A TECHNIQUE FOR ARTIFICIAL INSEMINATION IN SQUAMATES

Raymond Hoser (Snakebusters)

488 Park Road

Park Orchards, Victoria, 3114, Australia.

"Hard copy” published in January 2008 in the Bulletin of the Chicago Herpetological Society 43(1):1-9.

Abstract

This paper details the development and application of a simple to use method of artificial insemination in squamates of various taxa.

The benefits are obvious.

This includes that of breeding specimens not inclined to mate and the ability to transport semen, rather than reptiles across suburbs, cities, or even states and countries. In the Australian context this is significant as there are “six month rules” in most states, making breeding loans of actual specimens sometimes logistically difficult.

While most of our work has been with snakes, we have trialled the same techniques with lizards and found that what is reported here for snakes transposes to lizards as well.

In the USA context, the methods make hybrids between taxa simple to achieve, as it no longer requires the reptiles to mate.

Introduction (The problem)

All squamates bred at our facility (elapids, pythons and large skinks), are housed similarly and subjected to the same annual temperature regimes. This is spelt out in Hoser (2006) and Hoser (2007).

In summary it involves 7 weeks of cooling whereby the cage temperature is kept below 20 deg C at all times (with rare rises above this perhaps if the reptile is used in a “show” or “demonstration”). Temperatures do still have a slight diurnal cycle of up and down.

This is followed by a period of 12 hours a day “full heat” and 12 hours of “night”, after which the reptiles get 18 hours a day heat or all heat, in terms of the warmest section of the cage.

For us “cages” are in fact large plastic tubs.

The exact timing of the “winter” months at our facility broadly mirrors that of the wild, but our preference is to run the seasons about 8 weeks (2 months) ahead of wild counterparts here in Melbourne, Victoria, the end result being our snakes breed earlier.

For example in late 2007, our first breeding female (venomoid) Eastern Brown Snake (Pseudonaja textilis) laid a full clutch of 8 fertile eggs at end October, whereas the wild counterparts usually lay at end December. Two years earlier the same snake produced 10 eggs (9 fertile)(see Hoser 2006).

In spring of 2006 (broadly referred to here as July-November here in Australia), it was attempted to breed Jaffa (Collett’s) Snakes (Panacedechis colletti) at our facility. The snakes were two males aged 3 years and a four year old female.

Unless mating is being attempted, we usually house all our reptiles as “one per cage” as there are numerous husbandry advantages. When shipping our reptiles for demonstrations, often day after day for many weeks, reptiles are grouped with the only limiting parameter being that the snakes be of similar size, with taxa being irrelevant.

It is common for them to mate or attempt to mate when in transit, including snakes of different taxa (Hoser 2005b).

The males Collett’s Snakes had been held since hatching at the facility of Paul Fisher (Hoppers Crossing, Victoria) and the female raised for two years by Scott Eipper and then held by myself for about two years, being hatched by another breeder.

While one of the males attempted to mount and mate the female in late 2006, no actual copulation occurred. At the time it was thought simply that the male wasn’t trying hard enough, and perhaps the male may improve with age, as is often the case with snakes.

The following spring (2007), it was attempted to breed the snakes again and there was no success in mating. This time both males tried hard and yet the female successfully avoided copulation. She would flee as soon as the males attempted to mount her.

The cooling regime the previous “winter” had been particularly “brutal” in that the reptiles were kept colder for longer (8 weeks under 20 Deg C) and this reflected across the board in particularly vigorous mating activity across all taxa.

It is mentioned here that all the Collett’s had been made venomoid in late 2004, using the successful method detailed earlier that year by Hoser (2004). In fact all Hoser elapids referred to in this paper are as of 2007, long-term venomoids.

“Venomoid” means permanent surgical removal of venom glands from a snake to render it non-venomous (effectively) harmless to humans for the rest of it’s life.

Venomoids by definition are neither venomous or dangerous to humans and claims to the contrary are false.

Venomoids have the welfare advantage of not needing to be stick handled, pinned, tailed, or necked, (due to the removal of adverse safety issues), the result being better adjusted snakes that as a rule lose any urge to attack or bite human handlers, which is the complete opposite of what is often otherwise seen in venomous snakes.

For the purposes of this paper, the venomoid state is known to have no effect on fertility as long-term venomoids have been bred at our facility (producing normal healthy (and venomous) young), including Death Adders (Acanthophis antarcticus), Eastern Brown Snakes (Pseudonaja textilis), Tiger Snakes (Notechis scutatus), and Copperheads (Austrelaps superbus), which is all elapid taxa we held in the relevant period (2004-7) for which we had adult pairs, excluding our Red-bellied Black Snakes (Pseudechis porphyriacus).

This includes across several seasons involving the same snakes.

Mentioned here is that our Red-bellied Black Snakes (2 males and a female) mated and produced “slugs” in 2005/6, and hence by definition hadn’t bred, but at the time of writing this paper the female was noticeably gravid again (end 2007) and was expected to produce young either late in 2007 or early 2008 (also see later this paper).

In my experience, normally fertile female snakes that are either ovulating or about to, are happy to be mounted and mated and yet this above-mentioned female Collett’s was violently opposed to the idea.

It was as a result of this apparent exhaustion of ideas or means to encourage mating that I decided to “step outside the square” and attempt artificial insemination.

Materials and Methods

The theory was simple.

Get semen from the male and put it into the female.

After that, the spermatozoa should do the rest!

Analysis of AI methods used for other vertebrates such as cattle, dogs, sheep, humans and even birds, yielded two main methods to acquire semen.

One was “electro-ejaculation”, where by an electric shock causes ejaculation. For several reasons, the idea was thought not viable in terms of the snakes.

Getting hold of an “electroejaculation machine” was near impossible or cost prohibitive and then there was the problem of working out the voltage required to get semen but not kill the reptile by electrocution.

That is assuming the process was even possible for reptiles!

The alternative was to masturbate the snakes to get semen.

This was the preferred method of choice for most animals including bulls and horses, who are generally made to mate with a false vagina.

However the concept of masturbating snakes was unknown territory and one that I could get no guidance from by any veterinarians or others I thought likely to know.

Hence the idea of masturbating snakes to ejaculate semen was one that I had to develop from scratch.

In hindsight it was remarkably simple.

Over the last 40 years of keeping, breeding and observing snakes, observations of male snakes and mating snakes yielded certain pointers.

Most male snakes shed so called “semen plugs” which are essentially globules of dried and old semen that accumulate in the hemipenal pockets. Hence I knew that snakes oozed or released semen at times other than copulation.

This concept went further when it was realized that sometimes male snakes mount females, attempt to mate without success and then ejaculate semen over a female. Alternatively and worst case, would be a male snake mating that breaks off the copulation and has semen on the hemipenes.

This occurs naturally and if one looks at the picture of the everted hemipene in a male Death Adder on page 19 of Hoser (1989) you will see exactly this.

Knowing all the above pointers, it was thought that it may be possible to stimulate a known fertile male snake that has mounted a female to get excited and ejaculate semen in a quantity sufficient to be collected and transferred to a female snake.

Based on observations of the quantity and consistency of semen shed in semen plugs and observed when snakes ejaculated, it was decided to use a glass pipette or capillary tube to collect the semen and transfer to the female snake.

These come in various sizes and shapes and as it happens the first one trialed was the best. This was a “Kimax-51 1.5-18 X 100 mm straight glass capillary tube” that is sold commercially in 100 tube lots packed in small glass containers in boxes of various numbers. These are also the tubes of choice for aviculturists who use AI.

They suggested these as they are “non-Heparin coated” as Heparin is believed to be adverse to semen.

The pipettes were in the first instance hard to acquire as neither veterinarians, GP’s, hospitals or pathology labs routinely stock them. However science departments of most schools use these or similar and hence small numbers were readily available, including in various sizes and formats in order to confirm the best tube for the job, that being the one just named.

The first successful masturbation of a snake was when a male Eastern Brown Snake was seen mounting a female (who was already gravid from him).

The caudal region was stroked with my finger and the snake was visibly aroused and attempted to copulate with it.

As I rubbed the snake (just above the cloacal opening on the side of the base of the tail), the snake became stimulated and after loss than a minute he ejaculated semen.

This was gathered by sucking into the pipette and then checked on a microscope slide at 400X magnification and the sperm cells were readily visible.

Similar was attempted with the first male Collett’s Snake to attempt to mate the female and again semen was collectable from both hemipenes within 60 seconds.

The second male, also interested in mating the male, but not yet mounted her, was also used for semen collection.

In it’s case, the snake simply had it’s tail grabbed by myself as the snake sat in the cage. The relevant region was rubbed and again semen was yielded. This was the first snake actually masturbated without actually mating a female.

A Black-headed Python was seen trying to mate an Olive Python and it was removed from the transport box. The tail was rubbed (at the same place as for the elapids) and the snake yielded globules of semen within seconds.

In other words it was possible to masturbate a snake and get semen.

Taking the process further, two male Tiger Snakes held in cages on their own and simply resting were each removed and easily masturbated to yield semen.

With practice, acquiring semen from snakes (by rubbing the anterior caudal region above the vent) was easy, and for snakes that were apparently fertile, semen was now readily available.

In terms of the AI process, the theoretically hardest part of the operation was now complete.

In fact by the method devised, getting semen was now simple and routine and could be done with an un-aroused snake simply resting on it’s own in a cage.

Please note that in terms of the venomous taxa used, all were well-adjusted venomoid (no venom in the snakes) that are handled (by free handling only) for live shows on a daily basis.

Hence for these snakes, they have no handling stress or fear of human interaction with them.

No masturbated reptiles of any taxa ever attempted to flee or bite.

Masturbation of non-venomoid dangerously venomous elapids is not something that should be attempted, unless the handler is both experienced with the snake species, the snake itself is relatively tame and the handler is happy to countenance the possibility of a potentially fatal bite.

For the record, masturbating the snakes did not yield any signs of stress on the snakes.

The only obvious variables in behavior noted were the obvious movements downwards of the pelvic (or equivalent) region of the snake as they were stimulated, and a greatly increased frequency of tongue flicker as the snakes were aroused.

Transfer of semen to the female was via the pipette.

Acquiring semen

It seems that not all snakes produce semen all the time.

In our collection it became apparent that the snakes that mated most were those that produced most semen. The correlation was direct.

In terms of the Tiger Snakes, for which we held 6 adult males as of end 2007, the ones who mated the most all yielded copious amounts of semen readily, while I was unable to get semen from some who rarely showed interest in sex.

Please note that at the time this semen collection was done, the snakes were being held on their own, as we were trying to avoid breeding this taxon as we have trouble offloading the babies and hence for 2006/7 only one of four females was actually mated (with their mating season also including summer of 2005/6, due to our concern that these snakes store sperm for some time).

The same pattern was yielded across other taxa of snakes (elapids and pythons) as well as skinks, from whom we were able to get semen using the same method.

It should be noted however that lizards are far harder to stimulate than snakes due to their increased tail muscularity and other tissue present in the region that apparently makes direct hemipenal stimulation harder. (Please note the extreme care needed with tail shedding taxa to avoid any incidents of autotomy).

Notwithstanding this, it was possible to extract semen from all lizard taxa we hold, which includes Cunningham’s Skinks (Egernia cunninghami), Blotched Bluetongues (Tiliqua nigrolutea), Eastern Bluetongues (Tiliqua scincoides) and Shinglebacks (Trachydosaurus rugosus). Other lizards were able to be masturbated at other facilities to yield semen to be used for insemination including smaller skink species, Lace Monitors (Varanus varius), Gould’s Monitors (Varanus gouldi), Bearded Dragons (P. barbata and P. vitticeps), Gippsland Water Dragons and Robust Velvet Geckos (Oedura robusta).

Assuming the reptile (snake or lizard) is tractable, we found the best method to acquire semen was to simply hold the reptile in a way that is comfortable for it and to rub the hemipenal area (near the vent) with one finger with moderate speed.

You will know the reptile is stimulated as it pushes that region downwards, to give the angled position of vent region as seen as a snake attempts to copulate a female in the “natural” way. Most snake keepers are familiar with this positioning.

While the male may evert a hemipene if stimulated, as a rule this does not occur, and it is not necessary for semen extraction. For reasons not completely certain, snakes (and lizards) will ejaculate semen while the hemipenes remain retracted in the tail.

Hemipenal plugs and dried feces may be shed and this should be disregarded (discarded).

On some occasions, dried fecal matter may be around the cloacal region and this should be cleaned away with a wet cloth before masturbating the snake so as to ensure a “clean” semen sample is obtained.

As a rule, if the snake has semen, it should yield it within 60 seconds. Cooler snakes take longer to yield semen than warmer ones. The same applies for lizards, albeit on a slower timeline, although smaller taxa yield semen faster.

As a rule there is no need to check semen under a microscope for viability.

If the snakes or lizards have been cooled over winter according to the regime we use (see above), viable sperm seems to be a formality for almost all taxa, save for the inevitable small percentage that will never be fertile.

As part of the perfectionist system here, semen was checked under a microscope and images sent to Dr Barrymore Walters, an expert whose day job involves human vasectomy’s and microscopic inspection of semen samples.

While snake semen is different to that of human, he seemed to think what I sent him was OK and his judgment later proved correct.

Semen from the semen plugs in snakes was checked and found to be clumped, which is typical of dead and non-viable semen, indicating that snake semen does have a limited “shelf life” although it is hard to ascertain what that is.

As a rule, if a snake yields semen from one hemipenal pocket, it will yield from both and I found that the best way to collect semen was to masturbate both sides so the semen sat either just inside or just outside the ventral scale, form where it can easily be sucked into a capillary tube. As a rule, one tube is used for each side, enabling two lots to be gathered at a time.

Often the hemipenal region is massaged to assist in bringing the semen towards the vent for collection.

As a rule, it takes 5-7 days for a snake that has yielded semen to be able to regenerate semen again.

As a rule, snakes do not yield semen in smaller amounts when an attempt is made to extract semen in a period under the 5-7 regeneration period. Instead the snake yields nothing.

In other words semen seems to be yielded in distinct “loads”.

The semen that is viable and used is not the hardened material seen in dried hemipenal semen plugs.

Instead it is the milky white material that is obviously yielded at the time of masturbation and as an obvious result of it. To give an idea of the quantity, it is best to view a photo.

Sometimes masturbation of a snake will yield a semen plug followed by good whitish semen. In this situation the latter (whitish) material, should be used only.

Interestingly high sex drive snakes will still attempt to mount and mate females after being taxed of semen.

This is interesting because as a rule, once I have extracted a single load of semen from each hemipene, I am unable to repeat the process until the 5-7 recharge period has elapsed.

An important question to ask, is whether or not a snake that mates immediately after I’ve taxed it for semen is still able to pass viable semen (sperm) to a female at that time.

Refining the method of semen transfer

Aviculturists who do AI with birds said that they’d simply suck semen into the pipette and then place it into the ventral opening of the bird and blow out the semen inside the bird. They said in most cases, the spermatozoa did the rest and conception was the rule.

The same was done with the snakes, and we now know the same to be true for them as well.

However sometimes the semen was too viscous to be able to be blown out of the tube with success. I then found myself trying to blow out the semen without success and had to remove the tube from the uninseminated snake.

Narrower tubes were even harder to utilize than the originals and for the wider ones other issues arose, mainly in terms of sucking up the semen and then being able to blow it out, as opposed to just air. You see unless the semen blocks the tube, it will give a pathway for air to simply blow past it.

Due to the nature of the human mouth, it was far easier to suck fluid into the tube than blow it out. Hence the occasional difficulty of blowing the semen out of the tube into the female snake.

So the logistical problem had become how to quickly and effectively get the semen into the female snake.

The superior method developed was to get the male snake to the stage of yielding semen. At that point the capillary tube is placed in water to a point where a small amount of water is sucked in. This sometimes occurs automatically and other times you may need to do this by careful sucking.

Water is less viscous than semen and also known to be harmless to it.

Semen is then sucked into the tube and then a small amount of extra water, making the semen effectively inside the tube padded by water on either side.

By carefully sucking either end of the tube, you will be able to move the semen back and forth in the tube.

Once you get to the stage where the semen is easily moved, you should attempt the same by blowing.

Once you get to the stage where you can move the semen up and down by blowing the tube, and with relative ease, you are ready to inseminate the female.

The cloacal opening is opened sufficiently to allow the tube to pass through.

Usually this is simple, but if it is dry and tight, lubrication with water will solve the problem.

In summary the tube is inserted to a depth of just under about 3.8 cm in a 151 cm long snake and the tube blown to leave the semen in the female.

You will know that the semen is inside the snake when you notice the air back-up (from you) going into the snake, at which point the tube is removed and the vent held shut.

Assuming this is done properly, the semen will remain in the snake and make it’s way to the appropriate part of the female to fertilize ova or eggs.

In terms of the insemination of the female, there are other important pointers and notes. The tube will hit an apparent (soft tissue) “block” when pushing in an anterior direction. To give an indication as to the approximate depth of the “block” it should be about 3.8 cm into the snake if it’s a 151 cm total length snake (18 cm tail and 133 cm snout vent). This should not be pushed or pressured and the tube pulled back a few mm from this “block” point.

These measurements as given here are important as they can be scaled pro-rata up or down for larger or smaller snakes to give an indication as to likely and expected penetration depths.

If you find a “block” a substantially earlier than the indicated distance, then it will be caused by fecal material ready to be expelled. This should be removed before attempting insemination (see elsewhere in this paper for an explanation as to how this is done) in a manner that is simple and painless for the snake.

Assuming no fecal material in the relevant part of the snake then the insemination of the snake should be routine and trouble free, and success for the procedure assumed likely.

If there is trouble getting the semen to be blown into the female, it is often easier to blow and withdraw the tube at the same time. The backward movement creates a gap (void), which then creates a vacuum to suck out the semen sample.

When doing this, you may accidentally release semen either at the vent opening or even outside the snake.

If this happens, the semen (which usually presents as a sort of line of fluidy gunk), can usually be sucked back into the tube and the whole insemination process repeated.

Another mishap that occurs occasionally is that you may suck in semen to your mouth and then spit it out. Often this can be reused as well.

While none of this is sterile and there is an obvious germ transmission, no snakes inseminated this way have ever shown signs of illness and noting that reptiles cloacal openings are exposed to these germs in the normal course of crawling over things, this is not seen as an issue worth worrying about.

If the semen sample is degraded or lost before being implanted into the female, then the second one obtained (usual), can be used.

Alternatively if all runs to the theory and plan, it becomes possible to inseminate two snakes from one snake on one day.

The same as just described applies to lizards.

Measuring and maximizing success

All the above also assumes a pre-winter, winter, spring heating and cooling regime sufficiently similar to ours so as to get both male and female fertility cycles synchronized and viable.

How long until fertilization takes place after copulation or insemination is hard to ascertain and depends on variables such as “is the snake ready to breed?”, potential sperm storage and ovulation cycles.

However due to the fact my own breeding records have instances of snakes mating one day and conception being measurable effectively from that date (no measurable delay) and with no measurable differences in the development of young or eggs when born (excluding incubation-based temperature variables for eggs), it is reasonable to assume that active spermatozoa will travel to the correct parts of the snake well within 24 hours.

The significance of this known fact is when attempting to ascertain the likelihood of the insemination of the female snake being successful in fertilizing eggs/ova.

If the snake defecates within 24 hours of insemination, then semen may be released before it has time to work effectively.

As a rule of thumb, if this occurs, it’d be logical to repeat the insemination again, as soon as male semen becomes available (about a week if you only have one viable male).

To that end, it is possible to determine if a female snake is likely to defecate.

Females can be palpitated and an impending defecation determined in terms of likelihood.

While the advice may be to delay insemination until after defecation, there are other means to deal with this common potential problem.

In the first instance semen should only be collected if the female is deemed ‘clean’, feces free and ready in every other way. If the female is likely to defecate, a delay may be in order.

Having said this, in our situation the better method involved palpitating the female for feces.

If there was deemed a likelihood of defecation within a few days, the snake would be placed in luke warm (25-30 Deg C water) in a sealed container (with air holes only) to a depth sufficient to immerse the snake, but not too deep to drown it.

Usually such an environment will encourage any fecal material to be released within a few hours of soaking.

After this time the snake is placed back in it’s dry cage and after the snake and the relevant part of the body has dried out, the insemination is done.

By way of example, this regime was practiced with success when (successfully) inseminating a Bredl’s Python, that was made to defecate before it was inseminated.

The same also applies to lizards, noting that for them it is harder to ascertain major defecations.

It has been suggested that soaking of snakes and lizards to encourage defecation prior to insemination is advantageous in terms of maximizing success and if there is not a ready supply of semen to use repeatedly (as in you only have one chance of successful insemination), then soaking to ensure minimal risk of fecal disruption is the best course of action.

Just as snakes have no issue with sexing by “probing” if done properly and with care, the same applies with insemination of female snakes if done with due care.

Due to the variable of minor defecations, that may also inadvertently release semen, the advice for dealing with inseminated females is as follows.

The cage it is kept in for the day or two following insemination should be small, totally clean and one in which any new fecal material can be seen.

If any defecation is seen within 48 hours of insemination, then my advice is to re-attempt it on the basis that the first attempt may be a failure.

For our set-up all snakes and lizards are kept in what are usually very clean plastic tubs and because post insemination we ensure no fecal material is in the cages/tubs, any defecations post insemination are easily seen.

Storing semen and related issues

For mammals and other taxa, semen is often stored frozen and often for long periods.

Because our method involved immediate or near immediate insemination, storage has been a non-issue.

Sitting in a room at room temperature, globules of semen will dry up quickly, being noticeably drier and harder to deal with within minutes.

For this reason, masturbation of snake and subsequent insemination should be as quick a process as possible and as a rule can be executed within 60 seconds from extraction to insemination, assuming everything is at the same venue.

Semen in capillary tubes, takes a lot longer to dry out, due to the relatively small amount of contact between air and semen. Semen held at room temperature in a capillary tube, padded with water at either end will last for hours and apparently not degrade if stored in a sealed box, itself lined with moist tissue.

This is known, because semen stored this way for several hours has been checked under a microscope and found to be “normal”, enabling successful inseminations to have been done in various collections across our home state of Victoria, without the need to move reptiles.

This method has already enabled successful inseminations to be done in numerous reptiles in various collections involving reptiles that would otherwise either never have copulated, or alternatively never have even been in physical contact.

Improperly sexed reptiles and the need for routine probing of reptiles

An issue that has reared it’s head several times recently has been incorrectly sexed reptiles.

At our facility this has never been an issue. All squamates are probed and this method (if done properly) remains the easiest and simplest method to 100% reliably sex them.

Hence in terms of breeding, insemination and the like, we’ve always been able to go the males and tax them for semen, or impregnate known females.

Recently we supplied semen to other keepers and have struck some interesting obstacles.

Semen from a venomoid male Inland Taipan (Oxyuranus microlepidotus) was useless when it became clear that the “female” that had supposedly been probed as such was an obvious male. I saw the 2 metre snake with large tail and went through the motions of probing it as male.

In another incident, I used semen from one of my venomoid male Collett’s snake to inseminate a long-term captive “female” that had apparently eaten a smaller “male” some years earlier.

As for the Inland Taipan incident, the person was a long-term herpetologist of high repute, whom I had no reason to doubt.

I took the semen from the male and implanted it into the “female”. This part was apparently routine. It was only after placing the “female” back in the cage that I thought the tail of the “female” was too large. The snake was retrieved, probed and turned out to be a male!

When inseminating the (now known to be male) Collett’s snake, it was noticed that the pipette didn’t hit the same “block” as seen in all the female snakes. In other words it could be passed much further into the snake.

Hence it emerged that sexing errors in males can actually be diagnosed at the insemination stage as a second-best alternative to probing.

As a result of this incident, measurements were taken on my three Collett’s Snakes to give accurate indication as to where “blocks” seemed to occur when a pipette was inserted through the vent.

These were as follows:

Female: 151 cm total length, 18 cm tail, 133 snout-vent, 3.8 cm depth of pipette to “block” point.

Male (1): 169 cm total length, 20 cm tail, 149 cm total length, 5 cm depth of pipette to “block” point.

Male (2): 152 cm total length, 18.5 cm tail, 133.5 cm snout-vent, 5 cm depth of pipette to “block” point.

Similar data came from taxa such as Red-bellied Black Snakes and Tiger Snakes, hence giving a secondary means for sexing snakes and raising potential indicators of sexing errors if the pipette passes further than expected before reaching a “block”.

Mention of these sexing errors is done here as it serves as a warning to budding breeders who are not totally certain of the sexes of their reptiles.

It may also be medically significant to the reptile if a sexing mistake is made.

Probing snakes

Most herpetologists are familiar with probing of snakes as described by Hoser (1989).

The probes are usually devices of varying sizes, that are metal rods with a ball at the end. The sizes vary and the one used is that which is of appropriate size that can fit comfortably into the hemipenal pocket, with a ball at the end of sufficient size so as not to “spike” the end of the hemipene if it actually travels that far.

In terms of probing a snake, the operation is delicate and should be done with the utmost care and precision as it is easy to injure the soft tissue of the hemipenes, or corresponding tissue in females.

As a rule, the probe should not be inserted the full depth of the hemipenal pocket as it is rarely needed to accurately sex the squamate. I have seen many snakes probed by novices that have sustained injuries (sometimes eventually fatal) from probes going through one or other hemipene.

When probing, once the probe has been inserted to a point where it is clear the depth indicates the sex as male (say about 7 scales down for most snakes), there is no need to push the probe further.

Due to the fleshier tail in lizards, probing is often more difficult, due mainly to difficulty in finding the position of hemipenal pockets, but the underlying principals are the same as for snakes.

It is perhaps unfortunate the probes are routinely sold to persons without questions or training and who have no sensible training or experience in their use. This no doubt leads to many squamates being improperly probed by novices and potentially fatal injuries arising (the usual cause of death being untreated infections from injuries to soft tissue).

Worse still are recent instances of so called “snakehandlers” claiming decades of experience, which they do not have, doing so-called “courses” and teaching people, who then go away thinking they know what to do, and who in fact have been taught wrong or dangerous methods.

In the last 4 decades, I have seen countless cases of this and in the age of internet this problem has worsened.

Hence it’s appropriate to issue a generic warning about probing.

IF INEXPERIENCED, DO NOT PROBE A REPTILE.

DO NOT ACCEPT TEACHING FROM A PERSON WHO DOES NOT HAVE INDEPENDENTLY VERIFIABLE EXPERTISE.

DO NOT BELIEVE A PERSON’S CLAIMS OF EXPERTISE UNLESS THEY ARE VERIFIABLE.

While in theory, there is no shortage of capable and willing persons who will safely probe reptiles, it may take a bit of time and effort to find a person to do this at an acceptable time.

Most veterinarians have no training in probing reptiles. Of the small percentage who can and do, some will charge a fee to do so, but others will do it for free.

While a “free” probing is likely if only a small number of reptiles are involved, the situation may change for collections of dozens of reptiles. Note that the probing act usually only takes a matter of seconds per reptile if the practitioner has experience.

A number of (known to be experienced) private keepers and breeders will do it. For them it must be at no cost, as in most jurisdictions, including Victoria it’s illegal for a non-veterinarian (not licensed as a veterinary surgeon) to charge any fee for any service that can be defined as veterinary in nature.

However the majority of capable persons are reluctant to take time out to probe for persons they either don’t know, or have any compelling reason to do unpaid work for.

In other words, there is no short-term fix for the ongoing issue of unsexed or improperly sexed (guessed?) reptiles in private care in Australia.

In my own situation, I generally carry a set of probes in the car and also with our reptiles when doing demonstrations and probe individual reptiles for persons if and when requested and if and when it does not interrupt my own busy schedule.

It does not however solve the widespread issue of unsexed or improperly sexed reptiles at facilities across Australia.

A naming issue

While this paper has labeled the methods used as “artificial insemination”, the only artificial stage of the process is in terms of the acquisition of semen and then it’s implanting in the female.

An alternative name for the process is therefore “assisted insemination”.

For humans and other animals, Artificial insemination or “AI” sometimes refers to the actual process of conception (sperm penetrating egg and fertilizing it) in a non-natural environment such as a petri dish, as opposed to human movement of semen.

Viewing semen under a microscope

In terms of the methods described in this paper, it isn’t usually necessary to view semen under a microscope.

As already mentioned, if the reptiles have been subjected to a correct temperature regime over the preceding year (in terms of inducing mating and breeding) and there are no contra-indications, then it’s reasonable to assume that the males carry viable semen and sperm.

If intending to view semen under a microscope the recommendation is to retrieve and view semen without any delay, so as to avoid drying of the sample.

A day old sample or slide presents a very different view to a fresh sample.

Of note has been the strong variation in shape and form of spermatozoa in given taxa. The appearance of spermatozoa also varies depending on the resolution of the microscope and the preparation of the slide.

As a rule, stained slides make the individual spermatozoa easier to see and identify.

At lower resolution (say around 100 X), the semen gives an appearance of being striated in texture. The striations are usually caused the tails of the individual spermatozoa. At higher resolution (say around 400 X), the individual spermatozoa are delineated, including the head and tail. However due to the nature of a microscope’s focusing, most spermatozoa will not be visible, with the resultant view being a combination of heads and tails, with few if any complete spermatozoa being visible.

Most standard optical microscopes have resolutions of 100 X and 400 X.

In terms of slide preparation, the general recommendation is to smear the subject material very thinly over the slide, so as to yield a “single layer”, before placing on the cover slip. Dilution in water or dye (methyl blue), one droplet is more than enough, may assist for several reasons, including delaying the inevitable drying of material under the slide.

As it happens, none of these methods are mandatory in terms of observing semen. Semen is actually easy to place on a microscope slide and also easy to view due it’s light coloured texture (whitish to translucent).

If the spermatozoa present as evenly distributed, it is reasonable to assume that the sperm is viable.

If they are clumped, they are likely to be unviable or dead. Such clumping is seen in slides from old and dried hemipenal plugs often shed by snakes routinely and also shed prior to the yielding of fresh semen at time of masturbation.

If desired, and if only one snake (or lizard) is to be inseminated, it is routinely possible to use one semen sample to inseminate the reptile, and if this works according to plan, then use the other for microscopic analysis. This may give a better indication of likely success of the insemination.

Photos taken of microscopic material was done using a Pangor Microscope T-mount, specifically designed to enable SLR cameras to take photos on a microscope. The mounts are relatively inexpensive (under $100) and available from specialist retailers and fit on all major makes of camera, including in our cases, Pentax 35 mm SLR’s and Nikon digital SLR’s. The mount I own only fits over microscopes with relatively narrow eyepieces.

If taking photos, a microscope with it’s own light source (as opposed to a mirror to catch light) yields better photos.

Advantages of AI over “natural conception” in the captive reptile situation.

Obviously in cases where snakes won’t otherwise copulate, there is no contest in terms of comparing the methods of conception.

In terms of cases where a pair of snakes may naturally mate, use of AI has questionable benefits and it is here that a judgment call needs to be made.

If the mating is deemed likely, AI is redundant. This happened with two out of three pairs of Eastern Brown Snakes. Two South Australian Eastern Brown Snakes (a male and female) failed to mate and nothing I tried seemed to be able to induce a mating. The female was therefore inseminated and became gravid with fertile eggs.

Unlike for previous years, in spring 2007, I was unable to get either of my male Red-bellied Black Snakes to mate, so chose to use AI on the female.

Some weeks after the AI, the three snakes were taken to the Tuggeranong Hyperdome Shopping Mall in the ACT.

While being held in a box together, the two males apparently attempted a “three-some” as they tried to mount the female and one another. One of the males was removed and within minutes the other male had “locked up” as in commenced copulating the female.

This remained the case from 12 noon through the next on stage show at 1 PM, where the snakes were held and demonstrated as mating to the audience of many hundreds, and through the afternoon and into the night, at which stage the two snakes remained in a box in a nearby motel room.

In that case and with the benefit of hindsight, my early call to do AI on the Red-bellied Black Snakes was probably unnecessary.

Because it is routine to be able to tax a male snake for semen and successfully impregnate the female within minutes, with no pain or suffering for any party, AI becomes a compelling alternative for natural conception methods for a sizeable proportion of captive reptiles.

At our facility, for the 2006/7 season, most (but not all) breeding’s will be from AI, and we have had to get a new incubator to deal with the expected rush of eggs.

In terms of masturbating snakes to extract semen, this method is also useful to determine the likelihood of whether or not a given male snake will be inclined to mate.

Consistent failure to get semen (assuming you know how to properly masturbate the snake) and assuming that the snake hasn’t recently mated or been taxed for semen, has been shown to be a reliable indicator that the snake won’t mate or produce offspring (at least in the short to medium term).

This is a useful technique for potential reptile breeders with large collections of given species, who may be considering which individual snakes (or lizards) to cull from the collection.

Benefits (and negatives) of the methods described above

At our facility it manifests as more gravid snakes than would otherwise be the case. Using the method described in this paper, over ten snakes of various taxa (elapids and pythons) as well as numerous lizards from all Australian families (excluding pygopodids) have been successfully inseminated and either produced fertile eggs or young, or are due to by early 2008. Most would not have bred otherwise.

For the record, masturbating pygopodids for semen was trialed on a single male Delma inornata with success, but the semen was discarded.

Artificially inseminated snakes … the new venomoids?

While the first successful AI was done with venomoid Collett’s and Brown Snakes and in it’s first wide application, venomoids of various taxa were the main players, the method is probably usable for all squamates.

Because it is so unbelievably simple and effective, there is no doubt that some people will have doubts about the methods described here.

The saying “it’s too good to be true” will be used.

“Why hasn’t this been done on a wide scale sooner?”

Fuel to this fire will be added by the usual band of critics who complain about anything “Hoser”(see Hoser 2005a).

It will be like the non-stop rants and false comments about the alleged death, destruction, pain and suffering of the Hoser venomoids, the originals of which are still thriving, mating and breeding four years after their venomoid operations!

Again see Hoser (2005a).

Done judiciously and properly and in circumstances where reptiles may otherwise not breed, AI as described here has no known or obvious downsides and yes, I’ve been looking hard for any.

Having said this, I have a band of critics, who habitually criticize anything “Hoser”, and these inevitable criticisms are discussed here briefly and dismissed as frivolous.

There will be false claims on the internet of mass-wipe-outs of reptiles for the purposes of alleged AI experiments.

Put simply there has been no wipe out.

No reptiles have died yet as a result!

There will be claims of danger to the reptiles with AI.

If you get a kitchen knife and slice off a snake’s hemipene’s it will probably die. However in the real world of AI as described here, this has not happened and cannot!

AI in reptiles as first described here is in many ways like the venomoiding of snakes as described first by Hoser (2004) and later papers. It simply has no downsides!

The negative claims against the Hoser venomoids by the usual critics have long been shown to be false. Just as venomoids don’t roll over and die from a lack of venom, or from the relatively painless operation, the same applies for artificially inseminated snakes. There is no mass mortality or downside.

However in contrast to the venomoid operations, where the snakes do not regenerate venom (contrary to false claims otherwise), the sole purpose of AI is to get snakes to regenerate and reproduce themselves!

As it happens, AI in elapids in particular is best suited to venomoids and so to that extent, there is no doubt that this paper will get the hearts racing of my strongest critics.

“Hoser can Artificially inseminate Taipans and Death Adders and breed loads of them, but no one else can!”

No doubt there will be more claims of the “unfair competitive advantage” of the Hoser venomoids.

However there are a few claims that can (with humor) be truthfully made against myself.

“Raymond Hoser is a wanker”.

Yes, I masturbated some reptiles.

“Raymond Hoser” sucks cum”.

Yes, I have accidentally sucked snake semen into my mouth.

Hopefully these two admissions will make all the critics happy.

There is however one potential criticism of AI which has a sound basis of fact, even if is not agreed with by a given person.

Because AI enables transfer of semen from any reptile to any other, it makes the idea of hybridizing taxa simple.

Formerly unobtainable hybrids or rarer ones (as seen in some pythons, e.g. Female Water Python (Katrinus fuscus) X Male Jungle Carpet (Morelia cheynei) and Female Australian Scrub (Liasis amethistinus clarki) X Male jungle Carpet (Morelia cheynei) as seen in Hoser 1988) can now be effectively manufactured on call.

It is the removal of pre-mating isolation mechanisms that allow this to be possible and if there is opposition to hybridizing reptiles, this will most certainly manifest in the form of opposition to AI.

Having said this, Hoser (2005b) gave other previously unreported and unknown examples of a breakdown of pre-mating isolation mechanisms without any form of AI involving Death Adders, Copperheads and Tiger Snakes.

(Since that paper was published, we have witnessed male Brown Snakes and Male Copperheads try to mate one another, and also male Red-bellied Black Snakes engage in combat with male Eastern Brown Snakes at Tuggeranong Hyperdome, ACT in October 2007, immediately before the male Red-bellied Blacks were then separated, placed with a female Red-bellied Black and then both were seen trying to mount her in the above reported attempted “threesome”).

Hybrid reptiles are generally regarded as worthless here in Australia and in terms of pythons at least, they sell for less here than the “purebreds”.

In the USA the picture is mixed, where some of the more unusual hybrids such as “Carpondros” (Green Python (Chonropython viridis) crossed with Carpets (Morelia variegata et. al.) sell for huge prices.

With AI making these hybrids theoretically accessible to more people, it is likely that after an initial rise and spike in the number of hybrids, the novelty will wear off and AI’s main long term application will be simply for breeding rarer and harder to breed taxa or for facilities such as ours where the keeper does not hold large numbers of given taxa and has a greater dependence on single individual reptiles for breeding success.

The main features of AI as detailed here in it’s wider application will be even more captive breeding of reptiles, a corresponding drop in retail prices for private keepers and better still a direct reduction in pressure and exploitation of, potentially limited wild stocks.

Acknowledgments

Reptiles detailed in this paper were held and moved under various permits and acquired from several sources. In all cases, State Wildlife authorities issued permits and movement advices promptly when asked and as always this is appreciated.

Individuals who supplied snakes and lizards referred to in this paper, or held at the same time and inspected, but not referred to here include the following: Scott Eipper, Adam Elliott, Paul Fisher, Robert Gleeson, Ian Renton, Federico Rossignolli, Alex Stasweski, Drew Williams, Andrew Wilson, Peter Whybrow and several others. Several veterinary surgeons supplied equipment, advice and the like for the above reptiles, but as a result of criminal threats made against one in relation to the Hoser venomoids in 2006, their names are not published here.

End notes

All procedures described herein have been conducted with veterinary supervision. All snakes identified as “Hoser venomoids” as defined herein have been certified as such a number times by a licenced practicing veterinarian as permanently devenomized a number of times to satisfy demands of licencing authorities and other government entities (including for example Worksafe Victoria) in more than one Australian state.

To end October 2007, there have been at least 17 known “bites” (as in two fangs at least once and full skin penetration, in all cases), from the Hoser venomoids and in all cases, all were totally untreated and in all cases there were no symptoms or effects on the person. These bites, involving a total of 8 separate people, have included bites from adults of the following taxa, Inland Taipans, 2 snakes, 4 bites, Tiger Snakes, 4 snakes, 9 bites, Collett’s Snake, 1 bite, Eastern Brown Snake, one bite, Common Death Adder (Acanthophis antarcticus), 1 bite, Dajarra Death Adder (Acanthophis woolfi) 1 bite.

Since 1 July 2005, it has been illegal under Section 32 of the “Occupational Health and Safety Act 2004” to display or demonstrate venomous species of snakes in a public place in Victoria that has not had it’s venom glands surgically removed.

References

Hoser, R. T. 1988. Hybridisation between three different species of Australian Python. Litteratura Serpentium (Holland), 8(3), pp. 134-139.

Hoser, R. T. 1989. Australian Reptiles and Frogs. Pierson Publishing, Mosman, NSW, 238 pp.

Hoser, R. T. 2004. Surgical Removal of Venom Glands in Australian Elapids: The creation of Venomoids. The Herptile 29 (1):37-52.

Hoser, R. T. 2005a. Surgically enhanced venomous snakes. Venom glands out, silicone implants in! The creation of perfect exhibition snakes in the post HIH era. Crocodilian - Journal of the Victorian Association of Amateur Herpetologists:17-28, 5(2)(August 2005):17-28 (and covers),5(3)(November 2005):30-36.

Hoser, R. T. 2005b. Pecking orders in large venomous Snakes from South-east Australia … ecological and distributional Implications. Boydii (Journal of the Herpetological Society of Queensland), Spring: (33-38)

Hoser, R. T. 2006. Successful keeping and breeding of Eastern Brown Snakes (Pseudonaja textilis) including an assessment of previously documented failures and reasons for them. Crocodilian - Journal of the Victorian Association of Amateur Herpetologists 6(2) (August):16-28.

Hoser, R. T. 2007. Garbage Guts - Australian Tiger Snakes. Reptiles (USA) (March), 15(3):48-60.

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