Perinatal penile adhesions

Keywords: prepuce, equine, prepuberal, colt

A frenulum joins the prepuce and penis on the ventral aspect of the penis in all domestic animals except for the equine species. Usually, the frenulum In most animals breaks down largely under the effect of androgens (perhaps attempts at erection as well) after puberty. In horses this is not the case. No frenulum is present at birth and within days of foaling, a colt is able to extend his penis completely.

During fetal development the penis is of course adherent to the prepuce, dividing itself from that structure as organogenesis proceeds. 

In rare cases, exuberant adhesion between the prepuce and penis may actually occlude the preputial opening, even preventing urination. In one such case (see reference), gentle separation of internal lamina of the Prepuce and mild traction on the penis resolve the condition successfully

The images below show the penis and prepuce of a colt late in gestation.  Note the tenuous connection between the penis and prepuce at this time. At the time of foaling, the penis will have separated completely from the prepuce. 

Figure 1: Partial adhesion between penile and preputial mucosa (solid black arrow) in a newborn colt. The penis is being drawn to the right, the prepuce to the left. Image size: 3264 x 2448px

Reference

Canisso I.F. et al. 2020. Congenital phimosis causing preputial swelling in a newborn foal. Can Vet J .

61:247-250.

Pyometra in a pluriparous Standardbred mare.

Keywords: pyometra, equine, treatment.

Primary author: Dr Rob Lofstedt.  Dept of Health Management, Atlantic Veterinary College.  Additional authors:  Dr Anna  Potter (primary clinician & corresponding author: apotter@upei.ca ) and Dr Martha Mellish, both of the section of Theriogenology, Dept of Health Management, Atlantic Veterinary College & Dr Shannon Martinson of the Department of Pathology and Microbiology, Atlantic Veterinary College. Editors: Drs Rob Lofstedt (lofstedt@upei.ca) and Allan Gunn (algunn@csu.edu.au).

A 21 year-old nulliparous Standardbred mare in a teaching herd was euthanized because of respiratory pathology, lameness and a diagnosis of pyometra. Understandably, this mare had experienced numerous per vagina examinations, uterine cultures, uterine biopsies and at least one hysteroscopy.

Early in April 2019, an enlarged, fluid-filled uterus was palpable. On transrectal ultrasonography, this fluid was partially echogenic and suspected to be pus (see figure 1). Due to its size and dependency, it was not possible to delineate the uterus. It was also not possible to palpate the ovaries. Trans-abdominal ultrasonography revealed that the uterus was highly distended and at its cranial aspect, lay adjacent to the xiphoid process. Nevertheless (and typical for mares with pyometra) the mare was bright, alert and responsive and had a normal hemogram.

The mare’s ovaries containing no luteal tissue; not an unusual finding in equine pyometra. Pyometra in mares does not necessarily develop in a progesterone dominated milieu i.e. unlike the situation in cattle, cervical closure and myometrial quiescence due to progesterone is not required for the development of pyometra. In mares it is more likely that retention of  pus within the uterus is a function of deficient myometrial activity than cervical pathology (See LeBlanc et al 1994 and Troedsson 1999), especially in older animals. Occasionally however, cervical pathology is implicated.

Figure 1. Note the accumulation of moderately echogenic pus within the uterine lumen and follicles within the ovaries. This image also serves as an excellent example of  two common ultrasonographic artifacts; reverberation and enhancement through transmission (ETT). Image Copyright: Dr Martha Mellish. mmellish@upei.ca  Image size: 862 x 947 px.

During per-vagina examination, the mare’s cervix was found to be closed tightly. Cervical patency was only established after approximately 20 minutes of digital manipulation and only then, did pus drip from the vulva lips as shown in figure 2. This suggested that cervical pathology was indeed implicated in retention of pus within the uterus, perhaps in conjunction with poor myometrial tone as discussed earlier. As seen in figure 2, a large diameter stomach tube was eventually passed through the cervix to drain the uterus. Cytology and culture were performed prior to drainage of the pus.

Figure 2. The main image shows how pus dripped from the mare’s vulva after gradual dilation of the cervical canal. The inset shows how pus was then drained from the uterus. Image Copyright: Dr Martha Mellish. mmellish@upei.ca  Image size: 1513 x 1024 px

Several days after initial drainage, the uterus was  flushed repeatedly with saline (see figure 3). Then oxytocin was administered to facilitate the expulsion of any remaining fluid. In addition, 1000 mg of prostaglandin E-1 (misoprostol) in methylcellulose was applied to the cervix and cervical canal to facilitate dilation for drainage and further treatment.

Figure 3. Serial saline flushes showing the increase in clarity with each successive flush. Copyright: Dr Anna Potter apotter@upei.ca Image size 2000 x 1215

Pending culture and sensitivity results, the uterus was lavaged daily with lactated ringers or saline solution.  Two days after the initial drainage, culture results revealed Pseudomonas aeruginosa, sensitive to gentamicin, amikacin and enrofloxacin.  Treatment was initiated with 1g of gentamicin buffered with 10ml of 8.4% sodium bicarbonate infused into the uterus after uterine lavage, every 24 hours for five days.  On the fourth day of antibiotic treatment, an additional treatment of 100ml of 90% DMSO was added into the second to last flush for three days. These treatments were followed by twice daily oxytocin treatment at appropriate intervals.

Two weeks after initial dilation of the cervix and uterine flushing, ultrasonography revealed a uterus devoid of any free fluid, multiple small follicles in both ovaries and a corpus luteum. Uterine culture at that time revealed Citrobacter koseri and Streptococcus zooepidemicus. However, approximately a month days later, the uterus had re-filled with pus and uterine culture again revealed growth of Streptococcus zooepidemicus. Six days later i.e. approximately 48 days from the initial dilation of the cervix, approximately 2.5 liters of pus was drained and treatment with saline flushes, DMSO and oxytocin treatment were re-started. On this occasion, treatment lasted for four days; the antibiotic being penicillin, not gentamicin. On day four, another uterine culture revealed growth of Citrobacter koseri.

The mare was re-examined about three and a half months after initial presentation and summer rest on pasture. At that time, three liters of pus was drained from her uterus, followed by saline lavage.

In early November, 2019, re-examination revealed that a large amount of pus had accumulated in the uterus again. The mare was then euthanized and submitted for post mortem exam. Her pus-filled uterus is shown in the inset of Figure 4.

Figure 4. The pus-filled uterus of the mare seen during postmortem examination. The appearance of pus shown here is typical for equine pyometra. Image Copyright: Dr Shannon Martinson. smartinson@upei.ca  Image size: 1500 x 911 px.

Figure 5. It is probable that the cervical lesion seen here was implicated in the development of pyometra, together with general cervical cicatrization and myometrial compromise. The cause of the lesion was unknown and could not have been caused by foaling as this was a nulliparous mare. Image Copyright: Dr Shannon Martinson. smartinson@upei.ca Image size: 2002 x 1232 px.

Bearing in mind the difficulty of cervical dilation in this case and the absence of spontaneous drainage of pus before and after treatment, cervical pathology (seen in figure 5; probably fibrosis after cervical damage) was probably important in the development of pyometra in this case.

Editor’s comments: In light of the typical recurrence and usually dismal prognosis of equine pyometra, some may be critical of the handling of this case with regard to repeated attempts at physical drainage, antibiotic and anti inflammatory treatment. Consider however, that in recent years, the use of cervical wedge resection and intra-cervical stents have provided favorable outcomes in cases of equine pyometra. This suggests that even if one presumes the presence of myometrial inadequacy, attempts to dilate the cervix both physically and hormonally (using PGE analogs) may hold merit and should be considered in some cases. Of course, wedge resection and cervical stents should be considered as well.

The bacteriology in this case also deserves comment. The significance of any of the bacteria isolated is open to question. For example, a literature search of Citrobacter koseri is devoid of examples of infertility in mares so this can be presumed to be a contaminant.  Also, Streptococcus zooepidemicus is not only a common cause of endometritis in mares, it is a common commensal and contaminant of uterine cultures as well. Also, Pseudomonas aeroginosa is commonly isolated from soil samples and it therefore likely to be present on mares at pasture. The absence or presence of bacteria in single or serial cultures in this mare is also open to discussion; bacterial cultures vary in success according to sampling or culture methods. Essentially therefore, the bacterium or bacteria responsible for pyometra in this case remains a question.

Selected references:

Aguilar, J. et al. 2006. Importance of using guarded techniques for the preparation of endometrial cytology smears in mares. Theriogenology. 66:423-430

Arnold, C.E. et al. 2015 Cervical wedge resection for treatment of pyometra secondary to transluminal cervical adhesions in six mares. J. Am Vet Med Assoc. 246: 13540-1357

Blanchard, T. et al 1981. Comparison between two techniques for endometrial swab culture and between biopsy and culture in barren mares. Theriogenology 16: 541-552

Ismaïl, R. et al 2013. Methods for recovering microorganisms from solid surfaces used in the food industry: A review of the literature. Int. J. Environ. Res. Public Health. 10:6169-6183

Katila, T, 2016 Evaluation of diagnostic methods in equine endometritis. Reproductive Biol. 16:189-196

Krohn, J. et al 2019 Use of a cervical stent for long‐term treatment of pyometra in
the mare: A report of three cases  Reprod Dom Anim. 54:1155–1159.

LeBlanc M.M. et al. 1994 Scintigraphic measurement of uterine clearance in normal mares and mares with recurrent endometritis.
Equine Vet.J. 26: 109-133

Pasolini M.V. et al. 2015 Endometritis and infertility in the mare – The challenge in equine breeding industry–A review. Open access peer-reviewed chapter.

Rötting A.K. et al 2004 Total and partial ovariohysterectomy in seven mares. British Equine Vet. J. 36:29-33

Troedsson. M.H.T. 1999 Uterine clearance and resistance to persistent endometritis in the mare. Theriogenology. 52:461-471

The oviduct (uterine tube) revisited

Keywords: equine, mare, oviduct, uterine tube

The Nomina Anatomica Veterinaria refers to this structure as the uterine tube (to distinguish it from the oviduct of birds). However, that term is seldom used in either practice or publication. In both humans and animals, it is instead referred to as the oviduct. The oviduct in any species is amazing but even more so in mares. This entry substantiates that impression.

As shown in figure 1, the oviduct runs within the ovarian bursa, almost parallel to its margin but a full centimeter or more away.

Figure 1. The left ovary of a two year old mare, suspended under water. The ovulation fossa is invisible, pointing ventrally in the image. 2903 x 2054 px

Figure 1 is labeled above. Divisions of the oviduct (infundibulum, ampulla etc) shown here are those adopted from, and and described in: Aguilar, J.J. et al. 2012. Histological characteristics of the equine oviductal mucosa at different reproductive stages. J.Equine.Vet. Sci. 32:99-105. Note: The ovary in situ hangs from the mesovarian ligament so that the ovulation fossa faces ventrally. In this image, the ovary and bursa have been rotated as shown by the arrow in the small inset. The ovulation fossa is not yet visible despite this rotation. Image size: 1600 x 1041 px

The ovarian bursa can be likened to a lateral, low-drooping eyelid over an eye (the ovary). The long dimension of the ovoid-shaped ovary lies on a cranial-caudal axis. The cranial pole of this axis is slightly higher than the caudal pole. The infundibulum lies at the cranial pole of the ovary. As shown in figure 1, it is not attached to the ovary. In fact, it lies a remarkable distance from the ovulation fossa.

This anatomy never ceases to amaze the author. In essence, the infundibulum acts like a catcher's mitt in a baseball game, covering an ambitious area some distance from the pitcher's mound i.e. the ovulation fossa in this analogy. The baseball is of course, the oocyte. The infundibulum is well supplied with smooth muscle and engorged blood vessels during estrus, expanding the catcher's mitt. Yet the precise mechanism and magic behind the dependability of the catcher remains unknown and unseen. For those not familiar with baseball i.e. (insert nationality here), the author suggests consulting the book "Complete idiots guide to baseball and oocytes". It has been suggested that the fimbriae of the infundibulum sweep the surface of the ovary at the time of ovulation, picking up the oocyte in the process, moving it into the complex folds of the infundibulum. Although the frequency of loss of oocytes into the peritoneal cavity is unknown in mares, it probably does occur; it has certainly been documented in humans. Certainly, losses of oocytes into the peritoneal cavity is described in poultry, especially broiler hens. Interestingly, laying hens, selected for egg production are less prone to peritoneal loss of oocytes. In rodents and canids, oocyte-catching expertise by the infundibulim is less important than other species. This is because ovarian bursae in those animals surround their ovaries completely and are continuous with the infundibula themselves. This makes it impossible for their oocytes to escape into the peritoneal cavity.

Figure 2: An oocyte, 150µ in diameter is shown at the end of the yellow arrow. This simulates the appearance of a real oocyte shortly after ovulation. Note its size relative to the infundibulum and ostium. Also note the cloud-like mass around the oocyte. This is a simulation of the large cumulus oophorus that accompanies the oocyte into the infundibulum. The cumulus is lost within 6 to 12 hours after the oocyte enters the oviduct (personal communication; Dr Katrin Hinricks).  Image size: 1630 x 1236 px

When stretched out, the oviduct in a mare is a little longer than the human hand i.e. about 20 to 30 centimeters. It is a continuum with no distinct delineation. To facilitate functional descriptions however, it is divided into three main sections i.e. the infundibulum (L.< funnel), ampulla (L.< flask) and finally the isthmus (L.<neck of land between two seas) narrowing as it enters the uterus at a papilla that forms the uterotubal junction.

Figure 3: This figure includes a large section of the infundibulum and a smaller inset image of the isthmus. They are both at the same scale of magnification. Again, the author has modeled an oocyte in the infundibulum and in addition, an embryo in the isthmus. Both are visible beside black bars 150µ ling at the end of the yellow arrows. This was done to compare the size those structures with the histology of parts of the oviduct. The diameter of an equine oocyte is approximately 150µ; slightly larger than that a bovine oocyte (120µ). By day 6, the equine embryo is approximately 200µ in diameter. The white scale within each image is 500µ and the small bar adjacent to the oocyte and embryo is 150µ long. Image size: 4242 x 3090 px

The oviduct is of course, a conduit to transport oocytes, then embryos from the ovary to the uterus. But it is also an organ that performs the seemingly impossible task of (often simultaneously) transporting oocytes towards the uterus while promoting the ascent of spermatozoa from the uterus into the oviduct. Like other tubular organs throughout the body it has inner circular and outer longitudinal layers of smooth muscle that promote peristaltic movement. It also has within its mucosal lining, ciliated cells that also play a role in gamete transport. Presumably, peristaltic movements play a major role in transporting oocytes towards the uterus while cilia perform a major role with regard to the ascent of spermatozoa. However, the exact integration of these two propelling mechanisms has yet to be described.

It is known that spermatozoa ascend into the oviduct and bind to its epithelium, usually laying in wait for the oocyte to arrive after ovulation. It is possible for fertilization to occur when spermatozoa arrive in the oviduct up to 18 hours after ovulation (with post ovulation insemination) but it is far more common for spermatozoa to spend a day or two or even up to 7 days in the oviduct before ovulation. During that time, Ca⁺⁺ fluxes within the oviduct suppress capacitation while gaseous exchange and nutrition keep spermatozoa viable. True, it is more likely that an oocyte will be fertilized with close synchrony between insemination and ovulation but the ability of the oviduct to keep spermatozoa viable for long periods of time is still remarkable; far superior than any device contrived by humans.

As amazing as sperm preservation maybe in mares, it is overshadowed by the achievement of oviducts in other species. In some fruit bats in hibernation for example, spermatozoa can survive for weeks even months within the oviduct!

Capacitation and the release of spermatozoa from their binding sites on the oviduct epithelium is orchestrated by changes in the steroid milieu, especially increased progesterone production shortly before ovulation. The effect of the oviduct environment on spermatozoa is of critical importance in mares. Specifically (and peculiar to mares again) it is only in the oviduct that fertilization can occur. Therefore, unless spermatozoa are injected directly  into oocytes (ICSI), in-vitro fertilization in horses is seldom successful.

Figure 2: A 16 gauge blunted needle is introduced into the ostium of the oviduct from a two-year-old mare. Blue dye is then introduced to outline the convoluted shape of the oviduct. Evidence of the dye in the uterus is shown as it permeates through to the serosa at the site marked B. This image defines the oviduct clearly. However the anatomical divisions (especially the ampullary-ithmic junction) so glibly discussed in literature, are far from obvious. Image size: 3456 x 2557 px

The uterotubal junction too, is a remarkable structure in mares. Spermatozoa deposited in the uterus are swept up to the uterotubal junction and are found in the oviduct within a few minutes of insemination. Yet, it is virtually impossible in normal mares, to force either fluid or air from the uterine lumen into the oviduct. This is because the oviduct of the mare is unique amoung domestic species with respect to its uterotubal junction. In mares, the distal oviduct has a well developed muscularis which acts as a sphincter, making mechanical entry from the uterus difficult. One can introduce fine tubes into the oviduct from the uterus but otherwise the uterotubal junction in mares acts as a one-way valve preventing fluid ascent from the uterus. To some degree, this may explain the relative lack of oviduct pathology in mares compared to cattle.

Alter about six and a half days in the oviduct, incubating mainly at the ampullary-ithmic junction, embryos enter the uterus. In mares of course, single embryos are far more common than twin embryos. At that time, late morulas or early blastocysts can be collected by flushing the uterus. Before that time it is impossible to retrieve a fertilized embryo by flushing the uterus alone.

Although unfertilized oocytes are occasionally found in the uterus, this is unusual. In general, if an oocyte is not fertilized in mares, it will not reach uterus. This phenomenon is unique among equids. Therefore it is generally not important to determine if oocytes have been fertilized when they are collected for embryo transfer. It is now universally recognized that production of prostaglandin E₂ by embryos (not oocytes) causes relaxation of oviduct smooth muscle. This allows transport of the embryo into the uterus. When mares are examined postmortem, it is not unusual to find flattened, degenerate oocytes from previous cycles, caught within the oviduct.

Pathology?  Fibrinous masses that can be several mm in size, are often found in the oviducts of mares. Again, this is a phenomenon peculiar to equids. It has been suggested that they are pathological and may block the passage of  oocytes and embryos in the oviduct. However, these masses are found in 75% to 85% of mares (as reviewed by Tsutsumi, Y. 1979) therefore they are unlikely to be pathological. The origin of the masses is unknown but they may arise from fibrin discharged from follicles after ovulation, during the formation of corpora hemorrhagica. In that regard, it is also very common to see fibrin tags in and around the ovary in apparently normal mares. In fact, careful inspection of the images in this entry will reveal such tags. The author had the dubious privilege of spending many hours at an equine slaughter plant and saw such tags many times, often in young mares. The same can be said for para-ovarian (wolffian) cysts, sometimes reported as abnormal too. The vast majority of otherwise normal mares have these cysts.

Selected references:

Allen, W.E. et al. 1979. Evaluation of uterine tube function in pony mares. Vet. Record 105: 364-366

Arnold, C.E. and Love, C.C. 2013. Laparoscopic evaluation of oviductal patency in the standing mare. Theriogenology 79: 905-910

Bennett, S. 2002. Surgical evaluation of oviduct disease and patency in the mare. Proc. AAEP 48:347-349

Betteridge, K.J. 2000. Comparative aspects of equine embryonic development. Anim. Reprod. Sci. 60: 691-702

Brinsko, S,P. 1991. The effect of uterine lavage performed four hours post insemination on pregnancy rate in mares. Theriogenology 35: 1111-1119

Dobrinski, I. et al. 1997. Membrane contact with oviductal epithelium modulates the intracellular calcium concentration of equine spermatozoa in vitro. Biol. Reprod. 56: 861-869

Freeman, D.A. 1991. Time of embryo transport through the mare oviduct. Theriogenology 36: 823-830

Ghazal, S et al. Glob. libr. women's med., (ISSN: 1756-2228) 2014; DOI 10.3843/GLOWM.10317

Hinrichs, K. 2010. In vitro production of equine embryos: State of the art. Reprod. Domestic Anim 45: 3-8

Hunter, R.H.F. 1999 Ovarian follicular fluid, progesterone and Ca2+ ion influences on sperm release from the Fallopian tube reservoir. Gamete biology. 54: 283-291

Hunter, R.H.F. 2008. Sperm release from oviduct epithelial binding is controlled hormonally by peri‐ovulatory graafian follicles. Molecular Reprod. Devel. Incorporating Gamete Research 75: 167-174

Inoue, Y. 2013 Hysteroscopic hydrotubation of the equine oviduct. Equine Vet J. 45:761-765

Kenney, R.M. 1993. A review of the pathology of the equine oviduct. R. M. Kenney. Equine Vet. J. 25 (S15): 42-46

Leemans, B.M. 2015. Why doesn’t conventional IVF work in the horse? The equine oviduct as a microenvironment for capacitation/fertilization. Reproduction 152: R233-R245

Navara, K. J. 2015. Higher rates of internal ovulations occur in broiler breeder hens treated with testosterone. Poult Sci. 94:1346-1352

Rigby, S. et al. 2000. Oviductal sperm numbers following proximal uterine horn or uterine body insemination. Proc. AAEP. 46:332-334

Saltiel, A. et al. 1986. Pathologic findings in the oviducts of mares.  Am. J. Vet Res. 47: 594-597

Sieme, H. et al. 2003. The effects of different insemination regimes on fertility in mares. Theriogenology 60: 1153-1164

Smits, K et al. 2016 The equine embryo influences immune-related gene expression in the oviduct. Biol. Reprod. 36: 1-8

Tsutsumi, Y. 1979. Evidence of the origin of the gelatinous masses in the oviducts of mares.
J. Reprod. Fert. 57: 287-290

Weber, J.A. 1995. Relaxatory effect of prostaglandin E2 on circular smooth muscle isolated from the equine oviductal isthmus. Biol. Reprod. Monograph. series1: 125-130

An STL model of the equine uterus

Keywords: model, 3D, STL, equine, uterus.

The author has used a stereolithographic (STL) file to view and print equine uteruses and pelvic structures in 3D.

NEW: This, small file can even be downloaded on a cell phone. Pay no attention to dialog alluding to processing of the file. Go ahead and download it. With the App "Fast STL viewer" (see Google play store) already downloaded on your phone, click on this downloaded file (not the App itself!) and the uterus will appear on your phone as shown in the thumbnail above. Incidentally, the same file can be downloaded and viewed on your laptop. 

Viewing the model:

For STL viewing alone on your laptop the author suggests that you download this file. Click on it and your laptop (PC or Apple) will bring up a suitable program for viewing. An example of  such an interface is shown below. 

Printing the model:

Custom STL printing can be done through Axis Prototypes in Montreal after contacting the author.  Axis holds the right to a digitally engineered model of the tract and pelvis that is ready for printing. High quality prints such as those seen here are not inexpensive.  Printing a 25 cm model (caudal to cranial) printed from STS nylon will cost approximately $600 Can. ($460 US). 

Manipulating the model:

In the absence of a real uterus or a printed tract, this digital model serves to remind one of many practical aspects of equine reproduction  A short list comes to mind immediately. Some may wish to print this list so that it can be referred to as the model is manipulated. Better still, manipulate the model on a cell phone while reading the learning points on a laptop.

1.Rotate the tract for a view of the vulva lips and clitoris. Be reminded of its "winking" appearance often, but not exclusively, seen during estrus. Note the importance of the clitoris in diagnosis of CEM and the conformation of the vulva lips in infertility. 

2.Enter the vagina and move cranially until the external urethral orifice is encountered. Note that the hymen is normally found just cranial to the orifice but is absent here because this tract was modeled from that of a pluriparous mare. At this point, one should also contemplate the potential problem of urine pooling in the cranial vagina in older mares due to splanchnoptosis.

3.Traveling further cranial in the vagina, one can see the opening of the cervix, showing its dorsal frenulum. Consider the simple nature of the equine cervix compared to that in other species i.e. there are no transverse cervical rings and the cervix is easily dilated for artificial insemination, embryo collection and is even distensible enough to remove fetuses that are three or four months old. The same distensibility is certainly not encountered in ruminants and other domestic animals.

4.Turning to the left and right at a point just caudal to the cervix, one should be reminded that the vagina lies completely within the peritoneal cavity at this point. This is important because it permits one to penetrate the cranial vagina for standing ovariectomy in mares. It also means that if the cranial vagina is torn during foaling, evisceration can occur from this site.

5.Withdraw from the vagina and rotate the model so that the cervix is visible from within the peritoneal cavity. Note that the cranial portion of cervix is firmly suspended between the ilial shafts on either side, by the mesocervix i.e. the caudal portion of the mesometrium. This is significant because the cervix of a mare cannot be grasped, elevated and manipulated as it can in a cow.

6.Rotate and tilt the model so that the mesometrium, containing the uterine artery (middle uterine artery) can be seen clearly. Be reminded that a uterine artery can be forced back against the shaft of the ileum during foaling, rupturing the vessel, causing severe hemorrhage within the mesometrium. Unlike the situation in cows, there is no fremitus in this artery during pregnancy.

7.Rotate the model so that the attachment of the mesometrium the uterus along its lateral borders can be appreciated. Note in turn, how the mesometrium is attached to the abdominal wall on either side. This firm attachment affords substantial stability to the equine uterus and explains in part why uterine torsion is less common in mares than it is in cows. It also means that the equine uterus cannot be retracted during palpation as is done in cattle.

8.Rotate the uterus so that the intercornual ligament is visible. Although it is fairly well-developed. It is never used to retract the uterus  for reasons just mentioned.

9.Tilt the uterus so that its dorsal surface is visible, showing how the bifurcation of the uterine horns provides a T-shaped structure instead of the Y-shape characteristic of the bovine uterus.

10.Rotate the uterus up and down on it's transverse axis, noting the length of the uterine body in a mare compared to the very short body in ruminants. This has many implications. For example, in cattle is very important to enter a particular uterine horn for insemination or embryo collection. In mares by contrast, the long uterine body means that semen is simply deposited anywhere in the body and during embryo collection, the embryo is collected from the uterine body.

11.Rotate the tract and pan in and out over the ovaries, noting their large size in comparison to those of cows. Be reminded that although equine corpora lutea are three or four times the diameter of bovine corpora lutea, they cannot be detected by transrectal ovarian palpation. This is because equine  ovaries are surrounded by thick tunica albugineae, disguising the presence of corpora lutea. Therefore, to determine if a mare is ovulating (in the absence of progesterone assay) one must use ultrasonography

12. Noticeably absent from this model are the ovarian bursae. The author apologizes for that oversight and time allowing, will add that in his next life.

Unusual placentation in a twin pregnancy. 

Keywords: placenta, equine, mare, inverted, twin, complication

Companion to an entry on twinning in mares in LORI.

There is still some uncertainty as to how and why twin equine pregnancies result in abortion. Sharing the available endometrial area is certainly a significant factor. Although typical inflammatory reactions at placental interfaces do not characterize twin abortions, immunological rejection of one co-twin by another may also be an important factor. Figure 1 shows one of numerous permutations of twin placentas apposed to one another within the uterus; placental sharing of the endometrial surface may be almost equal. In others, sharing is dramatically unequal. Figure is a schematic representation of such a case. Indeed, the situation in this pregnancy.

In Figure 1 red arrows indicate an area of apposition between the two chorionic surfaces. In these placentas there was little if any, macroscopic reaction between the two conceptuses despite the intimacy of placental apposition. Approximately 40% of the chorionic surface from the smaller co-twin was invaginated into the placenta of the larger twin.

Figure 1: Apposition of twin placentas and the path of delivery for both foals. Size available: 1626 x 2172px

The green arrows in Figure 1 show where the chorioallantois of the larger twin ruptured; in the region of the cervical star. The chorioallantois containing the smaller foal (yellow arrows) appeared within the allantoic cavity of the larger foal, invaginated within a pocket of the larger foal's placenta. The smaller foal was then delivered through a rupture of two layers of chorioallantois; that of the larger foal and that of its own placenta. The lower part of Figure 1 shows the path (grey arrow) that had to be taken by the smaller foal to be born; through the placenta of the larger foal.

Figure 2. A schematic representation of the placentation in this case. In some cases, the smaller co-twin may die and become a mummified attachment to the placenta of a normal foal. Image size available: 2553 x 1886px

Often, the placentas of twin foals will lie side by side within the uterus and each foal will be born in a conventional fashion, through the rupture of independent chorioallantoic membranes adjacent to the cervix.

Equine anatomy from the seminiferous tubules to the ductus deferens. 

Keywords: seminiferous, tubule, testis, testes, epididymis, ductus, deferens, caput, corpus, cauda, spermatozoa, spermatozoon, sperm, efferent, stallion, Leydig, spermatogonia, Sertoli

For gross images of the testis and adnexa, see the LORI entry on reproductive aspects of stallion anatomy. This entry addresses the applied histology of the testis and its adnexa.

This is a substantial entry and should be viewed as a project in progress. Images will be added or replaced with time. The entry will improve under the scrutiny of colleagues.

Because of their combined use in common conversation, the author uses a combination of Latin and English terminology in this entry. For example, the terms caput, corpus and cauda are used in preference to head, body and tail. Although the term vas deferens appears about 35 times more frequently than the term ductus deferens in literature searches, this is no doubt due to its prevalence in human oriented literature. The Nomina Anatomica Veterinaria uses the term ductus to the exclusion of vas deferens. Therefore the term ductus deferens is used in this entry. In addition, eponyms are capitalized; a gradual editorial change throughout LORI. The author begs the understanding of purists.

Overview

Figure 1. A simplified overview of the testis, epididymis and ductus deferens. The cranial aspect of the testis lies on the left side of the diagram. Size available: 2478 x 1645px

Although seminiferous tubules join major rete tubules as shown, this image is a simplification. In reality, loops of seminiferous tubules actually end throughout the testes, joining tubules of the rete system throughout the testis. The word "rete" is derived from the Latin word for net. This is appropriate because the rete tubules form a net-like system throughout the testis.  Small rete tubules then run centripetally to major rete ducts as shown. Larger rete ducts then course cranially within a poorly defined mediastinum, to the efferent duct system.

Although the mediastinum carries a large central vein (see figure 4), the mediastinum is not nearly as obvious as it is in ruminants. Interestingly, there is no central artery in an equine testis; arterial supply is served via several peripheral arteries.

The rete is best developed towards the cranial pole of the testis where the total production of spermatozoa accumulates. As discussed later, the rete actually originates as part of the mesonephric system, not the primitive gonad. Therefore, rete tubules penetrate the testis during early embryonic development.

The label "Anastomosing rete tubules" in the illustration draws attention to the fact that parts of the rete tubule system fuse in that area. These rete tubules form sinus-like cavities (see figure 28) both inside and outside the tunica albuginea. The sinuses in turn, become efferent ducts in both the testis and caput epididymis. Downstream, the efferent ducts become a single tube, the epididymis.

Orientation of testis adnexa within the scrotum
In primates and ruminants the longitudinal axes of the testis is vertical. In those animals, the caput  epididymis lies dorsal to the testis, the cauda lies ventral to the testis. In horses and dogs the testes "swing" in a caudal direction during embryonic development so that the cauda comes to lie caudal to the testis, roughly on the same plane as the caput. In cats, camelids and pigs that movement is even greater, so that the cauda lies above the caput. See these approximations in figure 2.

Figure 2. Orientation of the testes in various species. Size available:1045 x 554px

In all animals the ductus deferens runs medial to the epididymis. In horses, this is on the dorsal aspect of the testes (see figures 2, 3 and 4). In the rare instance that one should perform a vasectomy on an equid (in zoo animals for example) this should be born in mind; a ventral paramedian incision is required to access the ductus on the correct side of the vaginal cavity. See this entry on vasectomy for clarification.

Figure 3. The origin of a section across the center of the right testis from a 26 month old Miniature horse stallion, viewed caudal to cranial. That section is enlarged below. Size available: 1747 x 1506px

Figure 4. A section across the center of the right testis from a 26 month old Miniature horse stallion, viewed caudal to cranial. Size available: 2765 x 2274px

For castration, local anesthetic is often injected directly into the testes to provide additional analgesia during general anesthesia. Indeed, studies show that blood pressure and heart rates remain lower in subjects so treated. See Haga et al. 2006.  Similar effects have been shown in dogs and cats. The artifact created by local anesthetic is clearly visible in this image.

Incidentally, a "closed" castration technique was used to prevent post operative evisceration in this case. This was fortuitous because the parietal vaginal tunic is still present in this specimen, showing an intact vaginal cavity. That would not have been the case if open castration had been used. In open castrations, the parietal vaginal tunic is transected as the incision is made through the scrotal skin, into the testis. In closed castrations, the scrotal skin is incised but the parietal vaginal tunic is stripped away from the tunica dartos to ensure that vaginal cavity remains intact. This allows for easy closure of the inguinal canal, preventing post-operative evisceration.

Puberty
During fetal development, Sertoli cells aggregate around clusters of germ cells in the genital ridges. These cells are known as gonocytes (the precursors of spermatogonia) and in linear clusters, they form gonadal cords. The testes, so formed, descend into the scrotum within the vaginal pouch. Gonadal cords, persist up to puberty when they become canalized and forming seminiferous tubules.

Remember to double click all the images in LORI to appreciate the detail under discussion.

Figure 5. Gonadal cords in a specimen taken from a ten month old Standardbred colt. Size available: 2496 x 1113px

Inside the cords, Sertoli cells surround gonocytes, shielding them from the effect of retinoic acid that is circulating in the interstitium. Retinoic acid is an oxidation product of Vitamin A. It is known to induce maturation of cells throughout the body, especially in the bone marrow and testes. In the testes, retinoic acid initiates the division of gonocytes during puberty. After puberty, retinoic acid controls the maturation and division of spermatogonia themselves. Were it not for the protection from retinoic acid afforded by the Sertoli cells, gonocytes would undergo meiosis prematurely. In the gonadal cords therefore, there is no meiosis or mitosis; only solid aggregations of Sertoli cells and gonocytes.

During the gradual process of puberty, gonadal cords canalize under the effect of testosterone from Leydig cells in the interstitium, forming seminiferous tubules. That effect is seen in figure 6. Seminiferous tubules do not mature simultaneously throughout the testis. Maturation usually starts at the center of the testis, progressing towards its periphery. Areas of canalized tubules can even be seen macroscopically as lighter areas of tissue on the cut surface of a testis.

Figure 6. Maturation of seminiferous tubules in a 10 month old peripuberal colt. Size available:  2384 x 1471px

Although a central-to-peripheral pattern maturation pattern appears to be common, testes may also mature in no fixed pattern or even peripheral-to-central in some individuals. The Standardbred colt samples for figure 5 was such an example. Because meiosis was not present in solid areas of the testes of this colt but was seen in canalizing areas, the author was assured that this was indeed a physiological effect and not an artifact due to injection. See figure 7.

Figure 7. Peripheral-to-central maturation of seminiferous tubules in a 10 month old peripuberal colt.  Size available: 1362 x 921px

In view of the fact that puberty initiates meiosis and at that time, the blood-testis-barrier is first tested against haploid cells, it is not surprising to see occasional infiltrations of lymphocytes within a peripuberal testis. Perhaps this formalizes the uniquely privileged immune system in testes. To the author's knowledge, this phenomenon has not be discussed in the literature. These infiltrations were visible in figure 7 and in sections from other peripuberal testes.

In peripuberal testes, where spermatogenesis is just beginning, the typical cell associations of spermatogenesis seen in mature testes are not yet evident. In canalizing tubules the process is irregular. In that regard, see figure 8, a section from the Standardbred colt mentioned above. The division between active seminiferous tubules and non-canalized cords is obvious

Figure 8. Division between active seminiferous tubules and non-canalized cords in a 10 month old peripuberal colt. Size available: 1919 x 1157px

The apoptosis that normally occurs amoung dividing spermatogonia in mature stallions is also noticeable in peripuberal testes. See figure 9. Pyknotic nuclei commonly seen amoung immature spermatocytes substantiated that impression. Also in figure 9, notice the inactive tubule lying adjacent to one that is beginning to show meiosis. 

Figure 9. Active and inactive tubules in a peripuberal stallion; apoptosis and the onset of meiosis. Size available: 2314 x 1268px

The mature stallion

Before the intricacies of spermatogenesis are illustrated and discussed, readers may be interested to see an overview of the seminiferous epithelium from a healthy six year old Morgan stallion taken during the breeding season. This is provided in the form of a virtual microscopy section. 

Cells in and around the seminiferous epithelium in mature stallions
What follows is an introduction to some of the cells visible in figure 10, a virtual microscopy section. The concept of stages of spermatogenesis is also introduced but a substantial explanation follows later in the entry.  After the ensuing discussions, one may wish to peruse the section again from a renewed perspective.

Figure 10. Click here (NOT on the image) to view the virtual microscopy section. Once the image appears in your browser window, use the scale above the image to select the desired magnification. Magnification may also be selected by placing the cursor over the image and rolling the mouse wheel. To move within the image, left click and drag to a position of interest.

Important! This virtual section uses Adobe Flash. Chrome is a great browser but it makes the use of Flash difficult. Google does this to promote HTML5.  If you have problems viewing the virtual image, use Firefox or Opera browsers. On Apple platforms, use Safari. After 2020, HTML5 will be adopted universally and this site will be amended accordingly.

Spermatogonia:
A complete discussion of spermatogonia is complex and beyond the scope of this entry. Suffice to say that these are diploid cells formed from gonocytes during puberty. After puberty, they form a population of spermatogonia along the basement membranes of the seminiferous tubules. Figure 11 shows spermatogonia undergoing mitosis (arrows); multiplying along the basement membrane of a semiferous tubule.

Image 11. Spermatogonia undergoing mitosis along the basement membranes of seminiferous tubules. Size available: 514 x 428px.

From these spermatogonia, a sub-population is recruited to become spermatozoa at intervals typical of the species in question. In the case of the stallion, spermatogonia are recruited from the same location in a seminiferous tubule at intervals of 12.2 days. In the parlance then, a stallion has a seminiferous cycle of 12.2 days.

Figure 12 shows two forms of spermatogonia present along the basement membranes in adjacent areas of the same seminiferous tubule. From known associations between cells during spermatogenesis (see later) these were identified as B-2 and A-2 spermatogonia.

Figure 12. Two forms of spermatogonia present along the basement membranes in adjacent areas of the same seminiferous tubule. Size available: 1556 x 872px.

Light-staining spermatogonia are recruited from constantly renewing populations on the basement membranes of seminiferous tubules. Although data are not certain in this regard, there are probably five successive generations of spermatogonia. After several successive A sub-type generations, two successive B generations form. B type spermatogonia have dark staining nuclei as shown here. The second B generation then divides to form preleptotene spermatocytes.

Sertoli cells: 
The recruitment, growth and fate of spermatogonia are controlled by Sertoli cells.

Sertoli cells play an important role in recruiting resting spermatogonia into an unforgiving, one-way path to spermatogenesis. The path is unforgiving because most of the unsuspecting recruits will actually undergo programmed cell death (apoptosis) and be phagocytosed by the Sertoli cells i.e. the very cells that have sustained them. Why this act of stepmother-infanticide occurs is not known but is presumed to be a result of genetic infirmity amoung the huge population of spermatogonia that are recruited from the basement membrane. Although good examples of apoptosis do occur occasionally (see figure 9)  apoptosis are not common in histological sections. At the light level anyway, phagosomes containing remnants of spermatogonia are not seen either. Readers may wish to convince themselves of these facts by searching for apoptic spermatogonia in the virtual slide. This phenomenon may reflect the phagocytic efficiency of Sertoli cells.

The cell walls of Sertoli cells are virtually impossible to discern using light microscopy. This is because their walls are in extremely close association with the cells that surround them. In the main image of figure 13, the author has attempted to illustrate Sertoli cell outlines by applying a transparent gray layer over two adjacent Sertoli cells. 

Figure 13. Putative outlines of two adjacent Sertoli cells Size available: 1526 x 948px.

Tight junctions between adjacent Sertoli cells are visible by electron microscopy. It is via these junctions that a seminiferous tubule is divided into basal and adluminal compartments. Spermatogonia (2n) are housed within the basal compartment. All the other cells in the lineage to spermatozoa occur within the adluminal compartment. These include double-diploid (2n x 2) primary spermatocytes, diploid (2n post-meiosis) secondary spermatocytes and haploid (n post-mitosis) spermatids and spermatozoa. The tight cell junctions are therefore an essential part of the blood-testis-barrier that separates the foreign antigens of haploid cells from the peripheral immune system. In figure 12, putative tight cell junctions have also been inserted by the author. In actual fact, these bear a strong resemblance to those seen on electron microscopy.

Note that spermatogonia, spermatocytes and spermatids are surrounded by Sertoli cells. Even sperm are initially surrounded by crypts in the apical region of Sertoli cells. Then, during the last days of spermiogenesis (the process of forming sperm from secondary spermatocytes) they hang by tenuous cell-to-cell connections with the Sertoli cells, within the lumens of seminiferous tubules. In humans and primates it is known that a single Sertoli cell can sustain between 10 and 25 germ cells and up to three spermatozoa. The author is unaware of similar data in stallions.

Sertoli cells are simply marvelous. To reiterate, they support and select generations of  spermatogonia undergoing mitosis to form primary spermatocytes. It is only as the selected spermatogonia become pre-leptotene spermatocytes, entering meiosis, that their surface antigens are recognized by Sertoli cells as beginning to differ from diploid cells. They are then admitted through the tight cell junctions into the adluminal compartment. After the process of selective destruction of most spermatogonia, Sertoli cells allow only the chosen few into that compartment.

Spermatogenesis consists of three phases: spermatocytogenesis, meiosis (including its mitotic phase) and spermiogenesis, which includes spermiation. The author has cut and pasted cells from images of various seminiferous tubules, inserting them into this image at equal magnification. Figure 14 shows their relative sizes, their appearance under light microscopy and their DNA content.

This sequence is never seen in reality because all of these cell types are never found simultaneously in a single cross section of a seminiferous tubule. The reason for this becomes clear later.

Figure 14. Cut and pasted cells from images of various seminiferous tubules. 
Size available: 844 x 742px.
Legend: Sc.gen = spermatocytogenesis. Sp.genesis = spermiogenesis

 

A quick reminder:  Meiosis consists of a meiotic phase followed by a mitotic phase. The DNA content of each new spermatocyte doubles in preparation for the pairing of homologous chromosomes. Then, double-DNA chromatids (including those of the X and Y chromosomes) cross over and exchange DNA for genetic diversity. All of this is accomplished during leptotene, zygotene, pachytene and diplotene.  After the exchange of chromosomal information in diplotene, the paired homologous chromosomes (still containing double the amount of DNA as a somatic chromosome) separate into single chromosomes. During diakinesis, the primary spermatocyte divides into two cells with half the number of chromosomes of a normal somatic cell, essentially making them haploid cells (n). However, these cells still contain twice the amount of DNA of haploid spermatids because their DNA was doubled in the very first stage of meiosis.These are secondary spermatocytes. Note their size in figure 14 and convince yourself that their nuclei are indeed twice the volume* of the spermatids. Secondary spermatocytes then undergo the mitotic phase of meiosis.  In somatic cells, doubling of DNA for mitosis is a time consuming process. However, during spermatogenesis, doubling of DNA has already occurred during leptotene. As a result, this mitotic event far more rapid during spermatogenesis that somatic cell division. Therefore secondary spermatocytes divide into spermatids in a very short time. This explains why secondary  spermatocytes are only visible for single stage in equine spermatogenesis. In some species, secondary spermatocytes are seldom visible on histology.  Diakinesis is also a rapid transitional phase. In fact it is not even long enough to be part of a single discrete stage of spermatogenesis in stallions. *Using the scale on this image and the formula relating radius to volume (V=4/3πrcubed), the author measured the approximate volume of the nuclei of primary and secondary spermatocytes, then spermatids. As expected, the volume of the nuclei halved in each successive stage.

Bear in mind that the DNA of a spermatozoon is highly condensed and its code cannot be translated into protein synthesis i.e. it is non-transcriptional DNA. Therefore spermatozoa are unable to "fend for themselves" and rely entirely on the kind hearts of Sertoli cells for this care. In fact, even as spermatozoa are leaving the maternal grasp of Sertoli cells, specialized connections stretch out to the spermatozoa, grasping them in the considerable slipstream of luminal fluid. Those connections tear back the extra cytoplasmic baggage (residual bodies) that they will not need on their outward journey. Indeed, it is known that  intraluminal fluid travels through the tubules quite rapidly because spermatozoa disappear quickly from the site of spermiation. This complex process is elegantly described in 2011 in a paper by O'Donnell and others (see references).

More on the amazing Sertolis: Sertoli cells produce androgen binding protein and bind testosterone at the high concentrations required for spermatogenesis and various other processes. They also aromatize testosterone into estrogen for important roles it performs in spermiation and later, to regulate fluid absorption in the head of the epididymis. Sertoli cells regulate the secretion of FSH and therefore its effect on the division of spermatogonia by producing the hormone inhibin. Sertoli cells also produce anti-Mullerian hormone and activin.  Other hormones have been identified as being of Sertoli origin as well. Sertoli cells also secrete that complex tubular fluid mentioned in the previous paragraph i.e.the torrent that tears spermatozoa from the nurturing grip of Sertoli cells. Admittedly this fluid will be changed considerably by the epididymis, but consider how specialized it must be to sustain the transcriptionally helpless spermatozoa en route to the outside world! Finally, bear in mind the chameleon-like nature of Sertoli cells with regard to the immune system: Although Sertoli cells are diploid and extend into the haploid environment of the tubule, they do not convey that immunological fact to the environment between the tubules. On the contrary, amoung their many talents, Sertoli cells also have a local immunosuppressive effects in the testes. Indeed, allografts in the interstitial tissue are usually well tolerated, while the same grafts are vigorously rejected elsewhere in the body, even in the epididymis. This is interesting because it indicates that immunological protection is less profound in the epididymis than the testis.

More on the cells of spermatogenesis

Pre-leptotene spermatocytes:

Note the pre-leptotene spermatocytes in figure 15. These cells are beginning to undergo chromosome contraction and double their DNA. They have larger and darker nuclei than spermatogonia but the chromosomes in their nuclei have yet to develop the ribbon-like appearance of leptotene.  The reader may recall that pre-leptotene spermatocytes are the first cells that are eligible to pass through the tight cell junctions of Sertoli cells en route to the adluminal compartment.  The wonder of inter-cell signalling that makes this possible is not yet understood.

Compare the nuclei of pre-leptotene primary spermatocytes to those of the leptotene/zygotene spermatocytes within the tip of a bend in a seminiferous tubule.

Figure 15. Pre-leptotene primary spermatocytes and leptotene/zygotene spermatocytes within the tip of a bend (see ring) in a seminiferous tubule. Size available: 1734 x 979px.

Leptotene spermatocytes:
Primary spermatocytes in leptotene < Greek: lepto (fine) & tene (ribbon or band) are characterized by having chromosomes that may indeed resemble a cluster of fine ribbons in the imagination of an histologist. More prosaically, they resemble thick, dark wet-mops.

Because the cell combination in figure 16 is in stage II of spermatogenesis, it is possible to say with some assurance that the ringed cells are primary spermatocytes in the leptotene stage of meiosis. If the resolution of the image permitted it, the shortened chromatids of each chromosome could be seen. The chromatids would also be thicker than otherwise, having just doubled their DNA content.

Figure 16. Stage II of spermatogenesis. The ringed cells are primary spermatocytes in the leptotene stage of meiosis. Size available: 1356 x 591px.

Zygotene, pachytene and diplotene spermatocytes:
Again, using the cell combinations present in a cross section of a seminiferous tubule and the expected appearance of various stages of spermatocytes in meiosis, one is able to identify cells with some certainty. In that manner, primary spermatocytes in the zygotene, pachytene and diplotene stages of meiosis have been identified in figure 17.

Figure 17. Primary spermatocytes in the zygotene, pachytene and diplotene stages of meiosis. Size available: 1572 x 941px.

Primary spermatocytes in the zygotene stage of meiosis are present with those in diplotene in only one stage of spermatogenesis in stallions i.e. stage III. The lower tubule in this image is in stage III, explaining the presence of both zygotene and diplotene cell types in that cross section.

A return to the virtual slide at the beginning of this entry will convince the viewer that spermatocytes in the pachytene stage of meiosis are very common in cross sections of the testis. This is because of the long duration of that stage of meiosis. In fact, in stallions, seven of the eight stages feature pachytene spermatocytes (See Fig. 18 below).

Seasonality and the effect of aging:
Although spermatogenesis declines during short-day seasons, all tubules show active spermatogenesis throughout the year in domestic stallions. The same appears to be true of donkeys, E. Przewalski and probably other equids (although data are scarce). This is in contrast to many wild ruminants where spermatogenesis may cease altogether in the non-breeding season.

Although spermatogenesis and the fertility of spermatozoa is maintained in stallions during short day seasons, there is a substantial decrease in spermatozoa production. Apparently there a relative shift in the numbers of one type of spermatogonium compared to another during seasonal transitions but that phenomenon is beyond the scope of this discussion.

As a stallion ages and maturity approaches (at four to five years of age) the diameter of seminiferous tubules and spermatozoa production continues to increase. The number of Leydig cells in the interstitium increase as well and continues to do so, even into old age.

An explanation of the stages of spermatogenesis:

Figure 18: The stages of spermatogenesis in stallions. Modified and used with permission. Authors: Drs L. Johnson and D. Varner TAMU. Size available: 1500 x 1115px.

Figure 19 shows a random section of seminiferous tubules, staged using the diagram guidelines shown in the diagram above.

Figure 19. Various stages of spermatogenesis. Size available: 2146 x 1140px.

Notice the difference between the various cross sections of seminiferous tubules. Although this holds little practical importance for practitioners, a knowledge of the stages of spermatogenesis it is potentially important for researchers. Failing all else, the author enjoys understanding why all the tubules in a cross section of a testis do not have an identical appearance. In some of the tubules shown in figure 19, spermiation is approaching, in others, only spermatids are present.

Notice that within each tubule, there is a set number of fixed cell associations based on the recruitment of spermatogonia, stage of meiosis, development of spermatids and finally, spermiation.  Figure 18 shows the eight sets of fixed cell associations referred to in stallions. By convention, each set of associated cells is referred to as a stage, with a Roman numeral used for each stage. Staging of spermatogenesis is not unique to equids. In other animals, fixed stages of spermatogenesis are also seen. For example, six in humans, eight in dogs, eight to 12 in various ruminants and 12 to 14 in mice and rats respectively.

In stallions, at any point along a seminiferous tubule, one of the eight sets of cell associations (cell stages) is seen repeatedly, every 12.2 days. These are "assembly lines" for spermatozoa where new spermatogonia initiate the assembly of spermatozoa from the basement membrane to the lumen by starting  new divisions. Some tasks in the assembly of spermatozoa take longer to complete (e.g. pachytene) than others (e.g. diplotene) so those tasks may be seen in more than one set of cell associations (stages) but each cell association changes every 12.2 days according to the drum-beat from the spermatogonia below. As mentioned earlier, that drum beat is known as the seminiferous cycle. Also to reiterate, a stallions has a seminiferous cycle of 12.2 days.

Note: The author intentionally repeats certain statements in this discussion; done to crystallize an understanding of this complex physiology.

After 57 days on the spermatogenesis assembly line, spermatozoa are produced at the lumen. They roll off the assembly line in that process known as spermiation. Adjacent to each assembly line, there is another assembly line, slightly ahead of behind the line in question. The presence of one assembly line being slightly ahead or behind an adjacent assembly line along a seminiferous tubule forms a sequence known as the spermatogenic wave. This concept is illustrated in figure 22.

The reason for the existence of a spermatic wave is not clear and to the author's knowledge, seldom addressed. It has been suggested that spermatogenic waves reflect be the inability of a single tubule cross section to accommodate continual spermatogenesis, preventing tubular clogging.  However the author has recorded very long sections of synchronous stages in the seminiferous tubules in rats, putting that suggestion into question. Bear in mind however, that the cross sectional area of a tubule increases as square of its radius therefore it is more efficient to supply nutrients to the contents of a small cross section than a larger one and intermittent spermatogonial division allows for a small tubule cross section. Finally, smaller tubule cross sections are geometrically more advantageous compared to large cross sections in terms of total spermatozoa production within a given volume of testes.

Each stage of spermatogenesis is only visible for as long as the shortest cell stage in that section of the tubule. As mentioned earlier, diplotene is a very rapid transition, so any stage containing cells in diplotene is visible for very short time and consequently form a small proportion (about 3%) of all the tubule cross sections in a testis cross section  By contrast a stage containing both pre-leptotene and pachytene spermatocytes persists for longer. Therefore this combination is seen in about 16% of all tubule cross sections. It happens to be stage VIII. See figure 20.

Figure 20. Stage VIII of spermatogenesis. Size available: 1546 x 969px.

Notably, only a small portion of any tubule is in stage VIII of the spermatogenic wave at any time; about 16%. This is the only stage during which spermiation is occurring. Therefore spermatozoa are produced by a mere 16% of a testes at any given time; two testes giving an amazing 5 billion spermatozoa every day!

Figure 21 is another example of a stage; in this case, stage II. It is visible in about 15% of all tubule cross sections.

Figure 21. Stage II of spermatogenesis. Size available: 1020 x 640px.

Figure 22 shows an idealized spermatogenic wave. In this case, cell stages are numbered from left to right along the tubule. Bear in mind that a seminiferous tubule is usually but not always in the form of a loop that attaches to the rete testis at either end. Some tubules are blind-ended. A spermatic wave may move in one direction or the other, depending on where it lies in the tubule. In most animals only one stage occupies a single section of a seminiferous tubule. However, humans, primates and others have no clear spermatogenic waves. Indeed, several stages often occupy a single segment of a seminiferous tubule. That state of disorder is infrequently seen even in stallions. In figure 22 these irregularities are indicated by red exclamation marks and are seen in reality in figure 23.

Figure 22. An idealized equine spermatogenic wave. See discussion for detail. Size available: 2154 x 1518px.

Note the arrow indicating how retinoic acid (a metabolite of  vitamin A) travels along a tubule, stimulating spermatogenesis. It moves between cells along the tubule via paracrine secretion. As discussed in the section on puberty, retinoic is known to be the primary trigger for spermatogenesis. Indeed, retinoic acid is that very drumbeat noted earlier in the recruitment of spermatogonia. Because retinoic acid initiates activity in each region of the tube, stage numbers are in reverse to the direction of retinoic acid travel.

Retinoic acid recruits a new wave of spermatogenesis in each section of a tubule; every 12.2 days. In doing so, it initiates the activity of at least a half dozen transcription reactions that lead to the recruitment of inactive spermatogonia

Apparently retinoic acid is not only taken up from the blood but is also formed by Sertoli cells. It has been stated that spermatogonia themselves produce retinoic acid but this seems unlikely; perhaps retinoic acid is present in spermatogonia merely as part of paracrine secretion along tubules.

Figure 23 shows how one stage of spermatogenesis may impinge on another. In this case, most of the  cells in the cross section belong in stage IV, yet some primary spermatocytes in zygotene (possibly leptotene) are present, close to the basal section on the left side of the image. This suggests that stage III may be starting to develop in that part of the cross section.

Figure 23. One stage of spermatogenesis impinging on another. Size available:  2176 x 117px.

Interestingly, one can destroy most of the cells in a seminiferous tubule in mice, leaving only Sertoli cells and early type-a spermatogonia. If this is followed by retinoic acid treatment, all the spermatogonia respond to retinoic acid simultaneously obliterating spermatic waves. As a result, spermiation occurs simultaneously along the whole seminiferous tubule!

The combined length of the seminiferous tubules in a stallion is over two kilometers (useful for pub trivia contests) therefore each testis contain many thousands of spermatogenic waves. Because the tubules are convoluted, different stages usually lie adjacent to one another in testes cross sections. This explains why a cross section in stage II lies adjacent to another in stage VI in figure 24.

Figure 24. A cross section in stage II lying adjacent to another in stage VI. Size available:  1903 x 1192px.

Leydig cells
Many readers will be aware that Leydig cells are the primary source of androgens (predominantly testosterone) in males. They are also the most common cells in the inter-tubular (interstitial) areas in testes. Figure 25 shows a cross section of a tubule in stage V.  Note the accumulation of lipofuscin in the Leydig cells in this image. Although lipofuscin is even seen in the Leydig cells of prepuberal stallions, it is generally interpreted as a sign of aging. Indeed, lipofuscin becomes more obvious in older stallions. Even macroscopically, testes become darker with aging.

Figure 25. Lipofuscin in Leydig cells and macrophages. See discussion. Size available:  2000 x 902px.

There are also large numbers of macrophages in the interstitial spaces, with lipofuscin being present in those cells as well as Leydig cells. There is an interesting but poorly understood relationship between Leydig cells and macrophages in the interstitial spaces because both appear to have phagocytic and steroidogenic activity! In adult stallions, interstitial macrophages can form close membranous interdigitations with Leydig cells suggesting that they even become a functional syncytium. In figure 25 therefore, it is not clear if the lipofuscin resides in Leydig cells alone, macrophages with Leydig cell-like activity or both cell types.

After spermiation
Following spermiation, spermatozoa are transported towards the rete testis within the seminiferous tubules.  They move along the tubules via the smooth muscle action of myoid cells (see figures 20 and 23). Eventually, these tubules join rete tubules as shown in figure 26. Close examination the rete tubules shows smooth muscle in their walls.

Figure 26. Junction of a seminiferous tubule with a rete tubule. Size available:  1613 x 875px.

Rete tubules course around the central vein of the testis towards its cranial-dorsal extremity. Some rete tubules fuse as they approach the anterior aspect of the testis. This can be seen in figure 27. In one case, several rete tubes lead into an efferent duct. Simple cuboidal epithelium identifies the rete tubules and pseudostratified columnar epithelium, efferent ducts.

Figure 27. Fusion of rete tubules with an efferent duct in the cranial dorsal extremity of a testis. Size available:  2269 x 1052px.

In most cases, rete tubes remain narrow until they reach the efferent ducts but occasionally rete tubes fuse into large, sinus-like collection sites with many criss-crossing trabeculae. One of these is shown in figure 28. An excellent scanning electron micrograph of such a sinus can also be seen in: Amann, R. P. et al. 1977 Connection between the seminiferous tubules and the efferent ducts in the stallion. Am.J. Vet. Res.38: 1571-1579.

Figure 28. Rete tubules and efferent ducts. In one case, efferent ducts have fused to form a sinus with trabeculae. The figure shows a white rectangle over the cranial portion of a testis, adjacent to the caput epididymis. This area included many rete tubules and efferent ducts. However, rete tubules and efferent ducts were also found outside the tunica albuginea. This tissue was obtained from a peripuberal stallion therefore spermatozoa were generally absent from the the duct systems. Size available: 2243 x 1589px.

Amazingly, seminiferous tubules and the rete-efferent systems develop from two extremities. Seminiferous tubules arise from the genital ridges (testes) while rete ducts and efferent ducts arise from the mesonephroses. In most cases these tubes meet end-to-end perfectly; the dream of every engineer on the chunnel project between England and France. In an embryo, the single tube that is the epididymis diverges from the mesonephric side into 10 to 15 efferent ducts and they in turn, diverge into hundreds of rete tubules. From the testis side, hundreds of seminiferous tubules straighten for a short distance then merge with ingrowing rete tubules. On rare occasions, perfect end-to-end anastomosis does not occur and after puberty, spermatozoa granulomas form in the affected areas, especially outside the testis where the interstitium is not as immunologically privileged as it is within the testis.

The epididymis

Within the epididymis: steroids, polypeptides and the autonomic system: Fluid produced by the Sertoli cells is mostly absorbed in the efferent ducts and the initial segment of the caput epididymis, increasing the concentration of spermatozoa. Fluid absorption is strongly mediated by estrogens with principal cells showing clear evidence of estrogen receptors. In fact, estrogen receptors abound throughout the epididymis. In stallions, more estrogen is produced by the testes than testosterone. Estrogen up-regulates oxytocin receptors in the cauda epididymis and modulate spermatogenesis via FSH secretion. Surprisingly, estrogens also modulate Leydig cell and Sertoli cell function! Finally (but probably not finally...) estrogens are known to be important in libido and erectile function in some species; likely the case in horses as well. So much for testosterone and being a "stud"! Smooth muscle increases in bulk and tone as one approaches the cauda epididymis. Both vasopressin and oxytocin receptors increase in the cauda too. This explains why spermatozoa numbers in ejaculates can be increased by oxytocin treatment before semen collection. The function of vasopression in the epididymis is unclear, causing contraction of the cauda in some studies but not others. Although it may be an oversimplification to say so, there is  general agreement that the parasympathetic system is largely responsible for erection and the sympathetic system, ejaculation. Therefore it is not surprising that sympathetic stimulation in rats is largely responsible for myoid cell contraction in the corpus and cauda epididymis. As expected, there is little or no contraction in the caput epididymis. Even after repeated ejaculation in rats, the spermatozoa in the caput are too immature for ejaculation. Similar data are not available in stallions but are presumed to be similar. The influence of the parasympathetic on epididymal function is not clear.

A quick glance at figure 29 shows that the epithelium in the caput  from a mature stallion is thicker and structurally different to that in the cauda. The structures in the image were taken from a mature stallion and are shown at the same magnification. As this figure demonstrates, the size of the lumen enlarges dramatically from the caput to cauda. Structures in the corpus epididymis are intermediate between those of the caput and cauda.

Figure 29.  Caput and cauda epididymis from a 6 year old Morgan stallion. The structures in the image are shown at the same magnification. The size of the lumen enlarges dramatically from the caput to cauda. Size available:  1913 x 1126px.

In stallions, the epididymis is about three quarters the length of a football field (more beer trivia) and although reports vary slightly, the usual time taken for sperm to move through this distance varies from 9 to 11 days.  Of that time, sperm spend just four days travelling through the caput and corpus. The remaining 5 to 7 days are spent in the cauda epididymis. The cauda epithelium secretes substances that promote quiescence and optimal storage conditions for spermatozoa, so it is essentially a storage organ. The caput and corpus hold only 20% of the extra-testicular spermatozoa while the cauda, approximately 65%. The remaining 15% are held in the ampullae and ducti deferentia. Despite the motility they may display in vitro, sperm in the caput and corpus are still immature and infertile. However, spermatozoa harvested from the cauda are indeed fertile, providing conception in both raw and frozen states.

The entire epididymis is lined with a pseudostratified columnar epithelium in which two cell types predominate; principal and basal cells. Figure 30a and 30b are cross sections of the caput epididymis from a 6 year old Morgan stallion.

Figure 30a: Caput epididymis from a 6 year old Morgan stallion. Principal and basal cells predominate. Size available: 2176 x 1136px

Figure 30b: Another view of the caput epididymis from a 6 year old Morgan stallion. Size available: 2500 x 1418px.

Principal cells in the caput epididymis are very tall but gradually become shorter towards the cauda .

Principal cells control the pH and oxygen tension within the epididymal lumen and provide energy for sperm maintenance. They are also the main source of proteins in the epididymis. Indeed, more that 200 proteins have been identified within the epididymal lumen. Some arise from the testis but many from the epididymis itself. These proteins coat sperm and are important in sperm/oocyte interaction. They also protect sperm from the immune system of the female. Others bind to sperm to prevent premature capacitation.

Note that the free surface of most principal cell bear tufts of very long, microvilli. Although they appear to be cilia at low magnification they are indeed villi, not cilia. The point here is that spermatozoa transport along the epididymis is due to smooth muscle contraction, not cilial action. In that regard, a thin layer of longitudinally oriented smooth muscle cells surrounds the caput, corpus and proximal cauda, becoming thicker towards the distal cauda and very thick in ductus deferens. See figure 35.

At their tips, the secretory products of principal cells are voided into the lumen within membranous extracellular vesicles, termed epididymosomes (see figure 33). At the bases of principal cells, there are tight adhering junctions between adjacent cells. These separate the haploid environment of the ductus from the diploid surrounding tissue; an extension of the blood-testis barrier.

Principal cells are by far the predominant cell type but figure 30b shows that halo cells, apical cells and basal cells also populate the epididymis. In the stallion, the author was unable demonstrate narrow cells such as those seen in rats and other species.

Under the effect of "cross-talk" between principal cells, basal cells and clear cells, bicarbonate absorption and hydrogen ion secretion occurs in the epididymis so that pH drops. This acidic environment prevents calcium channel activation that would otherwise trigger capacitation. It also maintains sperm quiescence. The fluid of the accessory glands are high in bicarbonate, increasing pH and reversing the quiescent state of sperm during ejaculation.

Halo cells are clearly visible in figure 31, from the corpus epididymis of a peripuberal colt. Although it is not possible to discern the difference between types of halo cells with ordinary histology, they have been identified as T lymphocytes, helper T lymphocytes and monocytes; all parts of the immune system in the epididymis.

Figure 31. Halo, principal and basal cells in the corpus epididymis of a peripuberal colt. Size available: 2375 x 2116px.

Apical cells secrete protons, maintaining a low luminal pH but the function of basal cells remains controversial. They are thought to be part of the blood-epididymis barrier and assist in cross-talk between principal cells. At best however, their function is poorly understood.

Principal cells in the caput epididymis (figure 32) are packed with vesicular secretory bodies containing protein. After breaking off into the lumen they will will form the epididymosomes seen in figure 33. Contrast has been enhanced in one region of the image to enhance the visibility of the secretory bodies. Long villi seen in most areas of the caput are absent in others.

Figure 32. Caput epididymis from a 6 yr old Morgan stallion. Note the principal cells packed with vesicular secretory bodies. Size available: 2562 x 1520px.

Figure 32. Caput epididymis from a 6 yr old Morgan stallion. Note the epididymosomes. Size available: 1798 x 1052px.

Clear cells (see figure 34) like apical cells, are responsible for pH balance and for phagocytosis of discarded midpiece droplets from sperm. To the author's eye however, there appeared to be relatively few clear cells in the epididymis, seeming too few to assist significantly with the task of midpiece droplet phagocytosis. Interestingly, phagocytosis of midpiece droplets is mainly attributed to principal cells, not clear cells. Yet even in clear cells, where millions of midpiece droplets are phagocytosed every day, phagosomes were not obvious. Colleagues may wish to comment on this puzzling finding.

Figure 34. Halo, basal and clear cells in the cauda epididymis of a 6 yr old Morgan stallion. Size  available: 1717 x 1053px.

Although phagocytosis of spermatozoa themselves has been documented in various sections of the epididymis in more than one species, epididymal phagocytosis remains a contentious subject. Indeed, some authors have not been able to document phagocytosis of spermatozoa at all. Certainly, phagocytosis of defective sperm is not the common system for eradication of defective sperm that it was once thought to be.

The ductus deferens

The ductus deferens shows many of the characteristics of the cauda epididymis with two remarkable differences; one is its relative absence of storage space and two, the massive development of smooth muscle required for ejaculation. Figure 35 shows a comparison between the smooth muscle layers of the cauda epididymis and the ductus.

Figure 35. Cross section of the adjacent cauda epididymis and ductus deferens of a 2 yr old Standardbred stallion. Note the massive development of smooth muscle in the walls of the ductus deferens at right. Size available: 1498 x 798px.

The ductus is not simply a pump for ejaculation, but a storage organ as well. It is clear that millions of sperm lie within the folds of the epithelium seen in figures 35 and 36. In fact, close to 10% of all extra-gonadal sperm reserves are found in the ductus and ampullae.

Although the epithelium in the ductus is more folded than the epithelium in the cauda, the epithelium of the ductus is similar to that in the epididymis, with principle and parabasal cells predominating. Villi are also present in some areas but not others. This suggests that metabolic activity still occurs within the ductus. This activity is probably related to sperm quiescence rather than maturation.

Figure 36. Ductus deferens from a 2 yr old Standardbred stallion with principle and parabasal cells predominating. Size  available: 2284 x 1295px.

Selected references:

Amann, R. P. et al. 1977 Connection between the seminiferous tubules and the efferent ducts in the stallion. Am.J. Vet. Res.38: 1571-1579

Amann, R.P. et al. 1979. Effects of age and frequency of ejaculation on sperm production and extragonadal sperm reserves in stallions.
J. Reprod. Fert. Supplement 27:1-6

Arrotéia, K.F. et al. The Epididymis: embryology, structure, function and  its role in fertilization and infertility. Embryology – Updates and Highlights on Classic Topics. www.intechopen.com. Edited by Luis Antonio Violin Pereira, ISBN 978-953-51-0465-0. Open access.

Bedford, J.M. 2015. The epididymis re-visited: a personal view. Asian J. Andrology. 17:693–698

Cardoso, F.M. and Godinho, H.P. 1982. Cycle of the seminiferous epithelium and its duration in the ebu, Bos indicus. Anim. Reprod. Sci. 5:231-245

Clemmons, A.J. et al 1995 Local initiation of spermatogenesis in the horse. Biol Reprod. 52:1258-1267

Clermont. Y. 1972. Kinetics of spermatogenesis in mammals: seminiferous epithelium cycle and spermatogonial renewal. Pysiological reviews. 52:198-236

Cooper, T.G. et al. 2002. Rebuttal of a role for the epididymis in sperm quality control by phagocytosis of defective sperm. J. Cell Science 115: 5-7

de Souza, A.P.B. 2017. The epididymis and its role on sperm quality and male fertility  Anim. Reprod. 14 :1234-1244.

Durairajanayagam, D. et al 2015. Sperm Biology from Production to Ejaculation in Schattman, Glenn L. et al. (Eds.) 2015. Unexplained Infertility, Springer. Chapter 5 pages 29-42, ISBN 978-1-4939-2140-9

Ehmcke, J. et al. 2005. Identification of the starting point for spermatogenesis and characterization of the testicular stem cell in adult male rhesus monkeys
Human Reproduction 20:1185-1193

Gregory, M. and Cyr, D.G. 2014. The blood-epididymis barrier and inflammation. Spermatogenesis. 4:e979619

Griswold, M.D. 2016. Spermatogenesis: The commitment to meiosis Physiol Rev. 96: 1–17.

Haga, H.A. et al 2006. Effect of intratesticular injection of lidocaine on cardiovascular responses to castration in isoflurane-anesthetized stallions. Am J Vet Res. 67:403-8.

Hai, Y. et al. 2014. The roles and regulation of Sertoli cells in fate determinations of spermatogonial stem cells and spermatogenesis. Seminars in cell and developmental biology. 29:66-75

Hedger, M.P. 1997 Testicular leukocytes: what are they doing? Reviews of Reproduction. 2:38–47

Johnson, L et al. 1978. Scanning electron microscopy of the epithelium and spermatozoa in the equine excurrent duct system. Am. J. Vet. Res. 39:1428-1434

Johnson, L. and Neaves, W.B. 1981. Age-related changes in the Leydig cell population, seminiferous tubules, and sperm production in stallions. Biol. Reprod. 24:703-712

Johnson, L. and Nguyen, H.B. 1986 Annual cycle of the Sertoli cell population in adult stallions. J. Reprod.Fert. 76:311-316

Johnson, L. and Tatum, M.E. 1989.Temporal appearance of seasonal changes in numbers of Sertoli Cells, Leydig Cells, and germ cells in stallions, Biol. Reprod. 40:994-999.

Johnson, L. et al. 1990 Staging equine seminiferous tubules by Nomarski optics in unstained histologic sections and in tubules mounted in toto to reveal the spermatogenic wave.
The Anatomical Record Part B: Anat. Record. 227:167-174

Johnson, L. 1991.Seasonal Differences in Equine Spermatocytogenesis. Biol. Reprod. 44: 284-291.

Johnson, L. 1997 Efficiency of spermatogenesis in humans and animals. College Station. Texas A & M University, Texas.

Kayalioglu, G. et al. 2008 Morphology and innervation of the human cremaster muscle in relation to its function. Anat. Rec. 291: 790-796.

Laserre, A. et al. 2001. Human epididymal proteins and sperm function during fertilization: un update. Biol. Res. v.34 n.3-4 Santiago Biological Research versión impresa ISSN 0716-9760.

McKinnon A.O. et al. (Eds) 2011 Equine Reproduction. Chapters 95 through 97 and 101. Second edition. Wiley-Blackwell. ISBN: 978-0-8138-1971-6

Naito, M.1. et al. 2008 The presence of intra-tubular lymphocytes in normal testis of the mouse. Okajimas Folia Anat Jpn. 85: 91-96.

Nicander, L. and Glover, T.D. 1973. Regional histology and fine structure of the epididymal duct in the golden hamster (Mesocricetus auratus) J. Anat. 114: 347-364

Nakanishi, Y. and Shiratsucho, A. 2004. Phagocytic removal of apoptotic spermatogenic cells by Sertoli Cells: Mechanisms and consequences. Biol. Pharm. Bull. 27: 13—16

Nieschlag, E. et al. Eds 2010 Andrology: Male Reproductive Health and Dysfunction. Springer Science & Business Media. ISBN: 9783540783558

O'Donnell et al. 2011 Spermiation. The process of sperm release. Spermatogenesis.1:14–35.

Online course in embryology for medicine students developed by the universities of Fribourg, Lausanne and Bern (Switzerland). Swiss Virtual Campus. Spermiogeic wave: http://www.embryology.ch/anglais/cgametogen/spermato04.html

Payal, M. et al. 2011. The blood-testis and blood-epididymis barriers are more than just their tight junctions. Biol reprod. 84: 851-858

Perez. C.V. 2013. Dual role of immune cells in the testis. Protective or pathogenic for germ cells? Spermatogenesis. 2013 Jan 1; 3(1): e23870.

Pozor, M.A. 2007. Evaluation of Testicular Vasculature in Stallions. Clin Tech Equine Pract 6:271-277

Phillips, B.T. et al. 2010. Spermatogonial stem cell regulation and spermatogenesis Trans. R. Soc. B. 365:1663–1678

Ramos-Ibeas et al. 2013. Most regions of mouse epididymis are able to phagocytose immature germ cells. Reproduction. 146:481-489

Shum, W.W.C. et al. 2009.Regulation of luminal acidification in the male reproductive tract via cell–cell crosstalk. J. Exp. Biology. 212:1753-1761

Silber, S. 2010. Sperm retrieval for azoospermia and intracytoplasmic sperm injection success rates – A personal overview. Human Fertility. 13:247–256

Soares, J et al. 2009 The seminiferous epithelium cycle and its duration in different breeds of dog (Canis familiaris). J Anat. 215: 462–471

Sutovsky, P. et al. 2001. A putative, ubiquitin-dependent mechanism for the recognition and elimination of defective spermatozoa in the mammalian epididymis. J.Cell Sci. 114:1665-1675

Swierstra, E.E. et al. 1974. Reproductive physiology of the stallion. 1. Spermatogenesis and testis composition. J.Reprod. Fert. 40:113-123

Svingen, T. and Koopman, P. 2018. Building the mammalian testis: origins, differentiation, and assembly of the component cell populations genesdev.cshlp.org. Cold Spring Harbor Laboratory Press

Toyama, Y. 1977 Actin-like Filaments in the myoid cell of the testis. Cell Tiss. Res. 177:221-226

Wistuba, J. et al 2003. Organization of seminiferous epithelium in primates: relationship to spermatogenic efficiency, phylogeny, and mating system. Biol Reprod. 69:582-91.

Zhou. W. et al 2018. Review article Characteristics of the Epididymal Luminal Environment Responsible for Sperm Maturation and Storage. Front. Endocrinol. https://doi.org/10.3389/fendo.2018.00059