Questions related to Frogs
I am studying the phagocytic activity of frog blood cells. After incubating frog whole blood with yeast cells, I had this microscopic image; Could be possible for frog erythrocytes to engulf yeast cell?
I am currently trying to make concentrated samples with a hundred frog embryos to make western blots.
Everything is fine during protein extraction. However, when I add the laemmi buffer, I begin to have aggregates that form. Once heated, my samples turn into blue chewing gum impossible to resurface.
Has this ever happened to anyone?
Hi, I have written a code for solving the second-order Helmholtz-Maxwell electromagnetic equation using finite-difference time-domain (FDTD) method. There is not much material available online for this second-order differential equation, except for its discretized forms. All the algorithms I've found online or offline are based on Yee's leap frog method.
I have been following the book "Computational electrodynamics" by Taflov and a PDF by Prof. John S, which is available online. When I use Yee's grid and implement the Total-Field Scattered-Field (TFSF) method, it works fine. However, when I try to implement TFSF on the code for the second-order Maxwell differential equation code then, it doesn't work at all. I'm following the same method to suppress the leftward traveling wave as mentioned in above books, albeit with slight modification.
If someone has prior experience with this and can help me, it would be of greatly appreciated.
I am researching a phenomenon that has many theoretical perspectives. If I have to unify the various sub-components of the phenomenon, I need to get out of the system to get a clearer vision...something like the 'boiling frog syndrome'. Can there be a theoretical explanation why it is better to be outside the system to bring about integration rather than being within the system?
I have attached a figure to enable visualization of my predicament. I will deeply value your insights.
Recently, my frogs and toads reported for the first time have been rejected by certain journals because the paper lacked genetic data. Now, the question is "Do we need to really get genetic codes for all species?" How can genetic data which comes as the outcome of laboratory works help us identify species in the field? Or we just keep the species name as just certain frog or toad (just example) and keep waiting until a genetic work is done? Don't species evolve and change their genetic content based on their evolutionary traits over time? Do we really trust the genetic codes for species identifications?
Are education systems "bribed" and extorted by the federal funding systems in place?
The centralized systems in education and federal funding appear to require political approaches to providing education.
Does this equate to extortion and forced propaganda in our education system, driven by a centralized force so powerful that the funding mechanism has created an indoctrination network?
Can a school/university get funding if it refuses to instruct the flavor of the day topics, such as CRT, Gender identification, and ESG?
Can students who have religious faiths choose to bypass indoctrination that violates their code of morality?
Can it be asked without condemnation, or are we now in a police state globally where asking these questions creates a digital profile that classifies the one asking as a threat to those in power?
Modern censorship, cutoff of access to social media sites such as LinkedIn, Facebook, Twitter, Meta, etc. are real and relevant examples of abuse of power already taking place.
It seems that federal funds are currently used, not to assist in financing a well-educated society, but rather, to extort institutions into programming a conformist society of minions to centralized command and control authoritarian rule.
I find this to be a troubling signal that shows that the education system has already fallen victim to being the frog in a pot of water that slowly becomes boiling water to kill the frog, with the frog never even knowing it.
I am trying to extract DNA from frozen newt skin swabs but have been getting really low concentrations (about 1-3 ng/µl). I am using the Qiagen Blood & Tissue Kit and just cut of the tip of the swab and let lysis go on overnight at 56°C and 400 rpm. I wonder if I should add a more rigorous shaking step? I know that I can't expect very high concentrations from this method, but I think it should be higher than what I have (e.g. Ringler, 2018: Testing skin swabbing for DNA sampling in dendrobatid frogs; she extracted 20-170 ng/µl in dendrobatid frogs). Does anybody have a useful hint what I could change to get higher concentrations? Thank you!
Ideally recorders that can be left out for weeks at a time, with frogs from tropical forests the target animals.
Pros and cons of passive acoustic recorders in the Neotropics.
Ideally for recorders to be left up for weeks at a time.
Frogs are the target animals.
Troglotrema a. is a parasite mainly found in European Forest Tilapia, which can cause massive lesions in the skulls of the animals, depending on the severity of infestation. So far, the parasite has been detected almost exclusively in dead animals.
However, during the rearing of orphaned polecats in the wildlife station of the Retscheider Hof (Bad Honnef), evidence of Troglotrema. a. was already found in juvenile animals by means of fecal analysis. However, infection by consumption of frogs seems unlikely at this age.
Vogel/Voelker (1978) describe the developmental cycle of T.a. as follows:
From the first intermediate host, the spring snails, the cercariae pass into the second intermediate host. Here the grass frog is indicated. Here the metacercariae develop, which then enter the wood tiltis through consumption and develop into adult sucking worms in the sinuses and frontal bone area. These in turn secrete eggs that are ingested by the spring snails.
Is a different development cycle conceivable or even known for Troglotrema acutum (Trematoda) than the one described by Vogel/Voelker (1978) (spring snail, frog, polecat)?
How do metacercariae move from flowing water to standing water (wetlands) to infect the grass frog?
How do the metacercariae ingested with the frog travel from the gastrointestinal tract to the sinuses/frontal sinus?
By what route do the excreted eggs of the adult sucking worm re-enter the headwater streams to be ingested by the headwater snail?
I’m new to the publishing world and I’m hoping to get some advice on the most appropriate model to use to statistically analysing my data please. To summarise the project, I deployed four AudioMoths (which are audio recording devices) at four different sites in the middle of the Davies’ tree frog’s breeding season (one AudioMoth per site). There is virtually no published research on the species and the frog’s daily calling activity has not been described. The aim of the paper is to describe the daily calling activity of the species (i.e., what time of day does the frog call at) and I am hypothesising that the frog will peak in calling activity in the hours just after sunset (i.e., between 7-9 pm).
My AudioMoths recorded 5-mins at the start of every half an hour in a day for a 10 day period. I used an automated sound recognition software called Kaleidoscope Pro to extract the frog’s call from all the audio data. I then calculated how many calls there were in each hour of the day (so there’s two lots of 5 min recordings per hour) and then took the average number of calls per hour from all 10 days for each detector (so each hour has a sample size of 10, 1 hour for each of the 10 days). I then combined the data from all 4 AudioMoths to get an overall average number of calls for each hour of each day (so each hour now has a sample size of 40). I then turned the average number of calls per hour to a percentage of calls per hour and created the attached graph with standard error bars.
What I’m hoping to get help with is how to appropriately test if there is a difference between the number of calls in each of the 24 hours per day? Would I use an ANOVA initially to see if there is a difference in the number of calls between any of the hours and then follow it up with a post-hoc test to find out where the differences are? Or would it be best to compare the percentage of calls per hour and use a chi-squared test? Or is there another better option? What problems arise when I’m comparing 24 different groups (because there are 24 hours in a day)? I’m mainly wanting to know how best to show statistically that there is a peak in calling between 7-9pm (if there is in fact a statistical peak)? If there is a statistically significant peak in calling between 7-9pm, then future field surveys can survey in this time period to maximise the probability of detecting the species.
I’d love to hear your thoughts on this if you have the time please! I’d also really appreciate to hear your explanation as to why you suggest the approach you do please?
This is a blood smear done to a post mortem frog after few hours in -2C temperature for an experiment. how can the changes in erythrocyte morphology can be explained?
I extracted both DNA and RNA from frog (X. tropicalis) liver samples using Qiagen Allprep DNA/RNA mini kit, and ran an electrophoresis agarose gel to check for DNA integrity, and a Bioanalyzer RNA nano chip to check for RNA integrity.
It seems like the DNA is completely degraded, the RNA looks quite good (the baseline anomaly was slightly higher than the threshold an had to adjust for that manually, but the profile looks good).
The DNA samples are in the wells 5 and 6 of the gel, and correspond to samples 1 and 2 in the bioanalyzer, respectively. The first 3 wells are brain samples with apparently good DNA.
Obs: The samples were thawed before for aliquot and using for other analyses.
I would appreciate any comments or thoughts.
There are many models which helps us to explain in some detail certain processes or phenomena or even to predict to some point how or why or what will happen to the organism if we did... animal testing, trying out a drug or a compound. there are many models, and this does not mean they are used dfor the same reason. Some organisms are better suited than others. How many models and why is this model appropriate? I would like to start by adding a partial answer. I hope we can make a detailed answer to the question, which fully explains the reason or the main aspects on the use of organisms for a model.
A related model may be the use of E. Coli for secveral purposes. some of them require the use of variants or stains, or some need the use og genetic engineering. Is this organism considered a model despite the transformations, or is it more related to the fact it is widely used? How many strains are there, why would we use a particular strain and is it related to the case of general use of model organisms? Are the more cases where we use organisms like this? Do we classify them under the model organism as a whole? Is there anything else you would like to add to the question?
I have a query regarding the most appropriate experimental design and statistical analysis for a research project. The project study area is located in a high altitude lagoon (Los Andes, Peru). The study subject is an endangered frog species (the Lake Junín frog).
The research question is: What is the impact of heavy metals, eutrophication and water level variation on the abundance and biomass of the Telmatobius macrostomus and T. Brachydactylu population?
After many field visits and literature research we've found out the 3 main environmental pressures on the frog population: (i) heavy metals from mining activities, (ii) eutrophication produced by untreated urban sewage discharge and (iii) water level variation to assure enough water for hydropower downstream. We have monitoring data (from secondary sources) on heavy metal concentration and some eutrophication indicators (N, P, DBO). For now we only have the resources to collect field data on water level variation, and the frog's biomass and abundance.
Currently we don't have resources to collect more data on heavy metal pollution or nutrient content in the water. Therefore, with the available data, we want to have some idea on what are the most relevant environmental pressures to:
- Know where to allocate more resources on monitoring and
- Evaluate some remediation techniques to improve the frog's habitat.
Thanks in advance for your comments.
ps. Feel free to contact me if any of you are interested in helping designing the study.
I have observed many infectious individuals of Euphlyctis sp. in various populations across Northeast India, while I have not observe such infection in other aquatic/semi-aquatic frog species. Can anyone tell me the probable reason as to why Euphlyctis sp. are more prone to infection?
I am trying to find associations between dorsal patterns (uniform, complex etc) of frogs and microhabitat selection (leaf litter, rock etc). I am using a 5x6 contigency table but more than 20% have < 5 expected values, because of this I have opted for the Fishers exact test rather than chi square. The following code was used that gave a significant result:
fisher.test(CT.MH.PAT, simulate.p.value = TRUE, B=100000)
I now need to find out which levels associate most with each other but I am struggling to find which post hoc test I should use. I have attempted to use the following:
But R doesnt seem to recognise these functions (I have devtools, tidyverse and MASS all loaded), is there a package I am missing?
If anyone could provide some assistance, or perhaps a good source I can read through! Thank you.
Kindly, name some reliable/popular field survey technique employed for amphibians (frogs) & terrestrial skinks (scincidae) in tropical forests. Thanks a ton.
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).
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
The goal is to identify different individuals in a population and observe their behavior.
I am trying to understan if there is a latitudinal cline in the degree of melanization of a frog, but I am not quite sure of which would be the best statistical analysis to perform.
I'm collecting road mortality data once per week along two different 3.5 km sections of road. I'm interested to know if there are taxa specific differences in road mortality.
As of now, I've simply pooled mortalities for each taxa (frogs, turtles, small mammals, etc.) from both sites and talked about differences with simple summary statistics (count of mortalities per group, proportion of mortalities attributed to each group, average count per survey). I'm not so much interested to compare between sites, but between taxa as a whole.
Is there a statistical test I can use to answer the question "is there is a difference in road mortality between taxa?"
Are the summary statistics I've been using sufficient to answer this question?
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!
I need an applicable molecular biomarker like expressing the genes in frogs who expose to chytrid fungus in different environmental stressors. Indeed, what kinds of genes I should notice in frog's body for my molecular analysis?
Probably a silly question but I am a complete novice in molecular biology. I have decided to attempt a phylogenetic analysis. I have managed to create a phylogenetic tree but want to confirm whether it is detrimental or not to align both types of sequences.
For some context I have collected 87 sequences from a frog family. Some of these species are fairly unknown so the choices for DNA sequences are limited.
I have got RAG1, Cyst and 16S rRNA sequences so far and my tree looks okay (as far as I'm aware).
There are a few examples of Asian frogs (Odorrana and Huia) with advertisement calls that contain either audible and ultrasonic components, or purely ultrasonic components, I was wondering if these types of calls are restricted to Asia or if there are other examples found elsewhere.
Hi, I'm analyzing some data on frog calls (model output in pic 1), and I'm running into trouble due to my linear mixed models not meeting the assumption of homoscedasticity. That's something I've run into before, but rather than the usual wedge shape that I have seen on other projects, this time there are three distinct clusters of points in my fitted vs. residuals plot (see pic 2). This pattern is resistant to all of the usual transformations that can be applied, and one person I have asked said that this pattern suggests that there is an additional variable that should be included in my model, but is currently unaccounted for (and he also suggests this may be the cause of my wonky QQ plot). Does this sound like a reasonable conclusion as to what is causing this clustered pattern in the residuals vs fitted plot? Has anyone dealt with this? And is there a methodical way to attack this problem? Or do I just need to try adding new fixed/random effects to the model (in a considered and methodical way)?
Model background: The dependent and independent variables are all continuous, and I've included one interaction term, and male identity as a random intercept.
Thanks for your time, and for any information/advice you can provide!
I am undertaking an undergraduate project where I'll be conducting some frog surveys and I was wondering what the best handheld GPS unit would be for this work that won't break me financially?
Additionally, how accurate are the GPS receivers in smartphones such as Samsung Galaxy S9's or tablets such as iPad's these days, and would it be acceptable to use one of these options instead?
The survey work will be in woodlands working in and around drainage channels so I am a bit concerned about the accuracy in this kind of environment.
Thanks in advance.
When some of the call properties show negative correlation with temperature . Is it necessary to perform temperature correction of such properties ? What are the steps involved in the correction.
Platz and forester, 1988 gives the formula D14 = Damb - (Tamb. -14.0) (-0.0974) . Can we apply our desired temperature here (Eg. 20 °C ? Or is there any better way to perform this correction
I am trying to design an experiment to obtain DNA from historical specimens of frogs (some of them were collected >100 years old.
Anyone experience with the manufactured DNA extraction kit from FFPE tissue samples? Could you share your experience in using this kit compare to the normal tissue extraction kits or the old-school methods (e.g., phenol-chloroform method)? Which one work best (e.g., produce large amount and high quality of DNA required for subsequent steps)?
Also, what do you think about this study?
And if anyone experience with the forensic extraction kit?
I really appreciate your help!
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.✌️
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
These families are best known for the nymphal stage, which produces a cover of foamed-up plant sap resembling saliva; the nymphs are therefore commonly known as spittlebugs and their foam as cuckoo spit, frog spit, or snake spit.
which one I must select from meta-heuristic algorithms (e.g., GA,Ant,Whale,Frog,Antlion, PSO, GSA, etc.) to Optimize the case
The role of whale or frog or lion ant algorithms ...etc... useful in solving a particular issue and how we decide which of them choose in our problems or which one will be effective in the problem
I am interested in these algorithms now and I hope you join to a research group, and work with them research in this area
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
Swammerdam (17th century) stimulated a muscle in a fluid-filled jar with a small-bore tube attached, measuring a slight decrease in volume, disproving the "balloonist" theory of muscle contraction. This finding is well-replicated in frog sartorius, but there is a recent claim (Clark & Demer 2016) that human eye muscles are different in that they increase as much as 18% in total volume when they contract to rotate the eye. I don't believe it!
Can anyone point me to contraction-volume measurements in vertebrate muscles, or any muscles that might be more like human EOMs, or to an expert who might know about this stuff?
What are the preferred tree species for the gray tree frog (Hyla versicolor) in New Brunswick, Canada? We are trying to clear a site of invasive glossy buckthorn and need to plant some trees to restore the site. What species would you recommend we plant?
It is a small site, about 9 hectares, but a very important site for H. versicolor.
I would like to analyse the shape of parotoid glands between two species by using landmarks. But because of assymetries between glands of an individual (i have observed assymetries in many of my samples) i m doubtful to try it. I am also inexperienced in this approach. So,could anyone inform me whether it is an appliable idea to analyse the shape of parotoid glands ?
I am trying to compare the abundance and composition of frog in different site (10 sites) over time. For each site, i sampled the abundance of each frog species and community of frog during 4 years. I would like to know if there is difference in the species abundance and composition between sites in between year.
I hope this finds you all well,
I will be soob have to extract the nuclear protein Nrf2 from zebrafish embryos homogenate. I have to saperate the cytosolic protein from nuclear protein, which is the Nrf2. Can anyone refer me to such a protocol b/c most of the protocols I have found are either performed in cells or in rodents. I am looking for a protocol that is performed in zebrafish (Danio rerio) or an African clawed frog (Xenopus laevis). If someone knows of such a protocol, I would truly and sincerely appreciate the help.
Thank you very much, I truly appreciate.
Good afternoon. Hope you guys are okay!
First of all, I would like to know what is the best way to prepare a frog finger (HAHA). The last time I prepared the same type of sample, I fixed on a carbon tape and coated with gold, but when observing on the SEM the frog finger was very dried out.
I would like to know if there is another way to prepare the same type of sample avoiding the dryness.
Thank you for the attention!
P.S: I attached some photos of the first analysis
During a recent field trip in Amazonian Peru, a colleague of mine and I observed stingless bees (Trigona sp.) preying upon frog egg clutches deposited on vegetation above a temporary pond. I would like to know if other researchers have observed this behavior and if you can share any published/unpublished account(s) on this topic.
Dear colleagues, I need to estimate the body temperature of very small frogs/toads (ca 20 mm SVL) with minimal disturbance, thus using an infrared thermometer. I screened the literature but can’t find any model suiting my needs (max 1°C accuracy, spot size of ca 5 mm, rugged enough to be used in field conditions, and preferably below 200 USD). A fixed emissivity of 0.95 is fine.
The model I have (Fluke 62max+) has a minimum spot size of ca 30 mm, which is way too large. If I understand well, the problem when the spot size is larger than the target is that the temperature inferred is an average of the temperature within the spot (so including the substrate around) and I need to avoid that. Does anyone have some experience with this and could suggest a good IR thermometer model for my purpose? Thanks in advance, Philippe.
I have sequenced a 7 kb part of frog mtDNA and have generated a bam file of the sequence. In IVG, I can visualize manually the per base read coverage but I need this information in an excel format (.csv or xlsx) so that I can load it into R. Please suggest me way/s to do it.
Is it possible to see if two ultrashort laser pulse (800 nm with 110 fs duration, and 1040 nm with 200 fs duration, 80 Mhz rep rate) overlap in both spatial and temporal domain with a SHG-FROG setup? Are there any specific crystals for this purpose?
What would be the best way to overlap the beams if this doesn't work?
The Ladder in the first well is biolabs quick-load 100bp and ranges from 100 to 1517 bp. I therefore don't understand how the bands present in wells 8 & 9 can possibly be? What could explain such large PCR products (or seemingly large)
This is a microsattelite locus for which the primers were developed using the genome of Pelophylax Ridibundus (European frog) and I'm experimenting with it's species specificity by running PCRs with other species of the pelophylax genus (wells 2-7). The 8th and 9th wells in question are in fact the P. ridibundis DNA samples and I'm very intrigued as to what may have caused such results.
Thanks for reading and any info is greatly appreciated :)
We documented a mass mortality event involving 2000+ sub-adult (almost fully grown) Ranid frogs at a brackish lagoon on Samothraki Island (Greece) just a few days ago (the Lagoon is called Koufki Lagoon, adjacent to Agios Andreas Lagoon SW of Kamariotissa port). Most dead and many living frogs were concentrated at areas of the lagoon that had slightly lower salinity - and water salinity may be rising (due to rapid drying). We have bio-samples and water samples and measurements. What we are interested in, is if anyone has seen this kind of event before in any kind of fresh or brackish waterbody? Any bibliographic support would be much appreciated.
Some frogs were reported the loss of ATP8 and ND5 genes.
Do you know any other vertebrate?
I found some paper said some fishes lost the ATP8 gene. But some researchers identified it was incorrect because the gene rearrangement was happened.
Of course the loss of ATP8 gene was normal in some in vertebrate .
But I need some examples in vertebrate.
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.
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,
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.
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: email@example.com / firstname.lastname@example.org
Hello folks. Need a hand identifying the species of this Anuran. The specimen came into the collections at the Oxford University Museum of Natural History with the rather unhelpful label of 'Spadefoot toat'.
As part of other work, I've done some Photogrammetry and CT-Scanning and appears to be female by the presence of eggs from the scans.
3D Model of Skin: https://skfb.ly/MXDH
3D Model of Skeleton: https://skfb.ly/RzyK
Any help would be greatly appreciated.
The pictures were taken in April 2016 in lowland tropical forests of the Corcovado National Park (Costa Rica). The second species (very large amphibian) might be Leptodactylus savagei.
The pictures were taken in lowland tropical forests of the Tortuguero National Park in Costa Rica. The first might be Hypsiboas rufitelus, the second Oophaga pumilo.