the bursa hole. IAJ, isthmic-ampullary junction; IU, upper isthmus, IM, mid isthmus, IL, lower isthmus. (B) Oviduct and uterus in situ showing the site of Ink injection; this image was captured from Supplemental Movie 1 showing the behavior of injected ink in the oviduct. (C, D) Diagrams of the structural relationships among the uterus, oviduct, ovary, and ovarian bursa. The ovary is completely encapsulated by the bursa membrane. Fluid within the bursa drains into the peritoneal cavity via the bursa hole. Note the presence of the bursa hole (arrow) in the mesosalpinx covering the top of the uterus. The thin-walled ampulla is where oocytes and spermatozoa meet during normal fertilization in vivo. Numerous motile cilia line the ampulla, infundibulum, and the inner aspect of the fimbria. The transitional region between the ampulla and the isthmus is the IAJ. The isthmus, except for its uppermost region, has numerous transverse ridges and pockets on its inner wall. In mice and many other animals, the isthmus serves as the sperm reservoir prior to fertilization in vivo [3, 25, 26, 27]. The UTJ is a thick-walled “straight” tube with longitudinal mucosal folds, connecting the isthmus to the uterus. Active contractions of the oviduct and rapid transport of luminal fluid The oviduct of estrous females displayed a very active contraction–relaxation movement. This movement could be seen clearly when a bolus of India ink (in PBS-BSA) was injected into the lower isthmus (Figure 3A and B; see Supplemental Movie S1 in real time). The ink bolus was quickly divided into smaller boluses by the repeated contraction–relaxation (segmenting) movements of the isthmus wall. These moved up (i.e. adovarian) and down. Because the upward movement was greater than the downward movement, all boluses moved steadily toward the oviduct's ampulla, some faster than others. This back and forth movement of each ink bolus continued until it reached the upper isthmus. In this region, the ink did not go back (i.e. toward the uterus) because of the unidirectional (adovarian) contractions of the upper one third of the upper isthmus. The most prominent adovarian motion of the oviduct was seen in the IAJ (Supplemental Movie S2 in real time). Ink quickly entering the ampulla hit the cumulus mass, stayed for a while but gradually moved to the end of the upper ampulla and to the ovarian cavity, and finally drained into the peritoneal cavity via the bursa hole (Figure 3C and D; Supplemental Movie S3). To our surprise, the ink put into the isthmus was transported to the ampulla at any time during the estrous cycle and even during early pregnancy (Figure 4; Supplemental Movie S4 comprising nine movies). However, the fastest ink transport to the ampulla occurred during the periovulatory period. We found that during estrous cycle (days 1–4) most of the ink injected into the lower isthmus was transported to the ampulla, whereas during early pregnancy (days II–IV) most of the ink remained in the lower and mid isthmus for a long time. Interestingly, nicardipine (20 μM) in PBS-BSA running through the perfusion collar did not prevent ink from ascending to the ampulla, even though this drug stopped the oviduct's muscular contractions completely. Apparently, the ink was able to ascend the quiescent oviduct along with the fluid secreted from the isthmus (see below). Figure 4. View largeDownload slide Time (in second) the India ink bolus took to travel from the lower isthmus to the ampulla. Each dot represents a single experiment. Note that the fastest ink (fluid) transport is seen during the peri-ovulatory period. Figure 4. View largeDownload slide Time (in second) the India ink bolus took to travel from the lower isthmus to the ampulla. Each dot represents a single experiment. Note that the fastest ink (fluid) transport is seen during the peri-ovulatory period. The entire length of the oviduct's isthmus secretes fluid continuously When both the IAJ and the UTJ were ligated and examined 1.5 h later, the entire length of the isthmus was seen to be swollen extensively. When the isthmus was ligated at four places as shown in Figure 5, all segments, in particular the upper isthmus with a thinner wall, were swollen extensively, indicating that the entire length of isthmus secretes fluid actively. Apparently, under normal conditions the oviduct fluid drains into the ovarian capsule, then into the peritoneal cavity via the bursa hole. We estimated that the isthmus of estrous females contains approximately 0.9 μl of fluid at any time and secretes about 2.2 μl of fluid per hour (averages of three determinations for each). Even when oviducts were immobilized by nicardipine, they kept secreting the same volume of the fluid. Ink injected into the lower isthmus of such oviducts ascended to the ampulla within 2–4 min, sometimes in more than 5 min. Figure 5. View largeDownload slide Isthmus immediately after 0 min and 30–120 min after four ligations of the isthmic region of the oviduct. Note that all segments of the isthmus expand with fluid accumulation. Figure 5. View largeDownload slide Isthmus immediately after 0 min and 30–120 min after four ligations of the isthmic region of the oviduct. Note that all segments of the isthmus expand with fluid accumulation. Both the oviduct's contractility and the upward flow of oviduct fluid are important for efficient sperm ascent and fertilization To investigate whether the oviduct's contractility is important for sperm transport and fertilization, we first immobilized oviducts using nicardipine, and then deposited about 100 live AcR TG spermatozoa in the lower isthmus of each oviduct. Oviducts bathed with PBS-BSA free of nicardipine served as controls. The results (Figure 6A) show that in the control (motile) oviducts, both live and dead spermatozoa, in particular live AcR spermatozoa, were rapidly transported from the lower isthmus to the ampulla. When live, “fresh” spermatozoa (which were mostly uncapacitated and acrosome-intact) were injected, more than 90% attached to epithelium of the lower isthmus and did not ascend to the ampulla. Within immobilized oviduct, on the other hand, none or only few spermatozoa ascended to the ampulla. When 100% AcR live spermatozoa were put in the isthmuses of intact oviducts and the eggs were examined 4 h later for evidence of fertilization, 94% of eggs were fertilized, whereas only 20% were fertilized in immobilized oviducts (Figure 6B), indicating the importance of oviduct's contractility for efficient fertilization in vivo. It should be noted that both immobilized and motile (control) isthmuses produced the same amount of oviduct fluid (∼2.3 μl/h). Figure 6. View largeDownload slide (A) Sperm ascent in normal (motile) and immobilized oviducts. About 100 TG spermatozoa, live or dead, were put in the lower isthmus. Five and twenty minutes later, the number of spermatozoa in the ampulla was determined. Note that inhibition of oviduct contractility by nicardipine results in a drastic reduction of sperm ascent to the ampulla. (B) Immobilization of oviduct by nicardipin results in a drastic reduction in fertilization rate. Mean ± SD. (C) Prevention of oviduct fluid flow into ovarian bursa/peritoneal cavity by ligation of the upper ampulla results in fertilization failure. Mean ± SD. Figure 6. View largeDownload slide (A) Sperm ascent in normal (motile) and immobilized oviducts. About 100 TG spermatozoa, live or dead, were put in the lower isthmus. Five and twenty minutes later, the number of spermatozoa in the ampulla was determined. Note that inhibition of oviduct contractility by nicardipine results in a drastic reduction of sperm ascent to the ampulla. (B) Immobilization of oviduct by nicardipin results in a drastic reduction in fertilization rate. Mean ± SD. (C) Prevention of oviduct fluid flow into ovarian bursa/peritoneal cavity by ligation of the upper ampulla results in fertilization failure. Mean ± SD. What will happen when oviduct's fluid flow from the isthmus to the peritoneal cavity is blocked? Will oocytes be fertilized normally? Figure 6C summarizes the results of experiments in which the upper ampulla was ligated to prevent the ascent of oviduct fluid to the ovarian bursa. When 100% AcR spermatozoa were put in the lower isthmus 15 min after ligation of the upper ampulla, about half of the oocytes were fertilized by 4 h. The oocytes were found either in the oviduct (3/5 animals) or both the oviduct and uterus (2/5 animals). In contrast, none of the oocytes was fertilized when spermatozoa were put into the lower isthmus 1 h after ligation of the upper ampulla. We found the oviducts very much distended by 1 h after ligation of the upper ampulla. India ink put into the lower isthmus along with spermatozoa did not ascend to the ampulla. Instead, both ink and spermatozoa descended to the uterus. Surprisingly, some oocytes also descended from the ampulla to the uterus. In one instance, we saw the ampulla rupture. In control (nonligated) oviducts, all oocytes were in the ampulla and all or most of them were fertilized by the time of examination. Discussion Role of the upward flow of oviduct fluid for fertilization in vivo As reported here, the entire region of the mouse oviduct secretes fluid very actively. The fluid is transported from the isthmus to the ampulla because of the predominantly one-way (upward) pumping action of the upper one-third of the isthmus, in particular the IAJ. The fluid eventually drains into the peritoneal cavity via the bursa hole. Because the distance between the lower isthmus and the ampulla is about 12 mm in the mouse and ink travels this distance in about 30 s in a to-and-fro manner, the fluid in the oviduct must move faster than about 400 μm/s. Although such upward flow of the oviduct fluid was seen during any time of the estrous cycle, it was most prominent during the periovulatory period. Such upward flow of the oviduct fluid must be important for fertilization in vivo. Prevention of upward flow of oviduct fluid by ligation of the upper ampulla after ovulation results in not only swelling of the ampulla, but also flashing of oocytes down to the uterus without fertilization. The swollen ampulla may even rupture. Role of the spermatozoon's own movement for fertilization in vivo Chang and Suarez [28] maintained that the spermatozoon's movement alone is enough for sperm transport from the isthmus to the ampulla to fertilize oocytes. This was based on experiments in which isolated oviducts of mated female mice were maintained in a medium containing the muscle-immobilizing drug nicardipine. Ishikawa et al. [18], who did similar experiments concentrating on the contractile role of the myosalpinx, considered that the spermatozoon's own motility is necessary for ascent from the mid-isthmus to the ampulla. Here, we found that (i) adovarian peristaltic movements of the isthmus; (ii) secretion and adovarian flow of oviduct fluid; and (iii) the spermatozoon's own active movement all contribute to efficient sperm ascent from the isthmus to the ampulla. A scheme of fertilization in vivo Based on information obtained from previous and this study, we propose that the following events likely happen in the female tract during natural fertilization in the mouse and common laboratory rodents. During and/or after coitus, millions of spermatozoa are deposited in the uterus, but only a small percentage passes through the UTJ. Those in the UTJ do not exhibit high amplitude tail oscillations, yet they are able to advance steadily to the lower isthmus of the oviduct [17]. Some spermatozoa reaching the isthmus may die there. They are quickly transported to the upper oviduct before draining into the peritoneal cavity (they are the so-called “vanguard” spermatozoa but play no part in fertilization) [29, 30]. Spermatozoa reaching the lower isthmus have not completed capacitation [17, 31]. They are held there, with heads firmly attached to the isthmic epithelium until they are capacitated and hyperactivated [5, 32]. It is unlikely that all spermatozoa are capacitated synchronously. Some must be capacitated and AcR faster than others [21]. Some might never be capacitated and simply die in the isthmus. Capacitated ones with lower binding ability to isthmic epithelium than uncapacitated ones [32], lift off the epithelium assisted by their vigorous (hyperactivated) tail movement [26, 33]. After swimming free for a while, they reattach to the isthmic wall. Adovarian oviduct peristalsis and fluid flow help to direct spermatozoa toward the ampulla. It is known that there are fewer spermatozoa around the oocytes in the ampulla during the progression of fertilization until almost all oocytes are fertilized [17, 34]. This does not necessarily mean that only a few spermatozoa reach the ampulla. It is more likely that both live and dead spermatozoa keep ascending the oviduct before, during, and even after fertilization. When a cumulus–oocyte complex is in the ampulla, live spermatozoa are trapped by the cumulus and enter the oocytes [21, 35]. Even after all the oocytes have been fertilized, spermatozoa that did not participate in fertilization drain out of animal's body or are phagocytized by epithelial cells [36, 37] or keep ascending the oviduct and drain into the peritoneal cavity (see Supplemental figure). On chemotaxis, rheotaxis, and thermotaxis of spermatozoa during fertilization in vivo How spermatozoa reach oocytes in the oviduct's ampulla has been the subject of much debate. Many investigators speculated that factors (e.g. progesterone) secreted from the oocyte and surrounding cumulus cells guide spermatozoa chemotactically toward the egg (e.g. [7, 9, 38]). Even though such chemoattractants might help orient spermatozoa toward oocytes in the immediate vicinity, it is difficult to conceive that the attractant could diffuse far down to the isthmus to guide spermatozoa. The oviduct's predominant fluid flow is upward (adovarian) rather than downward, as we report here. It has been known for many years that the presence of ovulation products (oocytes, cumulus oophorus, and follicular fluid) within the ampulla enhances sperm ascent from the isthmus to the ampulla (e.g. [39, 40]), but this can be explained by the sperm-trapping action of the cumulus oophorus rather than by sperm chemotaxis. Some other investigators have postulated that oviduct fluid flows down from the oviduct to the uterus and that spermatozoa swim rheotactically (upward) from the uterus to the ampulla of the oviduct [12]. Even though spermatozoa may swim rheotactically when observed using a microscope [41], it is most unlikely that spermatozoa within the lower isthmus of oviduct move toward the ampulla rheotactically, because the oviduct fluid is pumped up from the isthmus to the ampulla by adovarian peristalsis of the upper isthmus, in particular the IAJ. Other investigators have suggested that the temperature at the oviduct ampulla is slightly higher (1–2°C) than at the lower isthmus and uterus, and this temperature difference directs spermatozoa toward the ampulla [42]. Here, we bathed the mouse uterus–oviduct–ovary complex in running PBS with a constant temperature of 37°C. Because the temperature of the uterus, oviduct, and ovary is expected to be the same and fertilization proceeded normally under this experimental condition, it is most unlikely that any higher temperature of the ampulla is of critical importance for successful fertilization in vivo. Biological importance of the oviduct and problems to be studied further Even though the oviduct can be bypassed in assisted reproductive technologies such as in vitro fertilization and intracytoplasmic sperm injection, it is the oviduct where oocytes are normally fertilized and start to develop. Any errors in interactions between spermatozoa and oocytes and the development of embryos within the oviduct could have lasting detrimental effects on the well-being of offspring. The oviduct is not a simple tube; it provides an ideal environment for the spermatozoon's preparation for fertilization, fertilization itself, and the preimplantation development of zygotes. Several oviduct-related topics to be investigated further include (1) the origin and fate of oviduct fluid; (2) the effect of inhibition of oviduct fluid secretion; (3) the site and mechanism of sperm capitation, hyperactivation, and acrosome reaction within the oviduct, which are surprisingly still not yet fully understood [5, 43–45]; (4) the fate of surplus spermatozoa in the uterus, oviduct, and peritoneal cavity; (5) hormonal control of oviduct fluid secretion, oviduct peristalsis, and sperm transport in the oviduct; (6) confirmation of the proposed role of cell surface progesterone receptors of ciliary cells in the ampulla [46]; (7) the nature and function of “pacemaker cells” (interstitial cells of Cajal or telocytes) in the oviduct [47]; and (8) role of oviduct's peristalsis and fluid secretion in the transport of fertilized eggs and preimplantation embryos through the oviduct. These all undoubtedly play crucial roles in the coordinated movement of the oviduct before and during fertilization as well as in the development of preimplantation embryos. In this study, we have emphasized the importance of oviduct fluid flow from the lower to the higher segments of the oviduct for sperm ascent and fertilization. During the course of this study, we found that India ink injected into the ovarian bursa does not descend to the oviduct's ampulla (unpublished data), whereas cumulus-enclosed and cumulus-free oocytes as well as live and dead spermatozoa, if not all, are transported from the ovarian bursa down to the ampulla, apparently propelled by ciliary movement of the epithelia of both the fimbria and infundibulum. That mouse spermatozoa deposited into the ovarian bursa can fertilize oocytes in the ampulla [48] can be explained by ciliary movement of the epithelia of both fimbria and infundibulum. Fertilization of oocytes in the oviduct following intraperitoneal injection of spermatozoa is well known in various animals [3]. The differences among species with regard to the movement of spermatozoa from the isthmus to the ampulla as well as the role of the oviduct's pacemaker cells (interstitial cells of Cajal or telocytes; [47, 49, 50]) in sperm and egg transport must be the subject of future investigations. Supplementary data Supplemental Figure. An ICR female at D4 was caged with a TG male in the evening. Next morning, the formation of a copulation plug was confirmed. The mated female was sacrificed and the peritoneal cavity was washed thoroughly with 10 mL PBS (+) with 1% Triton-X and 0.1% BSA, which was thereafter centrifuged. The presence of the spermatozoa was examined under the microscope. Seven spermatozoa, of which six were separated heads only, were recovered from the peritoneal cavity after coitus. Supplemental Movie S1. A real-time movie showing ascendance of India ink injected in the lower isthmus. At 23:00 h of day 4 of estrous cycle. Supplemental Movie S2. A real-time movie of the isthmic–ampullary junction showing unidirectional (upward) pumping contractions. At 5:00 h of day 1 of estrous cycle. Supplemental Movie S3. A movie of India ink draining from the ovarian bursa into the peritoneal cavity via the bursa orifice. Supplemental Movie S4. Nine movies of oviducts displaying peristaltic movement at various times of estrus cycle (day 1–4 of estrous cycle) and early pregnancy (day 2–4 of pregnancy). Acknowledgment We thank Dr Hiroyuki Tateno for providing invaluable information and advice. We are grateful to Dr. Susan S. Suarez (Cornell University) and Dr. James M. Cummins (Murdoch University) for their invaluable advice. 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Journal

Biology of Reproduction – Oxford University Press

Published: Jul 1, 2019

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