INTRODUCTION
As in many arthropods, male ticks (Acari, Ixodoidea) transfer spermatozoa to the female encapsulated in an elaborate spermatophore (Nuttall and Merriman, Reference Nuttall and Merriman1911; Robinson, Reference Robinson1942; Tatchell, Reference Tatchell1962; Feldman-Muhsam, Reference Feldman-Muhsam1967a). The male deposits the spermatophore at the external genital opening of the female, but within 1–2 min, a part of the spermatophore (endospermatophore) everts down the female's vagina, carrying the spermatozoa into the female's uterus (soft ticks) or receptaculum seminis (hard ticks) (Robinson, Reference Robinson1942; Feldman-Muhsam, Reference Feldman-Muhsam1967a, Feldman-Muhsam et al. Reference Feldman-Muhsam, Borut, Saliternik-Givant and Eden1973). During the next few hours, the part of the spermatophore remaining outside the genital aperture – the ectospermatophore – desiccates, separates from the endospermatophore, and falls away from the female. Over the following 24 h, the spermatozoa undergo an extensive transformation within the endospermatophore, and then move (or are moved) out of the endospermatophore and uterus up into the oviducts. In argasid (soft) ticks, most of the spermatozoa congregate in expansions of the oviducts (‘ampullae’, Wagner-Jevseenko, Reference Wagner-Jevseenko1958), but a few penetrate further up into the ovaries. The site of fertilization has not been definitively determined, but it seems likely to be in the ovaries (Lees and Beament, Reference Lees and Beament1948; Oliver, Reference Oliver1974), although there have been assertions that it occurs in the oviducts (Goroshchenko, Reference Goroshchenko1965; quoted by Balashov, Reference Balashov1968). The mechanism of fertilization remains unknown.
While the transformation of the spermatozoa (by analogy to a much subtler phenomenon seen in mammalian spermatozoa, this is sometimes called ‘capacitation’) in the endospermatophore has been investigated in some detail (Samson, Reference Samson1909; Casteel, Reference Casteel1917; Wagner-Jevseenko, Reference Wagner-Jevseenko1958; Brinton et al. Reference Brinton, Burgdorfer and Oliver1974; Borut and Feldman-Muhsam, Reference Borut and Feldman-Muhsam1976; Feldman-Muhsam and Filshie, Reference Feldman-Muhsam, Filshie, Fawcett and Bedford1979; Shepherd et al. Reference Shepherd, Levine and Hall1982a, Reference Shepherd, Oliver and Hallb; Sahli et al. Reference Sahli, Germond and Diehl1985), many of the subsequent events in the career of the spermatozoa are controversial or poorly documented. This paper focuses on the nature of sperm motility in the argasid tick Ornithodoros moubata and the role of motility in the translocation of spermatozoa from the spermatophore to the oviducts. We specifically address the following questions. (1) How do the spermatozoa move? (2) Do the spermatozoa become motile in the spermatophore? (3) How do the spermatozoa exit the spermatophore? (4) How do the spermatozoa migrate up the female reproductive tract?
MATERIALS AND METHODS
Ticks
A stock of the African soft tick Ornithodoros moubata obtained from the NIH Rocky Mountain Laboratory (originally from Tanzania) was maintained on rabbits as described earlier (Shepherd et al. Reference Shepherd, Levine and Hall1982a). Only virgin females were used in mating experiments; virginity was assured by isolating last instar nymphs in individual vials after feeding. Adult ticks were mated simply by combining males and females in groups or pairs, in dishes or vials.
Dissections and observations
Female ticks aged 0 to 62 days after copulation were dissected under a physiological saline (9·0 g NaCl, 0·42 g KCl, 0·33 g CaCl2·2H2O in 1 L of H2O). After transfer to fresh saline, oviducts were slit longitudinally allowing their contents to flow out. Spermatozoa were then observed with phase-contrast optics and when necessary, counted under scored cover-slips. Endospermatophore capsules were carefully removed from uteri and then placed either in fresh saline, in tick sperm culture medium (41·4 mM NaCl, 41·6 mM KCl, 113 mM glutamic acid, 10 mM glucose, 10 mM citric acid, 50 μg/ml penicillin, 300 μg/ml streptomycin sulfate, 10 mg/ml bovine serum albumin, 50 mM HEPES-NaOH buffer at pH 7·0), or in fixative, as cited below.
To watch motility, sperm suspensions were mounted on a glass cover-slip as hanging-drop cultures and observed in phase-contrast optics or filmed by microcinematography. The major features of motility reported below were independently observed by at least 2 and usually 3 of the authors and the conclusions are based on hundreds of hours of observation over a period of about 10 years. Although suspensions of spermatozoa usually contained numerous particles naturally present in seminal fluid, observations of water currents were facilitated on occasion by adding particles in the form of dialysed Pelikan brand ‘Special Ink – Black’ or rabbit erythrocytes dissected from tick guts.
To count spermatozoa in endospermatophores, the capsules were emptied into 1 ml of culture medium. At least five 6-μl samples (withdrawn during stirring) were then transferred to slides, counted under marked cover-slips, and assessed for presence and type of motility. Accurate counts of spermatozoa in the oviducts were not attempted because spermatozoa tend to attach to and penetrate oviductal cells (Sokolov, Reference Sokolov1956; Wagner-Jevseenko, Reference Wagner-Jevseenko1958; Brinton et al. Reference Brinton, Burgdorfer and Oliver1974, and others): thus numbers of spermatozoa recorded from the oviducts were usually minimal estimates.
Spermatozoa and cover-slips over which spermatozoa had moved were stained for possible mucous secretions by the periodic acid-Schiff technique (Pearse, Reference Pearse1961), and by the periodic acid leucofuchsin, brilliant cresyl blue, and mucicarmine methods (Clark, Reference Clark1973).
Feeding experiments
From a population of females that had not fed or mated since their moult to adults 3 months earlier, 9 females were given a bloodmeal while the other 9 were not. Within 2 days after engorgement by the first group, all 18 females were mated. Three from each group were then dissected 1, 3 or 7 days after mating; spermatozoa were counted and assessed for motility as described above. One- and two-way ANOVAs were used for statistical comparisons of the 6 groups.
Transmission electron microscopy (TEM)
Endospermatophores were prepared for examination by TEM either in utero or after they had been removed from the uterus. All specimens were fixed for 2–4 h on ice with 2% glutaraldehyde in 0·1 M cacodylate buffer (pH 7·2) with 0·15 M sucrose added. Samples were rinsed within 2 h in 3 to 4 changes of the same ice-cold 0·05 M cacodylate/sucrose buffer. Specimens were post-fixed with 1% osmium tetroxide in 0·05 M cacodylate buffer (pH 7·2) for 1–2 h at room temperature (Resler, Reference Resler1981, adapted from Feldman-Muhsam and Filshie, Reference Feldman-Muhsam and Filshie1976). Best results were achieved when samples were dehydrated during agitation at room temperature through a graded acetone series for up to 4 h, and then infiltrated with agitation in a graded series of acetone and Spurr's low viscosity embedding medium (Spurr, Reference Spurr1969) for up to 80 h. Both thick and thin serial sections were taken through each specimen. For light microscopy, thick sections were stained with 1% toluidine blue in 1% aqueous borax. For electron microscopy, thin sections were stained with 2% aqueous uranyl acetate and lead citrate (Reynolds, Reference Reynolds1963). Grids were examined in a Philips EM 100 at an accelerating voltage of 60 kV.
Scanning electron microscopy (SEM)
Samples were fixed and dehydrated for SEM in a manner similar to that used for TEM. Following dehydration in acetone, all samples were critical point dried with carbon dioxide, mounted on stubs with conductive paint, sputter-coated with a gold-palladium alloy and examined on an ETEC Autoscan microscope at an accelerating voltage of 5 kV.
RESULTS
Characteristics of sperm motility
Mature tick spermatozoa have a unique morphology found otherwise only in anactinotrichid mites, to which ticks are probably closely related (Alberti, Reference Alberti1980). The spermatozoa are very large (ca 450 μm in O. moubata) and club-shaped, contain a large volume of cytoplasm, carry the nucleus at the posterior end, and lack a flagellum (Fig. 1). As many as 5 different kinds of movement have been described for tick spermatozoa (see Discussion section), but our observations indicate that these can be attributed to only 2 apparently different types of motility actively generated by the spermatozoa: (1) gliding locomotion of the whole spermatozoon, and (2) contortions of the anterior tip only.
Diagram of a mature spermatozoon from within the uterus of a female Ornithodoros moubata. The head – so-called because it is anterior during locomotion – contains numerous mitochondria in linear arrays in its anterior part. Currents along the head, as revealed by suspended particles, are indicated by the dashed lines along one side of the head. The cellular processes, which cover almost the entire surface of the spermatozoon and are thought to engender motility, are shown in the blow-up and along the lateral edge of the head, but are not indicated elsewhere. The length of the entire sperm is about 450 μm.
(1) Gliding locomotion
is generally seen, by light microscopy, as a steady translation (speed 4·7–6·9 μm/sec, n=4) of the whole spermatozoon along a surface with no accompanying deformations of the cell. When gliding, a spermatozoon looks like a resilient rod and appears fairly stiff, often showing irregular but persistent twists and folds when freely suspended (Fig. 2). Typically, a spermatozoon will move along a surface with only its head (as defined in Fig. 1) in contact with the surface, and its tail often curved into the medium in a C- or S-shape. Since the head moves straight along its long axis, dragging the bent tail passively in the same direction, the tail is obviously not contributing to the locomotion of the spermatozoon. But when the spermatozoon moves into narrow confines such as it encounters in vitro at the edge of a drop suspended on a cover-slip, the tail contacts the surface, the twists and folds of both head and tail straighten out, and the tail then follows the surface directly behind the leading head (Figs 3 and 4a). If the head then encounters an obstacle, it bends readily, even to the extent of doubling sharply (as much as 180 degrees) backward, and continues with the tail following exactly in its tracks (left spermatozoon in Fig. 4a–e). Occasionally, the head is slowed or stopped momentarily by an obstacle; at this point, the tail continues to move forward. As the head is encountering resistance, the tail first wrinkles into a series of fairly regular undulations (wavelength 7–10 μm in the top-most spermatozoon in Fig. 4b). Then a lateral loop appears which amplifies until the head is deflected or released (top-most spermatozoon; Fig. 4c–e), when the spermatozoon commonly begins moving forward again in a new direction. If a spermatozoon enters a cul-de-sac where the head does not easily find a direction for movement, the spermatozoon ends up bent many times, often forming a complex knot.
Mature spermatozoon moving freely within a drop of culture medium, showing irregular folds and bends.
Mature spermatozoon moving within the confines of a small drop of culture medium on a cover-slip, showing a straighter and smoother configuration. The large mitochondrial mass is clearly visible inside the head. The arrow indicates the edge of the hanging drop which the spermatozoon is pushing outward.
Motile spermatozoa in vitro, seen as a sequence of 5 frames from a film, each separated by about 15 sec. The head of the topmost spermatozoon (arrow in (a) indicates direction of motion) can be seen to stop as it encounters resistance along the edge of the drop (a–b); as its tail continues to push forward, undulations appear (4b), then it forms a growing loop (c–d), and finally the loop passes to the rear of the sperm as the head resumes locomotion (e).
These observations indicate clearly that although the head plays a dominant role in locomotion, the tail is independently capable of generating motility. For both head and tail, contact with a surface appears necessary for locomotion, even if the surface is only a water-air interface, as on the underside of a hanging drop. No evidence of chemotaxis or avoidance behaviour was observed – spermatozoa always moved directly forward unless they were obviously deflected by obstacles.
By dint of suspended particles, either incidentally or intentionally added, very active currents could be clearly seen in the vicinity of the spermatozoa, especially at the anterior end (as indicated in Fig. 1). Particles reaching the anterior tip were swept vigorously from the brim (for morphological detail of the brim, see Breucker and Horstmann, Reference Breucker and Horstmann1968 and Feldman-Muhsam and Filshie, Reference Feldman-Muhsam, Filshie, Fawcett and Bedford1979) backward to the widest portion of the head and, somewhat less often, further back to the beginning of the tail (Fig. 1). Even less often, particles were seen being propelled along more posterior portions of the tail. The currents at the anterior end were much more vigorous than can be explained simply as a passive consequence of the motion of the spermatozoon. Further, they appear to be independent of the head contortions described in the next section, as they were often observed in the absence of these contortions.
It seems likely that the superficial filamentous processes described by many previous observers (see Discussion section) are responsible for this gliding locomotion and also probably the currents. Although these processes could be seen fairly clearly at the 1000×magnification using darkfield and differential interference contrast (Nomarski) optics, persistent observations revealed no visible deformations of these processes in rapidly or slowly moving spermatozoa. Spermatozoa known to be either moving or still were fixed (some quick-frozen in −80°C petroleum ether and then fixed) for scanning electron microscopy; these showed no consistent differences in the disposition of their surface processes (Judy Perdue and Shepherd, unpublished experiments).
Gliding locomotion in some cyanobacteria and diatoms involves the formation of mucous trails (Walsby, Reference Walsby1968). While the surface of tick spermatozoa, especially in the head region, often appeared sticky (attached particles) and stained intensely with stains for mucoproteins, no staining indicative of mucous trails (see Materials and Methods section) could be seen on cover-slips over which spermatozoa had travelled.
(2) Contortions of the anterior tip
of the head (hereafter called ‘head-curling’) have been repeatedly reported (for references, see Discussion section). These motions include rotary oscillations, pulsations along the long axis, and unilateral bends, all restricted to approximately the anterior one-third or less of the head. Most commonly seen here were relatively gentle rotary oscillations of the anterior end of the head, though sharper bends were not uncommon. When pressing upward against a cover-slip such that the anterior end was seen in face view, the spermatozoa showed numerous fine spiral waves of contractions spreading outward (i.e. posteriorly) from the vicinity of the brim. The rotary oscillations were usually seen in spermatozoa gliding rapidly or slowly in a straight line, but also occurred in stationary spermatozoa. Conversely, oscillations could not be seen at all in some spermatozoa that were gliding freely forward. Thus, head-curling is not tightly coupled with gliding motility, as previously pointed out by Rothschild (Reference Rothschild1961) and Feldman-Muhsam and Filshie (Reference Feldman-Muhsam and Filshie1976).
The spermatozoa showed no other kinds of deformation that were obviously attributable to active motility of the spermatozoa. Regular undulations in the tail of the spermatozoa and abrupt changes in orientation of the sperm axis, as described by Oliver and Brinton (Reference Oliver and Brinton1973), Feldman-Muhsam and Filshie (Reference Feldman-Muhsam and Filshie1976), Wüest et al. (Reference Wüest, El Said, Swiderski and Aeschlimann1978), and Feldman-Muhsam (Reference Feldman-Muhsam1986), were seen in this study only when spermatozoa encountered an obstacle or a surface, as elaborated above. These movements appeared to be only passive reactions to encounters generated by the gliding motility or anterior contractions described above.
Motility in different regions of the reproductive tract
(1) Onset of motility in the endospermatophore
When the prospermia (immature sperm) are deposited by the male in the spermatophore, the future head of the mature spermatozoon is essentially an invagination into an internal vacuole of the cell. The prospermia then begin a transformation in which the future head breaks out of the vacuole to the exterior, and the whole cell literally turns inside out and elongates to twice its original length (for a fuller description, see references cited in the Discussion). Dissection of endospermatophores 3 h after mating revealed many spermatozoa which were only partially elongated, such that only their heads and part of their tails had emerged from the interior. Yet some of these already showed head-curling and/or slow gliding motility. Time-lapse cinematographic observations of spermatozoa maturing in vitro (as described in Shepherd et al. Reference Shepherd, Levine and Hall1982a) indicated that head-curling movements begin suddenly after the head is extruded, followed shortly in some spermatozoa by limited gliding motility. Such early movements apparently result in vivo in the mature spermatozoa becoming aligned with their heads all facing outward, abutting the inside wall of the endospermatophore (Fig. 5). However, extensive gliding motility in vivo occurs only inside females that have fed (see below).
Scanning electron micrograph of an endospermatophore capsule whose wall has been partially stripped away to reveal the mature spermatozoa inside, aligned with their heads pointing outward, abutting the inside wall of the endospermatophore.
(2) Motility in the uterus
Considerable numbers of spermatozoa, up to more than 500 in some cases, were found outside the endospermatophore within the uterus of fed females. These either showed gliding motility or were dead. Only in cases where endospermatophores were ruptured (accidentally during dissection) were non-motile living spermatozoa seen in the uterus.
(3) Motility in the oviducts
Compared with endospermatophore spermatozoa, living oviductal spermatozoa generally showed faster gliding and much more vigorous head-curling, though the characteristics of motility were basically similar. All spermatozoa in the oviducts were either motile or dead. Motile spermatozoa were still present as long as 62 days after mating. Dead spermatozoa (identified in the light microscope by Brownian motion inside the cells) were often shrivelled and electron micrographs showed degeneration in mitochondria and other subcellular structures.
Feeding as a trigger for sperm migration in the reproductive tract
Migration of the spermatozoa out of the endospermatophore and up the oviducts depends acutely on whether the female has fed recently. Initially, about 15 600±7600 spermatozoa (mean± S.D. for both capsules of 9 endospermatophores) are present in an endospermatophore (volume about 0·9 μl). In females fed before mating, more (probably considerably more) than 10 percent of these spermatozoa have migrated to the oviducts (Table 1) by 7 days after mating. Ovaries of these females contained 39–42 (n=3) mature eggs (vitellogenesis completed) 7 days after mating. By comparison, almost no spermatozoa migrate up the oviducts of females not fed as adults (Table 1). Correspondingly, the ovaries of these females contained no mature eggs at all (or any in the process of vitellogenesis). Also, spermatozoa within endospermatophores inside the uteri of fed females showed much more motility (P<0·005, 2-way ANOVA, F=13·92) than spermatozoa in unfed females (Fig. 6). Furthermore, the percentage of spermatophore spermatozoa that are motile increases significantly with time after mating in fed females (P<0·025, one-way ANOVA, F=8·24) but not in unfed females (P>0·05) (Fig. 6). Relatively few of the motile spermatozoa in the endospermatophores of fed females showed full gliding motility, even 7 days after mating. Since most spermatozoa further up the reproductive tract showed gliding motility, presumably spermatozoa which become competent to glide in the endospermatophore leave it rapidly.
Percent motile spermatozoa in endospermatophores of fed and unfed females over one week after mating (each bar represents mean of 3 ticks±standard error). Letters above bars indicate significant differences.
Number of spermatozoa in oviducts of 18 mated females
a Because many spermatozoa attach to and penetrate oviductal cells and would not be counted by this method, larger counts represent minimal estimates of sperm numbers.
Escape of sperm from the endospermatophore
Two principal routes for escape of the spermatozoa from the endospermatophore have been suggested: (1) through ruptures in the endospermatophore wall, or (2) through the thin neck left when the ectospermatophore breaks away from the endospermatophore (Fig. 7).
Diagram of a spermatophore, showing alternative points of rupture (a,b) on the neck. ectsp, ectospermatophore; endsp, endospermatophore.
(1) Endospermatophore wall
Transmission electron micrographs of the wall of recently formed endospermatophores show 2 layers: (1) an electron-dense outer layer about 0·2 μm thick and (2) an inner layer of fibrous material 1·0–2·0 μm thick (Fig. 8). Examination of 57 capsules taken from females up to several months after mating showed that the outer layer maintains its thickness and generally its integrity, but that the inner layer is variable in thickness and probably deteriorates with age.
Transmission electron micrograph of a cross-section of the endospermatophore wall, 9 days after its formation. OL, outer layer; IL, inner layer.
Because ruptures observed in the walls of endospermatophores could have been inflicted during removal of the endospermatophores from the uterus, a series of endospermatophores were fixed while still contained within the uterus, then embedded in plastic and serially thick- and thin-sectioned for light and electron microscopic examination. Of 9 pairs of endospermatophore capsules in 9 uteri of females mated 3 or more days earlier, only 2 of 18 capsules showed possible ruptures in their walls. Yet all females contained spermatozoa in their oviducts, and 7 of the 9 had considerable numbers (>100) of spermatozoa there. Evidently, spermatozoa in almost all cases left the endospermatophore by a route other than a hole in its wall.
(2) Endospermatophore neck
When the ectospermatophore separates from the endospermatophore, the break apparently occurs somewhat irregularly, either below (Fig. 7a) or above (Fig. 7b) the bifurcation of the neck, and sometimes as far up as the junction of neck and capsule. In either case, the breaks leave the endospermatophore open: when endospermatophores were dissected from the uterus and examined before fixation, motile spermatozoa could be seen exiting via these openings (Fig. 9). However, longer incubations of endospermatophores in salines or other solutions (including hyperosmotic ones) caused swelling of endospermatophore capsules and large masses of spermatozoa, including immature ones, began to exit the breaks in any orientation, tail-first or bent over. Thus, to ensure that the observations of spermatozoa actively exiting the breaks of freshly-dissected endospermatophores were not artifactual, endospermatophores were fixed quickly in utero and thick-sectioned. These almost always (9 out of 11 endospermatophores) showed only a few spermatozoa exiting at once and these were generally coming out of the breaks head-first, as would be typical of actively moving spermatozoa.
Scanning electron micrograph of spermatozoa exiting the break in the neck of an endospermatophore. Most spermatozoa have their heads pointed outwards, consistent with active locomotion out of the break.
Mechanism of sperm migration in the female reproductive tract
The above evidence suggests that active motility is the means by which spermatozoa escape the endospermatophore. The fact that virtually all living spermatozoa found outside the endospermatophore in the uterus and in the oviducts are motile suggests that the migration of spermatozoa up the female tract is primarily effected by the spermatozoa themselves. Other observers have observed contractions of the female tract (in other kinds of ticks) and inferred that these are the principal causes of sperm translocation. However, in the course of dissecting well over 100 O. moubata at various times after mating, no obvious contractions of the female tract were observed. Though we cannot exclude the possibility that the dissecting saline suppressed muscular activity, we feel that the latter is probably not an important mechanism of sperm translocation in this species.
DISCUSSION
Sperm motility
A number of attempts have been made to catalogue the types of movements shown by tick spermatozoa. The most recent reports or reviews by Wüest et al. (Reference Wüest, El Said, Swiderski and Aeschlimann1978), Oliver (Reference Oliver, Obenchain and Galunl982), and Feldman-Muhsam (Reference Feldman-Muhsam1986), list 5 types, though these 5 are not identical in each report. We feel that our observations allow us to discriminate between movements generated actively by the spermatozoa themselves and those which are passive consequences of the active types of motility. On the basis of our evidence, we suggest that the only 2 types of motility actively produced by tick spermatozoa are (1) linear, gliding motility and (2) head-curling.
Gliding motility
Of the principal modes of cell locomotion – amoeboid crawling, flagellar/ciliary swimming, and gliding movement – the mechanisms underlying the last are least understood (Heintzelman, Reference Heintzelman2006). While our observations do not elucidate these mechanisms for tick spermatozoa, we believe they add some new details and useful analysis to the numerous descriptions of the gliding motion of tick spermatozoa in the literature (most notably those of Christophers (Reference Christophers1906), Rothschild (Reference Rothschild1961), Oliver and Brinton (Reference Oliver and Brinton1973), Feldman-Muhsam and Filshie (Reference Feldman-Muhsam and Filshie1976), and Wüest et al. Reference Wüest, El Said, Swiderski and Aeschlimann1978).
Our observations indicate that the motive force generated during gliding is produced along most of the length of the spermatozoon, although much the strongest forces are produced in the head region. The head always takes the lead and is often the only part of the spermatozoon in contact with a surface. But when the tail does contact a surface, it generates force, as indicated by the fact that it continues to push the tail forward into waves and bends if the head is obstructed. Thus the motive force for gliding can be generated along most if not all of the spermatozoan surface behind the brim at the very anterior end. Rothschild (Reference Rothschild1961) questioned whether a substrate (surface) was necessary for gliding motility. While we did not observe any spermatozoa completely lacking contact with any surface, we did note that when the tail was not in contact with a surface, it appeared to be completely passive, in contrast to its behaviour when touching a surface.
The dominant role of the head in generating motility was also demonstrated by the powerful currents seen being generated at the brim, and extending back around the widest part of the head. However, the whole surface from the brim to the tip of the tail is densely covered with fine, ribbon-like processes that are about 11–12 μm long and 0·2–0·3 μm wide in O. moubata (Breucker and Horstmann, Reference Breucker and Horstmann1968; Pinkerton et al. Reference Pinkerton, Hall and Shepherdl982). These processes are finger-like eversions of the cytoplasm, connected to the cytoplasm at their anterior end but attached to the plasma membrane along the rest of their length (Feldman-Muhsam and Filshie, Reference Feldman-Muhsam and Filshie1976). Over 100 of these processes encircle the spermatozoon in O. moubata. Just because of their location, these cytoplasmic processes seem highly likely to be the source of motive force in gliding. Why, then, is the motive force clearly strongest at the head? Perhaps it is because most of the mitochondria are located in a highly organized cylinder in the head (clearly illustrated in Rothschild, Reference Rothschild1961 and in Breucker and Horstmann, Reference Breucker and Horstmann1968) and thus most of the power is presumably generated in the head. Further back along the tail, mitochondria that are dispersed in a layer just underneath the plasma membrane (Breucker and Horstmann, Reference Breucker and Horstmann1968; Feldman-Muhsam and Filshie, Reference Feldman-Muhsam and Filshie1976) would provide the lesser power needed in the tail. The store of energy for the mitochondria is probably the glycogen abundantly distributed along the length of the sperm (Reger, Reference Reger1974; El Said et al. Reference El Said, Swiderski, Aeschlimann and Diehl1981).
If indeed the cytoplasmic processes are the organs that generate gliding motility, how do they do it? Although the processes are near the limit of resolution of light microscopes, the various techniques used by us and others to observe sperm in the light microscope have not revealed any obvious mechanism. Scanning electron micrographs by some authors (Feldman-Muhsam and Filshie, Reference Feldman-Muhsam and Filshie1976; Wüest et al. Reference Wüest, El Said, Swiderski and Aeschlimann1978) have shown constrictions or undulations in the whole spermatozoon having a period respectively of about 1·25 and 5 μm, which these authors imply are natural and may be the mechanism of motility. However, undulations of such magnitude would be clearly visible whenever spermatozoa are observed gliding in the light microscope, but have not been reported by others or seen by us except when a spermatozoon encounters an obstacle obstructing its progress. Thus we believe that such undulations are a passive consequence of obstruction (as seen in Fig. 4b), and are not generated as an active form of motility.
Several authors have implicated in motility the bundles of filaments (diameter 5–6 nm in O. moubata) seen just below the plasma membrane in transmission electron micrographs of the spermatozoa (Breucker and Horstmann, Reference Breucker and Horstmann1968; Feldman-Muhsam and Filshie, Reference Feldman-Muhsam and Filshie1976; Wüest et al. Reference Wüest, El Said, Swiderski and Aeschlimann1978; Oliver, Reference Oliver, Obenchain and Galun1982). These bundles are sometimes strikingly arrayed in parallel with the cytoplasmic processes (Feldman-Muhsam and Filshie, Reference Feldman-Muhsam and Filshie1976), but it is difficult to see how they could be mechanically coupled with the processes, as they are separated from the external processes by the plasma membrane. Furthermore, as pointed out first by Breucker and Horstmann, (Reference Breucker and Horstmann1968) the filament bundles begin only at the widest part of the head, well behind the beginning of the cytoplasmic processes at the brim and behind the region where we observed the strongest currents. Thus the mechanism of gliding motility in tick spermatozoa remains unresolved.
Although an uncommon phenomenon, gliding motility has been identified as the principal means of locomotion in a number of unicellular organisms, including some prokaryotes (Merz and Forest, Reference Merz and Forest2002; Hoiczyk, Reference Hoiczyk2000) and various eukaryotes (Heintzelman, Reference Heintzelman2006), including diatoms (Poulsen, Reference Poulsen1999), apicomplexans such as Toxoplasma and Plasmodium (Keeley and Soldati, Reference Keeley and Soldati2004; Baum et al. Reference Baum, Richard, Healer, Rug, Krnajski, Gilberger, Green, Holder and Cowman2006), and a few green algae (desmids and Chlamydomonas flagella – Bloodgood, Reference Bloodgood1995). In the apicomplexans, motility involves a submembrane actomyosin system, coupled with transmembrane proteins which adhere to the substrate (Keeley and Soldati, Reference Keeley and Soldati2004). Movement of these adherent proteins within the membrane, analogous to capping phenomena, apparently propel the cells forward. Such movements are undetectable in the light microscope, which would explain how gliding motility occurs without visible deformation of the cell. Although the cell surface morphologies vary greatly among all of these kinds of motile cells and their phylogenetic relationships are distant, it seems at least plausible that tick spermatozoa are propelled by a mechanism similar to those proposed for the protists cited above. Tick spermatozoa do move at speeds (4·7 to 6·9 μm/sec – this report; 2·3 to 22·9 μm/sec – Rothschild, Reference Rothschild1961) comparable to the speeds (1 to 25 μm/sec) reported for these protists. The cytoplasmic processes seen in tick spermatozoa seem to be unique: some gregarines have cytoplasmic folds apparently responsible for gliding locomotion, but the morphology and ultrastructure of these folds are substantially different from those of tick spermatozoa (Feldman-Muhsam and Filshie, Reference Feldman-Muhsam and Filshie1976; Heintzelman, Reference Heintzelman2004).
Head-curling
Since spermatozoa can show these contortions without gliding and can glide without any visible contractions, it seems likely that the two types of motility are different phenomena caused by different mechanisms. However, the contortions, when present, occurred where we also observed greatest gliding power, and so perhaps the phenomena are not entirely unrelated. The subcellular basis of the contractions is obscure, especially as there are no subsurface filaments in the anterior part of the head where the movements take place. Electron micrographs presented by Feldman-Muhsam and Filshie (Reference Feldman-Muhsam, Filshie, Fawcett and Bedford1979) show an elaborate morphology in the region of the brim, which Feldman-Muhsam (Reference Feldman-Muhsam1986) plausibly suggests may be the source of the contortions. Any hypothesis for a mechanism should also explain the rotary and spiral nature of the oscillations and waves observed. The function of the contortions is even more obscure, as they do not seem to play a role in locomotion. Wüest et al. (Reference Wüest, El Said, Swiderski and Aeschlimann1978) suggested a tactile function, but produced no evidence.
Other types of motion
We also observed lateral bending of the tail, waves of constrictions and abrupt changes in orientation, as variously described by Oliver and Brinton (Reference Oliver and Brinton1973), Feldman-Muhsam and Filshie (Reference Feldman-Muhsam and Filshie1976), Wüest et al. (Reference Wüest, El Said, Swiderski and Aeschlimann1978), Oliver (Reference Oliver, Obenchain and Galunl982), and Feldman-Muhsam (Reference Feldman-Muhsam1986). But as elaborated above, all of these motions appear to us to be passive consequences of obstructions encountered by the spermatozoa during active gliding motility, not active types of motility generated independently by the spermatozoa. Rotations about the longitudinal axis, described by Wüest et al. (Reference Wüest, El Said, Swiderski and Aeschlimann1978) and Feldman-Muhsam (Reference Feldman-Muhsam1986), could be a passive product of the orientation of the filaments, whose longitudinal axis may spiral slightly around the spermatozoon (El Said et al. Reference El Said, Swiderski, Aeschlimann and Diehl1981).
Feeding as a trigger for sperm migration
The experiments detailed above provide data that support the assertions by Samson (Reference Samson1909) and Feldman-Muhsam (Reference Feldman-Muhsam1967b) that feeding is a necessary prerequisite to migration of the spermatozoa out of the endospermatophore and up the female reproductive tract to the oviducts. When females are not fed, the spermatozoa remain viable in the endospermatophore for a long time (up to 11 months in O. savignyi – Feldman-Muhsam Reference Feldman-Muhsam1967b). Our observations indicate that although the spermatozoa become somewhat motile in the endospermatophore during their maturation, a bloodmeal is essential for their migration out of the endospermatophore up into the oviducts. It seems likely that this induction of migration is chemically mediated, though whether the putative substance is a component of the blood meal or an endogenous message produced by the female is unknown. This induction of sperm movement adds yet another signaling event to the several now known to be associated with mating and engorgement in ticks (Kaufman and Lomas, Reference Kaufman and Lomas1996; Kaufman, Reference Kaufman2004).
Escape of sperm from the endospermatophore and translocation up the female tract
The route by which the spermatozoa leave the endospermatophore has been controversial. Several reports suggest that spermatozoa are released by rupture of the endospermatophore wall (Robinson, Reference Robinson1942; Wagner-Jevseenko, Reference Wagner-Jevseenko1958), perhaps facilitated by stretching and penetration of the wall by the spermatozoa (Brinton et al. Reference Brinton, Burgdorfer and Oliver1974). Other reports indicate that spermatozoa leave the endospermatophore capsules via the tubular openings formed when the ectospermatophore separates from the endospermatophore after formation of the endospermatophore (Feldman-Muhsam, Reference Feldman-Muhsam1964; El Said, Reference El Said1976). Robinson (Reference Robinson1942) claims, however, that in O. moubata these openings are sealed shut; as additional support, he says that capsules placed in distilled water swell osmotically.
Our serial sections of endospermatophores fixed in utero indicate that their walls remain intact during the emigration of the spermatozoa. This fact, plus the observation of motile spermatozoa exiting the broken neck of the endospermatophore, imply that the natural opening formed by the broken neck is the normal route of escape by the spermatozoa, not ruptures in the wall of the endospermatophore. Furthermore, the facts that (1) motile spermatozoa exit the neck generally headfirst, (2) only actively gliding spermatozoa (aside from occasional dead spermatozoa) are seen outside the endospermatophore and in the uterus, and (3) no contractions of the female tract were observed in over 100 dissections of mated female ticks, imply that the intrinsic motility of the spermatozoa is the principal mechanism by which spermatozoa are normally translocated up the reproductive tract in O. moubata. At the locomotory speeds we recorded (ca. 5–7 μm/sec), a spermatozoon could theoretically travel the length of the oviducts (ca. 5–7 mm) in less than 1 h, provided it moved without obstruction.
We believe this evidence calls into question the long-standing supposition, based largely on circumstantial evidence, that tick spermatozoa are passively translocated by the muscular action of the female reproductive tract (reviewed by Oliver, Reference Oliver, Obenchain and Galun1982). Although a thin muscle layer surrounds the oviduct in many species of ticks (Christophers, Reference Christophers1906; Robinson and Davidson, Reference Robinson and Davidson1914; Douglas, Reference Douglas1943; Till, Reference Till1961; Sonenshine, Reference Sonenshine1970; Brinton et al. Reference Brinton, Burgdorfer and Oliver1974), this may be used primarily to transport the eggs down the oviducts. If sperm motility is indeed the prime effector of sperm migration in O. moubata, chemotaxis need not be invoked to account for the orientation of the spermatozoa, as they have only one direction to go on exiting the endospermatophore – up the oviducts. The reason why many spermatozoa end up inside oviductal cells (Sokolov, Reference Sokolov1956; Wagner-Jevseenko, Reference Wagner-Jevseenko1958; Brinton et al. Reference Brinton, Burgdorfer and Oliver1974), and the locus and mechanism of syngamy, all remain unknown.
We are grateful for careful reviews of the manuscript by Kim Berlin and an anonymous reviewer. Part of this work was supported by NSF Grant #PCM 77-24933.
