Useful reptile care info

,,Multiclutching in Captive Lace Monitors,, by David S Kirshner




















,,Breeding Spiny-Tailed Iguanas Of The Genus Ctenosaura,, by Kelly Paul

A prerequisite for breeding spiny-tailed iguanas, of course, is healthy animals.Ctenosauraas young as 14 months have laid fertile eggs, but I wouldn’t recommend breeding spiny-tailed iguanas that are that young. I recommend waiting until they are at least 2 years old.

To condition your spiny-tailed iguanas to breed, they need to follow a natural daylight cycle. Starting September 1, begin decreasing their 12-hour light period by 15-minute increments every week. By December 1, their lights should be on nine hours a day. You can also skip turning on the lights for a day or two every so often, which will mimic a cold, cloudy day in nature. Yourspiny-tailed iguanas may experience some weight loss during the decreased light cycle; if it is excessive, increase the light cycle immediately.



Adult pair of Honduran black-chested iguanas (Ctenosaura melanosterna).

Continue the nine-hour light cycle until February 1, then begin increasing it by 20 minutes each week. By June 1 your animals should be exposed to 14 hours of light a day. May through August is the prime time for spiny-tailed iguana breeding and egg deposition. Males’ hemipenile bulges will be larger, as will their femoral pores that will be producing waxy excretions. Depending on the species, males may also begin dragging their hindquarters when walking. These are all clues that indicate breeding season has begun.

It is important to monitor your animals during breeding season. Males are usually larger and stronger than females, but either animal can be harmed during breeding. If you don’t keep a compatible pair of spiny-tailed iguanas together year round, during the breeding season you can introduce the male to the female’s enclosure or vice versa. Breeding usually begins with the male rapidly bobbing his head at the female. He will then approach her and usually begin flicking his tongue over her body. If the female is receptive she will lower her head and the male will grasp her by the neck. He will then hold her in position with a front and rear leg over her. His tail will begin to inch under hers, and at this point the female usually raises her tail and breeding will commence. They may remain in this position for up to 20 minutes. If the female is unreceptive she will run away, or she may push the male off using her tail.

Gravid females tend to bask more often. Gestation can take 45 to 70 days depending upon the species. Females of some species may exhibit a decreased appetite during the weeks leading up to laying eggs; others will continue to eat right up until they lay their eggs. Have a nest box ready and in the enclosure two to three weeks before the female is ready to lay eggs, so she has time to adjust to it being in her enclosure. The nest box length should be one to one and a half times the length of the female and 24 inches wide. The substrate inside, consisting of peat moss, coarse sand and slightly moistened top soil, should be 12 inches deep. Be sure it is well mixed so that the female can dig an egg deposition chamber without it caving in on her. When she is ready, she will dig a burrow in the nest box, turn around inside it and proceed to lay her eggs.

It can take several days for a female spiny-tailed iguana to finish laying her eggs. When she’s done, she will back-fill the nest. Immediately after laying, female spiny-tails will appear thin and exhausted. They should be soaked and will usually drink at this time. Within one to three days they should be eating normally again.

Clutch size varies with species, size and age of the female. Smaller Ctenosaura and younger animals lay approximately four to 10 eggs. Large, mature female Mexican (C. pectinata) and black (C. similis) spiny-tailed iguanas may lay 40 to 55 eggs. The eggs of most species are about the size of bearded dragon eggs.

I incubate all my Ctenosaura eggs at 86 to 88 degrees Fahrenheit with about 70 percent humidity. I have used perlite, vermiculite and sand as incubation mediums, though recommend vermiculite. Mix it with enough water so that if you squeeze it with your hand as hard as you can, only a few drops of water will fall out. I live in Phoenix, where it is hot and dry, and I check the moisture content of the incubation medium about every 15 days. If it is too dry the eggs will collapse, and if it’s too wet mold and fungal growth will develop. Following these guidelines, eggs should hatch after 65 to 80 days.

Kelly Paul is a hobbyist with a lifelong interest in reptiles. He has bred more than two dozen species, including six Ctenosaura. He has been a guest speaker at several events, including the International Herpetological Symposium. Email him

Breeding Spiny-Tailed Iguanas

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,,Assessing the health of wild-caught chameleons,, by Ardi Abate

By Ardi Abate, Chameleon information Network

Stressed chameleons attempt to drink from small pools of water on the floor of their cage at an export facility in Madagascar.

Stress is a major factor that will determine the long-term survival of wild-caught chameleons. Overcrowded cages, like that shown above, are not uncommon at every step of the importation process.

The dead chameleon pictured above was one of several dead animals observed at exporters' holding facilities.

Editors Note: It is my belief that the purchase of wild-caught chameleons is detrimental to chameleons -- the individual animals as well as the species which they represent. However, for those considering the purchase of a wild-caught animal, I have provided this article (with permission from Ms. Abate) in the hopes that it will better prepare you to maintain the wild-caught animals after the purchase. Make note, however, (as Ms. Abate establishes in her article) that doing things 100% "right" still will not ensure success with an imported chameleon. Also note that many imported animals are misrepresented as "captive raised", "farmed", "ranched" or other such vague terms. When purchasing a chameleon, it is best assumed wild-caught, unless you are purchasing from a well-established breeder. -- Michael Fry


The purpose of this article is to help buyers of wild-caught chameleons recognize the health problems that may result from transferring free-living wild chameleons into captivity. All too often I hear from chameleon owners who are distressed and perplexed by a decline in health, or the death, of an imported chameleon. Many of them were experienced in chameleon husbandry and provided adequate environments, food and water within the accepted guidelines of captive husbandry. However, the underlying causes of poor health and mortality in wild-caught chameleons may have begun long before being purchased by a new owner. We will examine some scenarios that can occur from the point in time that the chameleon was last living free, until the point in time that the chameleon is purchased by it's new owner.

Many wild-caught chameleons will not be subjected to all of the conditions described in this article and may be treated more gently and humanely at every stage of the exportation/importation process. This article is not intended as an indictment against exportation, importation or those engaged in this commerce, although I anticipate that it may be construed this way by some readers. I urge readers to view this information, some of which is speculative, in the light of identifying potential health problems and seeking timely medical treatment for newly-imported chameleons to maximize their chances of surviving as healthy captives.


I observed a few free-living chameleons with obvious health problems during my two field expeditions to Madagascar. The most remarkable was a male Ch. [F.] pardalis with all but one inch of his tail amputated. This animal's tail had been run over by a passing car, according to a man who lived in the area where I found this chameleon. It appeared to have healed without incidence as the wound was quite old, and he was in otherwise robust condition. I also encountered two chameleons with wounds near the mouth, one of which was badly infected; however both appeared to be in otherwise good condition.

I also saw several chameleons with the telltale raised outlines of parasitic worms under the skin. Other species of internal parasites are not outwardly visible like subcutaneous nematodes, but it should be assumed that wild chameleons harbor many different types of parasites. These chameleons were not in poor condition either; all were in good body weight, alert and strong. According to KLINGENBERG (1993):

In general, the parasites most successful in propagating themselves are those doing the least harm to their hosts. The perfect parasite would prefer a state of mutual benefit or a symbiotic relationship with their host. Many parasitic problems in nature are self-limiting.

Nature can be swift and merciless to the weak. Chameleons that are badly wounded or diseased are dispatched quickly by predators or harsh environmental conditions. This is the reason many wild creatures are adept at masking illness and injury until they are severely compromised, and chameleons are better than most at this preservation strategy. With this in mind, we will assume that most wild chameleons that are captured for the pet trade are outwardly healthy-appearing, robust specimens at the point of capture.


Chameleons are viewed superstitiously in many ways by the native peoples in their countries of origin. In my travels through Madagascar I often ask Malagasy people what they think of chameleons. The answers range from abject fear to reverence.

While these beliefs may change slightly from one geographic area to the next, one superstition seemed universal - Malagasy people do not want to touch chameleons! I got reactions ranging from wide-eyed shock and fear to uncontrollable giggling whenever I touched a chameleon or let it perch on my hand or arm. There were occasions when I asked villagers to guide me into the forests to search for specimens, but often I could not convince anyone to help me because of their fear of chameleons. Sometimes, one of the younger men would agree, but I learned quickly that I would have to instruct them on how to handle a chameleon without injuring it.

In 1996 I located a young male Parson's chameleon 15 feet high in a breadfruit tree. A young Malagasy man volunteered to bring it down so that I could measure and photograph it. He climbed to the branch, quickly pulled the poor chameleon from the branch, and before I could stop him, tossed it to the ground. After I inspected the seemingly uninjured, but visibly distressed chameleon (and recovered from my horror), my guide lectured the young man and showed him how to handle chameleons gently. We made it a point from then on to provide this instruction and give a demonstration of proper handling before we accepted help from anyone in locating chameleons. It was obvious, however, that most of them could not overcome their ingrained fear of touching chameleons.

I have not witnessed the collection of chameleons for the pet trade, but given the prevailing attitude of the Malagasy (and Africans) toward chameleons, I suspect that many of them are mishandled similarly to the Parson's chameleon in my story. Pulling a chameleon forcibly from its perch can result in bone fractures, muscle, joint, and tissue damage. Chameleons thrown to the ground from a high perch may suffer fractured bones, internal injuries or abrasions. Any swellings, but particularly of the feet, legs, ribs or backbone should be examined and x-rayed by a veterinarian to determine the extent of the damage and course of treatment.

Children may capture chameleons for the pet trade as a means of earning income and hold them for a period of time until the exporter comes around to collect and pay for them (FREED, 1998). It is unlikely that these animals are handled gently or kept under acceptable conditions, which may result in stress, dehydration, malnutrition or injuries.

Probably the most detrimental health aspect of capture is the resultant stress on a chameleon. The point of capture sets the stage for a multitude of health issues related to the long-term effects of stress. Stress can lower resistance to disease by suppressing the immune system. Stress also depletes stores of corticosterones (hormones that handle stress in the body) and reduces reproductive hormones to near zero levels. A stressed reptile will not and cannot reproduce (KLINGENBERG, 1993).


Chameleons may be collected in remote areas and transported to an exporter's holding facility hundreds of miles away when enough chameleons have been collected to comprise a shipment. In Madagascar we shot video footage of chameleons being placed in small cloth bags which were then tied shut and placed in a large cardboard box, layer upon layer, until the box was filled. This Malagasy collector lived in Sambava and the chameleons were destined for an exporter in Tamatave, necessitating air shipment.

Since no international flights originate in Tamatave, these chameleons would then have to be shipped by air to the capital of Antananarivo to be exported to foreign countries. There are no air-conditioned facilities at most of the airports in Madagascar and no temperature-controlled cargo holds on the smaller planes, which likely results in mortality due to heat stress during the hotter months of the year for chameleons packed for shipment in this fashion.

In the small airport in Maroantsetra, I saw two boxes labeled "Live Animals" destined for Antananarivo sitting on the tarmac in the open on a very hot day. I asked to inspect the contents of these boxes, and with the permission of the Mayor and Chief of Police, found twenty Uroplatus fimbriatus packed as I have described above. These leaf-tailed geckoes had been collected illegally and were released that day in a protected nature reserve.

It is impossible to estimate how many days or weeks from the point of capture this stage of the process may entail, but the stress of being shipped under the conditions I observed is undoubtedly detrimental to chameleons and likely results in significant mortality due to injury, suffocation, or overheating. In addition, the adverse effects of water and food deprivation and injury can occur to chameleons held over periods by collectors with inadequate facilities.


I toured three exporter's facilities in Madagascar in 1996 and 1997. The conditions varied in each one, and my observations were as follows:

  1. One facility was a large, open fenced field. Chameleons were kept in three large pens surrounded by sheet metal fences that prevented escape. Inside the pen were dead branches and dried palm fronds. Sections of bamboo on the ground provided shallow pools of drinking water. There were approximately 100 live Ch. [F.] pardalis from many geographic regions, Ch. [F.] verrucosus and Ch. [F.] oustaleti and 10 - 20 dead chameleons in each pen on the day I was there. The temperatures in November in this area were approximately 100°F [38єC] and the dried palm fronds provided the only shade available for these chameleons. There were several chameleons crowded into the smallest scrap of shade to escape the heat. I politely informed the exporter that the chameleons needed shade and water and that they appeared to be severely dehydrated. "There is no shade for them..." he said "...and besides, they will all be shipped out to America next week". These chameleons were dehydrated and thin. Those that survived exportation would require veterinary examination, intensive rehydration therapy, treatment for parasites and nutritional support. The long-term effects of dehydration can include kidney failure.
  2. In the second facility, chameleons are kept in wire-screened cages with concrete floors and dead branches for perching. The majority of cages were littered with fecal droppings, increasing the risk of parasite transmission and bacterial disease. There was no drinking water delivery system evident. The climate in this region is too hot and dry in summer, and too cold in winter for some species that were being held. When I visited there was adequate shade in most, but not all of the cages, and there were signs of dehydration in a few specimens, particularly the C. p. parsonii. Multiple chameleons of both sexes were overcrowded in these cages and there were a few dead and dying chameleons on the ground. It should be remembered that chameleons are asocial and usually will not comfortably tolerate the presence of other chameleons. Forced cohabitation in overcrowded enclosures elevates stress significantly and may result in injury from bites and scratches. Chameleons showing signs of injury from rough cage wire such as broken or amputated claws and toes, swelling, lacerations, bite marks or abrasions should be examined and treated by a veterinarian.
  3. The third facility had many well-designed, spacious, and heavily planted enclosures, although there were several rows of cages that were inadequate. This facility is open to the public for tours. For the most part, the animals I saw were in good condition, well nourished and hydrated. Six C. p. parsonii in one enclosure were in only fair condition - one large male lay dying on the ground and two other specimens appeared thin and possibly dehydrated. There was no drinking water delivery system evident. Insects were gathered by local children and provided to the chameleons in sufficient quantities to maintain adequate body weight. In my opinion, chameleons held in this facility for a short period under the conditions I observed in 1996 might not experience a significant decline in overall health, except for the ill effects of stress from captivity and low-level dehydration. Recent reports (1999) on this facility indicate that the current population may be overcrowded.


The United States Fish and Wildlife Service (USFWS) and the International Air Transportation Association (IATA) govern the packing and shipping regulations for chameleons arriving in the United States. Chameleons are listed on CITES Appendix II, which also governs how animals must be shipped. This is an area that has received a good deal of attention in the past year due to proposed changes to shipping regulations by the USFWS. One of the proposed changes stipulated that exported animals be examined by a veterinarian and certified healthy before they were shipped. In my opinion, this would be very difficult to achieve meaningfully in most countries. In underdeveloped countries like Madagascar it would be nearly impossible due to the lack of trained veterinarians and the financial resources for managing a program like this. Consequently, there is no veterinary inspection of chameleons that are shipped into the U.S. and it is unlikely that will occur in the future.

Upon arrival in the U.S., U. S. Customs or the USFWS inspect the shipments to verify that the numbers of specimens and the species match the CITES permits and that they have been shipped humanely. Legal action against the importer may result if the animals are dead or in poor condition. I have heard sporadic horror stories concerning the condition of some shipments in which mortality was very high. Delays in shipping, temperature extremes and the stress of being enclosed in a shipping container can have devastating effects for those chameleons whose health has already been compromised by poor management while in the care of collectors and exporters after capture.


I believe many importers work diligently to salvage as many chameleons in these shipments as possible. I observed one shipment of several hundred chameleons from Madagascar being tended to by an experienced importer. The chameleons were provided with copious dripping water and many lapped the water eagerly. A number of these chameleons were in poor condition. This importer did not have adequate room to house each chameleon singly. Each species was overcrowded in cages that still contained feces from the former inhabitants.

I saw another newly arrived shipment shortly after it was unpacked. This importer/retailer crowded more than eighty chameleons from Cameroon in a dark, glass-walled enclosure about five feet square. There was a strong odor in this enclosure and many of the chameleons did not have a branch to perch on. Some of the chameleons were thin and bore wounds and scratches.

Many importers sell their chameleons wholesale, others sell wholesale and retail. Wholesalers may keep chameleons a very short period of time before they ship them out to retailers, like pet stores. Some wholesalers attempt to rehabilitate the animals and assess their health status before they are shipped to retailers. Part of this rehabilitation may be to administer anti-parasitic drugs, typically fenbendazole (Panacur) and metronidazole (Flagyl). It is not common practice for importers/wholesalers to have individual chameleons tested by veterinarians for the presence of parasites via fecal, sputum and blood analysis. While Flagyl and Panacur are effective against a fairly broad range of parasites, there are many parasites that are not eliminated by these two drugs. All imported chameleons should have a fecal analysis performed by a veterinarian at a minimum. Blood tests and physical examinations should also be performed to determine the presence of blood-borne and subcutaneous parasites and the proper course of treatment prescribed by a veterinarian. Periodic follow-up exams are highly recommended.


Once chameleons arrive in the retailer's facility, they are available to the public for sale. Unfortunately, many pet stores do not have the knowledge or appropriate setups to maintain chameleons and are unprepared to provide buyers with proper enclosures or supplies, or educate them in the proper care of chameleons. By now, those chameleons who have survived being captured, held by the collector, shipped to the exporter, held by the exporter, shipped to the importer and finally, to the retailer, may well have experienced long term food and water deprivation, injury, inappropriate temperatures, exposure to pathogens in unhygienic environments, and the stress of being handled by humans, enclosed in bags and boxes, and attacked by other chameleons in overcrowded enclosures.

The parasites harbored with no apparent ill effects by the free-living wild chameleon take on a different dimension under these conditions. Parasite loads may increase dramatically, causing life-threatening, and irreparable damage to the stomach, small and large intestine, esophagus, organs and lungs. This can result in organ obstruction, loss of nutrients, blood loss, tissue destruction and the introduction of bacteria (KLINGENBERG, 1993). DR. KLINGENBERG (1993) wrote:

Captivity-induced stress tends to change the balance of the host/parasite relationship, which can result in disease. Such stress factors include crowding, inadequate heat or light, poor hiding areas (lack of security), substrate problems, altered diets, etc. All these factors suppress the immune system of the host and make the individual more susceptible to the effects of infestation.

To summarize, I sincerely believe captivity and captivity-related stress to be responsible for parasite infestations, self-limiting in nature, becoming pathogenic (disease causing) in captivity.

Because of the risk factors associated with captivity and the availability of relatively safe and effective compounds to treat both internal and external parasites, it has become my practice to eliminate all parasites in captive specimens. I feel we owe it to these captive creatures, who depend on us for their care, to eliminate the possibility of parasitic disease as a stress factor in their lives.

The buyer, unaware of what may have recently transpired in the life of the brilliantly colored chameleon pacing nervously in the pet store, buys what they may believe to be a healthy animal. As proof of health, sellers routinely assure buyers that a chameleon has been deparasitized and is eating. The new owner takes their chameleon home with high expectations that it will thrive and even reproduce. Under the best of conditions, a veterinarian examines the chameleon and minor health problems are treated successfully. Unfortunately, even these chameleons may fail to thrive due to maladaptation.

In my own experiences with several wild-caught chameleons a pattern began to emerge. I found that during the first few weeks to a month most chameleons consumed food eagerly and appeared alert and robust. Many looked comfortable, but wary in their new environments, however there was little other cause for concern from outward appearances. I have dubbed this the "honeymoon" period.

While it does not occur in every case, the next stage seems to determine whether or not the chameleon survives. Food intake often ceases and the chameleon appears depressed or nervous, and may pace the perimeter of the cage. It is at this crossroad where the chameleon either seems to accept captivity and continues life, or begins a downward spiral to death. I believe this to be the essence of maladaptation. Those chameleons that simply cannot tolerate their loss of freedom coupled with human interaction, or whose health has been irreparably damaged by the rigors of being transferred into captivity simply waste away. Probably the single most critical factor in maladaptation is the level of prolonged stress the chameleon has endured from the point of capture.

Reproductive females are more adversely affected by the rigors of captivity than males in my experience. I generally hold out little hope for the long-term survival of newly imported gravid females (if they are gravid at the point of capture or shortly thereafter), their eggs or offspring. I give most wild-caught male chameleons that are not visibly compromised about a 25% chance of expiring within 6 months to a year because of the possibility of maladaptation.

For chameleons with visible signs of ill health such as sunken eyes, serious injuries, bacterial or fungal infections, malnourishment, feeble grip, and heavy parasitic infestations, the prognosis is very poor.

"Nosey", a male Ch. [F.] pardalis shows signs of trauma which may have resulted from rough handling during capture, or from improper housing after being collected from the wild.


The Life with Chameleons story by Bianca Rice in CiN Journal No. 29 chronicled her experiences with an imported male Ch. [F.] pardalis she named "Nosey". This chameleon had been rescued from being euthanized in a pet store due to declining health by Bianca and her friend Mary Ellen McLoughlin, a veterinarian. Nosey's initial health problems were serious:

  • Weight loss
  • Malnutrition
  • Swelling near one eye
  • Bacterial infection of the sinus
  • Abrasion on the snout
  • Heavy Roundworm infestation

Dr. McLoughlin pursued an aggressive course of therapy for these health problems and gradually this chameleon was brought back from the brink of death. Recently, I received a follow-up letter from Bianca on Nosey's current condition. She wrote that he had developed a swelling on his lower jaw, edema, and that a blood test indicated that there were cellular abnormalities, anemia and elevated phosphorus levels. X-rays showed "...that an area on his back, that had always had a dime-sized discolored scar was in fact surrounding a broken spine. Perhaps this occurred upon capture." The photograph with the letter shows other scars on this chameleon on the rear leg and flank.


Wild-caught imported chameleons usually carry a much lower price tag than captive-raised chameleons. However, if the imported chameleon you purchase requires several courses of treatment for medical problems, the veterinarian costs may well run two to three times the original price of the chameleon.

Avoiding veterinary care and medical treatment may cost the chameleon it's life, which will certainly cost the owner their original investment in purchasing the chameleon. Here are some suggestions for safeguarding the health and longevity of a newly imported chameleon:

  1. Purchase imported chameleons from reputable sellers who offer a guarantee if you are dissatisfied with the chameleon, it is diagnosed by a veterinarian with serious health problems, or dies. Ask the seller how many days you have to make a determination before the guarantee expires. Remember that it may take several days to receive the results of some diagnostic tests. Understanding the seller's guarantee is especially important if you are purchasing a chameleon by mail order.
  2. Do not purchase chameleons with visible signs of ill health unless you are prepared to accept what may be a significant financial responsibility for veterinary care. For example: A recent treatment on a male Panther chameleon for the surgical removal (requiring anesthesia) of a heavy infestation of subcutaneous nematodes was $250.00. Three weeks later it is apparent that there are additional nematodes under this animal's skin and he will require additional treatment (cost to be determined). These treatments may have to be repeated 3 - 5 times to eliminate this parasitic infestation.
  3. Make an appointment as soon as possible after purchasing a chameleon with a veterinarian that specializes in reptiles for a physical examination and a fecal analysis, at a minimum. The examination may indicate that other diagnostic tests are advisable. Pursue the optimal course of treatment for all health problems. Get follow-up examinations every 3 - 6 months, even if no health problems are detected in the initial exam. (For information on veterinary care, review the article in CiN Journal No. 30.)
  4. Quarantine the new chameleon until diagnostic tests are completed and any health problems are treated and cured. Do not allow contact between the new chameleon and any other chameleons in your collection during quarantine, including mating. Disinfect your hands, food bowls and any utensils that come in contact with the quarantined chameleon. Do not recycle uneaten food from the quarantined chameleon's enclosure to another chameleon. Do not house more than one chameleon in a cage during quarantine. (For information on quarantine and infection control, review the article in CiN Issue No. 16.)
  5. Observation is critical in the daily assessment of the health of wild-caught chameleons, but keep physical handling and traffic in the chameleon's visual range to a minimum. Provide ample drinking water and ensure the chameleon is drinking. Keep records of food and water consumption on a daily basis.
  6. Large, airy, well-planted cages may help new captives acclimate. Chameleons that constantly butt against the walls of their enclosure attempting to escape may do serious damage to themselves in addition to the high levels of stress they are experiencing. Use soft mesh or PVC-coated wire for these chameleons.
  7. Consult a veterinarian immediately if :
    • Droppings are foul smelling, runny or mucus-laden.
    • The chameleon stops eating (anorexia) for more than 2-3 days.
    • Any swellings, sores or discolored areas are observed in the mouth or on any other part of the body.
    • Breathing is labored, or open-mouth breathing is observed. Mucus is present in the mouth.
    • There is a popping or wheezing noise when the chameleon breathes.
    • Food is regurgitated, or there is undigested food in droppings.
    • There is visible weight loss, sunken eyes, swollen eye(s), general weakness, inability to perch, or eyes are closed (sleeping) during daylight hours.

I have acquired a number of wild-caught chameleons over the years. The majority of these chameleons adapted well, did not have life-threatening health problems, and with a little persistence, parasites were eliminated. Unfortunately, I can remember vividly the ones who did not adapt and succumbed to serious health problems exacerbated by the stresses of captivity.

I urge everyone to provide the benefit of veterinary care to all chameleons, and to give special consideration to the wild-caught chameleons who sacrificed their freedom in the wild in exchange for the joy and wonder they bring to their human caretakers.

Opposing views, observations, questions and comments are welcome. Please call (858) 484-2669 or write to CiN, 13419 Appalachian Way, San Diego, CA 92129, email:

(c) 1999, Ardi Abate, Chameleon information Network. Copies of this article may be reprinted from this website by permission of the author and the Chameleon information Network.

,,Care sheet of Cordylus giganteus,, by SMITH, 1844

Cordylus giganteus SMITH, 1844 

Cordylus giganteus
Adultes Weibchen • Adult female
 Cordylus giganteus
Cordylus giganteus
Adultes Männchen • Adult male
 Cordylus giganteus
Benannt nach dem Lateinischen giganteus=gigantisch, da es sich um die größte Cordylus-Art handelt. Imposante Tiere mit kräftigen und effektiven Stacheln. Unvorsichtiger Umgang führt zu blutigen Händen. Im Terrarium einfach zu halten, aber nur schwer nachzuziehen, weil unbedingt die klimatischen Verhältnisse in ihrer Heimat nachgebildet werden müssen. Das geht fast nur im Freiland oder in einem kühlen Kellerraum. 
  1. Schutzstatus EU-Verordnung 338/97, Anhang B

max. 40 cm 

Südafrika: Oranje-Freistaat, Südost-Transvaal, West-Natal 

offene Grassteppen, meist in Kolonien in selbst gegrabenen Höhlensystemen, bis zu 3 m lange, 50 cm tiefe Gänge 

Sungazer or Giant Girdled Lizard 
Named after Latin giganteus=gigantic because it’s the largest member of the Cordylus group. Impressive animals with very large, hard, and effective spines. When handled without caution bleeding hands are rather the rule than the exception. Captive care is easy considering some basic rules, but breeding is hard since you need to meet the natural conditions. Possible only in outdoor cages or in a cool cellar room. 

EU Directive 338/97, Appendix B 

Adult Size 
max. 40 cm 

South Africa: Oranje Free State, SE Transvaal, W Natal 

  1. Environment open grassland, mostly in colonies, digging large cave systems with up to 3 m length and 50 cm

Um die wichtigsten Inhalte als PDF herunterzuladen und auf DIN-A4-Format auszudrucken, klicken Sie bitte auf den Thumb ... Cordylus giganteus To download the most important content as PDF and print on A4 standard paper please click on the thumb ...

Mindestens 150x80x60cm BTH. Braucht Höhlen, z.B. mit Sanitärrohren im Boden. Bodengrund muss fest sein, z.B. Fertigrasen oder grabfähiges Sand-Lehm-Gemisch (5:1). In Höhlen leicht feucht. Temperatur tags 22–28, Sonnenplatz 40°C, nachts 18–20°C, Luftfeuchtigkeit 50–60%. Tageslichtleuchtstoffröhren, HQL-Strahler, Wassergefäß, flache Steine. Lebenserwartung: bis 40 Jahre. 
  1. Futter Insekten, Nager, Fleischstücke, bei mir aber auch Weintrauben und in Fleischbrühe gekochte Nudeln, Mineralien/Vitamine (z.B. Herpetal
    Zucht und AufzuchtGruppenhaltung möglich, aber Vorsicht beim Füttern. Für jedes Tier eine eigene Höhle. Überwinterung unabdingbar. Weibchen bringt alle zwei Jahre 2–4 Jungtiere lebend zur Welt. 
  2. Literatur Langerwerf, B.: Keeping and breeding the sungazer (Cordylus giganteus Smith 1844). Reptiles 9 (6), 58–65. Boulder, CO,
    Schwier, H.: Der Riesengürtelschweif Cordylus giganteus – einzigartig und mysteriös zugleich. Reptilia 12 (3): 40–44. Münster, 2007. 
Captive Care 
At least 150x80x60cm WDH. Needs caves, made e.g. from plastic pipes in the ground. Ground itself needs to be solid, e.g. turf or clay/sand mix (1:5), keep a bit humid so they can dig. Temperature 22–28, sunny spot 40°C, night temperature 18–20°C, humidity 50–60%. Daylight fluorescent lamps, HQL spot, some flat rocks, water bowl. Life span: up to 40 years. 
  1. Food insects, rodents, pieces of meat, but also grapes and pasta cooked in bouillon, minerals/vitamins (e.g. Herpetal
    Reproduction and Rearing Groups possible, but every animal needs its own den. Hibernations seems to be necessary, but is no guarantee for success. Female gives birth to 2–4 youngs every two years in the wild. 
  2. Literature Langerwerf, B.: Keeping and breeding the sungazer (Cordylus giganteus Smith 1844). Reptiles 9 (6), 58–65. Boulder, CO,
    Schwier, H.: Der Riesengürtelschweif Cordylus giganteus – einzigartig und mysteriös zugleich. Reptilia 12 (3): 40–44. Münster, 2007. 
Cordylus giganteus
Cordylus giganteus: Die Art frisst auch ungewöhnliches Futter, hier eine Weintraube • species sometimes accepts unusual feeding items, captured here: a grape 
Cordylus giganteus
Da die Tiere auch in der Natur in Kolonien leben, funktioniert normalerweise auch die Gruppenhaltung • Since this species lives in colonies in the wild it is possible to keep groups 

,,A new leaf tailed gecko ( Uroplatus ebenaui ) species from northern Madagascar with a preliminary assessment of molecular,, from


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Zootaxa 3022: 3957 (2011)

Copyright © 2011 · Magnolia Press

ISSN 1175-5326 (print edition)




A new leaf tailed gecko species from northern Madagascar with a preliminary assessment of molecular and morphological variability in the Uroplatus ebenaui group



1Technical University of Braunschweig, Zoological Institute, Mendelssohnstr. 4, 38106 Braunschweig, Germany 2Département de Biologie Animale, Université d'Antananarivo, BP 906. Antananarivo, 101 Madagascar

3Center for Conservation Research, Omaha´s Henry Doorly Zoo, 3701 South 10th Street, Omaha, NE 68107, United States 4CIBIO, Centro de Investigação em Biodiversidade e Recursos Genéticos, Campus Agrário de Vairão, R. Padre Armando Quintas, 4485-661 Vairão, Portugal

5Zoologische Staatssammlung München, Münchhausenstr. 21, 81247 München, Germany 6Madagascar Biodiversity Partnership, VO Bis A, Manakambahiny, Antananarivo, Madagascar 7Corresponding author. E-mail:


Endemic to Madagascar, the genus Uroplatus of the family Gekkonidae consists of 13 nominal species of leaf-tailed geckos. These forest dwelling lizards are famous for their cryptic and odd appearance. We describe a new species of the Uroplatus ebenaui group from the Montagne d’Ambre massif in northern Madagascar. Uroplatus finiavana sp. nov., is morphologically similar to the sympatric U. ebenaui but differs in multiple character state expressions, among which are a longer tail and an unpigmentated oral mucosa. It also can be differentiated from U. ebenaui and all other Uroplatus spe-cies based on a high level of divergence in the mitochondrial ND4 gene and the nuclear C-mos gene, and no instances of haplotype sharing exist in these genes among the analysed species. The new species is relatively abundant at Montagne d'Ambre National Park where at lower elevations (ca. 700 m) it occurs together with U. ebenaui, without any signal of genetic admixture. Records of U. ebenaui in the mountains and forest blocks of northern Madagascar (especially in the Tsaratanana massif) actually represent other candidate species with distinct morphology and characterized by a high ge-netic divergence from the described species. Beside the description of the new species we discuss the geographic prove-nance of the holotypes of the nominal species and synonyms in the U. ebenaui group and provide further information on the phylogeny of the U. ebenaui species group including the first incorporation of Uroplatus malama in a molecular data set.

Key words: Uroplatus finiavana sp. nov., Uroplatus malama, ND4, C-mos, Gekkonidae, junior synonyms, Montagne d’Ambre, Madagascar


Due to its unique biodiversity and the continued decline of its biota as a result of human pressure, Madagascar is recognized to be among those countries with highest conservation priorities (Myers et al. 2000). Even so, knowl-edge of its vertebrate fauna is far from complete, and new vertebrate species have been discovered and described at an unprecedented rate in the last few years (e.g., Yoder & Nowak 2006; Glaw et al. 2010). Focusing on the herpeto-fauna, Madagascar harbors over 660 described species of which more than 95% are endemic to the island (Glaw & Vences 2007). This high species diversity is associated with a high degree of microendemism, and this high per-centage of range-restricted species is largely the outcome of a specific combination of climate, topography, vegeta-tion and historical events (Wilmé et al. 2006; Schatz 2000). One of the areas exhibiting high microendemism is Montagne d'Ambre, a volcanic massif covered by an extensive mountain forest that rises to an elevation of 1450 m, and forms an isolated rainforest patch surrounded by the otherwise dry habitat of the northern tip of Madagascar.

Established as a national park in 1958, and being among the seven most visited parks in Madagascar, Montagne d’Ambre National Park boasts over 75 species of reptiles and amphibians (Raxworthy & Nussbaum 1994; D’Cruze et al. 2008). The relevance of this massif for the harboring of new and microendemic species is illustrated by the recent descriptions of various new reptile species (e. g. Glaw et al. 2007, 2009).

The Madagascar-endemic genus Uroplatus consists of the unique nocturnal forest dwelling leaf-tailed geckos, all of which display bizarre shapes and appearances that camouflage these animals. Currently containing 13 reco-gnized species, the genus Uroplatus is represented by species that are distinguishable from other geckos by their rather flat or laterally compressed body, triangular head, and a leaf-shaped tail. Based on morphology (Glaw & Vences 1994) and molecular phylogenies the genus Uroplatus can be divided in several species groups: the U. fim-briatus group (U. henkeli, U. sikorae, U. sameiti, U. fimbriatus and U. giganteus), the U. lineatus group (U. linea-tus), the U. alluaudi group (U. alluaudi, U. pietschmanni, U. guentheri and U. malahelo), and the U. ebenaui group (U. malama, U. ebenaui and U. phantasticus). Since its discovery, the majority of taxonomic studies have focused on the morphology of the genus. The first phylogenetic hypothesis on the genus’ evolution was presented by Bauer & Russel (1989), who recognized six species. Recent molecular and morphological studies have however indicated that species diversity of the genus Uroplatus is underestimated, even though a comprehensive data set has not yet been established, and population-level data on genetic variation are largely unexplored (Glaw et al. 2006; Green-baum et al. 2007; Raxworthy et al. 2008).

One of the still unresolved questions is the status of the small sized of the Uroplatus ebenaui group that are found at Montagne d’Ambre, despite these being well-known by the scientific community for the past two decades. Glaw & Vences (1994) mentioned that the representatives of U. ebenaui occurring in Montagne d’Ambre exhibited differences from those of typical U. ebenaui. Additional evidence for their morphological distinctness was found by Böhme & Henkel (1995), and recently molecular evidence has supported the specific distinctness of this popu-lation (Greenbaum et al. 2007; Raxworthy et al. 2008). In the present work, we provide further molecular and mor-phological evidence for this differentiation. We show that two species of the Uroplatus ebenaui group occur sympatrically at Montagne d’Ambre and describe one of them as a new species.

Material and methods

Sampling. Specimens were collected during nocturnal surveys, using flashlights or headlamps to locate individu-als. Representative specimens were euthanized by injection of chlorobutanol solution, hardened in 90% ethanol and preserved in 70% ethanol. Locality information was recorded using Global Positioning System receivers. New specimens included in this study were deposited in the collection of the Zoologische Staatssammlung München, Germany (ZSM) and the Université d’Antananarivo, Département de Biologie Animale, Madagascar (UADBA). Historical type specimens used for the morphometric comparative analysis came from the Muséum National d’His-toire Naturelle de Paris (MNHN) and the Zoologisches Museum Hamburg (ZMH). FGZC, ZCMV and 2000-E refer to field numbers of F. Glaw, M. Vences and both respectively; URANO, JAR and UAMB refer to a survey done by E. Louis and F.M. Ratsoavina, ACZC to the field numbers of A. Crottini, and DRV to D. R. Vieites’ field numbers. MPFC acronyms are field numbers of M. Pabijan.

To facilitate effective comparisons, the description scheme of the new species follows that of Uroplatus malama by Nussbaum & Raxworthy (1995), and we used one adult male of Uroplatus malama (ZCMV 12128; to be catalogued in ZSM) to validate our interpretation of characters. Pictures of the living animal were taken at Andreoky in the Andohahela area, in southeastern Madagascar.

Definition of measurements (all in mm) and characters, shown in Figure 1 are as follows: snout-vent length (SVL); tail length (TaL); tail width (TaW); head length, measured from tip of snout to posterior end of the cranium (HL1); head length, measured from snout tip to small central head prominence (HL2), maximum head width (HW), forelimb length (ForL), measured from the proximal end of the humerus to the distal tip of the longest finger, hindlimb length (HiL), measured from the proximal end of the femur to the distal tip of the longest toe, length of medial posterior projection of the interorbital ridge (IRPL), neck triangle length (NTL), measured from the poste-rior edge of a small central head prominence (which is present in the U. phantasticus group species behind the interorbital ridge) to the point where the two lateral skin folds of the posterior head meet centrally on the neck (these ridges are well visible in Figure 8g). Morphometric data were analyzed using Statistica software (Statsoft) employing nonparametric Mann-Whitney U tests.


FIGURE 1. Measurements taken from Uroplatus specimens listed in Table 1, (a) head; (b) body and tail.

Molecular work. Two types of tissue samples were used in the molecular study (tail tip or muscle from the femur). All tissues were preserved in 96% ethanol for molecular analysis. Total genomic DNA was extracted from the tissue samples using proteinase K digestion (10 mg/ml concentration) followed by a standard salt extraction protocol (Bruford et al. 1992). Standard protocols for PCR amplification were used to amplify two gene segments: a fragment of the mitochondrial NADH dehydrogenase subunit 4 (ND4) using the primers ND4 5’-CACCTAT-GACTACCAAAAGCTCATGTAGAAGC-3’ and LeutRNA 5’-CATTACTTTTACTTGGATTTGCACC-3’ (Arévalo et al. 1994) and a fragment of the nuclear proto-oncogene mos gene (C-mos) using the primers CO8 5’-GCTTGGTGTTCAATAGACTGG-3’ and CO9 5’-TTTGGGAGCATCCAAAGTCTC-3’ (Han et al. 2004). We generated sequences using an automated DNA sequencer (ABI 3130 XL, Applied Biosystems) and aligned them using CodonCode Aligner (Codon Code Corporation) with the ClustalW algorithm. Altogether we analyzed 23 ND4 and 41 C-mos sequences of several individuals of the U. ebenaui species group, using samples collected between 2003–2010. Newly determined sequences have been deposited in GenBank (accession numbers JN038075-JN038137).

Bayesian inference (BI) searches were performed with MrBayes 3.1.2 (Ronquist & Huelsenbeck 2003). The Akaike Information Criterion (Akaike 1974) in MrModeltest v2.2 (Nylander 2002) selected a GTR+I+G model as the best fitting model to explain our ND4 data, and we consequently implemented this model in our BI analysis. We performed two runs of 10 million generations (started on random trees) and four incrementally heated Markov chains (using default heating values) each, sampling the Markov chains at intervals of 1000 generations. Stabiliza-tion and convergence of likelihood values occurred after three million generations. The first five million genera-tions were conservatively discarded and five millions trees were retained post burn-in and summed to generate a majority rule consensus tree (Figure 5).

Haplotypes of the C-mos fragment were inferred using the PHASE algorithm (Stephens et al. 2001) implemen-ted in DnaSP software (Version 5.10.3; Librado & Rozas 2009). Haplotype network reconstruction of phased sequences of the C-mos fragment (Figure 7) were performed using the software TCS, version 1.21 (Clement et al. 2000) based on the parsimony algorithm of Templeton et al. (1992).


Identity of the described taxa in the Uroplatus ebenaui species group. To confirm the taxonomic distinctness of the Uroplatus from Montagne d'Ambre (Figure 2), we first investigated the identity of the available scientific names in the U. ebenaui group. We reviewed the original literature and/or examined the type specimens of all nom-inal species in this species group and all their junior synonyms. Original measurements of specimens examined by us are given in Table 1. Specimens of the different species and candidate species in life are shown in Figs. 2, 3 and 8; other morphological and morphometric characters are reported in Figs. 4 and 5; the molecular differentiation among taxa is shown in Fig. 6 and 7; and the geographical distribution recorded for each species in the group as taken from the literature and own data is shown in Figure 9.



FIGURE 2. Photo in life of a male of Uroplatus finiavana sp. nov. from Montagne d’Ambre National Park, February 2004.





FIGURE 3. Uroplatus malamaencountered in May 2010 at Andreoky (Andohahela area), south east of Madagascar around 600–700 m a. s. l. (a) photo of a male and (c) the ventral side of its tail; (b) a female.


Uroplatus ebenaui —This is the oldest nominal species in the U. ebenaui group and was described by Boettger (1879) from Nosy Be (as "Nossi Be"). This species is characterized by (1) an almost straight line of dermal folds between the eyes (interorbital ridge) that connects the supraciliary spines, (2) a short (TaL/SVL 0.216–0.398, mean 0.317± 0.07) and narrow (TalW/SVL 0.047–0.088, mean 0.066± 0.02) original tail; (3) relatively short forelimbs (ForL/SVL 0.333–0.421, mean 0.382± 0.03); and (4) a blackish oral mucosa. Several of these features (such as the straight interorbital line) have been previously noted by Böhme & Henkel (1995) in their account of the holotype of U. ebenaui. Therefore, little doubt remains about the identity of this species.

Uroplatus boettgeri—This nomen was coined by Fischer (1884) based on a single holotype (ZMH R04356) from "Nossi Be" (examined by us). The short original tail of the subadult holotype (TaL/SVL 0.280), the presence of a straight line between the eyes, and the presence of a blackish oral mucosa, are in full agreement with Uroplatus ebenaui. Therefore, we continue to consider U. boettgeri to be a junior synonym of U. ebenaui.

Uroplatus phantasticus—Boulenger (1888) based his original description of U. phantasticus (including a fig-ure) on a single gravid female that lacked exact locality data (type locality: Madagascar) and that had no tail. This specimen was collected by Rev. R. Baron. Five years later, Peracca (1893) described an additional specimen from Andrangoloaka (catalogue number MZUT R.2548 according to Andreone 1991) with a short tail of 12 mm (which is possibly regenerated). Bauer & Russell (1989) mentioned the catalogue number of the holotype of U. phantasti-cus as being BMNH 1946.8.26.64 and stated "Andrangoloaka, 19°02'S, 47°55'E" as the type locality of this spe-cies, apparently in error.

According to Dorr (1997), Rev. R. Baron travelled extensively throughout Madagascar, but apparently toured in the northwest, north and northeast of the island no earlier than 1891, subsequent to the description of Uroplatus phantasticus. All Malagasy amphibians and reptiles collected by Baron and described by Boulenger are from type localities in the northern central eastern portion of Madagascar (e. g. Boophis albilabris), or in the southern central east (e. g. Madascincus macrolepis, Calumma gastrotaenia, Liophidium torquatum, Pseudoxyrhopus microps,Tam-nosophis infrasignatus, Mantella baroni; biogeographic regions according to Boumans et al. 2007). Consequently, we hypothesize that the holotype of Uroplatus phantasticus probably was collected in the same general area as the types of these species, and most likely in the southern central east of Madagascar.




FIGURE 4. The hemipenis of a paratype (ZSM 1133/2003) of Uroplatus finiavana, (a) sulcal view, (b) parasulcal view.

In northern and southern central east of Madagascar, the most common species of the U. ebenaui group is a comparatively large species characterized by a long and broad tail (TaL/SVL 0.622-0.756). However, from the area of Fierenana in the northern central east, another taxon with a very short tail (U. ebenaui-like) is also known. Addi-tional data are required to clarify the type-locality of U. phantasticus since the holotype is devoid of an intact tail; therefore, its tail length and width cannot be ascertained. Our morphometric analysis indicates that besides other characters, the long-tailed form is characterized by a rather relatively short head length. Using Boulenger's original measurements of the holotype (snout-vent length 58 mm calculated as total minus tail length, and head length of 17 mm), the relative head length is 0.293, which conforms well to the values we measured in the long-tailed form (HL/SVL 0.273–0.304, mean 0.291± 0.01). Despite variation and overlapping ranges of this character, most other species in the group have relatively longer heads, with average values of HL/SVL being 0.3. The taxa with longer heads include U. ebenaui, the new species from Montagne d'Ambre, and specimens from Fierenana. Based on this character, we hypothesize that the holotype of U. phantasticus is indeed conspecific with the long-tailed form, although this hypothesis needs further confirmation based on a closer morphological examination of the holotype, including assessment of the color of its oral mucosa.

Uroplatus schneideri—The original description of U. schneideri was based on a single juvenile holotype (MNHN 1914.4) from the Manjakandriana forest (Lamberton 1913), which evidently refers to the region around the village which is located on the national road between Antananarivo and Moramanga, a geographic region in which the long-tailed Uroplatus species (considered to be U. phantasticus) is typically encountered. The holotype (examined by us) possesses a relatively long and large original tail (TL/SVL 0.351), has a black oral mucosa and is from a region where U. phantasticus is widespread. Therefore, we hypothesize that U. schneideri is conspecific

with the long-tailed species from central eastern Madagascar, and we are confident that the synonymy of U. sch-neideri and U. phantasticus is warranted.


FIGURE 5. (a, b) Comparative graphs of relative tail length and relative tail width (TaL/SVL, TaW/SVL) of the specimens employed in our morphometric study. Specimens are separated by sex; graphs display same trend for the compared character. (c, d) Comparisons of the length of medial posterior projection of the interorbital ridge (IRPL) and neck triangle length (NTL), among described species and Uroplatus finiavana only.


FIGURE 6. Bayesian inference tree of the Uroplatus ebenaui group based on a 700 bp fragment of the mitochondrial ND4 gene. Only posterior probability values > 0.95 are displayed at nodes. In yellow the highly supported cluster of the newly described species. Adjacent to each clade, a picture of the oral mucosa is presented showing its pigmentation. Bars in grey scale tones color code roughly for group clades in relation to elevation.

Uroplatus malama—This species was described by Nussbaum & Raxworthy (1995). With a snout-vent length of 71 mm (male holotype) to 77.5 mm (male ZCMV 12128), it is the largest species of the U. ebenaui species group, with a tail long (TL 43 mm in the holotype, 56.1 mm in ZCMV 12128), wide (maximum tail width 12.5 mm in the holotype, 18.4 mm in ZCMV 12128, TL/SVL 0.606–0.724, n=2) and strongly serrated. This taxon differs from all the other species by lacking dermal spines on the head, body, limbs and tail base.

Species description

Based on our review of available names in the U. ebenaui group above, and our morphological and molecular com-parisons as reported below, we conclude that the species from Montagne d'Ambre represents a distinct and inde-pendent evolutionary lineage that differs from all nominal species of Uroplatus, and that no earlier nomen is available to refer to it. We therefore here describe it as new species.

Uroplatus finiavana sp. nov.

Holotype. ZSM 328/2004 (FGZC 625, adult male, hemipenes partly everted) collected at Montagne d'Ambre, 700–1000 m above sea level (a. s. l.) on 19–23 February 2004 by F. Glaw, M. Puente, R.D. Randrianiaina & A. Razafimanantsoa.

Paratypes. ZSM 1132/2003 (FG/MV 2002-2387, adult male); ZSM 1133/2003 (FG/MV 2002-2388, adult male); ZSM 1134/2003 (FG/MV 2002-2390, adult female); and ZSM 1135/2003 (FG/MV 2002-3083, adult male), all collected at Montagne d'Ambre, on 17–20 February 2003 by F. Glaw, R. D. Randrianiaina & A. Razafimanant-soa.


FIGURE 7. Haplotype network reconstruction of a 400 bp fragment of the nuclear C-mos gene for the analysed lineages of the Uroplatus ebenaui group. Haplotypes were inferred using the PHASE algorithm. The haplotypes of U. finiavana are in yellow. In green, blue and red are those of U. phantasticus and U. ebenaui, U. malama, respectively. Grey is used for the candidate spe-cies U. sp. 1–3 from Tsaratanana.

ZSM 322/2004 (FGZC 619, adult female), ZSM 323/2004 (FGZC 620, adult female), ZSM 324/2004 (FGZC 621, adult female), ZSM 325/2004 (FGZC 622, adult female), ZSM 326/2004 (FGZC 623, adult male), ZSM 327/ 2004 (FGZC 624, adult male), ZSM 329/2004 (FGZC 626, adult male), all collected at Montagne d'Ambre, 700– 1000 m a. s. l. on 19–23 February 2004 by F. Glaw, M. Puente, R. D. Randrianiaina & A. Razafimanantsoa.

Diagnosis. Uroplatus finiavana sp. nov. differs from all other taxa of the U. fimbriatus species group (U. fim-briatus, U. giganteus, U. henkeli, U. sikorae and U. sameiti) and U. lineatus by its smaller size (adult SVL 52–65 mm versus at least 85 mm) and lack of lateral membranous fringes on any part of the body and limbs; and from

Uroplatus alluaudi, U. guentheri, U. pietschmanni, and U. malahelo by its smaller size (adult SVL 52–65 mm ver-sus 69–81 mm), laterally compressed body with a vertebral keel, and more triangular head.

The new species is most similar to the other species of the Uroplatus ebenaui group (U. ebenaui, U. phantasti-cus and U. malama).

Uroplatus finiavana differs from U. malama by its smaller body size (SVL 52–65 mm versus 71–77 mm); shorter (TaL/SVL 0.42–0.65 versus 0.61–0.72) and narrower tail (TaW/SVL 0.05–0.14 versus 0.18–0.24), an unpigmented oral mucosa (versus blackish oral mucosa), and the presence of spines on the body, head, limbs and tail base (versus their absence).

Uroplatus finiavana differs from U. ebenaui by its slightly larger body size (SVL 52–65 mm, mean 58.2 mm versus 45–63 mm, mean 54.3 mm), longer tail (TaL/SVL 0.42–0.65 versus 0.22–0.40), longer forelimb (ForL/SVL mean 0.42 versus 0.38), posteriorly curved interorbital ridge (versus almost straight), the median posterior exten-sion of the ridge (1.6–3.0 mm versus 0.0–1.1 mm), more extended neck triangle (neck-triangle length 7.3–14.2 mm versus 1.7–3.4 mm), and by its non-pigmented oral mucosa (versus blackish oral mucosa).

Uroplatus finiavana differs from U. phantasticus in its slightly smaller body size (SVL 52–65 mm, mean 58.2 mm versus 52–76 mm, mean 60.3 mm), shorter (TaL/SVL 0.42–0.65 versus 0.62–0.76) and narrower tail (TaW/ SVL 0.05–0.14 versus 0.16–0.20), and its unpigmented oral mucosa (versus blackish oral mucosa).

In addition, U. finiavana differs from all other species of the Uroplatus ebenaui group (see Figures 6 and 7) and from all other species of Uroplatus (Greenbaum et al. 2007; Raxworthy et al. 2008) by its substantial degree of genetic differentiation.

Description of the holotype. Male specimen in good condition, with original tail attached to the body and par-tially everted hemipenes. Measurements and counts of the holotype: snout-vent length 59.0 mm, tail length 35.3 mm, maximum tail width 5.7 mm, and right forelimb length 25.5 mm. Head triangular in dorsal view, postorbital region (measured from posterior border of eye to anterior border of ear opening) of similar length to that of the snout (from anterior eye border to snout tip); snout sloping strongly and continuously downward anteriorly; snout depressed, short (1.3 times longer than eye diameter); canthus rostralis indistinct; eyes large, bulging slightly above dorsal surface of cranium, directed laterally, pupil vertical with crenate borders; ear opening very small (horizontal diameter 0.6–0.8 mm), its opening facing posterolaterally, but also posteroventrally (ear opening clearly visible in ventral view); nares laterally oriented; body somewhat laterally compressed, without lateral fringes; limbs well developed, without fringes, forelimb reaches beyond tip of snout and almost to the groin (forelimb length/axilla-groin distance 98%), hind limb reaches beyond axilla (hind limb length/axilla-groin distance 132%); tail 60% of snout-vent length, membranous borders of the tail narrow (maximum width on each side 1.6 mm) and completely absent from the distal tip of the tail. Nares separated from each other by eight small granular scales, from the first supralabial by one scale, and from the rostral scale by two scales; first supralabial taller than others; rostral entire, much wider than tall; mental scale very small, not differentiated from infralabial scales (total series of infralabials, left and right, plus the intervening mental scale yields a count of 35); no enlarged postmental scales or chin shields; dorsal and ventral scales of head, neck, body, limbs, and tail small, granular, juxtaposed and largely uniform in size, except for the irregular lines on the head and body which consists of series of slightly enlarged scales. Two curved lines starting at the posterolateral parts of the head converge in the neck forming a V-shaped pattern (neck triangu-lar line). A curved, moderately distinct and posteriorly directed line is present between the eyes and connects the supraciliary spines. Several spines on the posterior parts of the head, on hind limbs and a single pointed flap on the posterior portion of each upper eyelid; upper eyelid becomes broader as it approaches the parietal region of the head.

Coloration: All dorsal surfaces are brown in color except the tail which is light brown mottled with grey-brown and with few dark spots on the neck and one larger dark spot on the middle of the back after more than 6 years in alcohol. The mottling of the body is configured as a series of fine posteriorly directed markings along the dorsum. Tail is uniformly light brown. Two whitish spots present below each eye. Chin and throat beige with a dis-tinct blackish wide V-marking that merges together to form an indistinct dark longitudinal line that fades on the throat. Venter is light grey with few scattered small darker spots. Lower hind limbs and feet are slightly darker than other ventral surfaces. Postpygal portion of tail is marked with a whitish spot. Oral mucosa unpigmented.

Variation. The dorsal ground color varies from light beige (ZSM 323/2004, 325/2004, 326/2004, 327/2004) to reddish-brown (ZSM 324/2004), to dark-brown (ZSM 322/2004) and grey (ZSM 329/2004). Dark spots are most evident in those paratypes with light beige ground color, whereas the other specimens are largely uniform in color-ation and lack prominent dark spots.

The ventral side of all paratypes is similar to that of the holotype except for two characters. First, the V-mark-ing on the chin of the holotype is absent from two of the paratypes (ZSM 322/2004 and ZSM 329/2004) or is indis-tinct (ZSM 1135/2003 and ZSM 1132/2003) and also, the scattered dark spots are absent from three paratypes (ZSM 324/2004, ZSM 322/2004 and ZSM 329/2004). One of the paratypes (ZSM 327/2004) shows distinct lateral dark line starting slightly under the axilla and continues to the groin. The presence of a white spot in the postpygal area is not constant and is only found in three of the eleven paratypes.

Color photographs most likely showing Uroplatus finiavana have appeared in several publications, including those of Andreone (1991), Glaw & Vences (1994: color photo 252), Böhme & Henkel (1995: Figures 11, 12, 13, 14), Svatek & van Duin (2002), Glaw & Vences (2007: 378) and Schönecker (2008: 125–126). The same pattern as those encountered in the preserved specimens is observed in living animals. In some specimens, the markings on the head and the dorsum are underlined with darker coloration. Whitish spots below each eye are sometimes absent or are reduced to a single one.


FIGURE 8. Comparative pictures of the body and mouth pigmentation in living and preserved specimens of the different spe-cies of the Uroplatus ebenaui group. (a) female of U. ebenaui (ZCMV13013) from Nosy Be, November 2009; (b) male of U. malama (ZCMV12218) from Andreoky (Andohahela area), May 2010; (c) male of Uroplatus sp. 2 from Tsaratanana, June 2010; (d) female of Uroplatus sp. 3 from Tsaratanana, June 2010; (e) female of U. finiavana from Montagne d’Ambre, Febru-ary 2003, live and preserved mouth pictures are not from the photographed animal; (f) male of U. phantasticus from Ranoma-fana, March 2004; (g) female of Uroplatus sp. 1 from Tsaratanana, June 2010.


FIGURE 9. Map of Madagascar showing the distribution of the evolutionary lineages of the Uroplatus ebenaui group. Only selected localities: (1) those mentioned in the text, (2) those for which molecular data are presented herein or are available from us (unpublished), and (3) Andringitra Massif which is the southernmost record of U. phantasticus vouchered by a DNA sequence (Raxworthy et al. 2008). The Zahamena locality for U. phantasticus bears a question mark due to the high genetic dif-ferentiation of this population, which leaves its actual specific identity in need of confirmation.

Hemipenis structure. Because the holotype does not have a fully extruded hemipenis, we used one of the paratypes ZSM 1133/2003 with fully everted hemipenis for the description (Figure 4). In common with the other species of the ebenaui group, the hemipenis bears two lobes that become more obvious towards the apical region. The calyx displays a protuberance with a honeycomb appearance within which the sulcus spermaticus is concealed. The latter has a smooth surface. One to four folds separate the protuberance from the apex, which exhibits several papillae that wrap a single structured pedunculus in each side. A more in-depth description and comparisons with other taxa will be presented elsewhere.

Etymology. The specific name is derived from the Malagasy word "finiavana" meaning initiative. We refer to the fact that we took the initiative to name the species following years of this taxon having been recognized as likely being distinct. The name is used as a noun in apposition.

Distribution. In several herpetological surveys carried out in 1994, 2000, 2003, 2004, 2006 and 2009, the new species was encountered in the rainforest of Montagne d’Ambre National Park between 700–1350 m a.s.l. and in secondary forest fragments of lower elevation close to the town of Joffreville, where both U. finiavana and U. ebenaui occur sympatrically. This protected area is completely isolated from the major rainforest blocks of north-ern and eastern Madagascar and covers a surface area of 182 km2 (Nicoll & Langrand 1989; Raxworthy & Nuss-baum 1994). D’Cruze et al. (2008) reported encountering this new species in the adjacent forest (Réserve Spéciale Forêt d’Ambre) that has a lower elevation range of 400–850 m a.s.l.; but we cannot exclude the possibility that these records may partly, or entirely, refer to U. ebenaui. Records of U. ebenaui from the neighboring rainforest blocks (Anjanaharibe-Sud, Manongarivo, Marojejy, Tsaratanana and Makira) might be referable, by DNA sequences and morphology, to other taxa.

Habitat and habits. The holotype was collected in primary forest in Montagne d’Ambre National Park during a night survey. The species is nocturnal, arboreal and is remarkably common within the national park, especially around 800–900 m a. s. l., where it is possible to encounter 10 individuals during a single night’s walk (for instance on the path “Voie des mille arbres”). Remarkably, most of the encountered specimens have lost their original tail. One individual was observed feeding on a cockroach.

Morphometric differentiation. Relative tail length and relative tail width were recorded as ratios relative to SVL; these values were calculated separately for males and females. Figures 5a and 5b show that the same pattern is evident in both sexes. The new species, U. finiavana, shows intermediate values of tail length and tail width rel-ative to two other described species of its clade (U. ebenaui and U. phantasticus), and the differences are statisti-cally supported (Mann-Whitney U test, pooling males and females; P < 0.01). Figures 5c and 5d compare neck triangle (NTL) and medial posterior projection of the interorbital ridge length (IRPL), revealing only two clusters; U. ebenaui differs from U. finiavana and U. phantasticus in its smaller values for these two parameters. Significant differences only exist between U. ebenaui and U. finiavana for NTL, IRPL and relative ForL (P < 0.01). There is no significant differentiation among the three species in relative hindlimb length.

Molecular differentiation and phylogenetic relationships. The phylogenetic tree inferred from the Bayesian analysis of the mitochondrial ND4 gene is shown in Figure 6. We emphasize that this tree should be considered as preliminary as it is based on comparatively short sequences of a single gene. Rather than a reliable phylogeny, this tree wants to depict genetic similarities and differences among the specimens studied. On this basis, the morpho-logical differences observed between the new species and U. ebenaui from the type locality (Nosy Be) and U. phantasticus are supported by molecular data. The tree displays a subdivision of the U. ebenaui group into seven deeply divergent clades, with U. malama occupying a basal position. Besides the described species, the tree con-tains additional deep genealogical lineages, which we here classify as candidate species according to the scheme of Vieites et al. (2009). First, based on our initial assessment of identity of the nominal species, we assume that four of these are included in the phylogram: U. ebenaui, U. phantasticus, and U. malama, plus U. finiavana sp. nov. as described herein. Secondly, Uroplatus sp. 4 from Anjanaharibe-Sud is an Unconfirmed Candidate Species (UCS) due to its high genetic divergence but insufficient morphological data to assess its status. Uroplatus sp. 1, 2 and 3 can be considered as Confirmed Candidate Species (CCS) because of their concordant divergence in both molecu-lar and morphological data (e.g.: tail shape) (see Figs. 5 and 8).

Uroplatus finiavana is the sister species of the clade consisting of high elevation CCS Uroplatus sp. 3–4, and this sister relationship is strongly supported [Bayesian Posterior Probability (BPP)= 1.00]. The cluster of U. finia-vana and Uroplatus sp. 3 and 4 is the sister clade of Uroplatus sp. 2 (strongly supported relationship: BPP= 1.00).

The analyzed samples of U. phantasticus do not cluster as a monophyletic group. Uroplatus phantasticus from Anjozorobe and Ranomafana constitutes the sister species of U. sp. 1, although this sister-relationship is not strongly supported, and U. phantasticus from Zahamena is basal to the clade composed of U. finiavana and U. sp. 2–4, but also in this case the phylogenetic relationships are not reliably resolved. The monophyly of the clade con-sisting of U. finiavana, U. sp. 1–4 and U. phantasticus from Zahamena, Anjozorobe and Ranomafana is strongly supported (BPP= 0.98). The phylogenetic tree is somewhat concordant with altitudinal range data: U. ebenaui from the lowland occupies a basal position, followed by the CCS of U. ebenaui, both U. phantasticus lineages and U. finiavana from mid-elevations, and by the UCS U. sp. 4 and the CCS U. sp. 3 from the high mountains in a deeply nested position.

Among the analyzed taxa, exceptionally large genetic distances (uncorrected pairwise distances shown in table 2) in the ND4 gene are evident. The highest observed value is 37.5% and is observed between U. ebenaui from Nosy Be and U. phantasticus from Ranomafana (individual URANO4.29), whereas the smallest value among major clades is found between the candidate species U. sp. 3 and U. sp. 4.

Uroplatus finiavana is highly divergent from all other lineages of the Uroplatus ebenaui species complex. The uncorrected pairwise genetic distances of U. finiavana from U. sp. 1–4, U. phantasticus from Zahamena, U. phan-tasticus from Ranomafana and Anjozorobe, U. ebenaui and U. malama ranges from 20.8 to 34.5%. These large genetic distances corroborate the separation of the newly described species from all other lineages and known spe-cies included in this study. Moreover, relatively large genetic divergences are found between individuals assigned to the same species. For instance, the uncorrected genetic distance between U. phantasticus from Ranomafana and Anjozorobe is 6.6% and the uncorrected genetic distance between U. ebenaui from Nosy Be and Berara is 18.2%.

As our goal was not a detailed phylogenetic reconstruction but a test for concordant differentiation in two inde-pendent genetic markers for integrative taxonomic purposes (see Padial et al. 2010), we analyzed the nuclear C-mos sequences separately from the mitochondrial ND4 sequences, and used a network analysis to better visualize the low degree of haplotype differentiation among the focal taxa. The haplotype network reconstruction based on the C-mos data revealed no evidence of haplotype sharing between U. finiavana and other species and candidate species of the U. ebenaui group (Figure 7). This congruence between mitochondrial and nuclear data supports the status of the new species as a separate evolutionary lineage and indicates the likely absence of gene flow among different lineages of the U. ebenaui group. Individuals from Tsaratanana assignable to Uroplatus sp. 1–3 in the hap-lotype network (Figure 7) are in grey and haplotype sharing among some of these candidate species occurs. For instance, Uroplatus sp. 2 from Tsaratanana (ca 2000 m a. s. l.), Marojejy and Makira’s western slope have the same haplotype as Uroplatus sp. 3 from the Andrevorevo part of the Tsaratanana massif (ca 1700 m a. s. l.).


Vieites et al. (2009) found that undiscovered taxa are phylogenetically widespread among Malagasy frogs, and this seems also to be true for the leaf-tailed geckos, the genus Uroplatus (see Figure 6). Our work is congruent with pre-vious studies on Uroplatus, especially the molecular surveys of Greenbaum et al. (2007) and Raxworthy et al. (2008). By using morphological characters and the information provided by different genes, Raxworthy et al. (2008) identified several candidate species from different places around the Tsaratanana Massif.

Uroplatus finiavana sp. nov.has been found to be morphologically differentiated by numerous researchers (Glaw & Vences 1994; Böhme & Henkel 1995). Tail shape and length, and the lack of pigmentation of the oral mucosa support its distinctness from U. ebenaui and U. phantasticus. Tail shape and size, in particular, are known to be valid taxonomic characters for the recognition of nominal species of Uroplatus. These characters alone strongly support the status of U. finiavana as an independent evolutionary lineage (Padial et al. 2010). In addition, both mtDNA and nuclear sequences provide concordant support for the distinctive status of U. finiavana, which alone would suffice for defining this species as new, following the genealogical concordance method of phyloge-netic species recognition (GCPSR) (Avise & Ball 1990). However, the concordance between the molecular and morphological data is the most convincing argument and leaves little doubts about the specific distinctness of U. finiavana.

The species diversification mechanisms that have been responsible for generating the current pattern of high species diversity and microendemism in Madagascar are a matter of intense debate (Vences et al. 2009). Although no detailed conclusions can be drawn from the limited data available for the species of the genus Uroplatus, it is still possible to outline a plausible, hypothetical scenario of diversification in the U. ebenaui group on the basis of our phylogeny and taxonomic conclusions. The basal position of U. malama and of U. ebenaui in our phylogram indicates an initial split between southern and northern lineages. The ancestral lineage in northern Madagascar fur-ther diversified, with one lineage (corresponding today to U. ebenaui) becoming adapted to lower elevation habi-tats and one lineage colonizing the rainforests at mid-elevations (see Figure 6). These mid-altitude specialists dispersed, and the southern populations became isolated and diverged into the U. phantasticus lineages, whereas the northern populations diverged to become U. sp. 2 and U. finiavana. A further population of this northern clade also occupied higher elevations: the candidate species U. sp. 3 and U. sp. 4, representing the only nocturnal geckos in Madagascar occurring at elevations between 1500–2500 m a.s.l. This hypothesis requires further testing by addi-tion of more extensive DNA data from multiple genes which is currently underway (F. Ratsoavina, work in prog-ress), including partitioned analysis to take into account different substitution rates of codon positions and genes.

Several microendemic reptile and amphibian species occur in the Montagne d'Ambre massif: Uroplatus finia-vana, Brookesia tuberculata, B. antakarana, B. ambreensis, Furcifer timoni, Calumma ambreense, Calumma amber, Paracontias brocchii, Liopholidophis dimorphus, Rhombophryne matavy and Boophis baetkei. As men-tioned above, U. finiavana is commonly encountered within Montagne d’Ambre National Park where the habitat remains quite pristine. The only other known locality of U. finiavana is a secondary forest fragment outside the park boundaries near Joffreville, the closest village to the national park. Several individuals were encountered there in 2009. The latter forest is remarkably different from the habitat of the sampling localities inside the National Park, although we never observed the species in much degraded vegetation. Its broad distribution inside the pro-tected area, from 700–1350 m a.s.l. is suggestive of a rather high short-term probability of survival for the species. However, since its range is restricted to Montagne d’Ambre, and forests in this general area (e.g. in Fôret d'Ambre Special Reserve) are under strong pressure from deforestation, we propose that this species should be listed as Near Threatened in relation to the criteria of IUCN red list.


We are grateful to Angeluc and Angelin Razafimanantsoa, Marta Puente and François Randrianasolo for helping us with sampling and organization; Ivan Ineich (MNHN) and Jakob Hallermann (ZMH) for the loan of type speci-mens under their care; and Shannon E. Engberg, Gabriele Keunecke, Meike Kondermann, Runhua Lei, and Eva Saxinger for help with labwork and sequence analysis. We thank the Malagasy authorities and MNP (Madagascar National Parks) for issuing research and export permits, and to the team of MICET in Antananarivo for logistic support. Financial support was provided by EAZA to FG, the Volkswagen Foundation to FMR, RDR, FG and MV, the Deutscher Akademischer Austauschdienst to RDR, and to FMR by Omaha’s Henry Doorly Zoo.


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,,Keeping and breeding the Xenagama batilifera,, by Kamiel Hamers

Keeping and breeding the Xenagama batilifera

(this article is based on my own experience, combined with reading everything available about this species; by Kamiel Hamers)


The Dwarf Shield Tailed Agama (Xenagama taylori) is a small agamid originating from arid regions of Northern Africa and Somalia. The tail resembles a miniature "shield", hence the common name, Shield Tailed Agama. They are very hardy lizards which adapt well to captivity. One unique characteristic we have discovered is that they will dig a shallow tunnel and block the entrance with their tail at night in hopes of deterring any would be predators. The Xenagama taylori is a very personable lizard with many interesting habits and characteristics, many of which resemble the very popular bearded dragon (pogona vitticeps), which makes them an excellent choice as a new breeding project or a pet lizard.

Xenagama taylori grow to an adult size of 3" to 3 1/2" in length and weigh up to 20 grams. Hatchlings range from 5/8" - 1" in length and weigh as little as 3 grams at birth. Coloration varies from a dull sandy brown to a brilliant "brick red" body color with varying amounts of black speckling.   Small amounts of partial white spotting is noticeable on young specimens, but seem to fade with age. Adult males display a brilliant neon blue chin coloration when "fired up", usually during breeding behavior, male combat or a heightened state of alertness. Some females will also show varying amounts of blue chin coloration, but is very nominal compared to that of the male.

Housing & Substrates:

Communal cage set-up

We have tried several housing and substrate combinations and have had varying amounts of success with each. At first sand was considered to be the substrate of choice, due to the belief that this most resembled their natural environment, but was quickly discarded because of the lack its tunneling ability. Our next choice was a combination of cypress mulch and sand, where sand was placed on one end of the enclosure and mulch on the other. The lizards did not seem to prefer either end over the other, but we did notice they spent the night time hours buried under the cypress mulch. The problem with this type of set-up was their food items would bury themselves in the mulch and go unnoticed, and after several days and feedings the enclosure would have hundreds of crickets or mealworms running around stressing the lizards. We have found that garden soil dug from outside works the best. It has great compacting abilities, which allows for tunneling, is easily cleaned or replaced, and does not offer the food items a place to hide and go uneaten. Substrate should be between 3" to 5" deep as Xenagama taylori are great diggers, they are often seen digging multiple tunnels under and around the rock slabs and/or driftwood pieces supplied for basking. Their basking area usually consist of a large piece of driftwood and/or slabs of rock or brick. Temperatures at the basking site range from 90 - 110 degrees Fahrenheit which is supplied by an overhead lamp with a reflective shield. We use a 75 watt bulb placed 10" - 12" above the highest point of the basking rock. The cool end of the enclosure is approximately 20 degrees cooler, which allows for thermoregulation. Water is offered continuously in a small water dish about 1" deep, and is buried to where the rim of the dish is even with the top of the substrate. The lizards occasionally drink from the water dish, but we noticed they seem to prefer to drink droplets of water which form on the sides of the enclosure and the basking spots as a result of being misted every other day.

Rock slabs provided for basking.

Additional Lighting:

There are some concerns about the amount of UV-B and UV-A light requirements of shield tail agamas. At this time we offer little or no additional lighting except for an incandescent bulb used for basking. To date, we have not seen any ill effects from the lack of natural sunlight, although a vitamin/mineral supplement with vitamin D-3 is offered in hopes of fulfilling these needs. Long term deprivation of direct sunlight may prove detrimental, so some exposure is suggested, even if it consist of only a few hours per week.


The diet of the shield tail agama very much resembles that of the bearded dragon, consisting of small to medium crickets, mealworms, occasional super worms and a varied "green" leafy salad. Crickets or mealworms are offered daily in amounts which will be eaten over the period of the day. This is generally 3 - 5 food items each. We also dust the crickets/mealworms with a vitamin/mineral supplement such as "Miner-all" or "Reptamin" twice per week. Although, it is not known what importance leafy matter plays in their diet, finely chopped dark greens and vegetables are offered twice a week in small amounts and are misted with fresh water. Greens offered are: Collard greens, mustard greens, romaine lettuce and endive.   Vegetables consist of shredded yellow squash, zucchini and carrots. Sub-adult taylori and gravid females seem to relish the greens, but others tend to turn their noses at the offering.


Males (on left) can be identified by the enlarged femoral pores and a yellowish waxy substance present around the pores. Female on right has very small femoral pores which are barely noticeable.

Sexing of hatchlings and young juvenile shield tails is very difficult if not impossible. Sub-adult and adults can easily be sexed by examining the femoral pores present just above the ventral opening. A males femoral pores are very pronounced and secrete a waxy substance which is dark yellow in coloration. The waxy substance is not present on females, and the femoral pores can barely be seen.

Breeding & Egg Deposition

Male (left) showing typical blue coloration on chin.         Gravid female (right) shows increased girth as eggs develop.

First thought to be a solitary animal, we kept our lizards separate from each other except during breeding trials. The animals were brumated for a period of two months from November 15th to January 15th, where temperatures were kept at night time lows of 65 degrees and daytime highs of 80 degrees Fahrenheit, and a photoperiod of 8 hours. Food was offered once per week in smaller amounts than normal, with water available at all times. At the end of the brumation period, temperatures and daylight hours were slowly increased along with their regular feeding regimen. By February 1st they were back to normal feeding schedules and had a 12 hour daylight cycle.   Males were introduced to the single females one at a time, but no breeding activity was noticed. Thinking that they may be a communal breeder, we set up larger enclosures consisting of one male to four or five females. Breeding behavior was noticed almost immediately. The male would "fire up" his chin to the brightest neon blue we have seen yet, and commence to head bobbing erratically and "doing push-ups". The male will chase the females around the enclosure and literally "wrestle" the female while attempting to breed. The first time we witnessed this behavior, we thought we mistakenly placed two males in the same enclosure and they were fighting, but upon closer inspection we discovered they were actually in the act of mating. Males have also been seen copulating with multiple females over the course of a single day. From our experiences, multiple males or male combat is not required to induce breeding behavior.

Approximately two weeks after the first successful copulation, the females start to show signs of being gravid. The abdomen increases in size and bulges start to appear from the eggs forming inside. On several occasions, gravid females were pulled from the colony and placed in an egg laying chamber, which consist of approximately 10" - 12" of tightly packed soil, but failed to dig a nest and lay their eggs. So they were placed back in with the colony, thinking that they were not quite ready to lay. We then noticed that the gravid females started digging furiously after being sprayed with water during their every other day mistings. It seems that the females prefer to dig and lay their eggs after a simulated rain, so we started misting the egg laying chambers heavily to induce the females to deposit their eggs. The female will dig a deep tunnel, approximately 8" - 10" deep and deposit 5 to 8 small white eggs. After deposition, she will completely fill the tunnel and compact the dirt with her nose. Once finished there are no signs of any tunnels or eggs being deposited. After the female has deposited her eggs, we remove her from the egg laying enclosure and soak her in a container of water approximately 1/2" deep for 30 minutes so she can get re-hydrated, then place her in an enclosure by herself for a few days to recuperate from egg laying. After a couple days she is reintroduced to the colony. We have females which have already deposited their first clutch of the season become gravid for a second time, confirming the belief that they lay multiple clutches during a single season. The number of clutches per year is still unknown, but we believe they are similar to bearded dragons, and can deposit up to 4 or more clutches per season.

Egg Incubation:


Usually 6 to 8 small eggs are deposited in a 10" to 12" deep nest. Incubated at 82 to 84 degrees farenheit, the eggs will hatch after 45 to 50 days.

The eggs are carefully excavated from the egg laying chamber and placed in a perilite/vermiculite mixture and placed into an incubator calibrated to 82-84 degrees Fahrenheit. The incubator is kept at 100% humidity by keeping a container of water inside the incubator, and periodic misting with a spray bottle. After a 45 to 50 day incubation period the eggs start to darken in color and usually hatch within 48 hours.   The young xenagama are left in the incubator for 24 hours to allow the yolksac to be absorbed, and then moved to a small enclosure and kept on a paper towel substrate.   We mist the newly hatched lizards twice a day to keep them hydrated, and offer pinhead crickets after 2 or 3 days of emerging from the egg. Young Xenagama taylori grow fairly rapidly, and will double in size in the first two months. Size of the food items are increased as the young taylori grow, and greens are introduced at about one month of age. We keep the hatchlings and juveniles in communal set-ups identical to that of the adults. With the fast rate of growth, we believe sexual maturity is reached within the first year, but do not actually attempt breeding until well into their second year.

Captive born 2001 hatchlings at three months are approximately 1 3/4" long.

Although the exact husbandry of keeping Xenagama taylori is not yet completely known, we are well on our way to understanding this unique species.   Successful captive breedings will become more common and will help to promote the shield tailed agama as an exciting and interesting lizard kept by hobbyist. We urge hobbyist who have had success keeping this species to share their husbandry techniques, and help promote the species.

If you have any questions about or suggestions on keeping Xenagama taylori, please contact Terry McGleish at Glades Herp during normal business hours.

,,Breeding Uroplatus fimbriatus,, By Erik Strait

Breeding the Giant Leaf-tailed Gecko

Uroplatus fimbriatus has stringent care requirements

By Erik Strait

September 6, 2011

Uroplatus fimbriatus is a large, nocturnal gecko. It is the largest of the 12 known Uroplatus species, with an average total length of approximately 11 to 11.5 inches, with a snout-to-vent length of about 7.5 to 8 inches. Photo credit: Erik Strait

Commonly called the giant leaf-tailed gecko, Uroplatus fimbriatus was once imported into the United States by the thousands, but the number of imports is now very limited due to much-needed restrictions that protect wild populations. This is why its captive propagation is so necessary, and why this magnificent gecko should be kept only by experienced hobbyists, as its care requirements are more stringent than other popular pet geckos, such as leopard and crested geckos.

Uroplatus fimbriatus is a large, nocturnal gecko. It is the largest of the 12 known Uroplatus species, with an average total length of approximately 11 to 11.5 inches, with a snout-to-vent length of about 7.5 to 8 inches. It has a relatively flattened body, large eyes due to its nocturnal lifestyle, and very muscular legs it uses to leap from tree to tree within its tropical rain forest habitat along the eastern coast of Madagascar. Individuals can most often be found in trees, normally on branches or tree trunks within 10 feet of the ground. Wild leaf-tails appear to prefer smaller trees and branches that measure about 11.5 to 6 inches in diameter. Their cryptic body shape - they're also called "fringed" leaf-tails because of this - as well as pattern and coloration provides camouflage against branches and bark, and the geckos will shimmy around a branch or tree trunk to remain outside of a perceived predator's line of vision.

Leaf-Tail Care

Set up properly, giant leaf-tailed geckos can be prolific breeders. A hatchling or juvenile can be kept in an enclosure measuring 12 inches long, 12 inches wide and 18 inches tall. Single adults can be housed in cages measuring 18 inches long, 18 inches wide and 36 inches tall, and pairs or trios should be kept in a cage measuring 24 inches long, 24 inches wide and 48 inches tall.

I use paper toweling as a substrate for all my hatchling leaf-tails, to reduce the risk of accidental ingestion and impaction. It's easy to clean, too. I provide multiple layers to create some cushion, because Uroplatus are enthusiastic feeders that may bang their snouts against the substrate while pouncing at prey items. Substrate for juvenile and adult leaf-tails is a peat moss/soil/coco-fiber mixture, but coco-fiber alone can suffice, too.

Cage decorations and cover must be included but do not overcrowd the cage. The geckos should have space to jump to and from branches and the sides of the enclosure. I provide PVC for my geckos to perch on during the day and to climb at night. It's easy to clean and readily available. I also provide natural tree branches (be sure they are free from contaminants) and keep one or two large pieces of cork bark in the cage to provide other climbing and hiding opportunities, as well as rough surfaces to aid in shedding. Plants are also needed; I use large plastic plants with large leaves.Click image to enlarge

I provide PVC for my geckos to perch on during the day and to climb at night. It's easy to clean and readily available. I also provide natural tree branches (be sure they are free from contaminants) and keep one or two large pieces of cork bark in the cage to provide other climbing and hiding opportunities. Photo Credit: Erik Strait.

Giant leaf-tailed geckos like to be kept fairly cool. I keep the room's air temperature around 75 degrees Fahrenheit. Over the cages themselves, I use a 60-watt bulb that keeps the temperature at the top of each cage around 80 to 82 degrees. The geckos bask toward the top of the enclosure during mid day and move down to cooler locations once they reach a suitable body temperature. At night, I drop the temperature in the room to between the low 70s and high 60s.

I provide UVB lighting because even though Uroplatus are nocturnal, wild specimens do have some access to natural sunlight. My lighting schedule varies depending on the season - it's shorter during the "dry season" I use to induce breeding - but in general, I use a 12-hour-on/12-hour-off schedule.

Given the tendency of Uroplatus to dehydrate in captivity, misting and humidity are vital to their survival and breeding success. They will rarely drink out of a standing water bowl, but I provide one just in case. I mist them heavily every night, spraying cage decorations and the enclosure's sides to provide drops of water for the geckos to lick up. Misting, of course, helps maintain humidity, too.

Feed to Breed

A varied diet is crucial for leaf-tails to thrive. Every other night, I offer each adult four to six large crickets, as well as some additional insects (roaches, superworms, waxworms, etc.). To stop worms from burrowing into the substrate, I hand feed them to my geckos. During the breeding season, my females readily accept pinky mice, and these help the females to produce well-calcified eggs.

Calcium and vitamin supplements are also needed. I provide Miner-All with vitamin D3 to my geckos that are not kept under UVB, and Miner-All without D3 for the ones that are. I also dust insects with Herptivite multivitamin powder at least once a week.

Breeding the Best

When choosing breeding stock, always buy captive-bred leaf-tails, mainly because they are hardier and have better survival rates then wild-caught geckos. Only experienced keepers should consider adding wild-caught animals to a breeding colony. Not only will they require a quarantine period, they often arrive dehydrated and in rough condition. Without the proper supervision, they can quickly perish. Imports need treatment for both internal and external parasites, and because the medication can easily be overdosed, I recommend that only a veterinarian familiar with reptiles administer it. If you have no choice but to purchase wild-caught specimens, find a dealer who has had the leaf-tails for a few months, to ensure that they are eating and drinking, and that they have begun adjusting to captivity.

Breeding this magnificent species in captivity can be tricky. Keep them housed in a cage that provides a sufficient amount of room and that most closely replicates their natural habitat. For optimal breeding success, go with a breeding trio of one male and two females, or a male/female pair. Do not keep sexually mature males in the same enclosure. They will attack each other.

To initiate breeding I employ a cool-down period. Starting in September, I gradually drop the ambient temperature in my Uroplatus rooms to between 65 and 70 degrees. I also decrease misting to a lighter mist, which ensures that the geckos don't dehydrate while still simulating a dry season. I reduce the photoperiod to nine hours of light, after which any UVB lights, as well as the 60-watt overhead bulb on each enclosure, is turned off. I maintain these conditions for approximately three to four months, and then I gradually increase the ambient room temperature back to the usual 75 degrees, and slowly return misting and photoperiod to normal levels. These three variables - lowering the temperature, decreasing misting and changing the photoperiod from 12 hours to nine - have increased my success rate in producing leaf-tail offspring.

The geckos do not typically mate during the cool-down period; it is a time for them to rest from the previous breeding season. Mating activity generally resumes when I return the temperature, misting and light schedules to normal levels. Eggs may be laid from January until the beginning of September, prior to the next cool-down period. The majority of eggs are laid from March through August.

Gravid females will typically lay one or two eggs under the substrate approximately three or four weeks after a successful mating. Watch for signs of digging. It is crucial to extract the eggs carefully for artificial incubation. Doing so allows you to keep temperatures and humidity stable. It's safer for the eggs, too, and can prevent them from being overturned, oversaturated with misting water, and possibly damaged by feeder insects left in the adults' enclosure.

The egg (or two eggs) is incubated inside an incubator, in a plastic deli cup with a 1:1 ratio of perlite to water. I place eggs on cut pieces of foam to avoid direct contact with the incubation medium. This soft, 1-inch-thick, green foam can be purchased at any craft store; it is often used for chair cushions. I cut it into 2-inch-square pieces, and then scoop a divot out of the top of each piece. The egg nestles inside the divot so it doesn't rotate or come in contact with the moistened perlite.

If incubated at 78 degrees, eggs generally hatch after 90 to 120 days. I have found 78 degrees to yield the most successful hatchings.

As mentioned, the hatchling(s) can be kept in an enclosure measuring 12 inches long, 12 inches wide and 18 inches tall. They usually will not eat for the first few days while they absorb their remaining yolk. After a few days, I offer small crickets. I always watch new hatchlings to make sure they eat, and won't leave them to eat on their own until they seem to have gotten the hang of it. I typically offer a hatchling three or four crickets every night. Once a hatchling giant leaf-tail reaches juvenile status, at a length of about 5 or 6 inches, I offer food every other day and continue to do so from that point on.

Well Worth the Effort

I have worked with Uroplatus for more than a decade, and after considerable trial and error, I now know what works best when trying to keep and breed them. Captive-bred giant leaf-tailed geckos are still fairly hard to find, and those that do become available are often snapped up without ever being listed on any dealer price lists. As mentioned, the captive propagation of Uroplatus fimbriatus is vital to keep this magnificent species in private collections, as well as to help reduce the number of leaf-tails taken from the wild. I hope that this article motivates you to keep and breed these amazing geckos for yourself.

Varanus varius (SHAW 1790) Lace Monitor

Varanus varius (SHAW 1790)
Lace Monitor

 Varanus varius belongs to the subgenera Varanus. The major coloration on top is a deep blue with numerous scattered white, cream-colored or yellow singles scale. Some of these scales form groups and greater spots or areas. In young animals these bright areas are ordered between darker areas but this pattern of crossbands fades in age. In some animals alternately black and yellow ribbons appear on the body, the tail and on the limbs (belli-phase). This banded phase occur in an area of the northern New South Wales and the adjacent southern Queensland.
Normaly there are black bands seen across the snout, the chin and the throat. The tail is marked with irregular yellow crossbands. The head scales are moderate large and smooth, while the Supraocularia is uneven. The nostril is situated at the side, and is approximately twice as far to the eye then to the tip of the snout. Approximately 200 scale rows are around midbody. A clearly visible number of extended scales, which form a comb-like stucture is ordered on the inside of the base of the fourth toe. The tail is squeezed together strongly at the side unless at the base with a clearly visible double keel on the top side. It is approximately 1.8 times as long as SVL. The average total length is 150 cm but also animals which grew longer than 200 cm are described.

varius05 varius010 varius011

 Distribution, habitat and behavior

  The distribution of Varanus varius extends from the Cape York Peninsula in northern Queensland along the east coast of Australia to Victoria in the south. This monitor species also occurs in the southeastern South Australia (MERTENS 1942d, 1958, KEAST 1959, WORRELL 1966, BUSTARD 1968, SWANSON 1976, STORR 1980, STORR et al. 1983b, WILSON & KNOWLES 1988, HOSER 1989, COGGER 1992, EHMANN 1992).

In the wild, the Lace Monitor normally lives on trees. HORN (1980) however reported during his study, that this monitor frequently searches for food on the ground. According to his size carrion also is part of its food source (KENNERSON 1980, WARD & CARTER 1988). V. varius occasionally also feeds on bats (MANSERGH & HUXLEY 1985) and young birds, e.g. Tawny Froggmouth (KINGSTON 1980).

Keeping and breeding

HORN & VISSER (1989) described the behaviour of Varanus varius in captivity. Because of the size and the activity of thise monitor species a keeping in captivity is actually only possible for zoos or private persons, which can provide a lot of space.
In the wild the Lace monitor drops its eggs in termite mounts. After the termites have repaired her damaged mount again the eggs then have both, perfect incubation conditions (temperature and humidity) and the protection from eating enemies. A very interesting breeding biology is that at the end of incubation period the female returns to the termite mount, opens it, and makes it possible for the hatchlings to ermerge from the termite mount (BARTON 1991).
Because Lace Monitors are perfect tree climbers, you have to offer the animals an enclosure with the possibility to. Therefore some stable, vertical or horizontal tree-trunks are part of the basic furnishing. HORN (1991) used as ground cover a mixture of earth, peat moss, and gravel. Since V. varius is also a very good dugger, the layer of soil should should be fairly deep. An occasional humidification of the substrate for increasing the humidity has to be recommended. Because the distribution area of the Lace Monitor reaches from the tropical north of Queensland to the mediterranean climate of Victoria, the animals can accept a big variety of different climates. Unfortunately, in most of the time you do not know from which locality and from which climatic zone your animals originate. HORN (1980) gives a good summary of some of these parameters and of the biology of V. varius in the wild.
Till now, captive breeding of Varanus varius have been described already repeatedly. The most frequent successful breedings happened in Australia, where the animals were housed under free conditions (BREDL & SCHWANER 1983). HORN (1991) reported the first successful breeding outside Australia. The female dropped two clutches of eggs with 5 or 7 eggs respectively, always 4 weeks after observed matings, in a wooden egg-laying box. The eggs were incubated in a special incubator (Motorbrueter) without substrate in a steam saturated atmosphere (BROER & HORN 1985). At a temperature of 29°C altogether 11 young animals hatched after 226-243 days. The offspring measured between 32.7 and 37.2 cm and weighed between 28.4 and 40.8 g. The rearing of the babies did not cause any problems. They grew well at feeding with fresh born mice.
VISSER (1996) reported on a further captive husbandry with the offspring of the HORN-animals at the Zoo of Rotterdam. This is therefore the first captive husbandry in the second generation of this monitor outside Australia.
A special success is reported by K
RAUSS &HORN (2004). From on egg 3 offspring hatched. All babies survived and developed well. Only the body mass was a little bit lighter than in the other "regular" hatched siblings.


,,Abronia, Captive Husbandry and Breeding Notes,, by Jason Wagner


I’ve been keeping Abronia collectively for about 6 years, but have only been successful with breeding them for three years, from 2003-2006.  I owe a great deal of gratitude to a professor in the Pacific Northwest who teaches Biology and shares my passion for working with the Abronia genus, but he prefers to remain anonymous.  He was very helpful to me early on in providing basic husbandry guidelines for keeping this beautiful lizard species.  My goal in documenting these notes on keeping Abronia is primary out of respect for the animals.  I want to share my observations so that others who may have the opportunity to keep these lizards will at least have a reliable reference point from which they can work to gain there own experiences.  Abronia have adapted to a unique environment in the niche habitats they occupy in the wild, and as a result their captive care and maintenance does take some special attention to detail, if one is to become successful working with them.  I try to maintain a level of humility with the entire subject of captive husbandry, because in effect, all we are really attempting here is to mimic in as many ways as possible, what they would have in the wild (obviously without the dangers of predators and preferably without parasites).  Try as we might, there is no way to duplicate their native environment exactly, but we can get close enough to be effective in producing them on a sustainable basis and in my book that is an achievement worthy of the effort.

                    abronia taeniata   abronia graminea

                          Abronia taeniata, male                          Abronia graminea, male

Brief Natural History

Abronia lizards are found in several unique habitats but primarily inhabit cloud forests at high elevations in Central America, mainly in Mexico and Guatemala.  They are also found in some instances in Oak scrub and Pine forest areas, also at high elevations.  There are about 26 species known to Science depending on who you ask, and they are live bearing, producing a single litter each year.  Because they are not common in their native ranges and are limited to small territories, there are some species which may have already gone extinct due to massive land clearing for agriculture or charcoal.  In the wild they are mostly arboreal and usually live high in the trees, although I have found specimens on the ground and in rotting wood, or hiding in large clumps of moss.  The trees they live on are generally large oak varieties, and are typically covered with moss, ferns, orchids and most importantly…bromeliads.  They use the bromeliads as a micro-habitat for humidity, water, and shelter.  They are found at high elevations, usually between 4000-8000 feet (1200-2500 Meters).  At these high altitudes, there is a great deal of temperature variance between day and night, which is an important element to consider in captive husbandry, as I have found they will thrive best in a situation where they are provided with a strong night time temperature drop.  The temps will range from as low as 40F and can be as high as 90F, but those are generally the extremes.  In the cold season where they live, night temps can reach below 40 Degrees F ( 5 Celsius) for short periods of time and it may even snow on occasion.  During these very cold seasons, the lizards have been found to den together in hollow trees full of rotting mulch, where the temps are likely around 50 F (10+°C).  I assume that they don’t stay out in the open air exposed to such cold temps.  The same principle applies for the hot season and high temps.  They will seek out cooler shelters, like thick moss growing over a large rock in the shade…which may be 70 degrees F (20°C), when the ambient air temps are hovering around 90F (32°C). They are most active at temps around 75-80 F (25-28°C), and that is when you will find them basking or searching for food, typically in the morning hours from 9-11 am. On a mild sunny day, they can be seen basking later into the afternoon, usually in the trees. I believe its helpful to bookmark a weather website and monitor temps as well as rain patterns for Puebla, Mexico or Huehuetenango, Guatemala, as both of these climates support Abronia in the wild.  Cloud forests by definition are commonly a misty, foggy, high humidity area and it is in this niche environment where Abronia thrive. These areas typically get a lot of precipitation, and on a frequent schedule.  The mild climate also supports a large diversity of insects and my guess from captive observation is the Abronia take advantage of this.  Breeding usually takes place from September – December and 7-14 babies are born typically from April-June.

habitat of abronia

Typical niche habitat of Abronia photographed in Mexican highland forest on a sunny day.

Enclosure Design

All-Screen cages are the best choice, as they provide adequate ventilation (think of a tree dwelling species) and the advantage of offering natural sunlight.  Males should be kept separate, and will typically fight if kept together. Babies will also fight if kept too crowded.  I have only seen females fight twice and it wasn’t too severe, mainly a nip to establish territory (Abronia vasconcelosii).  I keep my animals in large screen cages, heavily planted with bromeliads and orchids, and offer plenty of climbing limbs or branches.  The most important thing I give them is a 2-4 inch layer of long-strand sphagnum moss as substrate on the bottom of the cage.  This provides moisture, and a cool area to retire from heat. The moss should be kept moist in some areas, not all wet all over, and should be dry in some areas. Occasionally I like to let the moss dry out completely, since that does happen in their native environment too…short dry-spells. The moss also has been documented as having a natural anti-bacterial property and was even used in World War I to transport organs, etc. UV light is thought to be important, and I would agree.  I have noticed my lizards always look more brilliantly colored when they have had extended exposure to natural sunlight.  Abronia graminea for example will typically fade to a grayish teal color if kept indoors under incandescent  lighting with little temperature variance over a long period.  In nature, they are usually a brilliant emerald green. I provide natural sunlight by moving the screen cages outdoors when the weather is good, but when kept indoors I offer a low emission UV light, that doesn’t put out a lot of heat. I use Arcadia compact fluorescents, UVB 7% in the hotter summer months and then switch to Active UV Heat bulbs (or similar) in the winter. These lizards will not do well if they are kept hot like a lowland tropical reptile. The high altitudes where they come from fluctuate a great deal in temperature, but they almost never go over 90 F (32 °C), and I would not allow them to be exposed to weather under 45 F (7°C) if I can help it. I have recently started keeping the cages on a Concrete slab, as I find that when the temps get too hot in the daytime, the moist moss resting directly on the flooring of the cage that is on the cool concrete, provides a cool temperature gradient that may be useful to the lizards. It is very important to provide shady areas within the cage if the animals are being housed outdoors. This can be achieved with broad leaf plants or cork bark slabs sitting at an angle in the enclosure.  They will die if they are left in a hot cage left in full sun with no shade obstructions, and have no way to thermo-regulate from the pounding direct heat of the sun through the screen.


Food and Water

I make a special effort to offer a variety of insects. The favorites are large black diurnal crickets, followed by grasshoppers.  They will also take green caterpillars, like hornworms, snails, spiders, meal worms, soldier fly larvae (phoenix worms – high in calcium content), roaches, etc.  I often feed my Abronia by hand or with tweezers, as they can sometimes be a little pokey in their eating habits.  When feeding lizards that are almost purely insectivorous like Abronia, I believe its important to be aware that they do not have the liver or kidneys equipped to deal with a high protein diet, for example like a snake that eats only mice. They instead will generally eat bugs that are full of plant matter themselves, like an arboreal caterpillar, grasshopper or katydid.  When I offer crickets, I make sure they have eaten a good variety of leafy greens like kale, romaine lettuce, dandelions, and then I supplement that with fruits and vegetables.  I also like to provide the feeder insects with a mix of ground up whole grains as substrate…essentially to “gut-load” the lizards food items with decent nutritional content.  I do not recommend that Abronia be fed exclusively on store bought crickets that have been raised on chicken meal, which is a high protein diet for baby chickens.  An occasional pink mouse is probably OK as a food item, but I have never offered one, and would definitely not over do that.  I believe the variety in the insect diet plays some kind of important nutritional role for captive lizards.

I almost always spray mist daily the plants, enclosure sides, and lizards themselves if they are out.  Additionally, 2-3 times a week, I will provide a dripper cup that drips about one cup of water over a plant in the enclosure so the lizards can drink at their leisure for a period of about 15 minutes.  I use tap water that has sat out in an open bottle for 24 hours.  Another watering method that I really like and am currently testing, is to use a “misty mate” pressure spray bottle. This high pressure system emits a fine mist over a period of up to 20 minutes and is excellent for simulating a cloud forest environment.

 Breeding behavior and raising young

The Adults will typically breed in their second year, when they are full sized and sexually mature.  I have seen breeding activity as early as August, and as late as November.  The pair may stay “locked up” for as much as 24 hours.  Babies are born in the months of April to June, typically.  My females have produced 7, 11, and 13 babies at a time.  In my experience, females will not eat their young, but I don’t trust the males or other adults. Gravid females will develop “chalk sacs” on the sides of her jowls before giving birth, likely as a calcium supply. She will also gain weight, noticeably. Perhaps the toughest part of keeping Abronia is raising the babies. They have all the same requirements as the adults, except they are smaller and have less body mass so they are more susceptible to over heating and death from high temps. I have found that anything over 85 F, is dangerous for them. They also may have little battles with each other over space so it is best to keep them in small groups, like 2 or 3 to an enclosure. I’ve tried keeping them separate in small “kritter keepers”, which seemed to work OK but they didn’t have optimum exposure to UV lighting or ventilation.  However that was a good way to ensure they were catching and eating their prey. Finding and offering small insects on a daily basis seems to be very important for the first 3 or 4 months. This can be challenging.  I’ve caught small grasshoppers by hand, and hunted for little green inch worms in the lawn, and used a net to do field sweeps, plus offer the normal crickets and smaller meal worms.  I’ve also recently discovered soldier fly larvae (marketed and sold as “Phoenix worms”) which are a good size and move slowly enough to be devoured by the young.  Plus they are high in calcium which is probably good at this stage.  My best experience has been to keep the babies exactly as the adults in full planted screen enclosures.  I had to use aquarium silicone to plug the small holes in the corners of the screen cages so the little crickets and bugs wouldn’t escape.  It’s important to offer plenty of small insects daily, but don’t overwhelm them so that the crickets are constantly bothering them at night, for example.

abronia  abronias

Miscellaneous Notes

The number one killer in my opinion of Abronia, is allowing them to get too hot.  If you don’t want them to breed, maintain them at room temperature constantly, and don’t drop the temps down at night.  There seems to be a correlation with temperature variance and breeding success.  Be wary of ants that can scout and mount surprise attacks, particularly with babies.  I watch temps closely and keep my animals outside for about 5 months of the year, which I think contributes to their overall health, and reproduction.

Abronia are rare in the wild, and with their small native habitats under a considerable amount of pressure, in some cases to the point of devastation, I feel it’s important to do what we can to preserve them in captivity when we have that opportunity.  The best solution would be to establish wild preserves in their native habitat where they can be safeguarded from the land clearing practices, etc.  But as an alternate method of conservation, I see considerable value in learning how to establish them successfully in a captive environment.

We express our thanks to the author of this article Mr. Jason Wagner. To find more information about the captive management of these lizards please visit

,,Phrynosoma Asio Husbandry,, by Jeff Judd

This is a large, hardy species of Horned Lizard (HL) that does well in captivity. It has a good appetite and thrives on a varied diet. These are very sociable Horned Lizards, interacting to each other with head bobs, arm waves, and tail wags. They are aware of their surroundings, and can recognize humans as a food source; they will watch in anticipation as food is being offered, and will eat from your hand.



GHLs should be housed in a glass terrarium. Overall, it should be long and wide but not very tall. Shallow tanks allow good ventilation and prevent heat from building up. The top of the substrate to the top of the terrarium should measure around 12 inches. The substrate level can be raised to accommodate a taller tank. The length and width needed depends on the number of GHLs that will be housed. However, the bare minimum for one or two adults should be 36" long x 18" wide.

Benefits of Large Tank:

No terrarium is too large. Generally, the bigger the terrarium is, the better. With adequate space, intense lighting can be provided them, as well as a wider range of temperatures for GHLs to properly maintain their preferred body temps. GHLs will be able to properly feed, and exhibit more natural behaviors. Also, you can create a more realistic landscape that will better resemble their natural environment.

What to add:

The bottom of the terrarium should be covered with three to six inches of sand. The sand should consist of various sized particles. Giant Horned Lizards purposely ingest specific sizes of these particles to aid in digestion. Rocks large enough for the GHLs to bask on should be placed under the heat lamp to provide basking sites. The cool side of the terrarium should contain plants or branches to provide areas for climbing. Suitable plants include many spineless cactus, succulents, palms, grasses, and non-toxic houseplants. Giant HLs are semi-arboreal, and during periods of inactivity, they will climb onto the plants or dead branches, or seek shelter under half-cut tree bark, cactus hollows, or sturdy rock formations.


The 160-watt or 250-watt mercury vapor heat lamps specifically designed for reptiles work very well for GHLs. They provide adequate levels of UV radiation as well as produce heat, which are both essential to keep GHLs healthy long term. Fluorescent lighting can be added if additional light is desired. The lights should be turned on and off by a 24 hour timer. They should be set to come on 13 hours a day May, June, July and August, 11 hours a day March, April, September and October, and then 10 hours a day November, December, January and February.


The terrarium should be placed in a room that stays between 65 and 75 º F year round. The heat lamp should be placed at one end of the terrarium while the other end remains unheated. The height of the lamp should be adjusted so that directly beneath the bulb the temperature is around 115 º F after the bulb has been on for an hour. This will provide a range of temperatures allowing the HLs to maintain their preferred temperature. No additional heating devices are needed after the lamp turns off, so the night temperatures should be falling between 65 and 75 º F.

Diet and Nutrition:

Giant Horned Lizards should be fed daily. They should be fed in the morning, or the late afternoon when they are most active. If they are fed in the morning, they should be allowed to warm up for about an hour beforehand. The insects should be placed in the same area of the terrarium at each feeding, because the Giant Horned Lizards will recognize a feeding area and look for food there. An individual’s appetite depends on its size, the time of year, and stress level. Generally, it is best to continue to feed the HL until it has no more interest in food.


In captivity, ants, crickets, or roaches should make up most of the diet with the occasional mealworms and waxworms. All of these insects are available commercially. Make sure the insects are from a clean source, otherwise they can harbor disease, which can be passed onto the GHLs. Crickets, roaches, and mealworms should be fed baby cereal and carrots before they are fed to the GHLs. The size of the insects is the most important aspect of feeding; they should be no longer than the width of the GHLs head. Feeding your Horned Lizards large insects can cause them to regurgitate it the next day, become very sick, or die.


Ants (Pogonomyrmrex barbatus and Pogonomyrmrex rugosus) should be included daily. The ants available commercially from the genus pogonomyrmrex are eagerly accepted by GHLs. They should be kept in a large jar with a lid that has very small holes drilled into it. It's important to only put 3-5 ants in the tank at a time. Dumping large numbers in will cause the GHLs to panic, often resulting in the ants biting or stinging them. Immediately remove any GHLs, then the ants if this happens. If an ant latches on with its pinchers, it must be removed by crushing its head with tweezers then slowly pulling it off. Offer ants until the HL stops eating or showing interest. Remove any that are not eaten with forceps or large tweezers.

Other Insects:

Crickets (Acheta domestica) and roaches (Nauphoeta cinerea) can be offered every other day. They come in many sizes, and do well with both hatchling and adult GHLs. They can easily be gut loaded and coated with vitamin and mineral supplements. However, crickets can be difficult to catch unless their hind legs are removed. Mealworms can be offered once or twice a week. They are difficult to digest. The number offered needs to be closely monitored. No more than 3 or 4 should be offered at one feeding. The freshly shed mealworms are best. These are white in coloring, and should be fed to Giant Horned Lizars when they are available. Waxworms are large and high in fat, which may be harder to digest in large numbers. No more than one or two should be offered to adults, once a week.


Giant Horned Lizards should receive a good reptile vitamin supplement and a good reptile mineral supplement twice a week. They are sold separately, so they must be mixed together. Add equal amounts of both in a jar, then put enough of either the crickets or roaches in for one feeding. You should shake the jar until the insects are coated with the supplements, then feed them to your Horned Lizards.

Water & Humidity:

May through October it rains very frequently in areas inhabited by GHLs. During this time, the whole terrarium should be misted twice a week with a hand sprayer. The substrate should be dampened, but not too wet and should dry out between waterings. The humidity should be kept between forty and seventy percent. Thoroughly mist the head and nostril area of the GHL until you notice that it is drinking water during each misting. November-April the terrarium should be kept dry. If live plants are used in the setup they can be watered by lightly misting the substrate around their base. The humidity should stay below 40 percent. Water should only be given to the Giant Horned Lizards once a month during this time.


Outdoor Exposure:

If outside temperatures are in the range of the above Climate Graph, GHLs can be placed outdoors. Behavior and health of certain individuals seems to improve when they are exposed to natural sunshine. The outdoor enclosure should be constructed of wood, screen, or fiberglass. All outdoor enclosures should be completely covered to prevent escape, as well as protect the GHLs from predation.

When placed in screen enclosures, nervous individuals should be watched closely. It doesn’t take long for one to rub its nose raw. Make sure the GHLs always have access to shade. Shade can be provided placing plants in the enclosure or by covering part of the top with wood. In some areas, small ants are a big problem. In a matter of an hour, thousands of small ants can mob and kill Giant Horned Lizards. In these areas, outdoor time should be brief and closely supervised.


GHLs mate from May to June. If successful, the female will lay eggs sometime in July or August, 60-70 days after copulation. She will become very restless and walk back and forth across the terrarium looking for a nesting sight. After a few days, she will usually choose a moist area near the heat lamp to dig a shallow pit 1 to 2 inches deep then deposit from 10-28 eggs. They must be retrieved shortly after they are laid, or they will spoil from the heat lamp. Turning the heat lamp off will extend the time the eggs can remain in the substrate. The female will be emaciated after the eggs are laid, and she will require plenty of food and water.

GHL Eggs:


Carefully remove substrate over the nesting site with a spoon, or your fingers until the eggs are unearthed. Use the spoon to transfer the eggs one at a time to a plastic container with a lid using 1 to 2 inches moistened perlite or vermiculite as the incubating medium. The lid should have 6 to 8 small holes drilled in it to allow air exchange. The medium should be moist but not soggy. This is about 4 parts of the medium to 3 parts water by weight. Too wet of a medium will spoil the eggs.


Place the container in an incubator set with the constant temperature between 82-86ºF. The eggs will hatch 11-12 weeks after they were laid. Water should not be added during the incubation periodunless the medium becomes dry to the touch and the eggs collapse farbefore they are due to hatch. Eggs ready to hatch will also collapse,so testing the medium is crucial. Usually a few eggs from a largeclutch will spoil during the incubation period. These eggs shrivel andare attacked by mold. The hatchlings usually emerge within 2 days afterthe eggs start to collapse. Do not remove hatchlings half-way out ofthe egg. They will stay in this position for a long period of time toabsorb the yolk and also to adjust to breathing. Removing them tooearly will often result in the death of the hatchling.

Raising Hatchlings:


Giant Horned Lizard hatchlings should be raised the same as adults, with only a few modifications. They should be set up in groups of no more than four so that their food intake can easily be monitored. If raising large numbers of hatchlings, the glass terrariums can be substituted with plastic containers about 36 inches long by 18 inches wide by 12 inches tall. The hatchlings should be fed twice every day.

Food & Water:

The main foods of P. asio hatchlings should consist of ants, very small crickets and roaches, and occasionally small freshly shed mealworms. Remember that ants can sometimes intimidate hatchlings. If the ants are not eaten right away, they should be removed. Putting the ants in the refrigerator for five to ten minutes will slow them down,making them easier for the hatchlings to take down. You should be giving the hatchlings water and vitamin and mineral supplements every other day.

This article is reprinted from web-page

,,Care Guide for the Genus Uromastyx,, by Douglas Dix

We've included a detailed care sheet for the most common Uromastyx species. Please take the time to read it before purchasing your Uromastyx.


General Husbandry / Breeding

Most Uromastyx species currently in the U.S. seem to have fairly similar requirements so I'll lump them together for the purposes of this care sheet. Where the various species differ in their care requirements, I'll so note in the text. Please also look at care sheets posted at The Uromastyx Home Page for additional information on a variety of Uromastyx issues. Also check out The Uromastyx Forums. These are useful forums for posting questions and sharing information concerning Uromastyx.  Be careful taking advice posted on forums as a fair amount of inaccurate information gets posted by well meaning but inexperienced individuals.   Use your common sense and get more than one opinion before making any major changes in how you keep your animals.


First and foremost, Uromastyx are heat lovers, the ultimate heat lovers! They must have a basking site that reaches between 110°F and 120°F (surface temp). No, that's not a typo, one hundred and ten to one hundred and twenty F! This is actually easy to produce with a Zoo-Med or comparable reptile basking bulb (reflector or flood type bulb) shining over a smooth piece of slate or other suitable rock. Adjust the height of the basking light so that it heats an area at least as large as the whole body of a basking Uromastyx and make sure the light is placed high enough to prevent the animals from accessing it. Do NOT use hot rocks or similar "in-cage" electric underbelly heaters. These will not suffice and can cause serious injury to your animals. An under-the-tank heating pad is ok but only for supplemental heat.  The overhead basking light is still essential. You're aiming for a general background temperature around 100°F in the warm end of the cage, and the mid 80's°F in the cool end of the cage. This permits your animals to self-regulate their body temperature. Night temps should be much cooler, typical of their desert homes. Most people shoot for the low 70's in the summer, the upper 60's in the winter. Along with the basking lights, we recommend installing a UV producing bulb such as Zoo-Med's Reptisun 5.0's, Arcadia High OutPut UVB or  Mercury Vapor basking bulb. The usefulness of these bulbs is still debated and some breeders feel they are a waste of money ($20 to $30 ea. for Zoo-Meds, $35 up for Mercury Vapors), but the jury is still out.  UVB initiates the conversion of vit. D3 precursor into active vitamin D3, and in theory these bulbs produce enough UVB to stimulate this reaction. However, for this to be reasonably effective, the bulbs must be mounted within a foot or so of the basking animal. Also these bulbs gradually loose the ability to produce UVB with use and thus should be replaced annually to biannually. Look for a change from bluish white to a clear white glow with age, faint blue tint = good, white = worn out).  Some breeders choose to simply add vit. D3 to the diet and dispense with the bulbs. This approach also seems to work, but which is more reliable is still unknown.  Regardless of how well UVB impact Vit D3 issues, strong UVB exposure does produce more intense pigmentation in captive Uromastyx.   For some species, this alone may be sufficient justification to use the UVB-producing bulbs. In addition to UVB, these bulbs also produce UVA, which has been suggested to increase appetite and give desert animals a "psychological" benefit. Again the jury is still mixed with some swearing the bulbs help and others equally convinced there is no noticeable impact on behavior or health.  We use a mixed approach.  We feed low levels of Vit D3 to our animals (by dusting w/ Miner-AL brand mineral supplement w/ D3) while using Arcadia Hi UV compact fluorescent bulbs in all our cages.

Our data on bulb-generated UVB's effects are mixed. Some Uros seem to do better with the bulbs while others show no detectable differences with or without the bulbs. We have noticed better coloration in many Uromastyx exposed to strong UVB from Mercury Vapor bulbs or Arcadia bulbs and Uro pairs in cages with these bulbs tend (by coincidence or not) to be some of our better breeders. Uromastyx do detect the difference between normal "man made" light and sunlight and are unmistakably attracted to sunlight. Uromastyx raised outdoors in unfiltered sunlight are the most dramatically pigmented of all.  Of the available commercial reptile UVB lights, the Mercury Vapors bulbs seem a better choice in open topped cages, however the cost and excessively short life span (most rarely last 6-8 months for us) make them impractical for us. The current wattages/sizes available also put out excessive heat and so are unusable in our solid-topped Vision and ShowCase brand cages. The fluorescent bulbs have minimal heat output and come in many lengths and are thus more user friendly. Their useful lifespan however is often comparable to the Mercury Vapor bulbs.  We use clear infrared bulbs for heat in our ground pens and standard silver-backed reflector-type bulbs for our oak cages and "Vision" / Showcase brand pre-fab cages as the heat sources.  We currently use compact Arcadia UVB bulbs for our UVB source but have been experimenting  with other brands of Mercury Vapor bulbs for our ground pens.

For most of the year, we are looking to produce a 13 hour day and 11 hour night time period for all species of Uromastyx.  You can shorten this by a few hours during the  winter, but only if you don't mind the possibility of them cycling and going through breeding behaviors in the spring.  This is NOT an option if you have multiple specimens of the same sex housed together. Going through a yearly full brumation cycle does not appear to be essential to the long term health of most Uromastyx.  Seasonal variation in day length and background temperatures is probably a good idea, but  for most pet owners, don't go below a 10 hour day during the winter or 68° F night time low temperatures.


Opinions vary on the ideal bedding. It's a common misconception that Uromastyx prefer sand and come from a sandy environment. In fact they tend to avoid overly sandy locales in the wild, preferring clay/sand or gravelly-loam mixes, rocky outcrops or other soils better suited to holding a burrow without collapsing. If you use sand, make sure it is a natural sand (rounded edges) like beach sand or washed playground sand. Man made sand (from crushing gravel) has jagged edges which easily interlock, leading to gut impactions in animals that swallow it. We personally don't like sand and restrict it's use to only in the nest boxes.

We've tried bark, which the Uros enjoyed but the excessive dust produced was unacceptable and picking out fecal pellets was far too labor intensive. We then tried rabbit pellets (alfalfa), but the problems were essentially the same as bark but with more odor. We finally switched to high quality wild bird seed (predominately millet) in the mid 90's and have been extremely pleased with the results. The Uro's can snack on the seed throughout the day, it's generally dust and odor free, and sticks to fresh fecal pellets, quickly drying them. Seeds which the Uro's crack before swallowing are digested while uncracked seeds pass whole, acting as much needed roughage. The fecal pellets can be quickly sifted out of the cage with a 1/4" mesh hardware wire sieve (easy to make from a cat litter scoop), allowing us to easily maintain a large number of Uromastyx without needing additional hired help. The seed is good for several months per cage, then with one final cleaning, can be fed to our other livestock (Fallow deer) or wild birds. If the Uros drag damp sand into the bedding or pile bedding in the nest box, the seeds sprout.  As a side note, we've also looked into using calcium carbonate sand (Calci-sand, T-Rex), but other Uro keepers have told us the dust produced is too great to be acceptable bedding. The fine dust has a tendency to get into the eyes as well, potentially causing significant irritation/injury.  It is particularly dangerous to use around hatchlings and juveniles, getting into the lungs and causing severe desiccation. It tends to clump when damp and form semi-hard masses which potentially could lead to intestinal blockages. There is also some concern calcium-based sands act like giant anti-acid tablets, upsetting the digestion process. Ground walnut shell has also been suggested as a good bedding and is advertised as an attractive, digestible, odor free, safe bedding by the manufacturers.   While we agree it's quite attractive, it is otherwise a nightmare of a product.  Walnut shell is composed primarily of lignin which in fact is not digestible by vertebrates, and the crushing process basically leaves most the resulting pieces with jagged edges. These edges have been indicated in the deaths of several Uromastyx - necropsies revealing their stomachs had been extensively lacerated by ingested bedding.   All in all, we strongly suggest avoiding it. Similarly, ground corn-cob is too dangerous to use. While it has smooth edges, it's extremely hydrophilic and if swallowed absorbs water from the gut and greatly swells. This can easily lead to fatal impactions and as Uromastyx don't normally drink water, even small amounts inadvertently ingested can easily dehydrate smaller specimens.  Bed-a-Beast (shredded coconut husks) are used by some with good success, but again fecal pellets have to be removed one by one by hand and it tends to be quite dusty. We've tried it in our nest boxes but it readily molds and attracts gnat flies which can harm newly laid eggs, so we've had to eliminate it.  The chucky version of it works well as bedding for our Agamas and Tortoises and at least for them is an attractive, low dust, no odor bedding.

Note for all these bedding, make the depth very shallow -say 1/4 inch max.  For most situations, making the bedding deep enough to burrow in greatly complicates their care.  Use artificial burrows or hide boxes to satisfy your Uro's desire to burrow.  For hatchlings/juveniles under 6 inches total length we recommend bare tank bottoms or butcher's paper. Hatchlings are much more sensitive to ingesting dry, hard material so it's best to avoid the problem.


Uromastyx are burrowers by nature and must be provided with some form of low shelter. In most of our breeding pens, we use patio blocks (8"x16" red cement bricks) and solid plastic boards (1/2" thick x 8"x18") glued onto 2"x2" boards to give a ground clearance of approx. 2".  The goal is to produce a shelter just high enough so that the Uro's can feel the top of the shelter while standing inside it. It's best not to place these directly under the basking area unless you also place a second one elsewhere in the cage.  For most cages we also add a nest box to simulate a burrow/sleeping chamber and the naturally higher humidity contained there-in. This is usually made from a Rubbermaid "Roughneck" brand 3.3 gal. or larger soft plastic tub. We then insert a piece of 3" diameter flexible plastic drain-pipe into the upper side of the box to act as the "burrow" leading to the nest/sleeping "chamber". The tube then extends approx. 18" from the side of the tub with the end touching the ground, preferably along the back wall of the cage. We prefer soft ABS drainage pipe as it's flexible, cheap and ribbed for easy footing. Fill the tub with only very slightly damp 50/50 sand/potting soil (preferably soil w/out perlite or added fertilizers or "water" holding pellets).  "SuperSoil" brand  potting soil is generally considered the best for terrarium use.



Uromastyx tend to  have a low tolerance for cohabitating with other Uromastyx unless reared together. Under most circumstances, two mature males may not be kept together! Sooner or later one will attack the other, possibly causing serious injury. While females of most species are more variable in this regard, many females also are intolerant of same-sex housing (Saharans and Mali's are occasional but not reliable exceptions). Many Uromastyx will tolerate and even prefer being housed with a member of the opposite sex, but exceptions exist even here (note: Moroccans, Orange benti, and Ornates in particular tend to be common exceptions to this rule). Females of all species tend to become very belligerent towards all other Uros, male or female, once they are bred and begin preparing a nesting site. Most are very moody the first few weeks pre-and post-laying and may need to be separately housed for several weeks or even months. The aggression can be subtle and easily missed if you're not around the animals throughout the day. Periodically examine your animals, noting their weight and the condition of the skin along their flanks. Individuals intimidated by others tend to gradually loose weight. Aggressive animals tend to bite others along the flanks, leading to distinct thickening of this area. If allowed to continue, this can lead to significant tissue damage or even death, even if the aggressor never directly breaks the skin.

If you wish to try to house a sexual pair or trio together, first setup the cage so that each individual will have access to separate basking, sleeping and feeding sites. Then introduce them to the new cage simultaneously. Uros are by nature territorial, and even calm animals tend to attack new individuals placed in their cage. A notable exception occurs between individuals of vastly differing size. In particular, large adults are very tolerant of sharing their cage with small juveniles. Note trios generally only work if you don't cycle them for breeding. Bred females are rarely tolerant of a second female living in the same cage.

As far as cage size, the larger the better for all but hatchlings. Our ground breeder pens run approx. 4' long by 2 1/2'deep by 2' high and house strictly pairs. We primarily use Vision brand and Showcase brand 4' cages for housing single individuals or breeding pairs off the ground.  If you wish to use a standard aquarium or terrarium as a cage, we'd strongly suggest not going smaller than a 40 gal. "Breeder" style long tank for young adults and no smaller than a 20 gal. (long version) for hatchlings. You should  cover the back glass with some background (dessert scene or whatever)  and at least initially, the two sides as well, leaving only the front open glass. This will help prevent the Uros from excessively clawing at the glass or running face-first into the sides of the tank if spooked. Placing cage ornaments (logs etc.) along the edges will also help in this regard. A better option would be to build your own cage out of A-grade plywood sealed with non-toxic polyurethane to produce a cage at least 4' long, 24" wide, 18" tall for an adult pair of Uromastyx.  Many breeders maintain their Uros in large steel or plastic livestock water tanks. This is an inexpensive means of housing them but the aesthetics are somewhat problematical for "in the home" housing!  Uros are active creatures and like to run around. Shoot for as large a cage as you have space for.

We keep our hatchlings 8 to 10 per cage in 4' Vision "low profile" cages (4' long, 24" wide, 14" tall). Larger than that and they tend to have more trouble finding the food or regulating their temperatures. Note hatchlings of some species such as  Moroccans tend to get very territorial after about 4 weeks of age and must have separate shelters throughout the cage. The more aggressive individuals may need to be housed separately (or at least moved in with much larger individuals) as soon as they start showing aggression.



Uromastyx are primarily herbivores, with a taste for insects on the side. Our primary diet is composed of  two main fractions.  The first is fresh leafy greens.  As of 2006, we are trialing Earthbound brand "SpringMix" as our base diet as we can buy it in bulk and it needs no further chopping etc. to be used.  It comes in 1 pound clear plastic resealable tubs and  for the most part contains a good mix of nutritious greens.  It is a bit too high in leaf lettuces so we mix in one chopped head of either Endive or Pok Choy leaves for each 1 pound tub of Spring Mix. We rinse it all in cold water, shake off the excess moisture then dust it very lightly every day with Miner-Al (I) mineral supplement. On alternate days we also dust it heavily with ground up Mazuri Tortoise pellets.  For specimens 1 year old or older we also add a handful of warmed up frozen mixed veggies (peas, carrots, corn, cut green beans)  to the mix once or twice per week. During the winter and early spring, this is our primary daily diet.  During the summer and early fall, we harvest home grown leafy greens and blooms and use that to replace about 50% of the Spring Mix.  The  greens/blooms we primarily use are dandelion greens/ blooms, clover greens /blooms, Rose of Sharon hibiscus blooms, nasturtium blooms, cats claw blooms (a late-season dandelion-mimic), viola (Johnny jump ups blooms), rose blooms, and fresh (not dried) alfalfa leaves / blooms. We try to feed a slightly different mixture of food items each day, alternating what greens we add to the base diet. The primary store-purchased greens we add are endive first and foremost, followed by Pok Choy, mustard greens, grated yellow squash, and collards. Spring mix is already very high in romaine and oak leaf lettuces so while those are fine as lesser food items, we don't want to add any more of those to the spring mix. We do not pre-chop the added leaves for hatchlings but simply tear it fresh into chucks for everyone. When available, we also periodically place cactus pads in each cage (Opuntia sp, commercially produced as human grade food, de-spined at the store). These last for days, allowing for periodic nibbling at will.

As stated above, we dust the food lightly but daily with Miner-AL brand mineral supplement. We use the indoor version which has low levels of vitamin D3 in it.  We're not convinced that the UVB levels from the commercial  UVB bulbs are completely adequate for natural vitamin D3 synthesis so the dietary D3 is our insurance policy. Other products on the market work as well, but for us Miner-All is the safest one. An alternate product usually available from  larger pet shops is Mardel Labs mineral supplement and no doubt several other brands should be ok as well. We personally really dislike the most commonly stocked pet shop product: Rep-Cal,  as we feel it is too high in D3 levels and it only supplies calcium, ignoring the other essential minerals. The excess calcium in the gut can then lead to excessive excretion of the other minerals in the diet, leading to other nutrient deficiencies.  What ever product you use, make sure it contains a well balanced list of minerals, not just calcium.

For yearlings on up, we also dust the food once per week with Uromastyx Dust. This is touted as a complete diet and looks good on paper, but we choose to use it only sparingly. We don't feel this is essential  if you dust the food with ground up Mazuri pellets, but it's a nice insurance policy. We're not sure it's safe for use in hatchings to sub-yearling, so we don't add it to the diet until the Uro's first birthday. Mazuri tortoise pellets have a longer track record and is well accepted by our Uros and we feel it serves a very similar function in the diet.  We use to feed it by moistening the intact pellets with water and adding it to the frozen mixed veggies, but this proved too problematic for us.  It tends to stick to the mouth, occasionally leading to bacterial infections and it spoils by the end of the day.  We now grind it up in a blender and use it dusted over the damp greens.  There is less waste this way and it non longer is a spoilage problem. 

If you house your Uros on anything other than bird seed, you can partially replace the Mazuri pellets with a dish of  dry "Pretty Bird" brand finch pellets or T-Rex tortoise pellets or Juvenile Iguana pellets (Uros housed on seed tend to ignore these pellets). These are a synthetic "seed" which has multiple vitamins added and is much better digested than most bird seed.  Hopefully if the Uros main diet is lacking in some minor nutrient, snacking on this will make up for it. Some breeders also like to add ground dried bean mixes to the diet. This mix is generally comprised of various soup beans to which low levels of a multi vitamin-mineral supplement is added then run through a coffee grinder.  The final mix is offered to the Uros in a shallow dish left in the cage.

It's wise to highly limit spinach, beet greens, Swiss chard, or true cabbage in the diet, and go easy on  broccoli, kale and collard greens ( the exception being the blossoms of these). These leaves either bind important nutrients or tend to induce metabolic problems over time. Peas have their faults as well but if you supplement with a balanced mineral supplement (especially ones containing zinc, manganese, magnesium  along with the more common additive calcium), the benefits out-weigh the potential harm as long as you use them sparingly in the diet.  In our experience, it's very difficult to reliably acclimate wild-collected specimens or underweight long term specimens without adding peas to the diet. In particular, we don't consider Sudanese or Orange or Rainbow benti to be successfully acclimated until they are eating peas. Insist on this when buying these 3 species, it will greatly improve your success potential with them.

While most Uros consume the occasional insect in the wild, these generally cause more problems than they are worth in domestic specimens.  On very rare occasion, we may offer an occasional superworm (Zoophobia sp.) to individuals that are slow to settle in. These are a great way to tame your Uros.  Many are easily addicted to superworms and will go to great lengths to procure them. Conventional wisdom suggests gravid females fed a slightly higher than normal amount of insect matter produce better clutches, but we have not found that to hold true.  Most commercially available insects are excessively high in phosphorous which causes the body to excrete calcium into the feces.  Be careful to supplement w/calcium whenever you feed insects and never feed more than just a couple per sitting and only a few per week at most.  Hatchlings in particular easily develop metabolic problems if fed too many insects.  All in all, we strongly suggest you avoid insects in the diet except under special circumstances (for example for individuals that are refusing to eat or refusing to tame down).


Opportunities to drink are a rare occurrence in the wild for most species of Uromastyx. Uromastyx solve this problem by producing metabolic water from their digesting food. As long as their bellies are relatively full, most are making more than enough water to meet all their needs. Thus we don't normally offer water to our healthy Uros. The exceptions are for newly acquired/shipped animals, individuals which haven't kept up a reasonable gut mass of digesting food, females which are near term-gravid or have just laid their clutch, and for fresh hatchlings. Individuals with near empty bellies MUST be offered drinking water on a regular basis. If a Uromastyx goes off-feed, their bellies slowly empty. As this progresses, their bodies tend to dehydrate. As they dehydrate, appetite is often further suppressed, resulting in a spiral down towards death. (Note: dehydrated animals have limited abilities to process proteins so NEVER offer insects or dry bean mixes to an overly thin, dehydrated Uromastyx. The burden on the kidneys and livers may prove fatal months down the road). Despite all I've stated above, there are still very few circumstances when it is acceptable to put a water bowl in a Uromastyxs' cage. If you feel an individual needs water, take him or her to a tub filled with approximately 1/2" of bath-water hot (100°F) water. It must be as warm as you can safely make it so that the individual stays near their optimum body temperature (105°F). Some will drink on their own, others can be enticed by dripping water on their snout. (Note: Saharan Uromastyx are prone to aspirating water into the lungs so be very careful when soaking them.  Put them very slowly into the tub and keep the water very shallow (1/4" max.)  Other Uro species seem much less likely to have this problem). Many unacclimated  individuals will not drink while being watched.  You must leave their line of sight. It's also wise to leave them undisturbed for a few minutes after drinking to avoid them regurgitating. Truly dehydrated animals may need to be tubed with a warmed electrolyte solution. See your vet if you are unfamiliar with this procedure. Using Pedialtye or even Gatorade or similar product instead of water for the soak is one way to supply these electrolytes.  Just make sure to rinse the solution off them and dry them well afterwards.  The hindgut is also capable of absorbing water, so use of dilute electrolyte/vitamin enemas may also be useful for seriously dehydrated individuals.  Individuals with intestinal problems (parasites or bacterial infections) may not be able to absorb water through the gut and will need to  be taken to a Vet for injections of a saline/glucose and  sterile water mixture (even ratio of each is usually best).  Note it's easiest to give this injection under the front arm pits - if placed just right, you hit what appears to be a lymph duct and you can easily inject several cc's of fluid without any backwash out of the injection site.  If you tube them orally, juveniles usually will hold down 2 cc, medium adults 5 cc, large adults 7 to 10 cc's of fluid.  While it's easy to give more than this, they will often regurgitate larger amounts a few minutes to hours later.  If given rectally, reduce these doses by about 1/3.

An alternative method to offer water is to take a small jar lid (approx. 1/4" deep) early in the morning, fill it with water and  place it in the cage (along a wall  in a corner).  Most Uro's have a higher tendency to drink in the morning, perhaps being programmed to seek potential dew at this time. This small amount of water should evaporate off during the day, causing no harm. We routinely have a lid of water in our hatchling tanks, but stop this practice once they go though the first  sheds (when 8 to 12 weeks old).  Note if you feed ground bean mixes, especially to hatchlings, a water dish can cause significant health problems. The Uro's tend to constantly walk through their water dishes. If they then walk through their bean dish, they essentially glue the powder to their bellies and toes. This can result in significant skin infections/lesions which can take months to clear up. Note feeding soft fruits can cause the same problem - the material easily glues itself to their bellies as they walk through it, resulting in significant infections if repeatedly left unattended.

A species exception to the no water rule are the benti and Saharan Uromastyx. While most Uromastyx species will commonly refuse offered water, both species of benti and Saharans will often accept the offering and drink heartily. While they can do fine without water as long as they keep a belly full of digesting food, since they readily drink, we offer them soaks more often than the other species. Note, we still don't keep a water bowl in their cages, we just offer more opportunities to soak in the tub.  Please be sure to dry them off afterwards as dampness will eventually lead to health problems.


Most breeders believe Uromastyx need to be put through some form of winter in order to sufficiently cycle to induce breeding and fertile egg production. The various species vary over how "severe" a winter they need, with Moroccans, Mali's, and Egyptians needing the coldest/longest winters, Ornates, Saharans and benti needing moderate/mild winters, and Sudanese needing the bare minimum of seasonal differences to successfully cycle. We've tried numerous approaches to wintering or "brumating" our Uromastyx, with widely variable results. Too warm or too short a "winter" and most species won't cycle, too cold or too long and mortality becomes a problem. The best solution for us has been as follows:

First, stop feeding Mali's, Moroccans, and Egyptians about 2 weeks prior to the start of your "winter". Cut severely back on the amounts but continue to feed Ornates, both benti species, Saharans and Sudanese Uromastyx. We mostly offer Romaine and endive at this time primarily for their high water content.  Avoid peas, beans, and any high protein foods. At the same time cut your day length to 10 hours of light but leave the cage temperatures close to normal during the day, while trying to keep no hotter than 70°F at night. After the first week, we drop day length down to 8 hours per day. All else stays the same. At the end of week 2, we drop day length to 6 hours per day, and try to maintain the cage temperatures around 60°F to 65°F for at least 20 hours per day. Then for at least 4 hours per day, we turn on the basking lights so that the cage temperature hits at least 80's °F, preferably 85°F,  for at least 2 full hours. The goal here is to stimulate the immune system to kick in and gut function to reactivate for at least 2 hours each day. Failure to do this will significantly increase your mortality rate, especially for Ornates, Sudanese, and the benti. If you haven't cleared the guts of your Mali's and Moroccans, they too risk suffering from gut paralysis and eventual necrosis. During this time we still offer limited food to the benti and Saharans  but generally restrict food for everyone else. Note we still have bird seed in the cage as bedding, so some feeding might be occurring, but except for the benti and Saharans, most Uromastyx will not seek food at this time. We continue this to produce a "winter" of approx. 6 to 8 weeks. We then bump the cage day temp to the mid 80's°F and day length to 8 hours per day for a week. The next week we go to normal daytime cage temps and 10 hours day length, the third week 12 hours, the forth week 14 hours. During this "spring" time , we try to keep our night temps near 70'°F. Most our Uro's will be up and basking by the end of "spring" week one and eating lightly by week two. By the third week most should be back to their normal activity levels. This system has worked well for us for several years running now and several other breeders use a very similar system with excellent success as well.

For egg laying, we use 3.3 gal. or 10 gal. Rubbermaid "Roughneck" soft plastic tubs, lids intact. We cut a 3" round hole in the upper corner of the long side of the tub, and insert a 2' to 3' section of 3" diameter drainage pipe (flexible, ribbed plastic, see photo above). We then cut a hole in the side of the pipe so that the Uro's have easy access out of the pipe and into the nest chamber. The insides of the nest boxes are half filled with a 50:50 mix of playground-grade sand and SuperSoil brand potting soil moistened just enough to allow it to hold a tunnel. We've taken to adding a handful of lime to the mix as well to lower the overall acidity of the soil. Our long-term established wild-collected animals and captive-breds use this setup without hesitation. Most of the newly imported Mali's have balked and buried their eggs in the bird seed, often under the basking spot. It appears that the nest boxes are best put in w/ the females PRIOR to the onset of the breeding season so that they can become accustomed to moving in and out of them and digging preliminary tunnels. As a side note, be sure to trim the toenails of your gravid females a few weeks before they lay. They are notorious for nicking their eggs while burying them.

We remove the eggs as soon as they are detected and place them on their sides in specially designed egg-holding cartons (these are a Deer Fern Farms "invention" (for  lack of a better term). These cartons are then placed in our incubator at 93°F.

Uromastyx eggs relay their fertility and viability status very clearly. Fertile eggs have a distinct red circle (the developing embryo) clearly visible at the time the eggs are laid. We orient our eggs so as to position the embryo along the side of the egg, but it's highly unlikely that this is necessary. Fresh eggs are somewhat water-balloon-like when laid, but good eggs usually firm up and whiten within a day or two at most. Eggs which are distinctly yellow or in which you can see the contents moving around inside in a two-toned pattern (milky yellow in a clearer yellow) are already in the early stages of disintegration and will not hatch. Dud eggs will begin to smell almost immediately and are often easy to detect in the incubator within 3 to 4 days. Duds also often keep a faintly oily look to them and rarely firm up.

In the past, we've incubated at temperatures ranging from 85°F to 88°F with so so success. This results in hatchlings in about 80 to 100 days.  Initial thoughts from various other reptiles breeders suggested that we were incubating too high.  However,  field data for Ornate Uromastyx now suggests we've actually been incubating too low.  Several of us thus tried 92°F to 94°F several seasons ago with excellent results. Thus while you may still see others list the lower temperatures as correct, we've decided to permanently change our temperatures to 93°F ±2°F.

At 93°F, incubation for most Uromastyx species should range closer to 55 to 65 days. The hatchlings are quite vigorous and ready to feed within a day or two. Treat them as you would adults, but slightly cooler and periodically offer water. Watch for signs of aggression. Dominant animals will significantly repress the growth of the other hatchlings housed with them. Siblings usually get along with each other (with one individual per clutch almost always being an exception). However intermixing already established clutches almost always leads to fighting. Sudanese and the benti Uromastyx must have drinking water available as described earlier until the first or second shed have been past. The other species do well with or without this extra water, as long as they keep their bellies full. Note hatchlings MUST be offered fresh fecal pellets from a healthy adult Uromastyx during the first few days post hatching. They need this in order to properly inoculate their guts and grow normally. Failure to do this will often significantly stunt their growth and increase their potential to suffer gut impactions early in life. The drive for them to eat this material wanes quickly, so you must do this as soon as possible. Crumble fresh fecal pellets into their normal food and watch to make sure each individual eats at least some of the fecal mass. Don't use a fecal pellet from an adult whom you've recently wormed or treated with antibiotics. Pick an individual that is obviously thriving and is free from an excessive load of parasites (not a lot of "rice"-like particles in the fecal pellet), but it doesn't have to be parasite-free. Parasite-free may not even be desirable - they jury is still out on that one. By parasites, we're strictly referring to nematodes. Other parasites such as coccidia etc. are undesirable at any levels.


Try not to offer hatchlings any dry foods for the first month or two. They easily get gut impactions from overly dry food lodging in the intestines. If you feel a hard mass in their bellies, try to induce drinking and later GENTLY massaging the mass to try to break it up. A warm water enema may prove necessary to hydrate the mass from both sides to free it up and allow passage. If you feed only moist foods and occasionally mist their foods, impactions should not be a problem. Hatchlings are also much more prone to metabolic bone disease from insufficient vit. D3 and calcium/trace mineral imbalances in the diet (or from excessive insect consumption). Avoid the temptation to feed insects, you are not doing them a favor! Getting your hatchlings off to a good fast start significantly lowers the incidence of problems down the line, especially for impactions.

Hopefully this covers the basic's you'll need to successfully keep and potentially breed your Uromastyx. Enjoy!

We express our thanks to the author of this article Mr. Douglas Dix. Original article you can find on his web-page

,,Uromastyx thomasi Oman-Spiny-tailed-Lizard,, by Thomas M. Wilms

Breeding Programme

Uromastyx thomasi

    Photo: Felix Hulbert

First Annual Report (2006)

Thomas M. Wilms

 1. Introduction

Uromastyx thomasi has been described in the year 1930 by PARKER on the basis of two specimens. The holotype originated from Bu Ju’ay, Rub’al Khali, Dhofar and is now in „The Natural History Museum“, London [BM 1946.8.14.43 (old number: BM 1930.6.30.2)]. Since its description only few has been published about this species (see also BARTS & WILMS 1997), including faunal lists (PARKER 1931, WERMUTH 1967, ARNOLD 1986, WELCH 1994) as well as publications on the zoogeography of Arabia (ARNOLD 1987) and on the phylogeny of the genus Uromastyx (MOODY 1987). WILMS (1995) compiled all available data on U. thomasi or the first time and prepared a distribution map for the species. At that time the knowledge on morphology, distribution and especially ecology was very limited. Ecological observations of U. thomasi have been published by ARNOLD (1980), WILMS & HULBERT (2000) and WILMS, LÖHR & HULBERT (2002). Uromastyx thomasi lives in coastal Oman. The status of the wild population is unknown, but preliminary investigations suggest, that this species is not common in wide parts of the range (WILMS, unpublished). The distribution area of U. thomasi has a length of approx. 600 km and a maximum width of 230 km. Obviously not all types of landscape (e.g. mountainous areas) represented in the natural range of the species are suitable for them as habitats. In the year 1998 the author was able to examine 19 preserved specimens in museum collections [„The Natural History Museum“, London (BMNH) and Zoologischen Forschungsinstitut und Museum A.Koenig, Bonn (ZFMK)]. Some more specimens are kept in the collections of he Museum of Natural History, Muscat, Oman; the University of Muscat, Al Khod; the California Academy of Science, San Francisco and the Museum of the Bombay Natural History Society. WILMS et al. (2002) estimated the number of Uromastyx thomasi known to science of not more than 30 specimens (until the year 1998).

2. Studbook population

The captive population of Uromastyx thomasi consists today (January 2007) of 33 specimens. The founders of this population have been imported to Germany in accordance with national and international laws in 1998 for breeding project. First captive breeding occurred in 2000 (WILMS et al 2002). The animals are beeing kept at six locations. Four locations left the breeding programme befor 2006 because of loss of all specimens. In 2006 breeding occurred at one location. The sex ratio within the population is: 12.4.17.

3. Imports

No imports of this taxon occurred since 1998.

4. Natural history notes

According to WILMS & HULBERT (2000) and WILMS et al. (2002) Uromastyx thomasi lives in stony and sandy plains with sparse vegetation. At the end of November the specimens where active from 11 a.m. to 4 p.m. The temperature of the surface and air temperature one meter above ground was measured. The surface temperature ranged from 37.5-51.1 °C, while the air temperature was 29.2-35.8 °C.  The body temperature of 24 specimens ranged from 33.1-39.4 °C. The temperature was measured within 10 and 35 min. after the first sighting of the respective animal (for details see WILMS et al. 2002). The temperature in the burrows was between 30.3-33.6 °C at a depth of 20-41cm. In four burrows temperature at a depth of more than 53 cm was measured. The data are 34 °C at  53.5 cm, 31.4 °C at  47.5 cm, 29.7 at  67 cm and 29.6 °C at 77.5 cm. The length of the burrows varied between 45 and 165 cm. One specimen played dead after capturing. A total of 38 faecal samples have been analysed and despite the fact, that six plant species were found in the habitat, only two different plant species could be found in the droppings: Indigofera sp. (Fabaceae) and Plantago albicans (Plantaginaceae). Remains of insects or other animals could not be found in the faecal samples (WILMS et al. 2002).

5. Reproduction in captivity

Fist captive breeding of Uromastyx thomasi occurred in the year 2000 (WILMS et al. 2002). The animals have been kept pair wise in enclosures with one square meter floor space. Substrate temperature was partially between 45 and 55 °C. Air temperature was between 35 and 40 °C at daytime and around 18 –23 °C at night time. Photoperiod was 10 hours. Courtship behaviour began late February. Pregnancy takes around 35 days and oviposition takes place between May and September. Clutch size is between 9 and 16 eggs. Animals hatched in August 2000 reproduced for the first time at an age of 24 month. The juveniles of the F2-generation hatched between 22-25, November 2002. In 2003 breeding occurred with one of the original (wild caught) pairs and a pair consisting of WC male and a CB 2000 female (F1). In total 58 Uromastyx thomasi have been bred in the years 2000, 2002, 2003 and in 2004. Unfortunately some of these CB U. thomasi experienced a high mortality at four locations, so that the number of specimens transferred into the ESF Studbook is much lower. Incubation period was between 81 and 101 days. The juveniles had an average snout-vent-length (tail length) of 45.7 (17,2) mm [Clutch 1] ; 49.8 (19,1) mm [Clutch 2] and 46,9 (20,2) mm [Clutch 3]. Mass was averaged 5,6 g [Clutch 1], 4,3 g [Clutch 2] und 4.87 g [Clutch 3]. Of the 34 hatchlings from 2000 only 4 specimens are females. Because of this fact we suggest, that in U. thomasi the sex of the hatchlings could be greatly influenced by the incubation temperature (temperature depending sex determination). Until today, there are no available data on the time of oviposition or hatching of wild Uromastyx thomasi. Only the collection dates of three very young juveniles in the Natural History Museum, London are known (BMNH 1973.403, BMNH 1973.2906 and BMNH 1973.2907), which were collected in November 1972 and March 1973. All three specimens are of similar size, which points to a elongated egg laying period of presumably 3-5 month. This fits with the observations on captive U. thomasi. The first clutch was laid 01.05 and the last one 23.09. Some of the captive females laid two clutches in one breeding season. If this is likewise possible in the wild is not known.

6. References

ARNOLD, E.N. (1980): The Reptiles and Amphibians of Dhofar, Southern Arabia. Journal of Oman Studies. Special Report 2: 273- 332.

ARNOLD, E.N. (1986) A Key and Annotated Check List to the  Lizards and Amphisbaenians of Arabia. Fauna of Saudi Arabia 8: 385-435.

ARNOLD, E.N. (1987): Zoogeography of the Reptiles and Amphibians of Arabia. In: Proceedings of the Symposium on the Fauna and Zoogeography of the Middle East. KRUPP, F.; SCHNEIDER, W. & KINZELBACH R. (eds.): 245-256.

BARTS, M. & T. WILMS (1997): Catalog of valid Species and Synonyms, Vol. 4, Agamidae, Leiopepidinae.- Herpprint International, Pretoria, South Africa: 398-418.

CUNNINGHAM, P. (2000): Daily activity pattern and diet of a population of the Spiny-tailed Lizard, Uromastyx aegyptius microlepis, during summer in the United Arab Emirates. Zoology in the Middle East 21, 2000: 37-46.

MOODY, S.M. (1987): A preliminary cladistic study of the lizard genus Uromastyx (Agamidae, sensu lato), with a checklist and diagnostic key to the species. – Proceedings of the Fourth Ordinary General Meeting of the Societas Europea Herpetologica, Nijmegen, Holland. S. 285-288

PARKER, H. W. (1930): Three new Reptiles from southern Arabia.- Ann. Mag. Nat. Hist., 6 (Ser. 10): 594-598.

PARKER, H. W. (1931): Some Reptiles and Amphibians from SE. Arabia.- Ann. Mag. Nat. Hist., 8 (Ser. 10): 514-522.

WERMUTH H. (1967): Das Tierreich (Agamidae). Lieferung 86, Berlin, 1-127.

WILMS, T. (1995): Dornschwanzagamen - Lebensweise, Pflege und Zucht. 130 pp. Offenbach; Herpeton-Verlag.

WILMS, T. (2001): Dornschwanzagamen - Lebensweise, Pflege und Zucht.- 143 pp. Offenbach; Herpeton-Verlag.

WILMS, T. & HULBERT, F. (2000): On the herpetofauna of the Sultanate of Oman, with comments on the relationship between Afro-tropical and Saharo-sindian faunas. Bonner zoologische Monographien 46: 367-380.

WILMS, T., LÖHR, B. & HULBERT, F. (2002): Erstmalige Nachzucht der Oman- Dornschwanzagame- Uromastyx thomasi PARKER 1930 - (Sauria: Agamidae: Leiolepidinae) mit Hinweisen zur intraspezifischen Variabilität und zur Lebensweise. Salamandra 38 (1): 45-62.

,,Observations on the Giant Sungazer Lizard, Cordylus giganteus, in Captivity,, by Gary Fogel

Bull.  Chicago Herp.  Soc.  35(12):277-280,  2000


Observations on the Giant Sungazer Lizard,  Cordylus giganteus,  in Captivity

Gary Fogel

Email:   kordylus@juno. com

In the past few years,  it has come to my attention that more sungazer lizards have been introduced into the commercial pet trade.   For this reason I thought it would be a good time to share my observations and interactions with this species,  in the hope that others might benefit from my experience in keeping these lizards.   As you may be aware,  there is still very little written on Cordylus giganteus,  compared to other,  more popular,  types of lizards.   The only articles I’ve seen in herpetocultural magazines have been Switak (1995) on the genus Cordylus as a whole,  and Donovan (1997) on sungazers specifically.   The Vivarium has never published an article on the genus Cordylus,  and popularized reptile books are general in their information,  often contradicting one another on the facts (e.g. ,  Bartlett and Bartlett,  1997; Mattison,  1983,  1991; Wynne,  1981; Zimmermann,  1983).

I acquired my first sungazers in 1989.   At the time,  information on these lizards was not easy to find,  so I really did not know what to expect from them as far as behavior and temperament were concerned.   I had experience keeping and breeding smaller cordylid species,  such as armadillo lizards,  Cordylus cataphractus,  and girdled-tailed lizards,  Cordylus warreni depressus,  but none with the majestic sungazer lizard.   My first job was to find written information currently available.   I did as much research as I could,  talking to other people and gathering information from various technical articles from the Field Museum library.

Having gathered verbal and written information,  the second step was to prepare a habitat enclosure for them,  as these lizards are really too big to be kept properly in an aquarium environment.   I modified a homemade wooden table,  adding pegboard walls for ventilation,  and two screen doors in the front,  one on each side,  for easy access.  The enclosure measured seven feet by four feet with two fluorescent light fixtures overhead,  each containing one full spectrum light and one black light,  approximately 15 inches from the ground.   I furnished a heating pad in the center of the cage for autumn and winter use,  and a large dog bowl for water.   Hiding places were provided in the form of two clay tiles,  24 inches long,  cut in half lengthwise,  to serve as burrows.   These lizards live in open grassland areas,  and hide in underground burrows,  rough ly three feet long,  either dug by the lizards themselves,  or dug by other animals and adapted for use by the sungazers.   The flooring of my enclosure was floor tile,  over which I placed artificial turf,  except for 12 inches at the front of the enclosure. 

This is where I placed the water bowl and I hoped was the place the animals would  use for their bathroom area (they tend to use the same spot repeatedly for this activity).   These lizards come from a temperate climate,  with low humidity,  where they hibernate in their burrows during the winter months.   I have never used a heat lamp in the enclosure,  as these animals prefer a relatively cool air temperature.   Extreme heat,  such as needed with Uromastyx and chuckwalla species,  could prove fatal for the sungazer.   Use of a small fan,  during the summer months, to help circulate the air,  has also proved helpful.   The cage was now ready for its inhabitants (Figure 2).

When I unpacked the shipment of sungazers,  I didn’t know what to expect.   Would they be aggressive and bite?  I was surprised to find them quite non-aggressive,  preferring to just lie there with their arms at their sides.   This is a defense posture they use in the wild,  as they are heavily armored from top to sides with sharp thornlike scales.   Anything biting them gets a mouth full of thorny protrusions,  not unlike biting a pincushion.   The other defense they use is to swing their armored tails back and forth at the mouth of their burrow.   This can draw blood if one is not careful.   Other than that,  I have never had any aggressive action taken towards me by the sungazers. They have never actively bitten me in defense,  only when I have gotten in the way during feeding.   They prefer to run and hide,  rather than attack,  unlike girdle-tailed lizards,  Cordylus warreni,  which will bite readily,  given the chance.


 The group consisted of one male and three females.   Males, incidentally,  are very easy to sex.   They have pronounced raised scales on the inside of their forelimbs,  which are very noticeable (Figure 3a),  as well as larger femoral pores on the hind legs.   Females have regular scales on their forelimbs (Figure 3b).   I do not know the reason for these raised scales on the males,  but males of other Cordylus species do not have them at all.

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 Figure 3.   A) Male sungazer,  Cordylus giganteus,  showing raised scales on the forelimbs.   B) Female sungazer,  showing normal forelimb scales.   Photographs by Carlos Sanchez.

 I  watched and waited for the usual lizard behavior patterns to emerge:  head-bobbing,  tail-wagging,  tongue-flicking,  etc. 
The sungazer lizards rarely or never exhibited these behaviors. They seemed oblivious to one another.   They chose their respective burrows; occupied by two animals each.   Over the years,  I’ve noticed that the male would change burrows from one side to the other,  every year or every other year. One year he would use the one on the left,  then move to the one on the right two years later,  for no apparent reason. The females would interchange with him,  depending on which side he occupied.   I’ve observed attempted copulation only twice in 10 years.   Of course this doesn’t mean that it hasn’t happened more frequently; perhaps I just wasn’t home at the time,  to witness it.   The first observed copulation was five years after I had obtained them.   It was in November and the barometric pressure had just dropped that day.   I heard scurrying inside of the cage and went to investigate.   I found the male chasing one
of the females slowly around the entire cage attempting to mount her from behind.   At one point he bit her head region,
and tried to position himself underneath her tail region.   I did document this with a photo,  but I didn’t get as close as I might have liked,  for fear of disrupting the action.   I couldn’t tell whether this was a successful breeding because of their position within the enclosure.   Afterwards,  they went back to ignoring each other.   Three years later,  I would witness this action again,  also in the autumn months,  but to my knowledge,  no young were ever born,  unless they were eaten after birth.   I have read that males can be cannibalistic towards any young (Marais,  1984).
In 1994,  I acquired six more yearling sungazers,  which were housed in two groups of three each,  until I could build
another large enclosure.   They were about seven inches total in length,  and unsexed at the time.   As they grew,  I discovered that the sex ratio was four males and two females.   Since I did not have the space for a lot of large enclosures,  I decided to put all six in another seven feet by four feet enclosure with an open top this time,  sort of like a big wooden box.   I also included six hide areas on one side of the cage,  using clay tiles again,  and 12 inches in length (Figure 4).   Housing them all together,  I was not sure how all the males would react to one another,  but then again,  this species interacts and behaves by their own set of rules.   When I first placed them all together,  I actually saw
tail-wagging and tongue-flicking,  as the two groups got to know one another.   The aggression towards one another srgrd34sub sided after one hour and they have been living as one group ever since.   I personally was surprised that four males would tolerate one another in an enclosed area,  but they are now about seven years old and I have not yet had a problem.   I do not know,  however,  at what age they reach sexual maturity.   They are not as large as my other adults,  so perhaps they are not yet sexually active,  but at seven years,  I would think that they are mature enough.

The water dishes for both enclosures are large enough for the sungazers to immerse themselves,  and I have noticed a rather unusual behavior.   In both cages,  these lizards like to sit in the water bowl,  much like a athtub.   I have used large water dishes for other species of Cordylus,  and they do not show this behavior at all.   The sungazers,  however,  seem to like their occasional dip in the water dish. I have seen other enclosures at zoos,  which use little dishes of water for the sungazers. Perhaps if they knew they liked bathing,  they would give them larger bowls. In the wild,  they do experience a rainy season,  so coming in contact with water is not an unheard of experience. Sometimes their burrows can become flooded out,  leaving these lizards to locate other,  drier burrows,  or digging upward in the existing burrow. They also do not like venturing far from the protection of the burrow entrance. I have left the cage door open on one occasion,  only to find that not one had even left the enclosure. Should they become startled; the sungazers literally fly into the burrow openings,  seeking refuge from impending danger. Should I open the cage door at this point,  they would whip their tails back and forth inside the clay tiles,  resulting in a hard,  hollow, thump-thump sound. Occasionally,  one can be heard digging inside its shelter,  at the end against the wall.

 Perhaps in part because these animals behave unlike other lizards,  breeding them remains a mystery.   They may only give birth every other year (van Wyk,  1988),  and the gestation period is unknown  - - -  it might last a year or even longer.   People who have claimed to breed them,  usually have done so only once,  leading me to believe that the allegedly bred sungazer was gravid when purchased.    If gestation can last a year or more,  then this scenario certainly seems plausible.   If someone had bred them after having kept them in captivity for two years or more,  I would be more inclined to believe that,  yes,  they actually had bred them.   Because they are notoriously difficult to get to breed,  I feel it is important for people purchasing them to know this fact.   If one is buying sungazers in hopes of making a small fortune off the offspring produced,  then one is in for a big surprise.   Many zoos and individuals have tried for years,  but breeding is still an elusive nut to crack.   It seems hibernation plays a key role in reproducing this species,  but if
you do hibernate,  are you willing to run the risk of a sungazer fatality as a result?  I have talked to someone in South Africa who did breed these lizards regularly by hibernating them in a refrigerator for several weeks,  in the winter months.   He also kept them outside the rest of the time,  utilizing natural sunlight in the process.   Bert Langerwerf maintains his reptiles in much the same way.   After many years of keeping sungazers outdoors at his Alabama facility,  with no young being born,  Bert Langerwerf finally did report breeding success this past year.

These days,  I find it shocking the obscene amounts of money that sungazers command in the commercial pet trade,
especially since the chance for breeding success is so slim. Should breeding occur,  they only give birth to one or two
young and the process could take two years.   This is quite a contrast from some species of commercially bred lizards,
which may lay 20 eggs each,  three times a year.  
At least if more people are buying and working seriously with sungazers,  then breeding may become more common-
place.   The more people contributing to the existing knowledge base,  the better for the animals’ continued survival in captivity.   It’s my hope that this article will inspire other sungazer keepers to share their experiences in print.

,,The conservation initiative for giant galliwasps at nashville zoo: a preliminary account,, by Dale Mcginnity




Giant galliwasps are diploglossine anguid lizards restricted to the Neotropics. They are rarely-seen skink-like lizards with snout to vent lengths (SVL) greater than 200 mm. Four species in the genus Celestus, collectively known as West Indian giant galliwasps, occur or occurred on Jamaica and Hispaniola. One, C. occiduus, is endemic to Jamaica: one, C. warreni, to Haiti: and two, C. anelpistus and C. carraui, to the Dominican Republic. Two species in the genus Diploglossus have maximum SVL less than 200 mm. D. monotropis is found from Panama through Colombia, and D. millepunctatus is restricted to Malpelo Island, located off the Pacific coast of Colombia.

         fghnfh7657 Diploglossus_lessonae_02

These giant galliwasps are impressive lizards, worthy of conservation resources as a unique group of animals. Most have extremely limited ranges and an apparent propensity for extinction. The Jamaican species, C. occiduus, is probably extinct (Schwartz, 1991). One of the species from the Dominican Republic (DR), C. anelpistus, was recently reported to be `at best exceedingly rare and at worst extinct' (Powell et al., 2000). The 1996 IUCN Red List of Threatened Animals listed C. anelpistus as Critically Endangered and C. carraui as Endangered. The status of the Haitian species is unknown, but its limited range occurs in a mostly deforested habitat in northern Haiti. The Central/South American species D. monotropis is considered rare throughout its range. The Colombian species D. millepunctatus has an extremely small range restricted to Malpelo Island, a 1-mile by 0.5-mile (1.6 by 0.8 km) barren island.

Giant galliwasps often have significance in the cultures of indigenous people, who often fear the lizards and consider them venomous. Lynn and Grant (1940) described how Jamaicans regarded galliwasps: `They are greatly feared by the natives and are the subject of many yarns and fables.' Currently, a widely held belief in Jamaica is that galliwasps are venomous and that if a bite occurs and the galliwasp reaches water first then the person dies, but if the person reaches water first, the galliwasp dies. The Haitian species is considered venomous by locals, and has significance in Voodoo religion (Needham, pers. comm.). Myers (1973) wrote the following about the giant galliwasp in Central and South America:

`People in northwestern Panama know Diploglossus monotropis by the name escorpion coral. The names used by Colombian Negroes living in eastern Panama are madre de culubra and madre [de] coral. Medem (`1968' [1969]) also recorded the last two names for Colombia. This ``mother of coral snakes'' is naturally believed to be poisonous and is something to be feared.'


The goals of this project are: (1) to conduct and support fieldwork on the endangered giant galliwasps; (2) to help educate people about these lizards; and (3) to developed standardized husbandry protocols for the groups.

Conservation plan

The conservation plan includes captive, field, and educational components. The captive history for giant galliwasps (primarily C. warreni) in the U.S. was less than stellar prior to 1999. Reasonable captive longevities (maximum recorded 12 years) for wild-caught adults were reported, and captive reproduction had occurred at several zoos (Bronx, Knoxville, and Milwaukee) (Lawler and Norris, 1979; C. Berg, pers. comm.; J. Behler, pers. comm.). However, the captive population became extinct in U.S. zoos by 1995, as the inability to raise offspring and the ban on exportation by range countries allowed for no recruitment. In 1998, the AZA Lizard Advisory Group (LAG) voted to add this project to its three-year action plan, and it was determined that a small group of giant galliwasps should be acquired for research to develop husbandry techniques.

The range and conservation status for the endangered and possibly extinct West Indian giant galliwasp species are poorly known. To the author's knowledge, no formal surveys have been completed for any of these species. However, Sixto Incháustegui has been collecting data on C. carraui in the DR for years. Fieldwork is needed, especially for the potentially extinct species, to determine their actual status. An educational component could both aid the fieldwork and help dispel some common misconceptions about galliwasps, which are often killed on sight.

Captive component

Development of a successful captive-management plan could help insure against future extinctions of populations or even species of giant galliwasps. The only known habitat of C. anelpistus, the Come Hombre forest in the DR, was being destroyed as the only four reported wild specimens were collected in 1977. The animals were sent to a zoo where they produced many offspring; but because successful captive management techniques had not been developed, all of the specimens died. A living C. anelpistus has not been documented in almost 20 years. The inability to successfully raise young giant galliwasps is the primary reason captive programs have failed in the past.

As C. warreni is the least endangered West Indian giant galliwasp species, it seemed the best choice for use as a conservation surrogate to develop captive husbandry techniques for the group. After nearly a year of respectful correspondence, the Director General of Haiti's Ministry of Agriculture, Natural Resources and Rural Development graciously provided permits for the scientific collection of 9.9 C. warreni. In July 2000, with funding provided by Nashville Zoo, Dr Don Gillespie, James Needham, and the author of this paper traveled to Haiti and collected a founder population of C. warreni. Due to the expert care and dedication of the zoo's lead herpetology keeper, Nicole Atteberry, over 300 offspring have been produced from the founder population. The mortality rate has been less than four percent and the oldest offspring are reaching subadult size. The three factors believed to be important for successful husbandry are: (1) intense UV-radiation created by utilizing self-ballasted mercury vapor bulbs; (2) a deep substrate, hot and dry on one side of the cage and cool and moist on the opposite side; and (3) a varied high-calcium and low-fat diet. The captive program will be considered a success when healthy F2 offspring are produced, hopefully in 2003 or 2004, at which time a studbook will be initiated for the group. In addition, the herpetology department at Nashville Zoo recently acquired a small group of Diploglossus monotropis, which will be utilized to develop captive husbandry techniques that may be utilized in the future for the more highly threatened D. millepunctatus.

Field and educational component

Fieldwork should concentrate on the endangered species, about which the least natural history information is available. For this reason, it was decided that fieldwork would concentrate on C. occiduus, C. anelpistus, and C. carraui. These species are represented by relatively few known specimens, indicating that they are extremely rare – two of them may be extinct. However, they may be more common than presently believed due to their presumed crepuscular and semi-fossorial lifestyles. Locating living specimens of any of these species will require some effort, as their relatively unknown localized distributions and apparent rarity may make them difficult to find. Some initial work has been completed in Jamaica.

George Shaw formally described the Jamaican giant galliwasp (C. occiduus) in 1802. It is the largest diploglossine anguid, with a maximum known SVL of 303 mm (Schwartz and Henderson, 1991). Sloane (1725) reported that this species occurred in marsh grounds on several parts of the island. Gosse (1851) wrote that `in the swamps and morasses of Westmoreland, the yellow galliwasp (C. occiduus), so much dreaded and abhorred, yet without reason, might be observed sitting idly in the mouth of its burrow, or feeding on the wild fruits and marshy plants that constitute its food.' In addition to plant material, this species was reported to eat fish (Schwartz and Henderson, 1991). The few preserved specimens are bleached, so few data are available on color or pattern (Schwartz, 1970). Sloane (1725) described a living individual as having scales on the back or upper parts of a brown color, with spots of orange color and an orange belly. In addition, Boulenger (1885) reported that this species was `brownish above, with dark brown spots or cross bands.'


In addition to traditional survey methods, it was determined that an efficient strategy to locate unknown populations of giant galliwasps would be to involve local people. This strategy could also help to educate the local people about the presence, lack of venom, and need for conservation of galliwasps. A poster was developed and produced for the project in Jamaica with a grant received from the Columbus Zoo Conservation Fund. A generous donation to the project by Rob Ferran funded the initial survey work in Jamaica. In January 2001, Dr Byron Wilson (University of the West Indies), Steve Conners (Miami Metrozoo), and the author conducted initial habitat surveys and interviews with local people, and distributed posters in wetland areas along the southern coast of Jamaica. In addition, a television spot about the project was taped and played over the entire island, and a newspaper article about it was published some time later. Three sites (the upper Black River morass, a small patch of swamp forest on the lower Black River morass, and Alligator Hole) were identified for more intensive survey work based on the results of the initial survey and feedback from local people.

In 1956, Cousens reported that a Jamaican giant galliwasp had not been collected in over one hundred years. Several authors have presumed that it is extinct, although no comprehensive survey has been conducted. Recently, Crombie (1999) reported that he believed that `. . . declarations that the species is extinct may be premature.' The Jamaican iguana (Cyclura collei) was thought to be extinct on mainland Jamaica in the twentieth century until 1970 when a dead specimen was found (Vogel, 1990). The first living specimen was found in 1990 (Vogel, 1990). If a large, diurnally active, terrestrial iguana could go unnoticed for almost 100 years in Jamaica near the capital city of Kingston, then a smaller lizard with a probable crepuscular or nocturnal and semi-fossorial lifestyle may still occur there in an area that Crombie (1999) described in relation to herpetology as follows: `. . . the single most extensive wetlands area in Jamaica is the Black River and its tributaries. With its large estuary and bay, in addition to broad inland freshwater swamps, this area remains very poorly collected and barely explored.

'Conclusions and the future

Two (C. anelpistus, C. occiduus) of the four species of West Indian giant galliwasps have been listed respectively as possibly and probably extinct. Surveys for these two species are critical. Collar (1998) stated that a `commitment to extinction' is perhaps most likely `when we declare species extinct too soon, sealing them off from further investigation.' He also suggested that these assumptions of extinction can be self-fulfilling.

Intensive surveys of the identified localities in Jamaica are planned for late 2002 or early 2003. Funding sources are being identified to support researchers in the Dominican Republic to conduct surveys for the endangered galliwasps in that country in 2003 and 2004. When captive-management techniques have been developed for Diploglossus monotropis, Nashville Zoo staff will work to acquire a small genetically viable captive population of D. millepunctatus. This population will be utilized for research and as a reserve population for this species, which could become extinct by a single catastrophic event due to its extremely restricted range.

Ideally, populations of the potentially extinct species will be found and habitat will be protected for them. Due to their fossorial lifestyle, specimens of these species may be found as their isolated habitats are altered by heavy equipment, as was the case for C. anelpistus in 1977. None of these specimens or their offspring survived longer than three years in zoos, and a living specimen has not been recorded since. If a similar situation occurs in the future, hopefully, the chances for survival will be enhanced.


I would like to thank Rick Schwartz, the Nashville Zoo Director, for his support for this project. Thanks are also in order for the herpetology staff and veterinary staff at Nashville Zoo for their help with the captive population of giant galliwasps. Thanks also to James Needham, Dr Don Gillespie, Dr Byron Wilson, Dr Robert Powell, Sixto Incháustegui, Jose Ottenwalder, Rick Hudson, Julian Duval and others who have contributed to the making of this project over the years. My special thanks to Dr Robert Powell, Sixto Incháustegui, and my wonderful wife Marcia for help with this article.


Boulenger, G.A. (1885): Catalogue of the Lizards in the British Museum (Natural History). Vol. 2. British Museum, London.

Collar, N.J. (1998): Extinction by assumption; or the Romeo error on Cebu. Oryx 32 (4): 239–244.

Cousens, P.N. (1956): Notes on the Jamaican and Cayman Island lizards of the genus Celestus. Breviora 56: 1–6.

Crombie, R.I. (1999): Jamaica. In Caribbean Amphibians and Reptiles (ed. B.I. Crother), pp. 63–92. Academic Press, San Diego.

Gosse, P.H. (1851): A Naturalist's Sojourn in Jamaica. Longman, Brown, Green, and Longmans, London.

Incháustegui, S.J., Schwartz, A., and Henderson, R.W. (1985): Hispaniolan giant Diploglossus (Sauria: Anguidae): description of a new species and notes on the ecology of D. warreni. Amphibia–Reptilia 6: 195–201.

IUCN (1996): 1996 IUCN Red List of Threatened Animals. IUCN, Gland, Switzerland.

Lynn, W.G., and Grant, C. (1940): The Herpetology of Jamaica. Bulletin of the Institute of Jamaica, Science Series 1: 1–148.

Lawler, H.E., and Norris, C. (1979): Breeding the Haitian giant galliwasp, Diploglossus warreni (Sauria: Anguidae) at the Knoxville Zoological Park. Proceedings of the Third Annual Symposium on Captive Propagation and Husbandry, pp. 73–79.

Myers, C.W. (1973): Anguid lizards of the genus Diploglossus in Panama with a description of a new species. American Museum Novitates 2523: 1–20.

Powell, R., Ottenwalder, J.A., Incháustegui, S.J., Henderson, R.W., and Glor, R. (2000): Terrestrial amphibians and reptiles of the Dominican Republic: species of special concern. Oryx 34: 118–128.

Schwartz, A. (1970): A new species of large Diploglossus (Sauria: Anguidae) from Hispaniola. Proceedings of the Biological Society of Washington 82: 777–788.

Schwartz, A., Graham, E.D., Jr., and Duval, J.J. (1979): A new species of Diploglossus (Sauria: Anguidae) from Hispaniola. Proceedings of the Biological Society of Washington 92: 1–9.

Schwartz, A., and Henderson, R.W. (1991): Amphibians and Reptiles of the West Indies: Descriptions, Distributions, and Natural History. University of Florida Press, Gainesville.

Sloane, H. (1725): A Voyage to the Islands Madera, Barbados, Nieves, St Cristophers, and Jamaica; with the Natural History . . . of the Last of these Islands. Vol. 2. London.

Vogel, P. (1990): Rediscovery of the Jamaican iguana (Cyclura collei). In Conservation of West Indian Herpetofauna through Captive Propagation (eds. B. Johnson and F. Paine), pp. 97–98. AAZPA, Wheeling.

Dale McGinnity, Curator of Ectotherms, Nashville Zoo, 3777 Nolensville Road, Nashville, Tennessee 37211, U.S.A. (E-mail: )

,,Keeping & Breeding The Dwarf Shield Tailed Agama (Xenagama taylori),, by Terry McGleish

The Dwarf Shield Tailed Agama (Xenagama taylori) is a small agamid originating from arid regions of Northern Africa and Somalia.  The tail resembles a miniature "shield", hence the common name, Shield Tailed Agama. They are very hardy lizards which adapt well to captivity.  One unique characteristic we have discovered is that they will dig a shallow tunnel and block the entrance with their tail at night in hopes of deterring any would be predators. The Xenagama taylori is a very personable lizard with many interesting habits and characteristics, many of which resemble the very popular bearded dragon (pogona vitticeps), which makes them an excellent choice as a new breeding project or a pet lizard.

Xenagama taylori grow to an adult size of 3" to 3 1/2" in length and weigh up to 20 grams.  Hatchlings range from 5/8" - 1" in length and weigh as little as 3 grams at birth.  Coloration varies from a dull sandy brown to a brilliant "brick red" body color with varying amounts of black speckling.   Small amounts of  partial white spotting is noticeable on young specimens, but seem to fade with age. Adult males display a brilliant neon blue chin coloration when "fired up", usually during breeding behavior, male combat or a heightened state of alertness.  Some females will also show varying amounts of blue chin coloration, but is very nominal compared to that of the male.

Housing & Substrates:


Communal cage set-up

We have tried several housing and substrate combinations and have had varying amounts of success with each.  At first sand was considered to be the substrate of choice, due to the belief that this most resembled their natural environment, but was quickly discarded because of the lack its tunneling ability.  Our next choice was a combination of cypress mulch and sand, where sand was placed on one end of the enclosure and mulch on the other.  The lizards did not seem to prefer either end over the other, but we did notice they spent the night time hours buried under the cypress mulch.  The problem with this type of set-up was their food items would bury themselves in the mulch and go unnoticed, and after several days and feedings the enclosure would have hundreds of crickets or mealworms running around stressing the lizards.  We have found that garden soil dug from outside works the best.  It has great compacting abilities, which allows for tunneling, is easily cleaned or replaced, and does not offer the food items a place to hide and go uneaten. Substrate should be between 3" to 5" deep as Xenagama taylori are great diggers, they are often seen digging multiple tunnels under and around the rock slabs and/or driftwood pieces supplied for basking.  Their basking area usually consist of a large piece of driftwood and/or slabs of rock or brick.  Temperatures at the basking site range from 90 - 110 degrees Fahrenheit which is supplied by an overhead lamp with a reflective shield.  We use a 75 watt bulb placed 10" - 12" above the highest point of the basking rock.  The cool end of the enclosure is approximately 20 degrees cooler, which allows for thermoregulation.  Water is offered continuously in a small water dish about 1" deep, and is buried to where the rim of the dish is even with the top of the substrate.  The lizards occasionally drink from the water dish, but we noticed they seem to prefer to drink droplets of water which form on the sides of the enclosure and the basking spots as a result of being misted every other day.


Rock slabs provided for basking.

Additional Lighting:

There are some concerns about the amount of UV-B and UV-A light requirements of shield tail agamas.  At this time we offer little or no additional lighting except for an incandescent bulb used for basking. To date, we have not seen any ill effects from the lack of natural sunlight, although a vitamin/mineral supplement with vitamin D-3 is offered in hopes of fulfilling these needs.  Long term deprivation of direct sunlight may prove detrimental, so some exposure is suggested, even if it consist of only a few hours per week.


The diet of the shield tail agama very much resembles that of the bearded dragon, consisting of small to medium crickets, mealworms, occasional super worms and a varied "green" leafy salad.  Crickets or mealworms are offered daily in amounts which will be eaten over the period of the day. This is generally 3 - 5 food items each.  We also dust the crickets/mealworms with a vitamin/mineral supplement such as "Miner-all" or "Reptamin" twice per week.  Although, it is not known what importance leafy matter plays in their diet, finely chopped dark greens and vegetables are offered twice a week in small amounts and are misted with fresh water. Greens offered are: Collard greens, mustard greens, romaine lettuce and endive.   Vegetables consist of shredded yellow squash, zucchini and carrots.  Sub-adult taylori and gravid females seem to relish the greens, but others tend to turn their noses at the offering.



Males (on left) can be identified by the enlarged femoral pores and a yellowish waxy substance present around the pores.  Female on right has very small femoral pores which are barely noticeable.

Sexing of hatchlings and young juvenile shield tails is very difficult if not impossible. Sub-adult and adults can easily be sexed by examining the femoral pores present just above the ventral opening.  A males femoral pores are very pronounced and secrete a waxy substance which is dark yellow in coloration.  The waxy substance is not present on females, and the femoral pores can barely be seen.

Breeding & Egg Deposition

male gravid

Male (left) showing typical blue coloration on chin.     Gravid female (right) shows increased girth as eggs develop.

First thought to be a solitary animal, we kept our lizards separate from each other except during breeding trials.  The animals were brumated for a period of two months from November 15th to January 15th, where temperatures were kept at night time lows of 65 degrees and daytime highs of 80 degrees Fahrenheit, and a photoperiod of 8 hours.  Food was offered once per week in smaller amounts than normal, with water available at all times.  At the end of the brumation period, temperatures and daylight hours were slowly increased along with their regular feeding regimen.  By February 1st they were back to normal feeding schedules and had a 12 hour daylight cycle.   Males were introduced to the single females one at a time, but no breeding activity was noticed.  Thinking that they may be a communal breeder, we set up larger enclosures consisting of one male to four or five females.  Breeding behavior was noticed almost immediately.  The male would "fire up" his chin to the brightest neon blue we have seen yet, and commence to head bobbing erratically and "doing push-ups".  The male will chase the females around the enclosure and literally "wrestle" the female while attempting to breed.  The first time we witnessed this behavior, we thought we mistakenly placed two males in the same enclosure and they were fighting, but upon closer inspection we discovered they were actually in the act of mating.  Males have also been seen copulating with multiple females over the course of a single day.  From our experiences, multiple males or male combat is not required to induce breeding behavior.

Approximately two weeks after the first successful copulation, the females start to show signs of being gravid. The abdomen increases in size and bulges start to appear from the eggs forming inside.  On several occasions, gravid females were pulled from the colony and placed in an egg laying chamber, which consist of approximately 10" - 12" of tightly packed soil, but failed to dig a nest and lay their eggs.  So they were placed back in with the colony, thinking that they were not quite ready to lay.  We then noticed that the gravid females started digging furiously after being sprayed with water during their every other day mistings.  It seems that the females prefer to dig and lay their eggs after a simulated rain, so we started misting the egg laying chambers heavily to induce the females to deposit their eggs.  The female will dig a deep tunnel, approximately 8" - 10" deep and deposit 5 to 8 small white eggs.  After deposition, she will completely fill the tunnel and compact the dirt with her nose. Once finished there are no signs of any tunnels or eggs being deposited.  After the female has deposited her eggs, we remove her from the egg laying enclosure and soak her in a container of water approximately 1/2" deep for 30 minutes so she can get re-hydrated, then place her in an enclosure by herself for a few days to recuperate from egg laying.  After a couple days she is reintroduced to the colony.  We have females which have already deposited their first clutch of the season become gravid for a second time, confirming the belief that they lay multiple clutches during a single season.  The number of clutches per year is still unknown, but we believe they are similar to bearded dragons, and can deposit up to 4 or more clutches per season.

Egg Incubation:

eggs  hatching

Usually 6 to 8 small eggs are deposited in a 10" to 12" deep nest. Incubated at 82 to 84 degrees farenheit, the eggs will  hatch after 45 to 50 days.

The eggs are carefully excavated from the egg laying chamber and placed in a perilite/vermiculite mixture and placed into an incubator calibrated to 82-84 degrees Fahrenheit.  The incubator is kept at 100% humidity by keeping a container of water inside the incubator, and periodic misting with a spray bottle.  After a 45 to 50 day incubation period the eggs start to darken in color and usually hatch within 48 hours.   The young xenagama are left in the incubator for 24 hours to allow the yolksac to be absorbed, and then moved to a small enclosure and kept on a paper towel substrate.   We mist the newly hatched lizards twice a day to keep them hydrated, and offer pinhead crickets after 2 or 3 days of emerging from the egg.  Young Xenagama taylori grow fairly rapidly, and will double in size in the first two months.  Size of the food items are increased as the young taylori grow, and greens are introduced at about one month of age.  We keep the hatchlings and juveniles in communal set-ups identical to that of the adults.  With the fast rate of growth, we believe sexual maturity is reached within the first year, but do not actually attempt breeding until well into their second year.


Captive born 2001 hatchlings at three months are approximately 1 3/4" long.

Although the exact husbandry of keeping Xenagama taylori is not yet completely known, we are well on our way to understanding this unique species.   Successful captive breedings will become more common and will help to promote the shield tailed agama as an exciting and interesting lizard kept by hobbyist.  We urge hobbyist who have had success keeping this species to share their husbandry techniques, and help promote the species.

,,Uroplatus pietschmanni Caresheet,, by Matt Coyne

KOV 5722 400x300


Uroplatus pietschmanni entered the pet trade even before being described by science. It has a variety of common names that include Corkbark Leaf Tail Gecko, Spiny Leaf Tail Gecko and Undescribed Leaf Tail. Corkbark Leaf Tail Gecko is the most commonly used name. Currently almost all of the Uroplatus pietschmanni offered for sale are Wild Caught. These are one of the rarest species of Uroplatus and often a pair will sell for $400+ United States Dollars. Captive bred animals are very rare and there have only been a handful bred worldwide to my knowledge. Because these geckos are so rare, single geckos should not be kept. An effort to breed this species should be made. Since this species is known to com form only a very small locality, it makes it one of the most vulnerable species of Uroplatus. In my experience hey are a very hardy species and many experienced Uroplatus keepers consider them one of the easiest Uroplaus species to keep. None the less they should not be kept by an amateur reptile keeper. The care sheet I have written is a description of how I keep my Uroplatus pietschmanni. I have successfully bred this species keeping them under the conditions I have described.


Curenntly they are only known to come from Amboassary Gara in Madagascar. This region is rumored to be slightly drier and warmer than other areas Uroplatus are found, with the exception of U. guentheri, but thanks to information from Patrick Schönecker this rumor appears to be unfounded.


Uroplatus pietschmanni is an arboreal and nocturnal species of gecko. The most amazing thing about them is their camouflage. They are arguably the most camouflaged of all the species of Uroplatus when in the right environment. In fact Uroplatus pietschmanni was first described in 2003 undoubtedly due to their excellent camouflage. Uroplatus pietschmanni attains a size of around 6 inches as an adult. They for the majority are more robust looking than the other leaf tails. Females are slightly larger than the males. Unlike U. sikorae, U. fimbriatus, and U. henkeli this species does not have any dermal flaps (flaps of skin on the outside of the body). While most of the other Uroplatus species appear smooth and sleek the general appearance of U. pietschmanni is bumpy and the whole body is covered in soft spines. Color varies greatly based on a number of different factors. During the day they are mostly brown colored with patches of green, black, white. At night they are almost a uniform ash grey color. When stressed, cold, or gravid they become dark and sometimes an almost black color. Their most distinguishing feature is the light colored stripe from the eyes down to the tip of the snout. This stripe is undoubtedly to break up the outline of the gecko and suggests that they reside on lichen covered trees. Males and females are easily sexed. Males have a large bulge at the base of the tail and females have none. Females also develop calcium sacks at the base of the neck.


If you are buying Wild Caught geckos you are going to have to deal with a number of issues. Most imports come in a little skinny and dehydrated but generally seem to look pretty good unlike many of the other Uroplatus species. Acclimation is very simple with this species and most specimens seem to bounce right back and get accustomed to their new home pretty quickly. When you first get your gecko keep it in a dark and cool area for about a week. Try to disturb it as little as possible. Make sure you spray it two to three times per day when you first get it. I would also recommend quarantining all geckos individually for a month. This will help prevent the spread of disease and or parasites and allows you to nip any problems in the bud before they spread to your whole collection. Imports will often have red mites on them. These need to be treated as soon as possible. When you first get your gecko make sure to thoroughly check the whole body including the two pads a favorite hiding spot for mites. I use Reptile Relief on mine which works extremely well. Treatment of internal parasites is best left to an experienced veterinarian.


Large screen and glass cages seem to work well and are preferred over glass. Keep in mind these are an arboreal (live in the trees) species so the cage should be vertically oriented. Screen is easier for the geckos to cling to and also allows for better ventilation which in turn keeps mold from growing. I think that the importance of good ventilation is often downplayed with Uroplatus. My cages have glass on the front, back, and bottom with screen on the rest. Plants should be included in the cage. Broad-leaf, sturdy plants with thick stems seem to be preferred over pothos and ficus. I recommend leaving the plants in the pots because not only does it make cage cleaning a whole lot easier but females seem to like to lay their eggs in the pots. Slabs of corkbark and sticks of arm width are placed so that the geckos have access to the whole cage via the corkbark and sticks. For substrate I use Coco-fiber. Other substrates such as peat moss and green moss can be used but in my experience they break down rather rapidly.

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Variety is the key. Crickets and/or roaches make a good staple food but other prey items should be offered if possible. Crickets should be dusted every feed with a calcium supplement and every other feeding with a vitamin supplement. I feed mine moths and grasshoppers during the summer. Snails are taken with special relish by this species but are only accepted by females. Snails are also a great way to boost calcium intact for breeding females. A food cup seems to work well for some of the geckos but others will refuse to eat from it. These geckos will not drink from a dish, at least in my experience, and need to be sprayed. I spray my enclosure very thoroughly once a day.

Note: Make sure any field collected prey comes from areas where pesticides and herbicides are not used.


A heat gradient is provided in the cage with it being around 73-74F on the low end and 75-76F on the high end. I also use a spot lamp for a basking location which reaches a temperature of the mid to high 80’s depending on the season. Temperature fluctuates several degrees depending on season. Night time temps are around 65F but can drop to the low 60’s to high 50’s without a problem. I use a small nocturnal heat light during the winter when it gets really cold which they really like, it also seems to boost their activity and food intake. The main thing to remember with these geckos is to make sure they don’t get too hot. The ambient cage temperature should not enter the high 70F or 80F degree range. On the hottest days of the year the ambient have temperature for me is around 75F-76F.


Humidity is around 65-70%. I don’t strive to keep this species really wet and for the majority of the day their cage is relatively dry. I spray my cage very thoroughly once a day. Since my cage is screen it usually dries out relatively quick.


An incandescent lamp for heat and a full spectrum UV fluorescent light should be used. It is not clear if UV is needed for calcium absorption in this species but it never hurts, plus it will help the plants in the cage grow. Also even though this isn’t a very colorful species of gecko they show their best color under UV light. These are nocturnal species of gecko so a day time and night time should be simulated. Keep in mind that although these are nocturnal they are not cave dwellers so keeping them in the basement with absolutely no light during the night time is not a good idea. Make sure there is some light at night in the room the geckos are kept in.

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Uroplatus pietschmanni seems to be one of the most difficult species of Uroplatus to breed. There have been very few bred of this species and this is one of the reasons I do not recommend this species for beginner reptile keepers. I keep mine in pairs of one male to one female. Some people have experimented in keeping them in groups in various arrangements of males to females but this does not seem to aid in breeding this species. Females lay 2 eggs at 2-3 month intervals. Breeding time seems to be from late summer to spring. Mating involves lots of tail waving plus vocalizations and may appear violent but no damage is actually done. A neck bite is sometimes implied. Gravid females develop a darker and a noticeably different body shape. It is still unclear as to what triggers breeding but an attempt to mimic the seasons of Madagascar is advisable. From the months of May to October, Madagascar is slightly cooler and drier with the other months being more wet and warmer. Mating has often been observed with no egg production. The eggs are white and about the diameter of a dime. Females will not bury the eggs if an egg laying location is provided. A laying location can simply be a small pile of leaves or moss in the corner of the cage. If you keep the bottom of your cage clear besides the egg laying location females will most certainly lay only in that area. This saves you from searching for eggs every time you clean the cage Eggs should be incubated at around 73-75 degrees during the day and 70-72F degrees at night in moist vermiculite. However the eggs themselves should not be in contact with moist vermiculite. Eggs hatch at around 110 days of incubation but can take longer than even 150 days to hatch. The higher the temperature you incubate the eggs the quicker they will hatch.


Hatchlings are approximately 2 inches in length and accept 1-2 week old crickets. Fruit flies are too small and are not accepted. They should be sprayed several times a day to keep them from becoming dehydrated. Hatchlings should be kept in a small enclosure that is not clear. They do have very good visual perception and if kept in a glass or plexi-glass cage will spend countless hours trying to crawl through it. I keep my hatchlings in large cottage cheese or yogurt containers that have been washed out and modified to accommodate them. They can be housed together. Hatchlings should always have food available to them. Besides crickets the hatchlings will also accept small moths and grasshoppers.


These geckos should be handled only when necessary. Occasionally taking them out to take a few pictures or to examine them is okay but frequent handling is not advisable and will lead to a stressed, unhappy gecko.


Uroplatus pietschmanni is a very enjoyable species to work with. Since they are such a rare species I would only recommend them to an advanced reptile keeper. There is still a lot to learn about them and I hope to be adding to this car sheet in the future.

,,Introductory Care and Reproduction of Varanus prasinus, the Emerald Tree Monitor,, by Boelen's Python

KOV 9548 640480

17 October 2011 | Boelen's Python

Natural History and Taxonomy

The Emerald Tree Monitor (also called the Green Tree Monitor) is a medium-sized arboreal monitor lizard that is brightly colored with green and transverse black banding. This species has a highly developed sense of smell and extremely acute vision. V. prasinus spends most of its time perched high above the canopy basking and foraging for food.

It is found in New Guinea and several small adjacent islands, on a few islands within the Torres Strait between Papua New Guinea and Australia, and on Cape York Peninsula in Queensland, Australia. It thrives in lowland environments, including tropical evergreen forests, palm swamps and plantations. Varanus prasinus was first described as Monitor viridis by John Edward Gray in 1831; however, his original specimen was lost. This species was later re-described by Schlegel eight years later as V. prasinus using the found specimen.

The Emerald Tree Monitor is a member of the Varanidae family, which includes a variety of interesting lizards that are commonly referred to as monitor lizards. Monitors vary greatly in size, from about eight in. (20 cm.) long for the pygmy goanna up to ten feet long (3 m.) – the length of the largest lizard of all, the Komodo Dragon of Indonesia.

Captive Care

The Emerald Tree Monitor is highly arboreal and needs lots of vertical space to feel comfortable. It is highly prone to stress and if all captive husbandry is not done correctly, it will not feed properly and could eventually perish from stress-related issues. Single animals can be housed in large vertically-spaced enclosures measuring at least 5 ft. x 3 ft. x 3 ft (1.5 m. x .9 m. x .9 m). Lots of vertical branching and limbs should be placed in the enclosure to allow for much needed climbing and foraging behavior. I recommend using several large broad-leafed plants and some smaller foliage for concealment and humidity. Flat cork slabs can be fixed to the back of the enclosure to allow for vertical access and exploration. This also will allow for arboreal air plants to be fixed to the walls.


One of the most important considerations when attempting to keep this lizard is hydration. Misting multiple times a day is crucial. The animal also should be allowed to gain access to arboreal-placed water dishes in the enclosure. Fresh water should be provided daily.

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Varanus prasinus lives almost 90% of its life high in the canopy. I highly recommend using ultraviolet lighting for this species. I suggest using several Zoo Med 10.0 fluorescent lights fixed to the top of the enclosure. Multiple basking sites with varying intensities also should be provided. Ambient temperatures for the enclosure should be around 78-84° F. (26-29° C.). Isolated basking temperatures for this species should be in the 90°-100° F (32°-38° C.) range. The temperature should not exceed this upper limit and care should be taken when setting up the lights so that the lizard cannot come into contact with them. Varanids can and will burn themselves while basking.


In the wild this species feeds on a variety of arboreal insects, centipedes, spiders, crabs, birds, and small mammals. In captivity it will readily accept small rodents into its diet. It is thought, however, that too much of this type of prey item can lead to unhealthy, obese animals. I recommend offering insects such as Dubia roaches and crickets dusted with calcium three times a week. Offering small pink mice in moderation also is OK.  There have been reports of V. prasinus feeding on a variety of plants and fruits, but in my opinion this is a rare occurrence.


This species can lay as many as three clutches throughout the year. Captive clutches have been laid in January, March, April, November, and December. Clutches consist of up to five eggs. In the wild, the female deposits her clutches of eggs in arboreal termite nests. The eggs hatch between 160–190 days, typically from June to November, after which the young eat the termites and the termite’s eggs. Although this species has reproduced in captivity numerous times, it still can be challenging to have them breed. It appears that this species responds well to heavy periods of rain during the breeding seasons.

Introducing a female to a male typically will result in the male following the female and engaging in tongue flicking and attempts to copulate. The male will approach the female and try to pin her down and maneuver himself so that he can wrap his tail around her to position himself for copulation. It is not uncommon for a male to invert its hemipenis during this time. Copulation can take place from several minutes to several hours. This breeding behavior can occur for several weeks. After breeding behavior has ceased, I recommend removing the male so no unnecessary stress occurs and the chances of the male eating the eggs is reduced.

Depositing the eggs can take from 30-60 days. Multiple nesting areas should be placed inside the enclosure. This will give the female time to explore and chose which nest box she prefers. Once the eggs are found, they should be incubated in a 1:1 ratio by weight of vermiculite to water. Incubation temperature should be 84°-86° F. (29°-30° C.) and the humidity should be as high as possible.


The Emerald Tree Monitor is a truly fascinating species of small monitor to keep in captivity. It can prove to be very entertaining and rewarding as a captive. By following the simple recommendations in this article, you can help them flourish in your care.

,,The Art of Armadillo Lizards (Cordylus cataphractus): Fifteen Years of Captive Observations,, by Gary Fogel

Bull. Chicago Herp. Soc. 38(6):113-119, 2003

The Art of Armadillo Lizards (Cordylus cataphractus): Fifteen Years of Captive Observations

Gary Fogel

kordylus@juno. com

This article is a look back at my personal experiences over the past fifteen years during which I have been keeping and breeding armadillo lizards, Cordylus cataphractus. For you scientific types, I’m afraid this will all be hearsay and anecdotal. Written reports on these little beasts are few and far between. As far as the general population is concerned, many people might not even know that armadillo lizards exist. Even in this day and age of Internet knowledge, no one in America seems to have written any definitive article on the armadillo lizard. After years of waiting for someone else to do it, I have finally decided to do it myself. Here within these pages should be everything you ever wanted to know about Cordylus cataphractus, as my life and theirs have been forever intertwined into one. Let’s start back at the beginning, shall we?

 For me, the odyssey began back in 1985, in the pages of a small book called Lizards in Captivity by Richard H. Wynne,  under the chapter for Cordylidae. There was a short paragraph describing the armadillo lizard and its basic needs in captivity. Of course there were no photographs, just the printed word, leaving the rest to one’s imagination.

 In August 1985, the Chicago Herpetological Society, which I had just joined that June, would have a speaker who would shed some light on this subject. This was John Visser, a South African wildlife biologist, whose topic was the herpetology of South Africa. His slides at this meeting gave me my first glimpse of an armadillo lizard. To me they looked almost wooden in appearance, their scales hand-carved out of some brown balsa wood, with a triangular shaped head, heavily armored from head to toe. After seeing what these clumsy, comical fellows looked like, I knew I had to obtain a few for my collection.

 At this point in the herpetocultural timeline, we had no monthly reptile magazines to look through for animal ads, no monthly reptile shows to buy animals from, and the Internet and personal computers were still in their infancy. We had animal lists, which were mailed out by various dealers throughout the U.S. (less than half a dozen) which, from time to time, might have an animal you might be interested in. One such list from California did offer the aforementioned armadillo lizards for sale. After some hesitance, I called and purchased two. I was keeping primarily geckos at the time, so these were a departure for me into uncharted territory. It was a decision that I have never regretted.

Armadillo lizards occur naturally in South Africa. They are diurnal creatures, reaching an adult size of somewhere between seven to nine inches in length. They live in social groups amongst rocky outcrops, wedging themselves between the cracks and crevices of the rocks, much like a North American chuckwalla does for protection from predators and the elements. These lizards have a long life span --- twenty-five years or more. The surrounding temperatures can get very hot in the summer and cold in the winter, when armadillo lizards will naturally hibernate. They can be a light brown to dark brown in coloration and are sometimes referred to with the common name of golden armadillo lizard. The underbelly is yellow with a blackish pattern, especially under the chin. They are one of comparatively few live-bearing lizards --- they do not lay eggs like most other lizard species. They are insect eaters and have an interesting defense, in that if frightened, they will grab their tail in their mouth and roll into a ball. This behavior is remarkably like that of the mammalian armadillo, which explains the common English name for these lizards. And just as it does for the mammal, this defensive posture enables the lizard to protect its soft underbelly from predators, exposing only its armored back.

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      Figure 1. Armadillo lizards showing typical defense postures. Top photograph by Carlos Sanchez; bottom photograph by the author.


Such was the basic information I had to go on before obtaining my two. When they did arrive, they looked identical in size and shape and I found out that sexing them was somewhat of a mystery.


I housed them in a 30-gallon aquarium with a water dish, hot rock and flat pieces of shale with small rocks glued to the bottom, like little legs, to raise the rock up about one inch from  the floor for them to hide under. I used Astroturf on the floor with more pieces of flat shale, which they would utilize for waste elimination. The two animals were fed crickets and given a reptile multi-vitamin powder. During this time I had yet to start using any fluorescent-type lighting, for fear of burning down the house, so they had only natural light from the windows. I kept them like this for over a year with no ill effects. I tried to find a few more to buy after that, but even then they were scarce. South Africa, it seems, was not commercially exporting reptiles into the pet trade anymore; so except for shipments rerouted to be exported from a different country, you didn’t see a lot of these animals offered for sale. It wasn’t until 1987 that I located three more from a place in Minnesota. When these animals arrived, I placed them in the 30-gallon aquarium with my original two from 1985. After a few days of observation, I learned that my two original lizards were males, something I had suspected all along, since they were identical in body shape and size.


 It seems the introduction of the three new lizards caused the two males not to be best buddies anymore and to now look upon each other as rivals. I would notice that one would chase the other inside the aquarium, while the three new ones got along with each other. The new armadillo lizards were also smaller in body weight and had narrower triangular heads than the two larger males. Could it be that these three were all females? Well, that was exactly the situation, as I later found out. I separated the two males, leaving one in the 30-gallon tank with one of the smaller ones and put the other male into a 50-gallon tank with the remaining two. At this time I started using fluorescent lighting, which I would manually turn on and off daily. No further aggression was noted between any of the armadillo lizards. Since no dealers had any Cordylus cataphractus to sell, it was at this point that I started placing ads in various regional herpetological society monthly newsletters requesting them. This is how I started obtaining my main collection.




Figure 2. The armadillo lizard on the left is a male; the other two are females. Note the larger triangular head of the male in comparison to both female heads. This is a reliable way to sex these lizards. Photograph by Carlos Sanchez.


As people responded to my ads, I would buy them one or two at a time, never knowing what sex I was getting, because no one could sex these lizards with any certainty. I’m sure this was part of the reason people were willing to part with them; breeding lizards commercially was just beginning to take off, and people didn’t want to deal with an animal they couldn’t breed. After seeing a few more armadillo lizards, I determined that females do have smaller, less markedly triangular heads than their male counterparts. Both sexes, however, had femoral pores on the hind legs, thereby confusing a lot of folks who thought they had males because of this feature. They both produce a waxy secretion although it’s more prominent in males. I was never one to probe or pop a lizard’s hemipenes to determine the sex, as I have heard of too many cases where the final outcome was incorrect. I would then house these animals in groups of one male and two or three females, to the best of my sexing abilities. Extra males would be housed alone, although I have housed a few males together without any ill effect. It all depends on the personality of an individual lizard.




Figure 3. Both male and female armadillo lizards have femoral pores on the hind legs. These are roughly equal between the sexes in size and number. The male is on the left, female on the right. Photograph by Carlos Sanchez.


Breeding armadillo lizards, I’d been told, was a daunting  task  - difficult at best. I decided not to handle them excessively and to keep them as wild as possible. My enclosures were in the living room / dining room area of my house and not in a separate animal room. Soon we tolerated each other’s movements. Mine, as I moved through the room so as not to startle them, and theirs, as they made intermittent scuffles that I had to investigate. It took a few months until they felt secure enough to come out from under the rocks to bask under the fluorescent lights. Even then if I walked briskly past, they would all scurry for cover and hide - a domino effect from one cage to the next.  It was in 1989 that I experienced my first live birth. It was December and I remember looking into the aquarium and seeing a smaller version of a head peeking out from under a rock. I did not know that the female was even gravid at the time, so it was a complete surprise! To this day after achieving dozens of live births, I still get an adrenaline rush when I look into an enclosure and see a smaller version of an adult basking or looking out from under a rock. The young are identical to the adults, except smaller in size. They are quite large when first born; total length up to 2.5 inches. Although most literature states it can be one or two young, I have never had a female give birth to more than one lizard at a time. And the scientific literature has recently caught up to this fact (see Flemming and Mouton, 2002).




Figure 4. Note an adult female armadillo lizard (7 inches total length) in comparison with a day-old baby. These babies are rather large, compared to other lizards, when first born. Photograph by the author.


Only once have I actually been able to witness this occurrence of live birth. I was able to photograph the process, but not very well, as it was over in a matter of minutes. I had eight photos left on my roll of film and just enough time to focus and shoot, so of course the photos were over-exposed, but at least I caught the birthing process on film, perhaps for the first time ever for this species. The female came out from under her rocky hiding place into an open area of the enclosure. I knew she was gravid, as she was somewhat swollen on her sides. She just lay there as her sides undulated a bit. She then raised her tail and hind portion and proceeded to give birth. During this time, the lizard would open her mouth as if to utter a silent scream. The baby came out in a thinly membraned embryonic sac. The baby’s body was folded in half inside of this, tip of nose to tip of tail, much like a lizard positioned in an egg would be, except flatter. It then broke the sac and stretched its jaws wide open to get its first breath. The female, having finished the birthing process, scurried off under a rock, and the baby did the same. I was very surprised that the lizard came out into the open to give birth, rather than do so under the cover of a hiding area.




Figure 5. An adult pair of armadillo lizards in mating position during the breeding season. Copulation can take up to a few minutes at most. Photograph by the author.


Once a female has given birth, her job is over and the juvenile is on its own. Since 1989, I’ve produced as few as two to as many as six babies every year. Not every female lizard will always breed from year to year. I’ve had certain groups stop breeding for as long as a five-year period, then start producing young again after this absence. Breeding season for my captive lizards usually runs from the end of January through March. This is the time that I have noticed the most activity between the sexes, with males actively pursuing females in their attempt to copulate (Figure 6). I have determined that gestation for these lizards must be at least four to six months, since most young are born between September through December.


 I have read that these lizards can be cannibalistic towards the young, but I have never seen this to be the case in captivity. There have been many instances where I’ve found the babies and adults alike, basking on top of one another in harmony. During feeding time when crickets are tossed into the cage, an unforeseen baby lizard will dart out from under a rock, past an adult lizard, to chase down a food item almost as big as he is, in an effort to subdue his prey. Once I find the captive-born baby lizards, I place them in a separate enclosure. Sometimes a baby lizard will not readily eat on its own. If small crickets are ignored, I’ve found that waxworms will usually be eaten without problems. The advantage of feeding baby armadillo lizards in a group is that when food is offered, a feeding response kicks in and when one lizard begins feeding the others usually will follow suit. Care should be taken to insure that all the lizards are equally fed, so that one individual does not eat most of the food, leaving the others still hungry. I’ve found that one or two feedings per week is sufficient to ensure good health for adults, perhaps more for the baby lizards. I also have noticed that these lizards are scavengers, searching the ground for items to consume. This was discovered when I placed a small rubber piece under a rock to help keep it level. One day it disappeared, only to turn up a few days later, partially digested, on the floor of the cage. Some of the lizards were nibbling at the Astroturf also, leading me to believe that they might eat plant material. I experimented with throwing small pieces of kale into the enclosures. The result was that some of the lizards did indeed eat the plant matter. Some, of course, choose to ignore the kale and stick to their insectivorous diet. So some armadillo lizards will accept plants as well as insects for their dietary needs, although I would never recommend a diet of just vegetables for these lizards. Armadillo lizards will also accept small pinkie mice, but I worry that an exclusive diet of these would result in too much dietary protein for the animal. One interesting behavior I have witnessed on more than one occasion, is that of armadillo lizards gnawing on large rocks. They would just try to take a bite out of one, attempting this behavior for a minute or so. Perhaps they are trying to consume rock material to aid in digestion, as some lizards are apt to do. Maybe they are trying to consume rocks for mineral supplement needs, or as part of their daily scavenging routine. I have never witnessed this behavior with any other type of lizard I have kept in captivity.


 Armadillo lizards are extremely social animals, living inlarge groups in the wild. For this reason I raise up each year’sbabies in a family group, so to speak. Males seem to becomesexually active around the third year of maturity. Since Iusually house all previous offspring together from each year, they grow up and get along without incidents, usually until this third year. This is when you will notice if more than one male is present in a group, as one will become the dominant alphamale. Although they live in social groups in the wild, introducing a new lizard into an existing group in captivity may have negative consequences. If a few days after the introduction an animal is constantly chased and harassed, it is best to remove this animal; otherwise it will be bullied to death.  I’ve had instances where I tried to introduce a new female to an existing pair of armadillo lizards, only to have the male constantly pick on the new female. I then tried a different pair only to find that, this time the existing female was the aggressor towards the new female and not the existing male.


Another example shows that males aren’t the only ones who can display aggressive behavior towards a newly introduced animal. I had a group of three females whose male had died, so I wanted to introduce a new male. In the meantime, one of these females had claimed the alpha spot and she would chase the existing females around. The day I introduced a new male, she came charging out to confront him, but he held his ground against her and eventually she went back to submissive from dominant behavior.




Figure 6. A group of juvenile armadillo lizards warming themselves on a heating pad inside the enclosure. Photograph by the author.


Armadillo lizards display the typical lizard communicating skills of head-bobbing, tail-wagging and tongue-flicking, when confronting cagemates or another unfamiliar armadillo lizard. These lizards have very powerful jaws, which can be very hard to open. When they do fight, they can clamp down on legs and toes, biting them off and definitely drawing blood. They can bite down on the sides of the body and start rolling or twisting their opponent’s torso. This can cause internal damage to the animal. Although they do not readily bite in captivity, they will sometimes open their mouths to try to bite their tail. I did not witness this behavior until about a year after I had obtained my first two armadillo lizards. I had them both at an educational show, when all of a sudden one just curled up into a ball! Once they do this it is almost impossible to make them let go, until they are good and ready. I have seen them hold this position for almost up to an hour. When it happened the first time, I removed the lizard and placed him into a bag to make him feel secure until he finally let go of his tail. Because of this behavior, armadillo lizards are more reluctant than many other lizard species to dispense with their tails. This is one of the features that drew me to these lizards initially, as most of the geckos I had been keeping previously would drop their tails quite readily. If a portion of an armadillo lizard’s tail is lost, the regeneration will occur in segments, much like the original tail. I have individuals who almost never bite their tails, and others who will perform this behavior on cue, should I desire to demonstrate this ability to someone. This is an instinctual behavior, but you certainly do not want to stress out an animal by forcing him to perform this defense over and over.


 In the course of keeping armadillo lizards I have gone through various cage incarnations, from aquariums to hand constructed units, which resemble bookcases. Since these lizards are rather clumsy at climbing, my enclosures are all open at the top. The three sides are 12 inches tall laminate board and the front is Plexiglas (Figure 8). As long as no large rocks are placed along the edges, there is no possibility of escapes. Armadillo lizards do like to climb rocks, on top of which they will bask under the lights. My units are five feet long by two feet wide. I use a four-foot fluorescent lighting fixture for each enclosure with one full-spectrum bulb and one blacklight bulb. These fixtures hang twelve inches from the floor of the cage. After years of using these bulbs, I’ve wondered as to the validity of such lights, since I don’t replace them every year. Because the UVB output has surely declined with the aging of the bulbs, my theory is that these types of lights are more beneficial in triggering a breeding response than with aiding calcium absorption. As long as a vitaminmineral powder is used and the food items are fed a high calcium diet, I believe that full-spectrum lighting over regular fluorescent lighting does not make a big difference. I have never used a heat lamp since in the summer months it would get too hot, and in the winter months, I prefer to keep these lizards cool, to mimic their natural seasonal change. I utilize a heating pad in each cage, which is turned off during the spring and summer months. I also shorten the timers on the lighting cycle to mimic the natural lighting outside.


 I like to keep my cage furnishings simple for cleaning purposes, yet esthetically pleasing. I include several flat rocks for hiding places and various larger rocks for climbing. An Astroturf rug covers most of the floor, except for the front of the enclosure. This whole area I leave exposed for a six-inch width, and this is where the lizards defecate. Since it is right in front for the length of the enclosure, it makes cleaning up on a daily basis extremely easy. I want each enclosure to look the same; I feel that this reduces stress levels should I have to move any animals around. This type of cage design seems to work best for the lizards, as well as my needs. I use a very low, flat, water dish, about an inch in height. This makes it easy for armadillo lizards to drink out of it. If it were much higher, the lizards might not even know it was there. Before I switched to the lower type bowls, I had one lizard that used to lick the side of the water bowl, sensing that water was there, but not possessing the brainpower to approach the dish from the top of the bowl. I have had only one bad experience using a low-sided water bowl. One morning I noticed an object in the water, only to discover that during the evening, a baby armadillo lizard must have been born and wandered into the water by accident and drowned. I had placed a small rock right next to and level to the top of the bowl a few weeks earlier, to aid the lizards in drinking more easily. The end result for me was an accidental death. I hope that such an unfortunate occurrence will never happen to anyone else. Always leave the area around a water dish clear of any objects.


 Medical problems have been minimal, but there have been a few that I’ve encountered throughout the years. Loss of digits is common, as the toes are very small and delicate, and a single bite during mating or fighting can easily amputate one or more. I have several lizards missing various digits, but with no ill effect to the animal. Mouth infections are the most aggravating problem to treat. You can tell if an armadillo lizard has this if, during routine feeding, the animal runs around chasing its prey, but then does not try to bite it. If you look into this animal’s mouth and see a white pus-like area, it has a mouth infection. If left untreated, the infection usually spreads to the eyes and ears, resulting in an oozing swollen eye or tympanum, as the infection works its way throughout the head area. Treatment consists of cleaning the infected area together with the use of an injectable antibiotic until the symptoms have subsided. My most severe case of this involved a female who just would not open her mouth no matter how hard I tried to make her. Eventually I had the idea to stick the eraser end of a pencil in between her jaws so she could not keep closing her mouth. Upon doing this, she clamped down so hard on the eraser that she cracked her bottom front jawbone in half! These lizards have very powerful jaws and this proves it. That is why injections are my preferred choice over any oral medication. Not only did I have to treat the mouth infection, now I had to treat a broken jaw as well. I also determined that she was gravid at this time. To help speed the healing process, I decided to house her in a screened aquarium and stick it halfway out the window like an air-conditioning unit, to expose her to natural sunlight. The part of the aquarium inside the house provided the hide area and the shade, so she could venture out into the sun when she felt the need to. When all the medication was done, her jaw mended, although a bit crooked, and she eventually gave birth. I still have her in my collection to this day. This is just an example of how tough and hardy these little lizards can be.


 The second medical problem I have encountered is a femoral pore infection. This causes the rear legs to swell up and accumulate pus behind the femoral pores. The lizard will continue to be alert and eat regularly as if nothing is wrong, but if no medication is administered, the back legs will continue to swell until the lizard has trouble walking. To treat this type of infection, injections must be given and the pores cleaned out by gently squeezing out the pus and bacteria until the infection subsides and heals. I’m not sure what causes this, but it may have something to do with not enough rough substrate to continually rub up against, thereby keeping the femoral pores cleaned out naturally. These pores do secrete a waxy substance in both males and females that could build up and clog the pores. Since encountering this type of infection, I’ve included rougher climbing rocks to help scrape off these pores on a periodic basis. I have not had any recurrences since this was done.


 The last medical problem is a common one of the occurrence of arasites. This again involves a lizard that is not eating, but shows no interest in food, whatsoever. If this happens, first I’d check the mouth to make sure it is not a mouth infection. If that looks all clear, then a stool sample is taken to determine what parasite is present and what treatment is in order. After a week or two the lizard is usually back to normal and eating regularly. Sometimes in the colder months armadillo lizards will go off feed for a few weeks, as they would be hibernating in the wild at this time. If this happens, just keep an eye on the animal to make sure it’s not losing any weight and that it is still healthy. Eventually it should begin eating again when it feels ready. Make sure to check on a lizard if it is hiding all the time and never comes out. It could have an infection or be stressed out.


This has occasionally happened to me. By the time I had noticed there was a problem, it was too late to save the lizard in question. The key to good husbandry is to always observe these lizards. You become aware of their habits and movements so if something doesn’t seem quite normal it can be taken care of immediately.


 Armadillo lizards are not as common in the U.S. as many other types of exotic lizards. This is due to the fact that they are not offered commercially through the pet trade with any regularity, nor have they been since 1989. Another reason is that these lizards are not a moneymaking commodity for a lot of reptile breeders. They are slow to reproduce and only net one offspring each year, per female, if you’re lucky. Compare that to your more acceptable, double clutching lizards, who lay dozens of eggs each year, and you can see why most breeders will not bother with a lizard that is not going to contribute substantially to profits. People do keep them in this country, but not in any great numbers. Few American zoos, even, have bred them in captivity. You normally will not see them at any of the reptile shows held around the country, because nobody is really captive breeding them, at least for general sale to the public. Your best bet in obtaining this type of lizard, is the same way I purchased mine years ago - by placing various ads and hoping someone responds. You can also network through the Internet to see if there are any available. Once you do find one, there is no guarantee as to how old it is going to be or what sex. Remember, I still have my first two male armadillo lizards from 1985, and they were adults then, so who knows how old they really are at this point? Also, they are still viable in the reproductive department, mating and breeding to this day. In this day and age of instant gratification you may have to be patient in obtaining these lizards. I had to wait two years before I found a few more to add to my original two.


 Let’s now review the basics in keeping these lizards in a captive environment.


Cage enclosure:


A large aquarium will do for two or three of these lizards, although I like to give them as much room as possible so that they can behave in a more natural manner. It’s always more fun to see them run around --- something that they can’t do in a small aquarium. I’ve found in a few of my large enclosures, males will usually stay to one end of the cage, with females grouping on the other. It is very important to include several hiding areas throughout the cage and let the animals choose which one is best for them. Flat rocks, propped up an inch off the ground work best. Armadillo lizards like to be able to wedge in between rocks naturally, so they like to be able to feel that roof of rock on their backs. If you were lift up one of these rocks, you would notice that the lizards arch up their backs in order to feel where the top of their hiding place is. Avoid a hide area with too high a ceiling, as the lizard may get stressed if not able to hide properly.




Figure 7. A typical cage enclosure, five feet long by two feet wide, with a Plexiglas front. Photograph by the author.


The drinking area should be shallow - one inch or so high - so they can find water without a problem. Large rocks should be included for climbing and basking purposes. You can use a rock or sand substrate, but they will probably eat it, which may lead to medical problems. Astroturf works for me, although a few have tried to eat that too. I recommend full-spectrum


lighting in combination with blacklight lighting, kept as close to the lizard as possible (12– 15 inches is best). You can start out with this lighting and judge for yourself how beneficial, or not, it is. Some sort of heated area (heating pad, hot rock) is needed in the autumn and winter months, especially in the colder states in the U.S. Use common sense with these items. If the area gets too hot, it may be necessary to diffuse the heat with ceramic tiles on top and underneath. I use a twelve-inch heating pad on top of which I put a twelve-inch ceramic tile. I have never yet had an armadillo lizard burn its underbelly lying on one of these.





A room temperature of 70– 85°F is fine, perhaps a bit cooler in the winter, as they can certainly tolerate it. I have always worried about my lizards getting too hot, rather than too cold. The room that they are now in can get up to 100°F in the summer. I keep a small air conditioner in one of the windows to keep the temperature down when this happens. A fan of some sort to circulate the air is also desirable. Although they can tolerate high temperatures, too much might stress them out. In the winter they are still active with the cooler room temperatures, (65 to 70°F) as they utilize the heating pads during the day at this time of year.





The basic staple diet consists of crickets, superworms, butterworms and waxworms; the latter two have a higher fat content so I use them sparingly. I actually have two groups who would rather eat superworms than crickets. They will only eat crickets if they get good and hungry enough. I feed my lizards once a week, making sure that everyone gets fed. When they are in groupings of four to six, you have to see to it that certain lizards aren’t hogging the food items. Always put in only enough food items that all will be eaten, and watch to make sure this is happening. This is especially true for superworms, for if they crawl away somewhere; they will morph into a large black beetle that the lizards do not like to eat. You will probably find one somewhere outside the cage enclosure, perhaps running across your carpeting at the most inopportune time. Always use a vitamin-mineral powder on your food items. The smell of this helps the lizards to better locate the crickets and it gives the lizards the essential vitamins they may otherwise be missing in an indoor, captive environment. Also, feed the insects you intend to use as food items twenty-four hours before you give them to the lizards. Kale, collard, turnip, and mustard greens work well because they have a high calcium content. By feeding your insects, you’ll find that they will live longer and you won’t have to keep getting them so frequently.


 Armadillo lizards are one of the most easily kept lizard species. Their longevity in captivity and relatively few special requirements make them an excellent choice as a captive lizard pet. Unfortunately, they are scarce in the pet trade and command a high price as a result.


 I hope this information is beneficial to anyone who currently keeps armadillo lizards or who may just want to learn more about them. They are an overlooked, little known species who, I think, are one of nature’s most interesting lizards. May they never go extinct!





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Curtis Books.


Flemming, A. F., and P. Le F. N. Mouton. 2002. Reproduction in a group-living lizard, Cordylus cataphractus (Cordylidae), from


South Africa. J. Herpetology 36(4):691-696.


Fogel, G. 1994. Live birth of an armadillo lizard. (Photo essay) Bull. Chicago Herp. Soc. 29(1):8-9.


Patterson, R. 1987. Reptiles of southern Africa. Cape town, South Africa: C.Struik (Pty) Ltd.


Rogner, M. 1997. Lizards, Volume 2. Malabar, Florida: Krieger Publishing Company.


Switak, K. 1995. Girdle-tailed lizards. Reptiles Magazine. 3(6): 8-10,12,14,16,18,20,22,24, October


Sorensen, D. 1986. Interview on armadillo lizards. Wisconsin Herpetological Society Newsletter 7-11, November.


Stanek, V. J. 195?. Introducing dragons. London: Spring Books.


Wynne, R. 1981. Lizards in captivity. Neptune, New Jersey: T.F.H. Publications


Zimmerman, E. 1983. Breeding terrarium animals. Neptune, New Jersey: T.F.H. Publications





,,Spearpoint Leaf-tailed Gecko (Uroplatus ebenaui),, by


Introduction: The spearpoint leaf-tailed gecko belongs to an unusual group of nocturnal geckos native to the island of Madagascar. All geckos belonging to the genus Uroplatus have cryptic coloration and patterns to mimic different parts of their environment. The spearpoint leaf-tailed gecko mimics dried leaves. This species varies in color from dark chocolate brown to light tan. Some geckos are even red, burgundy or orange in color. Most have a reticulated pattern covering their body to some degree. Small fleshy projections or spikes jut out from their body and are particularly noticeable over their eyes and on their head. They are the smallest Uroplatus species, generally maturing to just under 4 inches (10 cm) in total length. Uroplatus ebenaui resemble the closely related satanic leaf-tailed gecko (Uroplatus phantasticus), but lack a long leaf-like tail. Instead, the spearpoint leaf-tailed gecko has a short angular, almost diamond-shaped or spear-shaped, tail. When threatened, some spearpoint leaf-tailed geckos will drop their front arms from the perch they are on and hang from their back legs to very accurately mimic a dried leaf.

Male geckos can easily be distinguished from females by the presence of their hemipenal bulge. Some have also suggested that tail size and shape can be used to sex spearpoint leaf-tailed geckos, but looking for a hemipenal bulge is more reliable. Most people agree that male spearpoint leaf-tailed geckos can be housed together provided the cage is large enough; however I have no personal experience housing multiple males together so I’m not able to agree with or dispute this claim. I strongly encourage those interested in Uroplatus to purchase geckos in male-female pairs or trios for the purpose of breeding, rather than simply for display or pets.

Starting with healthy stock is important. Whenever possible, purchase geckos that were born in captivity because they will generally be in better health than their wild-caught counterparts. Unfortunately, it can be difficult to locate captive-bred geckos because nearly all spearpoint leaf-tailed geckos available in the pet trade are wild-caught. Avoid purchasing a gecko that is active during the day, has sunken eyes, open wounds or sores, is missing a tail, or is being kept in poor conditions. I would suggest that each individual wild-caught gecko be quarantined in its own simple enclosure for one or two months regardless of how healthy it appears when purchased. This will provide the keeper time to monitor each individual’s weight and appetite, as well as prevent diseases from spreading between geckos. Uroplatus have a reputation for arriving from Madagascar with large parasite problems so it may be beneficial to take a fecal sample of each gecko to a veterinarian and then medicate accordingly. Wild-caught Uroplatus also often arrive severely dehydrated, so it’s important to provide plenty of water to new arrivals by misting the cage frequently, or even placing the gecko in a hydration or rain chamber for a few hours.

Cage: Spearpoint leaf-tailed geckos are not very active during the day but at night they will use all of the space that is provided to them. A 15 gallon high aquarium that measures 20 inches long by 10 inches wide by 18 inches high (51 cm by 25 cm by 46 cm) is large enough for one male-female pair of geckos. Smaller cages can be used for housing single animals. It’s important that the sides of the enclosure are made of a material that will help maintain a high humidity level, so screen cages should be avoided unless the ambient humidity in the room they are kept is already high. A tight-fitting cover is also important. Screen covers can be used if plastic wrap or glass is taped over half or more of the cover to help keep the humidity level high. If the cage is made of glass or other see-through material it can be beneficial to cover all but one side of the cage with an aquarium background or black poster board.

The main components of their setup should include a substrate, hiding spots, and perches. Paper towels work well as a substrate, particularly in quarantine and isolation cages because they make it easy to collect fecal samples. They are cheap, easy to clean, and easy to replace. It is important to replace them on a regular basis to prevent unwanted bacteria and mold from growing. Another substrate that can work well is a safe soil such as coconut husk fiber (bed-a-beast, eco earth, forest bed, etc.). Foam rubber also is suitable as a substrate, and can be used both in temporary and permanent setups. Substrates such as gravel, moss, and small pieces of bark should be avoided. It can be beneficial to place a layer of leaf compost or dried leaves over the substrate. Oak and magnolia leafs both tolerate high humidity levels and are good choices. Take care to prevent the substrate from becoming too wet. If the substrate is squeezed in your hand and more than a few drops of water come out then it is too wet.

                                       spearpointcage01  spearpointcage03

Hide spots, perches, and other cage décor can consist of live or fake plants, branches, driftwood, dried leaves, and pieces of cork bark. It’s important to provide perches that are thin enough for the geckos to hold on to. There should be at least a few small branches or perches that are about the width of the gecko's limbs. I’ve found that bird perches made of manzanita wood work very well. Some pet stores and terrarium supply companies also sell manzanita branches specifically for reptiles that don’t have the metal screws on the end like the bird perches. Cork bark slabs make great hiding spots when positioned at a vertical angle against the side of the cage. If live plants are used, purchase them from a garden center or terrarium supply company that does not use pesticides or fertilizers that could be harmful to reptiles.

Established, healthy spearpoint leaf-tailed geckos can be kept in living terrariums. See the article about tropical terrariums for information about creating one.

Temperature and Humidity: Spearpoint leaf-tailed geckos need to be kept cool. A range in temperature from 68°F to 75°F (20°C to 24°C) during the day with a drop to between 60°F and 70°F (16°C and 21°C) at night generally works well. They will not tolerate warm temperatures and often die when exposed to those above 80°F (27°C) for extended periods of time. The best way to keep the cage cool is to keep it in an air conditioned room or cool basement. Occasionally, it may be necessary to cool the cage by placing an ice pack on top of it. Supplemental heating is rarely required, however if it is needed I would suggest using a low wattage infra-red light bulb as opposed to a heat pad or heat tape.

Maintaining high humidity is as important as maintaining cool temperatures. The humidity level in the forest vegetation and scrub layer in Madagascar is constantly high and this should be duplicated in captivity. The humidity level in the cage can range from 80% to 100%. This can be accomplished by spraying the cage with water once or twice a day and restricting ventilation.

Water: A shallow water dish can be provided but is not necessary as long as the cage is misted with water on a regular basis. Leaf-tailed geckos generally prefer to lap water droplets off of plant leaves, glass, and other smooth surfaces as opposed to drinking from a bowl of standing water.

Food: The majority of a spearpoint leaf-tailed gecko’s diet should consist of crickets. In addition to crickets, they can be offered wax worms and small silk worms in a shallow dish. Small moths and other flying insects can also be offered. Some have suggested that fruit baby food will be accepted by leaf-tailed geckos but I have never observed my geckos showing any interest in baby food. Offer adults between three and six food items per gecko two to three times a week at night. If there are live feeders in the cage the next morning, the geckos are being offered to much food at once. Feeder insects should be coated with high quality vitamin and mineral supplements every two to three feedings. Juvenile geckos should have their food supplemented at every feeding.

Last updated 03.19.08

Online Resources
Fauna Imports: Uroplatus
The Herp Venue: Spearpoint and Satanic Leaf-tailed Gecko Care
Nature's Dead Leafs and Pez Dispensers: Genus Uroplatus
Of Another Color: Uroplatus
Uroplatus phantasticus Care Sheet by Robert Gundy


Uroplatus sikorae (sikorae and sameti) - Mossy Leaf Tailed Geckos By Hervé Saint Dizier, Caen, France (abridged)

Class: Reptilia, Order: Squamata, Sub-order: Sauria, Infra –Order: Gekkonomorpha, Micro-order: Gekkota, Family: Gekkonidae, Tribus: Gekkonini, Genus: Uroplatus

The genus Uroplatus is endemic to Madagascar and the coastal islands. The nominal subspecies inhabits the remaining patches of the Eastern rainforest at an average altitude of 3,000 feet. These luxuriant primary rainforests have a very damp climate with rather cool temperatures (average 70°F ,mean range 35-80°F) . There is an alternance of a “dry and cool” season and of a “rainy and warm” one (November-March) ,mating taking place during the first weeks of the latter . The subspecies sameiti ,dwelling on the coastal island of Nosy Bohara ,is exposed to a constantly damp climate (hygrometry above 80%)and to slightly higher temperatures ,in the low 80s . It is thus a little more tolerant to heat peaks in the vivarium .These geckoes rest on tree trunks during the day ,their legs stretched alongside the body and their heads generally oriented downwards . Their dermal flaps all along the body and legs give them a perfect and shadeless camouflage ,along with their basic brown-grey colour enhanced by lichen- or moss-like patches with various hues of green ,grey ;even yellow ,orange or red on some individuals .Their patterns are unique and there is no individual perfectly similar to the other .Furthermore ,The colour and pattern of the young seem totally independant from those of the parents (F.CAVY ,pers.comm.).Their mimicry with rough bark is absolutely stunning ,males tend to be more bark-like than females .They are active at night ,jumping on moving prey in a quite spectacular manner .

Uroplatus sikorae is a mid-size uroplatus (average SVL 4 to 5 inches ,total length up to 8 inches for the sameiti subspecies ).It has a broad mouth enabling it to swallow large prey items ,a long ,rounded snout ,big protudent eyes with a yellowish mottled iris pattern and a vertical cat-like pupil .The eyes are often lined with a bright yellow circle .Endolymphatic calcium sacs are commonly seen in captive individuals .The tail is flat ,leaf-like ,made of dermal tissue and counting for about 40% TL .They can throw off the tail completely ,always the entire tail but never a part of it as some other geckoes do ,in order to escape some threat ,but it never regrows as the original .The belly is often spoted with black tiny points and brick red hues are not uncommon .The body has a “triangular” section being dorsoventrally flattened ,legs are stout enough to enable it to jump quite far and fingers end up with rounded “adhesive” structures made of microscopic hair called setae .

Sexing the adults is unproblematic,as the males display huge hemipenal bulges .There are no femoral pores on this species .

U.sikorae sikorae and U.sikorae sameiti can be distinguished by the colour of their buccal mucous membrane :it is black in the nominal subspecies and pink for U.s.sameiti .It is said (SVATEK and VAN DUIN ,2001) that some individuals in the area of Montagne d’Ambre (Northern Madagascar )reach lengths of 9 inches ,thus building up a “giant morph” .

Longevity in good captive conditions may reach 7 years .


Uroplatus sikorae is a very delicate species ,especially as regards humidity and temperature .It simply does not stand high temperatures ,and a fast death can be expected if exposed to more than 80°F for only a few hours .Wild-caught specimens are also heavily loaded with a broad variety of parasites .Each newly bought specimen should be individually quarantined for a month at least and rehydrated with frequent spraying of the vivarium. These animals are particularly subject to stress ,and handling them is definitely not a good idea .They are good display lizards but handling should be strictly limited to necessary operations like a veterinarian treatment .Too much stress unavoidably kills them on the short term .

I underline that males are not aggressive towards others of their gender ,thus they can be housed together with females without any risk of aggression .


Glass tanks taller than they are long are perfectly suited for Uroplatus .I kept a trio in a 32x16x32 terrarium. Ventilations should be sufficient to prevent dampness stagnation and the apparition of mould and rot ,but care should be taken not to allow the vivarium to dry up too much during the day .

The substrate is made of a bottom layer of small clay balls or vermiculite to retain humidity and drain excess water at the same time .Expansed coconut fibers ,or sterilized peat ,makes up the second layer .As uroplatus vividly dive on prey ,no stone ,nor hard or sharp item should be present in the enclosure . Plants like bromeliads ,orchids ,live moss ,small ficus species should also be used to retain humidity and to provide the animals with suitable egg-laying sites .A neon tube with 5% UVB is ,in my opinion ,useful at least for the plants and I am convinced that the animals receive a non-negligible UV dose while resting on tree trunks at daytime in the wild .So it seems accurate to provide them with UV light in captivity too .It will also create a very localised “basking spot”,appreciated by gravid females for instance .Day-and night rythm should be 13 hours of lighting and 11 hours of darkness in he “hot and wet” season ,the reverse during the resting “dry” period .

Heating devices are perfectly useless if the vivarium is kept inside ,they are on top of that dangerous for the animals .If the reptile room is very cold ,8 to 15W heating carpets arranged vertically outside the reptiles environment can be used .The whole installation should also be kept away from everything that dries up the atmosphere like radiators .Above 80°F ,the Uroplatus face a thermic stress ,deadly temperatures begin at about 85°F .The ideal day range is 69-73°F and 60-67°F at night during most of the year ,and 78°F is fine in summer .

An Uroplatus exposed to high temperatures is unavoidably condemned to death .

The vivarium should be sprayed twice daily ,in the early hours of the morning and in the evening as lights are switched off .Spraying must be abundant but it is not a good idea to turn the substrate into a swamp .Hygrometry should slowly go down during the day and come to a peak again at night .Uroplatus with insufficient humidity soon dehydrate and meet severe shedding troubles.


Uroplatus sikorae are mainly insect-eaters .Dead prey will never be accepted ,as far as my experience goes .For lean animals or gravid females ,a living pink mice a month is benefic .They also love small snails and can thus absorb the calcium from snail shells ,which is good for pregnant females or just after egg-laying .The best size for snails is around ½ inch (shell diameter) .

All food insects are properly gut-loaded and given every two days .I don’t use a feeding dish ,I prefer leaving the insects wandering in the vivarium or giving them with tweezers .They are generously coated with Miner-All I on every feeding and every fornight I add extra vitamins in a small amount .Bimaculatus ,grasshoppers ,cockroaches ,waxmoths larvae ,morios are equally accepted.Uroplatus are voracious eaters once acclimated.


As I previously said ,courtship begins shortly after the beginning of the rise of temperatues and humidity .A resting period in the winter month (for the North hemisphere ) is therefore necessary ,as well as for the welfare of the animals .Mating is gentle and I did not witness any biting from males .
There are 3 to 5 clutches a year ,and amphigonia retardata enable the females to be fertilized for several clutches .Eggs are either buried in the substrate ,or in the pots holding the plants ,and are rather soft-shelled ,about ¾ inch long ,and bright white .Female roll them between their hind feet to make earth particles adhere to their surface. Clutches occur at a 4-6 week intervals and females should be well-fed (snails ,pink mice )and supplemented with high doses of calcium afterwards .Eggs ,without being turned ,are transfered with care into an incubator.Young females tend to lay a single egg but two is the common rule .The incubator is made of a plastic cricket box filled with small clay granulate or vermiculite ,laid on a weak-powered heating device (carpet or cable ) ,15W is sufficient .Eggs are half-buried in the substrate which must be very wet .Heating is stopped at night for 10 hours and during the day the incubation temperature vary from 72 to 78°F and a constant hygrometry of 85%-90% .It is harmful for the eggs to receive drops from condensation.Babies hatch after an average incubation duration of 75-90 days .Their basic requirements are the same as for adults but they are extremely fragile ,measuring around 2 inches (SVL ) and should be transfered in 8x8x12 inches individual terrariums for a better monitoring of their health and growth .They are not offered vitamins until 3 months old and their basic diet is made of small crickets coated with Miner-All I and sometimes tiny snails offered every evening .They are even more vulnerable to the lack of humidity and high temperatures than the adults ,so it is better to keep them under 75°F .


Since October 2004 ,all Uroplatus species are considered as endangered by the Washington Convention ,thus being classified as Annex II of the CITES treaty .


Uroplatus pietschmanni (Cork Bark Leaftail Gecko)

This is a caresheet given to me, original author unknown - all credit to whomever wrote it with my thanks.

Corkbark Geckos (U. pietschmanni):

Along with U. henkeli, U. pietschmanni are amonge the easier Uroplatus geckos to care for.  While cool temperatures and humidity are important, as they are with all Uroplatus, U. pietschmanni can tolerate a wider range that some other species.

TEMPERATURE:  Temperatures in the high 60s to high 70's are best with a day/night fluctuation.  My U. pietschmanni have tolerated nighttime lows to 60 degrees and daytime highs to 84 degrees with no ill effects, but these extremes should not be prolonged.

HUMIDITY:  Overall, U. pietschmanni like a humid cage, but not a dank, moist one.  The substrate should never become water logged or muddy.  because this will lead to bacterial infections.  The cage should be thoroughly misted at least once in the evening, and the surfaces of the cage and its furniture should be allowed to dry during the day so that there is no standing water left.

I keep a pair in a well ventilated cage (36H x 20 x 20 inches with a screen on the top and on one side).  The cage has several inches of topsoil in the bottom and is well planted, so it holds humidity.  I mist the cage thoroughly~~very thoroughly~~ every evening.

Water droplets remain until morning, but during the day the cage dries so that no water droplest are visible.  This regime seems to be working well.

FURNITURE:  They like thick branches to climb and sturdy broadleafed plants.

FOOD:  In my experience, Uroplatus geckos will only eat bugs that have legs, ie., no worms, grubs or caterpillars like mealworms, waxworms or silkworms.  My U. Pietschmanni eat mostly crickets, and also receive occasional roaches and moths.  I have found that new imports like moths.  They must recognize them from back home.  They are hardy eaters and don't refuse food.  Since I have laying females, I lightly dust nearly every feeding with a calcium supplement, and I add a vitamin supplement about once per week.

BREEDING:  I have found that females breed and lay fertile eggs when pairs are housed individually.  I recently moved two pairs into a large community cage and now most of the eggs are infertile.  This change could be for a variety of reasons.  All I can say is that I had better breeding success when the pairs were housed individually.

A female will lay two eggs about every 6 weeks or so.  The eggs are brittle, white spheres (amazingly large).  I have had hatching success in the following incubator:  plastic container with small holes for ventilation; moist vermiculite to sustain humidity;  sponge on top of the vermiculite with dimples cut into it in which the eggs rest.  Even this arrangement sometimes proved to be too moist as moisture permeated the sponge from the vermiculite.  So, I cut a piece from a plastic bag with the same dimensions as the sponge and placed it between the sponge and the vermiculite.  Hence, the eggs like a humid environment, but direct contact with moisture quickly drowns them.

Eggs hatch in about 3 months.  After a few clutches the female should be separated from the maile and given a couple months to rest. 

HATCHLINGS:  The hatchlings can be kept like the adults but in a small cage so that food is easily accessible (5 gallon glass terrarium with screen top).  The hatchlings shed within their first day of life outside the egg (sometimes within the first hour).  They can eat small crickets (3/16 to 1/4 inch.)

PERSONALITY:  Females tend to be calmer than the males, and will occasionally eat a cricket from my fingers.  Males are more anxious.  At night, U. pietschmanni are more active and perky than most other Uroplatus and occasionally vocalize.  They are not a shy gecko.  I don't recommend handling them, but they become accustomed to their keeper and don't disappear into the foliage just because you are in the room.