Science topic

Amphibians - Science topic

Vertebrates belonging to the class Amphibia such as frogs, toads, newts and salamanders that live in a semi-aquatic environment.
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I would like to open a discussion regarding Benzocaine 20% oral administration and its potential as an effective and humane method for the euthanasia of small reptiles. Finding both practical and humane methods of euthanasia that can be used in the field is challenging but essential.
Benzocaine, similar to MS 222, is an accepted euthanasia agent for fish and amphibians under the AVMA GUIDELINES FOR THE EUTHANASIA OF ANIMALS: 2020 EDITION, it is effective and rapid. However, a paucity of literature is available on its efficacy in small reptiles. There appears to be several advantages and disadvantages to the Benzocaine oral administration method. But let's hear from you. Has this method ever been accepted by your ethics committee for small reptiles? And if so, what were some of the comments from the committee? For those that have applied this method, have you ever witnessed vomiting or regurgitation after administration? What were the dosages given, how was death confirmed and after how long? What are your institutions' preferred methods of euthanasia for small reptiles?
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It is not a nice thing to hurt those who are smaller and weaker than us. I like small reptiles and would like to protect them. Therefore, my principal question to you: why euthanasia is needed at all? Couldn't another research direction be devised? Please...
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Ideally recorders that can be left out for weeks at a time, with frogs from tropical forests the target animals.
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AudioMoths - https://www.openacousticdevices.info/audiomoth - great sound quality, relatively inexpensive. Depending on the recording duty schedule and the size of the memory card, they can be left out for a few weeks. Unfortunately, they are very difficult to find right now because of chip shortages.
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Pros and cons of passive acoustic recorders in the Neotropics.
Best brands/models?
Ideally for recorders to be left up for weeks at a time.
Frogs are the target animals.
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The best program I used to record the sounds of different types of amphibians is Avisoft-SASLab Pro
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Hey everyone, I was wondering if much research has been done on the affect of lunar cycle on the activity of amphibians. I am aware of Deeming's 2008 paper where capture of newts correlated with the lunar cycle. I am interested to know if any other such correlations have been found in species around the world.
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I have published a lot in this area, since 2009. Each amphibian species has a different response to the lunar cycle, depending on ecology. I have also done a review of this topic. Be careful when looking at studies, that they have used a variety of sites over at least 4 years to make sure the result is meaningful. In one or two years only, any correlation with lunar cycles can be co-incidental. Most of my research has been done at a large number of sites an / or over many years. Bufo bufo in particular breeds at the full moon.
Grant, R. A., Chadwick, E. A., & Halliday, T. (2009). The lunar cycle: a cue for amphibian reproductive phenology?. Animal Behaviour, 78(2), 349-357.
Grant, R., Halliday, T., & Chadwick, E. (2013). Amphibians’ response to the lunar synodic cycle—a review of current knowledge, recommendations, and implications for conservation. Behavioral Ecology, 24(1), 53-62.
Jarvis, L. E., Grant, R. A., & SenGupta, A. (2021). Lunar phase as a cue for migrations to two species of explosive breeding amphibians—implications for conservation. European Journal of Wildlife Research, 67(1), 1-11.
Arnfield, H., Grant, R., Monk, C., & Uller, T. (2012). Factors influencing the timing of spring migration in common toads (B ufo bufo). Journal of Zoology, 288(2), 112-118.
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I am looking for literature that deals with amphibians in urban areas. Are there studies on threshod values for toxological substances at which amphibians are no longer able to survive? Which species are more sensitive which are less? For example, at what concentration are acidification and contamination by heavy metals such as cadmium, copper, lead and zinc a problem? What about pharmaceuticals and pesticides? There are a lot of studies for non-urban areas, are the results transferable?
Studies in German are very welcome.
There is a review, but I think it‘s not sufficient for answering this specific question:
Toxicological Threats to Amphibians and Reptiles in Urban Environments (2008)
Maxine C. Croteau, Natacha Hogan, Jennifer C. Gibson, David Lean, and Vance L. Trudeau, In: J.C. Mitchell, R.E. Jung Brown, and B. Bartholomew (editors). Urban Herpetology. Herpetological Conservation 3: 197–209
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I have done herpetological surveys in several urban areas, most notably in Milwaukee, Wisconsin (USA). Several species are highly tolerant to heavy metal contamination such as Family Ranidae; Lithobates clamitans, L. catesbeiana, L. pipiens, Family Bufonidae; Anaxyrus americanus. Treefrogs (Hylidae) were less tolerant and more common near the periphery of the city (Psuedacris crucifer, Hyla versicolor). See attached report.
(PDF) Milwaukee Estuary Area of Concern Wildlife Population Assessment Report (researchgate.net)
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Hi there! I am interested in understanding (or making myself more confused, whatever) the evolutionary pathways of coprophilic habit in dung-inhabiting fungi. Taking into account the dung of herbivore animals as a substrate to dung fungi growth, we have few "candidates" to be dung-producers with some requirements to early dung fungi (e.g. the amphibians Ichthyostega (I don't know if it was an herbivore or omnivorous, appearing about ca. 375 million y/a in Devonian, once the first tetrapod herbivores made their first appearance in the fossil record near the Permio-Carboniferous boundary, ca. 300 million y/a.). Terrestrial plants made their first appearance ca. 450 million y/a, with a well-accepted role of arbuscular mycorrhizal fungi in this process. So, my question is about if there is some study dealing with the evolutive process of dung-inhabiting fungi, presenting some consideration where and how, in the evolutive process, this ecologic habit firstly appears? Any considerations are welcome! Thank you!
If you want to help me with this question with more details (or more questions ¯\_(ツ)_/¯ ) or papers/books, feel free to also send me an email at: calacafjs@gmail.com.
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Also kindly check the following very good link:
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I've tried the Qiagen RNeasy Mini kit on chicken blood samples but got no RNA.
Started with 1ml fresh whole blood collected on ETDA, after adding the 600ul lysis buffer the lysate was SOOO VISCOSE, and so hard to pipette even after trying to homogenate it with "Syringe & needle 20 guage co.9 mm"
Knowing that the whole blood of birds, reptiles, and amphibians contains large quantities of protein that must be separated from the nucleic acids to perform a successful extraction, But I don't know how ?!
- should I start with less quantity of blood or what should I do
- the pic shows the pellet size before I discard the supernatant & adding the lysis buffer
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The avian blood is very concentrated, so it would be better to study with low volume of blood. Remind that avian erythrocytes contain nucleus. So, I also suggest you to dilute the blood samples with RNAse free water before isolation. You can find enclosed protocol. Good luck!
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I prefer the recording of one, or a few individuals, not a large chorus.
It is for use in a playback experiment.
Thanks!
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Hi Gabriel
For call recording data, I would recommend looking at a place like amphibiaweb.com
E.g.
or possibly better GBIF
E.g.
Places like that will 1) give some info about the biogeographical origin of your call (potentially useful) and allow you to document the source with minimal effort.
Enjoy!
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Kindly, name some reliable/popular field survey technique employed for amphibians (frogs) & terrestrial skinks (scincidae) in tropical forests. Thanks a ton.
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Hello,
besides the previous answers, and integrating some of them, I list some survey techniques used for amphibians and most terrestrial reptiles, inclusive of skinks, in tropical forests, but also in more general contexts.
Active methods
· Visual detection along fixed transects (not the best in tropical environments, but applicable in some context)
· Visual detection without fixed transect, freely scouting a specific area
· Acoustic encounter along fixed transects (amphibians)
· Acoustic encounter scouting a specific area (amphibians)
· Sporadical-opportunistic observations and acoustic records
Other active methods (captures):
Amphibians
· Hand-capture
· Dip-nets
Skinks
· Hand-capture
· Grabber
· Noose
The use of binoculars can be applied in some environmental contexts and cameras are often essential, as photographs of detected or captured animals are an evidence for verifying species identifications.
Obviously, a general knowledge of the potential presence of some species in the investigated area must lead to examine the zone keeping into consideration the general ecology of each taxon:
- specific forest type (e.g., zones where small areas of primary-secondary dry forest, transitional dry to moist, moist forest, human altered forest are close to each other)
- specific habitat (e.g., trees, poles, tree holes, small rivers, and waterfalls, breeding sites)
- best season
Naturally, each point should be examined regardless of the knowledge of the potential presence of some species in the investigated area (hiding places for some amphibians and skinks, poles for tadpoles, etc.).
Other similar, obvious considerations are as follows.
In all cases, the above mentioned techniques are employed in different ways based on:
- forest type
- season and/or the weather conditions
- hour of the day
(e.g., clearly, for amphibians these techniques aren’t employed in tropical dry forests, during the dry season and in full daylight).
To maximize the success of a survey, some artificial environmentscan be used, such as:
· Artificial covers (amphibians)
· Shelters (amphibians and skinks)
· Basking substrates (skinks)
If the transect techniques is used, each transect can be settled basing on the presence of one of these artificial environments.
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Passive methods (amphibians)
· Pitfall traps (eventually with drift nets)
· Funnel traps
· Bottle traps
· Artificial cover traps
· Microphones (vocalizations ;-)
· Camera-traps (very rarely)
Passive methods (skinks)
· Pit-fall traps
· Pipe-trap
· Camera-traps (very rarely)
Here again cameras are essential, as photos of captured animals are an evidence for verifying species identifications.
A general, again obvious, remark is to record the location, date, time, and micro-habitat of each record.
A conclusive short remark is as follows.
There aren’t fixed rules to plan a survey, even though sometimes it’s recommended to involve, if possible, 3 to 6 people for 3-5 days in each survey. The number of surveys and their temporal distance depending on the specificity of the study.
Last but not least, if aiming at creating an erpethological checklist of an area:
Op­portunistic records by local people
Finally, op­portunistic records of various species encountered by local people are useful to create a more exhaustive checklist of the species of an area.
General references
Bennett, D. (1999). Expedition Field Techniques - Reptiles and Amphibians. Geography Outdoors.
Heyer, W. R., M. A. Donnelly, R. W. McDiarmid, L.-A. C. Hayek, and M. S. Foster (1994). Measuring and monitoring biological diversity. Standard methods for amphibians. Smithsonian Institution Press, Washington DC.
Rödel, M.O, Ernst, R. (2004). Measuring and monitoring amphibian diver­sity in tropical forests. I. An evaluation of methods with recommen­dations for standardization. Ecotropica 10: 1–14.
Wilkinson, J. W. (2015). Amphibian Survey and Monitoring Handbook. Pelagic Publishing, Exeter.
Simply three case studies in tropical forest environments
Costa-Campos CE, Freire EMX (2019). Richness and composition of anuran assemblages from an Amazonian savanna. ZooKeys 843: 149–169. https://doi.org/10.3897/zookeys.843.33365
Mira-Mendes CB, Ruas DS, Oliveira RM, Castro IM, Dias IR, Baumgarten JE, Juncá FA, Solé M (2018). Amphibians of the Reserva Ecológica Michelin: a high diversity site in the lowland Atlantic Forest of southern Bahia, Brazil. ZooKeys 753: 1–21. https://doi.org/10.3897/zookeys.753.21438
Rödel MO, Glos J (2019). Herpetological surveys in two proposed protected areas in Liberia, West Africa. Zoosyst. Evol. 95: 15-35. https://doi.org/10.3897/zse.95.31726
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Anomalous phenotype; polydactyly, adactyly, hyperregeneration, trematodes & mutations are observe in amphibians. Similarly, beside the the aforementioned reasons what are other associated phenomenon responsible & may suspected to influence such developments in terrestrial skinks (Scincidae) as well. Please suggest, thanks a ton.
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Although I am not a specialist in the field of amphibians, I know for sure that my colleagues carried out this research in Russia. One of the reasons for this is environmental pollution. This was primarily observed on the territory of large cities. Subsequently, we found that the reason for everything is the high content of heavy metals.
Regards, Sergey
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It intrigues me that, the poisonous amphibian (frogs) evolutionary process might play pivotal role and especially genetically have to do with their morphological appearances in poisonous frogs alluring appearances than the regular frogs whether tropical forest or temperate. Any other specific reasons or detail classifications for such existance of differences in amphibians. Elaborate, please (Thank you).
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I agree with Ghassen Kmira. Another benefit is that they are advertising for reproduction as they can afford to be colorful. You could turn it around and ask why aren't all frogs extremely colorful? Most frogs depend on camouflage to avoid predators. Poisonous frogs don't need to hide from most predators. Now I'm wondering if poisonous frogs are as loud as other frogs, since their coloring makes it easier for potential mates to find them. Also most poisonous frogs live in rain forests, where the flora around them is also very colorful.
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I'm interning at a non-science conventional organization, where I'm conducting studies on a wetland that's inhabited by an endangered species of snake and a threatened species of frog. I'm trying to better understand how the flow of nutrients from the surrounding groundwater/run off sources effect the soil and surface water, which ultimately affects the development of amphibian development and habitat.
So, because it's an unconventional place to be doing these studies, their facilities aren't comprehensive. I am able to do water nitrate and phosphate tests using TNT kits, but I was wondering if there was any low cost/practical soil phosphate and nitrate testing methods? Whether it would give me a general/semi-accurate result. I had the idea of mixing DI water and the soil, then letting the particles settle and using the surface water to do a regular water analysis. Does anyone have any experience doing that?
Appreciate any help
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Hi. I hope the following link could help you very much:
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Background: In my proposed study area, I have identified four pond types: 1) Ephemeral (dries at least once per year), 2) Matrix (part of pond dries dries, but other part remains permanent), and 3) Permanent (has not dried). I have four of each pond type, totaling 12 ponds total. I will be sampling for the presence of fish, amphibians, and reptiles eight times each year across three years. I would like to assess how pond type influences the occupancy of certain species and how the presence of fish influence the occupancy of certain species.
Question: Is four ponds per site type enough of a sample size?
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In many conservatories around the world, as well as in many organic-farms, insecticidal soaps (potassium salts of fatty acids) are widely used to combat aphids, mealybugs, mites etc. They are considered safe to mammalians and are prioritized instead of chemicals.
Very little to no information can actually be found whether the soaps may be toxic to amphibians. Can anyone help us on this matter? An eductaed guess would tell me that the thin film created by the sopa on aquatic enviroment as well as, presumably, on the skin of the amphibians would cause significan damage.
Many thanks!
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I looked up the technical fact sheet and although there was no specific mention about amphibians apparently it is toxic for fish and aquatic invertebrates and the EPA requires it not to be applied to water or to contaminate water sources with it. I think this supports your hunch.
"Scientists concluded that potassium salts of fatty acids are slightly toxic to cold-water and warm-water fish (1)."
"Potassium salts of fatty acids are highly toxic to aquatic invertebrates."
"The EPA requires all product labels containing this active ingredient to state that the product is not to be applied directly to water and the user is not to contaminate water by cleaning equipment or disposing of wash water that contains potassium salts of fatty acids (1, 11)."
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Our team has been trying to locate and collect the pineal gland in the American bullfrog's brain (Lithobates catesbeianus) for some time now. However, we don't have a standardized protocol for that. Therefore we're not so sure that the tiny tissue we're collecting is actually the pineal gland. Does anyone have any protocols, pictures, or videos for pineal gland collection in amphibians? I really appreciate any help you can provide!
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It used to collect the glands of Telmatobius and Hyla frogs (now Hypsiboas). After euthanizing the specimen, I proceeded to extract the lower jaw, exposing the palate. Then, with scissors, I peeled off the epithelium from the mouth, exposing the base of the skull. With scissors, again, I made vertical cuts on the lateral extension of the prootics and anteriorly on the anterior tip of the parasphenoid. After that, I inserted one of the scissors blades through the foramen magnum, horizontally, to make a horizontal cut of the lateral walls of the brain box, on both sides. After that, with forceps, I would lift the ventral covering of the brain cage (parasphenoids and parts of the prootics and exoccipitals). In this way, the ventral base of the brain was exposed. Generally, the hyophysis did not adhere to the bones , and it remained attached to the brain. The pituitary gland is clearly noticeable, differentiating itself from the rest of the nervous tissue, when the material is fresh, due to its great vascularity; it is of oval shape. If it was extracted with fine-tipped forceps, the forceps were passed underneath of the "grain" of the pituitary, which came out easily, tearing the stem of the pars intermedia, but both anterior and posterior lobes remained intact, as he could see later by histological sections. I hope that those memories may help you in any way. If you have questions, please contact me.
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What are the parasitology guides that you are recommending to identify the helminth infections of wild animals? Especially helminth infections of amphibians.
Please mention links of guides/books which are available to refer online or titles of the books that you know.
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I invite you to see this paper.
Good luck,
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I am doing species niche modeling of an amphibian species. I have 151 presence observations, after filttering points that were too close (< 1 km). I am using data presence from GBIF, iNaturalist, and presence data shared by colleagues. However, I am worried about sampling bias, because it is an opportunistic kind of approach. Therefore, I am trying to find a way to correct sampling bias to better our models.
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When the sample size is large, naturally the error will get minimized. Further, if you collect data on relevant foraging variables, they should be subjected to tools such as PCA/ DFA, which will result in niche segregation.
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I want to measure the metabolic rate and water loss in amphibians, usually frogs and salamanders, that have different life histories. And, I want to build a chamber that I can use for aquatic and terrestrial species or life stages. I want them to work for both because the idea is to take measurements in the field and the system itself is already large.
I am starting to work with physiology, so any advice for the chamber or how to measure metabolic rate is welcome. Thanks!
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Hello Zuania,
In your case, I think it would be interesting to build a chamber that can measure respiration both in water and in air. You can check papers on mudskipper or crabs which studied bimodal respiration, you may find advices for chamber design.
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Does anyone have access of this paper/book chapter? Found this citation in the book Amphibians and Reptiles of Nepal. Based on the citation format, it looks to be a book chapter, and the language seems to be Uzbekistan (please correct me if I am wrong)?
Any help will be greatly appreciated!
Panfilov AM, Eremchenko VK. 1999. Novye dannye po karyiologii shesti vidov szinkov (Sauria: Scincidae) Evrasii. Nauka i Novye Tekhnologii, 1999(2), 61–67. Bishkek
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Dear Kai, Bishkek is in Kyrgyzstan (capital). I visited once when checked a proposal to work for the Issyk-Kul Lake hydrobiological lab. As well the Journal changed its name twice. This article doesn't exist in digital form, so I recommend you to find a colleague from the country and ask him for a favor!
Andrey
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Hello,
Amphibians eggs are surrounded of a jelly coat constituted by one to several layers depending on the considered species. The composition of these layers depends on the considered layer and species. The functions of this layers are multiple. Indeed, the jelly coat is for instance known to be involved in fertilization, to avoid polyspermy, to act as a sperm chemoattractant and to play an important role of barrier against UV, infection and contaminants. But, the role of barrier against contaminants is not so clear. Indeed, the complex structure of the jelly coat provides to it an affinity to specific molecules. Therefore, certains molecules are stopped by the layers while others penetrate to the embryos.
In spite of this, most protocole for testing effect of toxicants on amphibians suggest to dejelly the eggs before the exposure. From my point of view, this recommendation lead to a loss of ecological relevance and I'm wondering if dejellying should not be reconsidered.
How about you ? Is there something that I don't see ?
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Hi Laurent ,I agree that removing the jelly reduces ecological relevance in the toxicity testing. However, one reason could be to create a worst-case (tier 1) laboratory experiment with enhanced exposure. For increasing realism, the next step could be to test eggs with jelly (tier 2). This of course would assume that jelly protects the embryo by lowering exposure.
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Only supplier i've been able to find (bioMérieux) apparently only stock the tubed version (100) meaning an eight week wait to get much cheaper softpack swabs. Does anyone know of alternate suppliers in Australia or can suggest comparable product (MWE 100 or 113 appear most popular in the literature). 
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I´m looking for papers showing raw abundance data for different groups of animals, plants, and bacteria. I would be interested in studies on: mammals, birds, reptiles, amphibians, insects, fishes, annual plants and trees.
Any recommended paper will be of great use.
Thank you so much !
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The ATLANTIC and NEOTROPICAL series data papers published in Ecology has data about many groups....see https://esajournals.onlinelibrary.wiley.com/doi/toc/10.1002/(ISSN)1939-9170.AtlanticPapers
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Looking at Plethodontid salamanders, they have a gular fold, a characteristic seen in reptiles often as the dewlap for threat displays, but I haven't found any explanation for it being in amphibians. What are some of the behavioral uses of gular folds in amphibians?
Consequently, we discussed the ability of ancestral tetrapods --originating in Sarcopterygian fishes-- having articulation in their wrists, making them the first animals to be able to do push ups which I then want to connect again to threat displays in reptiles. Are there any soft tissue samples of ancestral tetrapod gular folds? Would this be an ancestral characteristic?
Is there some sense to this, or am I over-extrapolating?
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If you look at salamander feeding videos from Duban lab you might see some movement of the gular fold. I think it would be difficult to assign function to this feature in terms of the study of adaptation. Because there is so much convergence and paedomorphosis in Caudata it might also be difficult to study the origin of the feature.
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I am trying to model nutrient uptake and turnover from a long-term experiment. Currently I found a reference to a turnover rate for N used in Burton and Liken's classic study; however, when I asked the authors for the source of their estimate they (1) could not recall, and (2) did not provide a reference in the original paper.
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Dear all,
I am sorry to answer to your question with another question, but I think that you all could actually give me a very good and practical advice here.
Using 13C and 15N, I want to assess changes in diet composition in tadpoles and newt larvae after being exposed to a source of pollutant (with theoretically no direct effect on the consumers). The hypothesis is that the pollutant would affect the density of the sources (not the signature).
My questions are:
-Can we in this case, collect prey and consumers at the same time without really considering tissue turnover of the tadpoles and newt larvae?
-How much would you wait from the application of the pollutant in the water before collecting the animals (considering that in my situation, after max 4 weeks from the application, they will probably develop in adults).
-On the other hand, if you would collect sources and consumers with a delay, how much time would you expected to be necessary between the source and the consumer sampling?
Thank you all, and sorry again for invading your question.
Best regards
Alessandro Manfrin
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Eryops marks the Permian age.
Scleractinia marks the Tertiary period.
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Regardless of where and when they occur in the fossil record, the best biostratigraphic marker fossils are common, easy to identify, and have a short duration. Marine invertebrates, such as a coral, are typically widespread and easy to identify, but persist for long periods of time. On the other hand, vertebrates like Eryops are typically areally restricted and rare, but have shorter durations in the fossil record. If you are fortunate enough to have both in hand, I would say that Eryops is more useful because of its shorter duration in the fossil record. Its taxon range zone would provide more biostratigraphic precision.
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We have been attempting to radio-track Litoria aurea, however, our site is intersected by a large electricity easement, therefore, we are struggling to pinpoint any signal over the static. Does anyone have any tips on how the interfering noise can be minimised or eliminated? Thank you.
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I quite agree with Peter M Narins , of using a directional antennae like a yagi. With an omni directional or whip antenna, you must move to determine the direction of the higher signal level. But, If you use a Directional, handheld or vehicle-mounted Yagi antenna, you can follow the direction of the strongest signal to the noise source. This will greatly reduce the amount of time and travel distance required during the activity.
Radio Direction Finding techniques typically offer the best and most efficient approach to locating most power-line noise sources. A handheld Yagi works at VHF and UHF within a specified frequency range. Not only are VHF and UHF antennas typically smaller, but direction headings are more reliable. An attenuator is required between the antenna and the receiver if the receiver does not have one. The operator must be able to pinpoint the source on the structure down to a component level. You also can use a hot-stick-mounted device to locate the source. An ultrasonic dish is quite helpful in pinpointing the source of an arc. An unobstructed direct line-of-sight path is required between the arc and the dish. It is only useful for pinpointing a source once it has been highly localized and is ideally suited for pinpointing the arcing hardware once the offending pole has been isolated.
hope this will help you in your research.
Best Wishes,
Adarshjit Das
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I have found a protocol outlining how to count the number of leucocytes (attached file) but I am also hoping to look at the relative number of lymphocytes, monocytes, neutrophils, basophils and eosinophils in amphibian blood. Has anyone had experience with doing this successfully on a NC-3000?
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For my research class this spring, I would like to study amphibians (specifically salamanders) if possible. My limiting factors are time (March 1-mid April) and that the research must be field-based (no capturing for lab studies). Would I be able to find breeding or migrating salamanders during this timeframe with reasonable effort?
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Kenyon Tweedell had some great advice regarding timing. Just a bit of extra advice... If you want to study salamanders in a short period of time, cover boards can help. To set these up, you place thin pieces of wooden boards directly on the soil (remove any duff and replace over the board to camouflage.) Salamanders will be attracted to the cover/moisture provided by the boards. Another technique would be to do eye shine surveys or timed visual encounter surveys to determine the number of amphibians you encounter in a specified time. But make sure you have any necessary permits to work with vertebrates if you are going to alter their behavior at all (check with your instructor.) Good luck and happy herping!
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Hello everyone. I represent a small group of students from Maastricht University in the Netherlands. We are currently working on a project regarding toe-tapping behavior in frogs and toads (see https://www.youtube.com/watch?v=gl_A4UosQjw). We are collecting as much information as possible regarding it in order to try to shed some light on this very understudied phenomenon. If you've ever observed it and could spare a few minutes of your time to help us in our research, please fill out our questionnaire or share your knowledge with us. Any input is greatly appreciated.
Thank you for your time and happy herping.✌️
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Hi Nikola,
This is a very interesting topic. I first noticed this phenomenon when on a field trip to Thailand in June, 1995 with my then graduate student Thom Ludwig and my then post-doc, Jakob Christensen-Dalsgaard. We saw several females of Polypedates leucomystax toe tapping on the dense vegetation on which they were perched. We reported this in several publications, including these:
Narins PM (1995) Comparative aspects of interactive communication. In: Active Hearing (Å Flock, D Ottoson, M Ulfendahl eds.) Elsevier Science Ltd, Oxford, UK 363-372.
Narins PM (1995) Frog Communication. Scientific American 273: 78-83.
Narins PM (2001) Vibration communication in vertebrates. In: Ecology of Sensing (F Barth, A Schmidt eds.) Springer-Verlag, Berlin 127-148.
Christensen-Dalsgaard J, Ludwig TA and Narins PM (2002) Call diversity in an Old World treefrog: Level dependence and latency of acoustic responses. Bioacoustics 13: 21-35.
Narins PM (2019) Seismic communication in the Amphibia with special emphases on the anura. In: Biotremology- Studying Vibrational Behavior II (PSM Hill, R Lakes-Harlan, V Mazzoni, PM Narins, M Virant-Doberlet, A Wessel eds.) Springer-Verlag, Heidelberg pp 277-292.
Hope this helps!
Peter
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I am interested to hear about the concern of adult amphibian marking by elastomers (visible implant elastomer). I have seen an article that concludes that this technique is reliable, and others that conclude otherwise (e.g. "VIE tags, as individuals were correctly identified only 18.4% of the time" Brannelly et al. 2014).
Has anyone used them in thick skinned species, such as those of the genus Rhinella?
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I've tried tagging dark-skinned salamanders with VIE, which cause the same problem as thick skinned anuran species.
One of the solutions I found was to inject the elastomer on the ventral side, at the basis of the limbs. The skin is thinner, which makes it easier to see the tag?
Another solution I used was to inject the VIE in the palm of the hands and feet. As with the ventral side, the skin is usually thinner there, which makes tags more visible.
With Rhinella, fingers are probably long enough to even allow coding schemes using finger locations, as described in this paper :
( Technology meets tradition: A combined VIE-C technique for individually marking anurans, Hoffman et al, 2008)
Here, they use a combination of clipped toes and VIE tagging, a double-tag system enabling to identify animals even if one of the tags has failed (it is unlikely that both fail at the same time). Hope you'll find this paper useful !
Good luck with your research,
Benjamin
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I have a relatively small budget but I would like to purchase a hydrophone that would enable me to call and describe the vocalisations of frogs underwater. The animals I hope to record are in glass 20 L glass enclosures
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Just a word of caution when recording with a hydrophone in a closed (20L) container. Any vocalizations by the frog will reach the hydrophone via the direct path and via various paths after reflections from the tank. The hydrophone will measure the sum of the sound arriving via all the paths. Ideally, you would want to measure the sound pressure level via the direct path only. There are at least two ways to make the correct measurement. 1.Measure the frog vocalizations in a natural pond (without glass surfaces) to minimize reflections. 2. Measure during a discrete time window that includes the earliest arriving wave (the direct wave) and then stop the measurement before any of the reflected waves arrive. The larger the tank, the easier this is to do. I hope this helps.
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Hello, I am a caver and a biologist (just by passion, not professional). In summer 2018 I went caving to Magagascar in Namoroka Tsingy. I didn't ask for a collecting permit so I only took photos from the fauna.
I have some from amphibians inside the caves.
See joined file. Can you help me to determine those specimens ? Even only the family would be a great help.
Futhermore, I will go back to that area next july.
Is it usefull I take more pictures ? Is it usefull I collect some amphibians ? If yes, how can I have a collecting permit ?
Great thanks,
Josiane Lips
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I would start with the iNaturalist community resources. https://www.inaturalist.org/
They have excellent algorithms for species of all kinds but are particularly good with animals.
It is possible to build projects and store image files for identification from the community of ecologists.
They currently have over 2,000 observations of 200+ species of frogs:
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Species: Lissotriton vulgaris (please confirm; photos attached)
Subspecies: ?
Sex: ?
Total Length: 10 cm
I found this smooth newt yesterday (18.12.2019, 21:30 local time) in the outskirts of Vienna (Austria) on a stone walkway in a garden.
Environmental data (at time of observation):
Air temp: 2°C
Humidity: not measured, very likely 100%RH: foggy and very wet since sunset (no rain)
No wind
Weather during the day: 9°C and sunny around noon/in the early afternoon, no wind, no rain
In the same garden, newts have been found in previous years in springtime (in a water meter shaft 1.5 to 2 m below ground level). There is a pond in one of the neighboring gardens; the urban housing area is located within 1 km of Donau-Auen National Park.
According to Jablonski (2013) and Kaczmarek et al. (2018), winter activity in the species is unusual in this part of Europe. For this reason, I include here as much information as possible. The environmental conditions were similar to those reported in both papers.
The animal was not moving when I found and subsequently observed it for 10 min. Upon touching it (to remove from walkway), it moved its limbs lethargically. Due to a further drop in temperature (to ca. 0.5°C), I took it in overnight.
What is the best procedure to maximize its chances of survival? Where should I release it today (open/vegetated area in the garden)? Which time of day (weather forecast: 5°C in the morning, 13°C at noon, 10°C in the evening)?
Literature:
  • Jablonski D (2013) Unusual observation of the winter activity of Lissotriton vulgaris from south-western Slovakia. Folia faunistica Slovaca 18: 301-302.
  • Kaczmarek JM, Piasecka M, Kaczmarski M (2018) Winter activity of the smooth newt Lissotriton vulgaris in Central Europe. The Herpetological Bulletin 144: 31-32.
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And I would release it in the evening (if the temperature is around 10°C) and place it under or next to a dead wood or some other structure to hide (but I think this advice comes too late)
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We wish to quantify the amount of pesticides presence of pesticides in different landscapes to assess if they have an impact on amphibian communities.
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There are a number of color-strip tests for the more common chemicals, e.g. organophosphates, atrazine... but other than the specific pesticides you would like to quantify, of course it is important to consider accuracy/precision required for reporting, of which field kits will likely not be able to match the certified LC or GC methods. Nevertheless, there are kits:
Abraxis has a wide range of ELISA assays: https://www.abraxiskits.com/products/pesticides/
Neogen has a color disk kit for quite a few classes: https://foodsafety.neogen.com/en/agri-screen-tickets
Another company with ELISA assays:
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I am working on a biogeographic study, in which to assess the latitudinal diversity gradient of amphibians in the Malay Peninsula. Thus, I’m required to examine the effects of utilizing different distribution data such as IUCN range maps, GBIF, modeled distribution, and a combination of those data, in mapping species richness in the Malay Peninsula.
My question is, what kind of statistical analysis that is suitable to compare the performances of these different distribution data in mapping species richness?
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So, it is geographical scale of species richness you have to do. But take care patterns of moutain-scale patterns & what about fresh-water as you are studyinf amphibians? I'm attaching a book on species richness. As well I agree with Andrew that in this case ou can use some indices of diversity (may be better to say "commonality). All known indices of commonality are divided into two groups depending on whether, they take into account or ignore the number of negative matches (d). The greatest value in environmental jobs are indexed in the formula which includes the number of positive coincidences.
It is offered a huge number of indices of commonality, but more often in biocenological, faunistic and biogeographic studies are used indices of Jaccard and Sørensen – Czekanowski. These coefficients are equal to 1 in case of complete coincidence of the types of communities and equal to 0 if the samples are completely different and do not include the common types. In comparing should understand that standard mistake of methods is normally higher than statistical mistake.
Indices of community that take into account negative matches, usually when comparing collections when is known full species lists. The use of this group of indices in biogeographic studies were subjected to serious criticism. Limited use have indexes that take into account negative matches, due to their large dependent from a rare species that can not get into the sample.
I know minimum 10 indices based on comparing the qualitative data. For quantitative is very useful Sørensen for the data with abundance!
Andrey
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Briefly, I need a cost and time effective method - which completely exludes the possibility of contamination between samples - in order to get ranaviral DNA from the liver of EtOH stored tadpoles.
Criterions are the following:
- For DNA extraction I will use Wizard® Genomic DNA Purification Kit Protocol (Promega).
- I don't have bead beater or tissue lyser machine just an Eppendorf thermomixer.
- Large sample size (approx. 500 samples).
Many thanks in advance.
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With what you have available, you may try to space yourself if you do come back to doing it manually. Patience is key. Also manual homogenizing will best prevent any instance of contamination. I believe you have disposable pestles which saves you the problem of having to disinfect each time you use one. I hope this helps if it becomes necessary.
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Does anyone know where I can find a Government report on Biodiversity from the Cayman Islands? Something with reported numbers on species of Mammal, fish, reptiles, amphibians, and birds.
Thanks!
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Hello,
I plan to measure AChE esterase on amphibians and I read that Butyrylcholinesterase can interfere in the measurement.
If I measure the transformation of Acetylcholine iodide, what am I measuring? Acetylcholinesterase activity or both acetyl- and butyrylcholinesterase activities ? Should I measure AChE activity with a BChE inhibitor for stating that I'm measuring AChE?
For me, I should but I've read a lot of articles that don't mention any use of the inhibitors, so I'm a bit confused.
Cheers,
Laurent
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To add to Peter Jackson's well explained answer, if you measure AChE usind acetylthiocholine as a substrate, you do not have to take measurement of butyryl cholinesterase activities into consideration.
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I am assessing Amphibian diversity in selected wetlands in Kampala, Uganda. I am using the Urban-rural design. I purposively selected three wetlands that run from the City to the outskirts. For each wetland stretch I randomly chose three sites to represent urban and rural categories. For each site I recorded amphibian species, habitat and micro-habitats in which they were. Therefore, am trying to analyze whether amphibians prefer certain habitats or micro-habitats to others. Thank you. I will be grateful for your guidance.
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I've always though the best way to answer this question is to capture the amount of microhabitat available to the frogs in addition to their use. Then you can see if they use some habitat more or less compared to what is available to them. That is how I answered a similar question. The methods and stats are described in Beard et al. (2003) Journal of Herpetology.
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I am studying Ethiopian amphibians now and I need the software to analyse the phylogenetic relation ship of amphibians.
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perhaps Phylip
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I observed remarkable sperm morphology diversity and chromosomal dysploidy in the spermatogonial cells of an Ichthyophis and Gegeneophis caecilian (legless amphibian) collected from dung pits from Western Ghats. Inputs from you people is of great help in drafting the discussion part of my manuscript on the impact of dung on caecilian amphibians.
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I don't think so. Those changes you saw are not neoplastic
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Hello, I am trying to generate primary cell cultures from different tissues of adult zebrafish (including gills, skin and muscle). I have tried several methods for cleaning the whole fish (hypochlorite, ethanol) and the tissues pieces after dissection and before digestion. Nevertheless, after plating I can see not only cells but also bacteria. I have tried different antibiotic combinations including penicillin/streptomycin, gentamicin and primocin but I still get a high amount of bacterial contamination. Any tips on how to improve this? is this normal?
P.S- No many publications mention the contamination of primary cultures with the own fish's bacteria. I have only found this one from 1976 - THE USE OF ANTIBIOTICS IN THE PREPARATION OF AMPHIBIAN CELL CUL TURES FROM HIGHLY CONTAMINATED MATERIAL,
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The majority of bacteria in fish are in the external mucous coat or the gut. So one approach is to minimize these microbial populations by changing fish husbandry prior to collection:
  • 1-2 days before collection, stop feeding fish.
  • Place fish in dedicated tank of sterile aquarium water supplemented with broad-spectrum antibiotics.
  • Fish density should be very low (much lower than usual) and the water should be well-agitated with bubblers.
  • Once anaesthetized, rinse fish again in sterile system water, and briefly spray/wipe with a surface decontaminant such as 70% ethanol, topical hydrogen peroxide, or diluted iodine.
  • Dissect the fish in a pre-cleaned surgical area such as a UV-sterilized containment hood, an autoclaved dissecting pan, or a clean bench covered with a sterile drape.
  • Autoclaved or flame-sterilized instruments are advisable.
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I would like to ask if you have any information about aberrant caecilians/impact of agrochemicals on caecilian amphibians. Any suggestion or links would be highly appreciated.
With kind regards
Venu Govindappa
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May this one (on general morphology of caecilians) be of an interest for you?
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As you may know there are several compilations of classic papers in Ecology (e. g. Foudations of Ecology). I am trying to find such a volume or classic papers about morphological abnormalities in Neotropical frogs.
Please share your opinion and/or any sources. Thanks
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Your welcome!
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Hi,
I'm currently doing a PhD in Switzerland. Swiss legislation on the euthanasia of laboratory animals just changed. Now, euthanasia of amphibian's larvae have to be done using liquid nitrogen. My problem is that I have then to measure molecular biomarkers and I don't know if it's possible after such a euthanasia method. Does anyone already tried it, maybe on fish?
Best wishes,
Laurent
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Hi Christoph,
Thanks for your reply. I'm thinking the same but that's would be great to hear something from someone who already tried. That way, if it's not or hardly possible, we could argue for a dispensation from the veterinary services.
Best,
Laurent
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One could conclude from this article that results from most amphibian monitoring programs aren't useful due to lack of statistical power and extreme temporal variability in results. Do you agree? Would resources spent on monitoring better serve conservation in other ways?
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Having worked on some monitoring programs from the management/government side and also from the academic side, I can tell you that your question is constant concern. All things being equal, monitoring is worth it. If you can't detect subtle trends, hopefully you can detect a crash. Monitoring can be designed to help inform management as well. With no monitoring data, it's more difficult to justify spending money on other conservation actions. Finally, management often proceeds on the best available information, not statistical significance. So if a negative trend starts to appear, even if its not significant yet, you can justify more funds to look into it, or start management that may reverse the trend. At the end of the day, some information, no matter how weak, is often better than no information.
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Hi colleagues,
I am trying to test how pH may effect the behaviour of the skin of an animal. To do this, the first step is to understand what is the normal condition. How could I do it? In particular, it is an amphibian.
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Dear Giorgio, am only familiar with how to measure the PH of chicken checking the meat quality.And it can be measured using PH meter. PH is related to the amount of glycogen. When PH is greater than 6.2, it means meat is darker and the quality is low.
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I'm working on the osteology of fossil anuran amphibians. Please let me know has the radioulna some diagnostic characters in these animals (i.e. is it possible to determine this bone up to species/genus level etc.)? I will be very thankful for your comments.
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Я определяю эти кости как Anura: строение их у современных представителей из Восточной Европы очень сходно. Иногда кажется, что какие-то конкретные кости более вероятно относятся к тому или другому семейству, но я не могу объяснить это морфологическими особенностями.
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I am interested in the phylogeny of vertebrates, i.e. the phylogenetic relationship among fishes, amphibians, reptiles, mammals and birds. I find a cool article published in 2003 (see below), and want to know the recent advances in this field.
Could you provide any more recent information on the phylogeny of vertebrates?
Thanks.
AxelMeyer, RafaelZardoya. Recent advances in the (molecular) phylogeny of vertebrates. Annual Review of Ecology, Evolution, and Systematics. 2003, 34: 311-338.
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New facts or new interpretations are presented every year. (Some of the new advances, however, soon proved to be erroneous.) As a starting point safe, I suggest a good textbook – e.g., Vertebrate life (Pearson, 2013, 9 ed.), by F H Pough et al.
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Amphibians normal blood parameters...
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Dear Dr.;
You can find good information in the following book:
Veterinary Hematology and Clinical Chemistry, Mary Anna Thrall, Glade Weiser, Robin Allison, Terry Campbell, 2nd Edition, John Wiley & Sons, 2012
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I am searching for ways to promote lizard and amphibian capacities as a profit from the restauration of their habitat in private gardens.
Therefore I need evidence of these actions
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Hi all,
do you know a local example of common toad (Bufo bufo) or fire salamander (Salamandra salamandra) population extinction (or close to extinction) probably due to road mortality ? so far I found this paper: Cooke, A. (2011) 'The role of road traffic in the near extinction of Common Toads (Bufo bufo) in Ramsey and Bury', NatCambridgesh, 53, pp. 45-50.
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Hi all,
Could you guide me/suggest an article that helps in preparing chromosomes from long preserved tissues of Beetles, Geckos, Amphibians etc.,
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If DNA can isolate then fragmented chromosome also can be. In fish karyome database in NBFGR, India...you can have check once for your query. Regards
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Amphibian conservation demands that less specimens are collected for parasitological investigations. Is there a means of detecting species of helminth parasites within a host without having to sacrifice the host? How can we make parasitological investigations of amphibians more sustainable?
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Following
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Relevant for my research on amphibian nematode disease in Southern Africa
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Plz you may also read this attachment
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Hi,
I'm wondering what molecular biomarkers can we get from amphibian saliva ? I'm sure we can get hormones but what else ?
Cheers,
Laurent
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Hey Laurent,
have a look at the paper " The use of buccal swabs as a minimal-invasive method for detecting effects of pesticide exposure on enzymatic activity in common wall lizards" from Mingo et al. (2016).
They analysed GST, GR and AChE in wall lizards. I would guess that this will also work for amphibians and several other biomarkers. We took some Bufo bufo saliva samples to test this last year, but haven't analyzed them yet.
Best,
Christoph
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I sequenced one mitochondrial genome of frog and found three control regions. It is very interesting that three different tRNAs are existed in three different control regions, respectively. I failed to find some references with similar phenomenon. Do anyone know some references or would you like to tell me the paper?
I found some references, which own one control region with one tRNA in it. Most researchers used the TDRL model to explain this phenomenon.
Do you have any good suggestion to explain it?
Thanks.
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Dear Dr. Chen,
Thank you for your information.
Jiayong
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Hello ResearchGate community,
Our lab has recently adopted GBS SNP data to gain a better insight into population dynamics of endangered non-model species in Australia (mainly amphibians, geckos and marsupials).
For most projects, after gaining a first insight into current population structure, we would like to understand whether the observed structure is a result of recent or ancient splits. As most of us are relatively new to the world of GBS and SNP data, we would really appreciate if you could point us to the best methodologies for doing so.
Any help is much appreciated.
Kind regards,
Lorenzo
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I am by no means an expert in this but will share what I have come across. I believe BEAST, in particular the extension SNAPP, provides a way to generate phylogenies using allele frequencies. Then you can fix divergence dates on some of your nodes and estimate them for the rest of the tree. BEAST* also offers an option that exploit the actual haplotypes (not just SNPs) to estimate divergence. See Saglam et al. ( ) for an example.
I also received some advice to use MCMCtree to do something similar, which is part of the PAML package. Good luck.
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Dear all,
In literature, we may frequently face with the term "stream breeding/spring breeding" amphibian. There is likewise the term of "stream dwelling/spring dwelling" amphibian. I need to know which taxonomic groups are stream dwelling and which one are spring dwelling.
Any idea would be appreciated.
Forough
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Micrixalidae is a family of 'Torrent Frogs' as they are found only in the fast flowing streams of evergreen forests of Western Ghats. And in one species, there is a report of tadpoles from the sand beds of forested streams.
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I have got five species of adult trematodes and three species of metacercariae from a single individual frog. How can it bee linked to its ecology/ecological parasitology?
At the same time many species of frogs from the same habitat are free from parasitic infection.
I have had enough experience in Parasitology (especially trematodology) but very poor in ecological parasitology. Seeking expert advices/opinions/comments from eminent scientists/researchers on the above.
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Parasite colonization inside the host and way of life would vary according to their feature.
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Dear everyone,
I have a dataset on the migration pattern of amphibians, including the angle at which the movements were initiated, and the distance covered.
I am interested in finding out if the angle described by the movements (i.e. directionality) is related to landscape features. To do so I need to use circular statistics as binary encoding "towards" and "away" from key features is not precise enough.
Would you be able to recommend a methodology to do so, and/or a (free) software that can be used for such an analysis.
Thank you in advance,
Amael
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Thank you very much, Max! I will try Oriana first, hoping it is straight forwards!
Amael
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Amphibia on the tree
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Hello,
It's Kaloula pulchra and it's an interesting observation because the species is not often found on trees. Other species within the genus (K. borealis) also climbs trees in fall, for unknown reasons (see link below), and other species preferentially live on trees ( Blackburn et al. 2013. Evolution 67:2631–2646). If it's a (natural) recurrent observation it could be interesting to think about a natural history note to report the behaviour.
Best,
Amael
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I'd appreciate advice about acoustic recorders for undertaking surveys for a range of taxa, particularly birds, frogs and microbats (i.e. audible and ultrasonic). The units will be left in situ for days/weeks and will need to withstand a range of environmental conditions (e.g. deserts and wet tropics). I've previously used Song Meters. Is anything better.....that's not considerably more expensive? If not, which Songmeter model would be best? Thanks in advance.
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Hi, you can use the 4th generation Song Meter SM4 which is a compact, weatherproof, dual-channel acoustic recorder capable of capturing large amounts of data from wildlife such as birds, frogs and aquatic life.
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Dear all,
 
We are working on Amphibians in Cameroon especially the giant frog and other critically endangered species living around Littoral and South west region of Cameroon. We want to know if I can have some partners to promote the conservation of this species who is captured and eaten by many communities as their protein source. We collect information on the diversity of this species, phenotypical and genotypical identification, habitat, mode of feeding, treats, sensitization and training of communities adjacent to areas where Amphibians are found.
For more information, visit our website www.abirsd.org, or contact: petercoolpetercool@yahoo.co.uk / info@abirsd.org
Thanks
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Thanks Dr Bakwo for your contribution. i will contact them and give you feeback
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I am looking for experts suggestions, which have experience on thess species groups distributions, therefore capable to point out species from high andean and paramos ecosystems that might also be point of interest to develop niche models. 
With these models we expect to assess, through high resolution climate information, changes of distribution under climate change scenarios. 
I hope i made myself clear.
Thanks beforehand. 
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Hi Johana, you got several great ideas already. To contribute with just a thought, I would recommend to chose a species that is suitable to be accessed in the needed frequency and numbers to develop a sound based model. Some charismatic XYZ species may be very interesting but if you find them in low numbers or they are present only during a season, they may not be the best source of data to develop your model.
Models are as good as the information they were based on. So, thinking in niche modeling for climate change assessment, I would choose species among insects, rodents, plants and a-like, that were locally known to have strict environment limits and are accessible in large numbers to dilute the noise from individual divergences. Therefore, more probable to show geographic/demographic/fitness/phenotype shifts due to climate change.
Kind regards!
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Hi. I am searching for literature about mutation rates in mitochondrial genes (besides those for ND2 from Crawford 2003), particulary for amphibians. Does somebody has references about it that could recommend me?
Thanks in advance
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Before putting priors into your BEAST xml file, I would use MEGA to test if your any of your genes are clock-like. If they are you can use a strict clock model when you enter the rate. If they are not clock-like you should use a relaxed clock model. Use BEAUTI to create your BEAST xml file.
Most importantly, the rates most often reported in literature are the pairwise divergence rates. In BEAST you need to enter the rate along a single lineage. This means simply dividing the rate by 2.   
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It is already well known that the world wide amphibian dying has its reason for one part in the baby birth control pill, where the one active substance from it comes into the enviroment over the urin if there is  no biological clarification stage- and this makes after a message from a big pharma industry not only amphibian barren, unfortunately all animals- therefore this could be one of several reasons for world wide bee dying too? I asked a professor for plants and just got a short answer, that they found already antibiotica in plants, therefore it is probably that there is more from active substances from pills in plants all over the world- here a link for it in water:
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did research on this matter.
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I am studying two insular populations of an amphibian species. These insular populations are located in continental islands, which got separated from the mainland following the sea level rise during the Holocene (approximately at 8-10 kya).
I am using an ABC framework to test which demographic scenarios are most supported. Broadly, I am testing two competing scenarios; (1) a scenario of genetic isolation following sea level rise (i.e. vicariance). This scenario implies that these populations already inhabited these islands before their formation due to sea level rise; and (2) a posteriori colonization of these islands from the mainland by few individuals. I am assessing this in DIYABC 2.1 with microsatellite data.
However, I am also interested in testing a third scenario: (3) vicariance but continuous gene flow from the mainland after the formation of islands. DIYABC does not mode explicitly model gene flow, although admixture events can be created. The issue with these admixture events is that they require the creation of an additional population resultant from the admixture of two existing populations. 
To better illustrate what I want to model, please find attached a pdf representing, in a simplistic way, the demographic scenarios. Consider the insular populations N1 and N2 and the mainland population N3. There are three scenarios represented, but the scenario "Vicariance + gene flow" is the one that I cannot include in DIYABC. In the figure I just represented gene flow from mainland (N3) to one insular population (N2).
Do you know if there is a clever way to model this in DIYABC? If not, which software do you recommend to accomplish this using microsatellite data? 
Thank you very much for your help.
Best,
André
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I use Hudson's ms and R for ABC analysis. You can modify some code I have made available at: http://github.com/mnavascues/microsatABC-IM
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I work with amphibians and I can only find antibodies for my gene generated against mammalian sequences. I have contacted as many companies as possible and there is no epitope that is a perfect match for my gene. Will an antibody recognize a protein that isn't a perfect match?
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I agree with Michael and Gertrudis that this is variable, difficult to predict, and should be determined experimentally.
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Hello,
I have fixed and obtained frozen sections of salamander embryos for fluorescence microscopy. The sections are 8um thick and the embryo used for the attached images was kidney bean shaped (sorry not super familiar with the development; helping someone else with this project), maybe stage #24 according to this chart: http://www.virginiaherpetologicalsociety.com/amphibians/amphibian-development/amphibian-development.htm
In the attached images, I only rehydrated the section and stained it with Hoechst. I then mounted using a glycerol-tris-n-propyl gallate mounting medium. Unfortunately, there is a lot of autofluorescence in both the TRITC and FITC channels and it's pretty bright. I didn't see nearly as much autofluorescence when I mounted in PBS alone, but I was hoping for a more permanent mounting medium. My experimental sample will require viewing the TRITC and DAPI channels so I really need to minimize this autofluorescence.
So my question is, what is the stuff autofluorescing and why does the glycerol make it worse? I would guess that it is yolk and did run across some info about it being autofluorescent but it would be nice to be certain. 
Thank you!