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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.
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Unfortunately, systems that were innocent, such as education, can become doctrinal or ideological elements in the hands of certain governments.
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Hi, I have written a code for solving the Helmholtz-Maxwell electromagnetic equation using finite difference time domain (FDTD) method. There is not much material available online for this equation. All the algorithms available online or offline are based on Yee's leap frog method.
I have followed the book Computational electrodynamics by Taflov and a PDF by Prof. John S that is available online. Now when I follow Yee's grid then TFSF work very well but when I try to implement it on my code then it doesn't work at all. I'm following the same method to suppress the leftward traveling wave as discussed in these sources albeit with slight modification.
If someone has done it before and can help me on this then it would be of great help.
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Hello. Isn't the M-H equation the same as the Wave equation? That equation is discussed in Taflove 2nd Ed., Section 2.5. When you implement the Wave Equation and the FDTD equations, without separation into TF-SF, do you get identical results? What additional info about the Wave equation are you looking for? Why do you want to extract the SF (I assume SF=Field without Scatterer-Field With Scatterer)?
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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!
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Why is the lysis so long? It should take 10 min.
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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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I want to work on the effect of nitrate fertilizer on toad (Bufo bufo)
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Interesting question
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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?
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(The trematode Troglotrema acutum and nematodes of the genus Skrjabingylus are parasitic helminths infecting nasal sinuses of mustelids. Despite different infection routes of these parasites, their occurrence becomes evident due to their destructive lesions of the bone structure of the head).. cited from: Evidence of Troglotrema acutum and Skrjabingylus sp coinfection in a polecat from Lower Austria... DOI: 10.1515/HELMIN-2015-0011
Best regards
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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?
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Hi there Lachlan,
I don't think you want to do ANOVA or similar linear models since you're using time-dependent data (the number of calls at one hour depends on the number of calls the previous hour). I think your approach of averaging throughout the days is good, but maybe there's no need to average within the hour. Why not have 48 points instead of 24?
Regarding the analysis, you need to do something called time-series analysis, which can get quite complicated (https://stats.stackexchange.com/questions/30640/a-list-of-common-time-series-tests). Instead of going deep into time-series statistics, I propose you perform pair-wise t-tests for each step (0 vs. 0.5, 0.5 vs 1, etc.). There will be significant differences if the frogs start/stop singing. Looking at your graph, that'll be significant at four different times (18-19; 20-21; 5-6 and 7-8). If the difference is (t1 - t0) is positive, the frogs singing more, if it's negative, they're singing less. Using the significance and differences you'll be able to describe their singing behaviour.
Just a note, you might want to consider plotting the counts in a circle plot (polar coordinates), since the end and beggining of the plot are the same hour (24/0). Here you can see some examples in R. http://rstudio-pubs-static.s3.amazonaws.com/3369_998f8b2d788e4a0384ae565c4280aa47.html
Good luck!
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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?
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As far as I understood correctly from the description of this case. We see an erythrocyte that has frozen - it has a white lumen in its cell, the ice has violated the integrity of the cell. And normal red blood cells that did not have time to freeze.
Regards, Sergey
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Hello,
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.
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Thank you for your answers!
Artur Burzynski
, I expected that the RNA would be degraded due to thawing the samples (ubiquitous RNAses) while the DNA (usually considered less prone to degradation) would still be viable, so I was wondering what would be the cause for seeing the exact opposite.
As I am more interested in extracting the DNA for methylation analysis, I was wondering if it could be something related with the extraction kit that I used, so if I use another method, maybe I could still save the DNA from those samples.
But I was not able to formulate a reasonable technical or biological hypothesis for this.
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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?
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interesting. I did not know. Keep going!
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Dear colleagues
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.
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Dear Dalia, Kindly follow the references, although I think you need more papers for comparing the contamination and pollutions, by the way , you can ask me to send other articles for your discussion part!
Anyway, hope they would be helpful:
1- Distribution and Accumulation of Heavy Metals in Surface Sediment of Lake Junín National Reserve, Peru; Available online:
and it is in research gate !
2- The heavy metal contamination of Lake Junín National Reserve, Peru: An unintended consequence of the juxtaposition of hydroelectricity and mining
3- The History of Mining in Cerro de Pasco and Heavy Metal Deposition in Lake Junin Peru ( it related in 2012 and I attached it)
4- Protected Area Profile Perú Junín National Reserve ( It has been attached)!
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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?
Thank you.
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Chytridiomycosis is an emerging infectious disease of amphibians caused by an aquatic fungal pathogen, Batrachochytrium dendrobatidis (Bd). You can refer the link given below for more information about Chytridomycosis and its causes, how many species are infected by it, where it is found and how it is spreading etc.
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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:
fisher.multcomp()
pairwise.fisher()
chisq.posthoc.test()
pairwise.t.test()
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.
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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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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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The goal is to identify different individuals in a population and observe their behavior.
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Hi. I hope the followimg website could help you:
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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.
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Selection of appropriate statistical method depends on the following three things: Aim and objective of the study, Type and distribution of the data used, and Nature of the observations (paired/unpaired). https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6639881/
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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?"
or
Are the summary statistics I've been using sufficient to answer this question?
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Agree with the views of Dr Zaal Kikvidze and Dr David Dawson
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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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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?
Thank you
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I don't work on this but someone in my department does. She said that it varies among species. There are several gene expression studies concerning this topic. Check out Eskew et al., 2018 and Rosenbum et al. 2008
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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).
Thanks:)
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Maybe you could use Partitionfinder (https://www.robertlanfear.com/partitionfinder/) to test if it's ok to run your analysis as a unique sequence or if it would be better to partition it based on different substitution rates. Some algorithms (like the one on RAxML) can construct a tree based on partitioned data with reliable results.
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Has anyone used a drone equipped with a microphone/camera to study the acoustic behavior of canopy species of frogs and/or birds?
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Hassan some drones are fairlu silent and especially when hovering. It is possible. I prefer a drone drop and pick up of a song meter to record for longer time periods. That also eliminates the drone noise or possible drone interactions with species.
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How should the initial population be taken in the shuffled frog leaping algorithm?
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The algorithm should generate the initial population in the search space
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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.
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Hi Davi Lee,
Fascinating stuff!
I always thought that glass frogs would be good candidates for ultrasound production.
What kind of recorder are you using, Davi Lee? With a sampling rate of 44 kHz, you will necessarily be limited to 22 kHz upper limit in your recordings, as you point out. We are using a 2-channel digital recorder from Sound Devices (model 722) and a GRAS ultrasonic microphone. The recorder has adjustable sample rates up to 192 kHz. I like this recorder very much. It can record directly onto its compact flash card and then the data can be stored on the internal 40 GByte hard drive. The recorder is small and lightweight, but a bit expensive; the microphone is also high-priced. If you can borrow one for your work, this would be a good plan. Hope this helps.
Peter
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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!
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You somehow seem to confuse homoscedasticity with the fitted values. Clusters of the fitted values only mean that not all fitted values are equally frequent. This happens when the model function has segments that are more "horizontal" (parallel to the predictor dimensions) then other segments. For these "flat" segments, all fitted values are very similar, leading to a cluster in the fittes vs. residual plot (in your case it should be the interaction of continuous predictors allowing for such a "flat" region in the response surface). Homosecdasticity is a feature of the residuals. The variance of the residuals should be similar, independent of anything else, including the fitted values. This means that the scattering in the vertical direction in the residuals vs. fitted plot should be similar along the horizontal direction. It is irrelevant if and where the values on the horizontal direction cluster.
In your plot I don't see a severe violation of the assumption of homoscedasticity.
The QQ plot shows that there is in fact some more or less considerable information in the data that your model is not aware of. Hard to say if this unused information is relevant to your problem.
I would make (many different) plots of the fitted function together with the observed data to get an impression how the model looks like, if it is reasonable and where it has problems to describe the data well. Added variable plots can also be helpful here.
Possible remedies are: i) adding interactions, ii) modelling non-linear relationships.
i) You could try to fit a model with at least all pair-wise interactions
~ (bout_progress + bout_length + WAMP_t + numbr_chucks)^2
or even with all triple-interactions
~ (bout_progress + bout_length + WAMP_t + numbr_chucks)^3
to see if this may be due to a missed interaction.
ii) If variables are included "just" to adjust the effects of other variables, you may consider to use splines rather than linear relationships for these variables (include package splines, then use ... + ns(x, df=3) + ... to include a nonlinear relationship of the variable x in your model. "df" is the number of degrees of freedom of the spline and has a similar meaning like the order of a polynomial (like poly(x,3)), but splines are more flexible (e.g. a spline with 3 d.f. does not need to be symmetric, but a cubic polynomial must be symmetric).
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G'day all,
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.
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Please see some of the papers we developed at gps-test-site.uga.edu/.
In general, a recreation grade GPS (inexpensive GPS, watch, or phone) has an average accuracy of 6-10 m. A better GPS ($1000-5000) should have an average accuracy of 2-3 m. None of these are "sub-meter" accurate on average.
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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
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Hi Vineeth,
In the paper
Narins PM (1995) Temperature dependence of auditory function in the frog. In: Advances in Hearing Research (GA Manley, GM Klump, C Köppl, H Fastl, H Oeckinghaus eds.) World Scientific Publishers, Singapore 198-206, the authors present examples of how temperature afects frog call parameters. The following is one example from this paper of how to calculate the effect of a 10 degree C temperature change (the thermal Q10) for a frequency shift of s octaves:
In all frog species tested to date, increasing the temperature results in an upward shift in the CF and a concomitant reduction in CF-threshold of the tuning curves for fibers innervating the amphibian papilla (low- and mid-frequency fibers). In contrast, basilar papilla (high-frequency) fibers appear to have temperature-independent CFs and CF-thresholds.
Temperature induced shifts of CFs varied from 0.08 octaves/degree C for low-frequency fibers to no CF shift for high-frequency BP fibers. The frequency shift/degree C may be expressed as a thermal Q10 dB value to simplify comparisons with other temperature- dependent processes (Eatock and Manley,1981). The thermal Q10 for a frequency shift of s octaves may be calculated by:
Q1O = e**s (10/delta T)ln2 where ** means "raised to the power of."
Hope this is useful,
Peter
PS- If you want just want to compare frog calls recorded at different temperatures, then it is useful to shift all the temporal parameters to a common temperature (say 20 degrees C) for a valid comparison.
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Hey folks,
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!
Best regards,
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Three methods of DNA extraction from FFPE samples (for each method) were tested: salting-out; Phenol-chlorophorm methods and EasyPure® commercial kit. Two common methods have been used to measure quality and quantity, DNA purity / concentration is measured for quantity using a spectrophotometer (absorbance) and quantity using an agarose gel analysis.
The best concentrations and purity of DNA were obtained by salting-out method and Phenol-chlorophorm . The BCL-2 gene was amplified by PCR to evaluate the equality of extracted DNA. There was no amplification for salting-out and phenol chlorophorm methods. Instead, the BCL-2 gene was amplified only in 2 of 18 samples extracted by EasyPure ® FFPE kit.
Size: We used the same size of tissue for the three protocols, 10µm.
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How to calculate ecological risk assessment of heavy metal pollution in environmen??
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Is frog a model organism for metal toxicity assay? How much heavy metal-containing meals do frogs consume? What does the literature say about effect of heavy metals on frogs? How many people consume frog meat? People in terms of cultures. These are some of the areas of your research that you must tidy up. My answer is Yes! you can. However, your ecotoxicological model will be localized.
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For my experiment I have to extract DNA from frogs. I wanted to use a CTAB method but I have a RNA contamination. How can I remove this whitout using RNase or other enzymes? It has to be as cheap as possible!
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Use RNAaes step in the DNA extraction protocol
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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 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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I want to know if a given environmental variable has an effect on the range of two species/two groups of species. Specifically, I am interested in testing if one species has a "preference" for higher/lower ranges over the other (for example, do blue frogs prefer colder temperatures than red frogs?). I have occurrence data for my species as well as environmental data in a grid over the area. Can anyone point me in the right direction?
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Dear Kyle,
If you are familiar with R, I would suggest to explore such potential differences simply using boxplot() and kruskal.test() before trying anything more sophisticated. You will need pairwise comparisons after the kruskal-wallis test if you have more than two groups/species. Have a look at the R package conover.test. https://cran.r-project.org/web/packages/conover.test/
Cheers,
Rainer
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Reproduction by Parthenogenesis
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Yes
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Spittlebug nymphs
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.
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how to isolate proteins from spittlebug's foams
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which one I must select from meta-heuristic algorithms (e.g., GA,Ant,Whale,Frog,Antlion, PSO, GSA, etc.) to Optimize the case
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many thanks Orlando Grabiel Toledano López for your contribution
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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
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Thank you very much for your contribution Orlando Grabiel Toledano López
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Dear colleagues, I need to perfom a thyroidectomy on living early tadpoles in order to observe how it will affects the development of axial skeleton. Сertainly the tadpoles must survive after operation! Thanks for assistance!
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Dear Dr. Crawford, are you sure that two VERY will be enough?
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Dear all,
After some disappointing orders from Nasco in the US, I am looking for alternative suppliers for Xenopus laevis frogs! Any recommendations?
Thank you!
Marco
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Xenopus Express
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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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Frog from East Kalimantan
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Please have a look at the following references regarding occurrence of Rhacophorus harrissoni in the East Kalimantan province of Indonesia.
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Good afternoon, Im a livestock engineer and I have been looking for some information on frog culture . Did I help? And what specific information available ? I really appreciate it is an area that has not been explored in my country
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Dear Daniel, im a biologist from turkey. İ have experience about frog culture if you look my researchgate profile you will see my frog culture publications.
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I am wondering how long it takes the common south american toad, Rhinella arenarum to hatch.
It would be helpful to know the temperature as well.
Also, if it is known, how long does it take for this species to metamorphose.
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Well i dont have exact idea about hatching period of south american toad but it takes about 3 to 14 days and temperature could be in between 17 to 28°c, i hope this little information will be helpful to you. You can refer to this link http://dougwechsler.com/toad/toad_life_cycle.php @
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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?
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Re Stephen M Levin: Wouldn't an auxetic muscle decrease in volume when it shortens? In any case, it's actual measurements I'm after. Thanks for your interest.
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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.
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Do you have any information regarding the historic forest composition of the site or region? You could also compare your site to Hyla versicolor sites in Maine, USA. Since H. versicolor has such an extensive range spanning multiple biogeographic regions of the US, it inhabits a wide range of forest types and isn't likely associated with any particular species of tree, but may be more associated with deciduous trees in general. I would try to research the predominating tree species in sites as close as possible to your site and think about forest composition further south using the biogeography of H. versicolor as a guide.
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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 ?
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Christopher James Evelyn I am thinking about both of them but mainly focusing on the positional status. Thanks for advices, i got them.
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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.
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Hi Tokouaho,
It sounds like you should generate a diversity measure for each site (eg. such as the Shannon or Simpson index, or Bray–Curtis dissimilarity), then plot these metrics against time to look for trends. The non-parametric Kruskal–Wallis ANOVA test can be used to test for differences in species richness and relative abundance across sites. Non-metric multidimensional scaling (NMDS) plots would be useful to visualize species-richness patterns among sites based on species composition. There are other ways of getting at these type of questions, but these methods should be a good starting point.
Good luck!
Danny
ps. check out the attached files for more details on assessing temporal changes (Magurran et al. 2010) and spatial differences (Ndriantosa et al. 2017).
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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.
Sincerely,
Omar
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Hi,
I am including a reference to an article which I think might be useful. Kindly have a look.
Using hyperLOPIT to perform high-resolution mapping of the spatial proteome, Nature Protocols volume12, pages1110–1135 (2017)
Thank you.
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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
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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.
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Thank you, Rainer and Alexandre, for sharing this information. Interesting! I have also seen Trigona bees feeding on carcasses of different animals. In the upcoming months, I will share our observation of Trigona bees preying upon frog eggs ( in a short manuscript currently in review).
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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.
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Thanks all for your valuable input! I got the FLIR E8 and will do several tests while in the field.
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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.
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I have found an answer to this problem. Using galaxy the mpileup option lets you generate a file showing variants in vcf format. Viewing the data in galaxy will show you the raw read depth per base (DP). The output file will be in vcf or bcf format but it can be converted into tabular. I loaded the tabular file in R and created a .xslx file as an output. The raw read depth per base can then be accessed via excel file.
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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?
Thanks,
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Hello --
If you just want to check spatial and temporal overlap of two beams, all you need is a crystal supporting sum frequency generation (SFG) of the two input beams. You can search online for the correct crystal and phase matching angle for these wavelengths, or you could calculate them yourself very easily using some free software like SNLO's Qmix function. In any case, oftentimes the required crystal cut isn't too different than one you might use for SHG (like in your FROG), so you can often use a crystal you already have (e.g. BBO or KDP) and just tilt/rotate it a little bit to get the desired angle. (Be sure to pay attention to your desired polarization to know if you should be targeting type-I or type-II phase matching! Also, if you reuse a crystal, be aware that some crystals might be coated specifically for use with certain wavelengths, so be aware you might incur some extra losses by using unintended wavelengths.)
Once you've got the right crystal set at the desired angle, align both beams to the crystal using your favorite IR card/viewer in order to ensure spatial overlap. For the beampaths themselves, measure out as carefully as possible the propagation distance of your 800nm and 1040nm beams to get their lengths starting as close as possible to each other. (It would also be wise to use a photodiode and an oscilloscope to directly view the arrival time of your two different pulses, which should easily let you get them overlapped to well within a foot/nanosecond of each other if you've got a diode and scope with halfway decent bandwidth.) In one beam, incorporate a roof mirror on a delay stage so you can easily add and subtract propagation distance without changing the pointing (and therefore the spatial overlap); this will allow you the fine tuning needed to achieve temporal overlap at the fs level (imagine that, for a 200 fs pulse, your path length needs to be accurate to within a few tens of microns (200fs * c (0.3um/fs)) at the very most). To find the temporal overlap at first, adjust the micrometer of the delay stage to the end of its range, and then manually push the stage back and forth, quickly scanning multiple times across the full range while looking at the output for flashes of blue/violet (the sum frequency should generate ~450nm light). If you see the flash, eyeball the position of the stage where you get this temporal overlap, and adjust the micrometer to the position until you can fine tune the temporal overlap with your delay stage, and you're done!
(If you don't see the blue flashes, any number of things could've gone wrong: Do you still have spatial overlap? Do you have the right crystal and polarization? Are the path lengths still too mismatched? Are the intensities of your beams too low to generate the nonlinearity? Is there an underlying synchronization problem? Is the individual jitter in timing/pointing/path length/etc. too large to overcome?)
(All of this assumes that your two beams share a common source/are mutually coherent; if not, none of this will work under normal circumstances.)
(An SHG-FROG is usually designed to accept a single input of a pre-designed wavelength and pulse duration, so the purpose of the device is not specifically to look at overlap of two different beams.)
Good luck!
~Eric
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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 :)
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I believe what you have circled is, as Oualid pointed out, genomic DNA, not pcr product. When you start out with a large sample amount you often end up with a large quantity of starting material and, if not diluted, often shows up as a high MW smear on the gel.
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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.
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I would rule out Batrachochytrium dendrobatidis as temperatures in that brackish waterbody are probably way above the temperatures Bd can handle. But as I am already mentioning temperature: Salinity looks obvious but if that happened during a very hot summer temperatures could have suprassed the CTMax of the species, especially because adult and subadult frogs have lower thermal tolerances than tadpoles. Here is some lit:
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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.
Thank you!
Jiayong
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Dear Zhang professor,
About missing the protein coding gene, I have concluded in my previous paper in scientific report. I focus on this aspect in discussion. Please check my paper, The mitochondrial genome of booklouse,Liposcelis sculptilis(Psocoptera: Liposcelididae) and the evolutionary timescale of Liposcelis.
Thanks!
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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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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.
Paul
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looks like Pelobates cultripes (western spadefoot or Iberian spadefoot)
Best regards
Alfonso
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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.
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Dear Gregor ,
what is the estimated size of the tree frog?
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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.
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Great. Thanks for this addition!
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my name rouland, working in university of pakuan departmen of biology. now i studying about frogs. i have a big problem, as a beginner biggest problem was identified frogs sound. so i have a idea to analyze every type of frog sounds and colleting base on similarity of wave sound so i can have what kind of wave which same frogs. the hard one is searching software to analyze the sounds? so what kind of software suit for my research? thnks
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There are two excellent alternatives (free software):
For R users, the package "seewave": http://rug.mnhn.fr/seewave/
The Brazilian biologist Marcos Gridi Papp developed Sound Ruler: http://soundruler.sourceforge.net/main/
Both are excellent options.
Of course there are commercial software (most of them with free "light" versions)
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We had generated mitochondrial genome of frog using Ion Torrent PGM platform. The genome is missing two tRNAs namely t-His and t-Ser. what could be the reason for this? Can anybody suggest a method to find out the missing tRNAs.
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I have checked NCBI and usually tRNA-his and tRNA-ser come next to each other in the mitochondrial genomes of frogs. if you can find any closely related species from the NCBI organellar genome database (http://www.ncbi.nlm.nih.gov/genome/browse/?report=5) then you can take the sequences and run a blast against your contigs, you might possibly find both the sequences on the same contig.
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Herpethology
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Hello Adrian,
well, it depends how large the area is, that you want to research. And also if the species has individual caracteristics. We once did a quantitative research (unpublished) with Bufo calamita with photographs of the bellys, which have individual patterns.
If the ravine is a large area, I could imagine to make a sort of grid with statsitic relevance and you search detailled in specific fields of the grid.
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Specifically, I curious is anyone has experimented with different dye concentrations, staining duration's, and how long the larvae remained distinguishable from non-stained individuals. The literature has all sorts of nasty stories about dye concentrations that work on one species and causes 100% mortality in another, something I'd like to avoid if possible.
Cheers,
Paul
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I don't know if it is still usefull, but I use VIE for larvae, and they retain it after metamorphosis. Applied for several tropical species and worked just fine for over 5 years already.
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I am trying to investigate the effect of high calcium concentration on isolated frog's heart and I am wondering if it's advisable to modify the amount of calcium chloride salts of the Ringer's solution to investigate such behavior.
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