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This species is encountered in Kokrajhar town of Assam, India. It is thin walled bamboo. As per my knowledge it seems to be Schizostachyum sp. I request the peers to kindly help me identify the same.
we are using circular pitfall traps (regular yogurt cups) since many years, and unfortunately, sometimes we encounter mice and lizards as by-catch in our samples.
These animals fall in the pitfall traps, are not able to climb out of it and drown...
Researches (e.g. see link attached) and personal reflections resulted in the following possible easy-to-install preventive measures:
1) covering the pitfall traps with a thin, wide-meshed iron grid, so it is too thin to "grab and climb" for invertebrate, but slippery enough for insects and spiders
2) using funnels with a slippery surface, so small vertebrata can't enter, while insects can fall into it
I personally like more solution 1, but I wanted to ask you to share your experience :-)
We aim mostly on spiders, centipedes, millipedes, beetles and other surface insects.
Thank you and greetings,
I recently began to work with viviparous Neotropical skinks of the genus Mabuya. Specifically with females at different stages of gestation. My tutor and lab colleagues have studied them for a long time and a recurring comment is related to the difficulty of field sampling.
The standard method of catch is by hand, but these lizards are very quick moving through the litter, and their smooth-scales covered skin makes them difficult to hold. Also, in the most advanced stages of pregnancy, these lizards stops feeding so funnel traps will likely be less effective.
I have little experience in catching, and I am planning some field trips to obtain some specimens (especially to learn about the field work). I would like to try different catch methods, hoping to make it easier to obtain research material.
I will be very grateful for any suggestions or advice you can provide.
I've been googling around for quite some time trying to find papers pertaining to pilot studies on ecological/wildlife/biodiversity assessment research, but with little success. Most of them pertain to medical research only. Shall be grateful.
I'm in the process of buying new equipment to record the vocalizations of wild mammals, and have used Marantz's solid state recorders with great success in the past. However, it seems that their PMD range of recorders are not easily available anymore. I've looked around and the Tascam DR-1 portable solid state recorder seems like a good option, but I have no experience with that product.
I'd appreciate any advice on the best, portable recorders to be used in the field. Nothing too fancy (I don't expect ultrasonic vocalizations, for example), but I'd love to hear from you about the pro's and cons of different recording tools that you've used. I would need the recordings to be of good enough quality to analyze properly in programs like AviSoft.
I have a long-term slow loris research project. We have been using the very expensive Biotrack Sika and six element yagi. Unfortunately, despite working in a non-arduous anthropogenic landscape and keeping equipment clean and dry (although it rains frequently), the break downs have become financially non-viable and we are looking for a more commercial radio receiver that is more affordable. I used Icom in the past but they seem largely discontinued. Does anyone have any experience with Uniden? It is to track small animals within a few km at a frequency of about 150. Key is cost efficiency as our project budget is very low!
Nowadays, CRISPR take a lot of attention as new revolution in biology, on another hand we have some hot topics such as antibiotic resistant, nanobiotechnology (hydrogels and drug delivery) and imaging development technologies etc.
So tell me your opinion about the next revolution in biology? (considering of all fields of biology)
And are we ready for that?
I am looking into the assemblages of Silphidae between different habitats and within each habitat as well.
I'm currently looking for modern field techniques in catching wrasses and would like suggestions or tips, as some papers that I have read can be quite vague with their capture process (i.e., tools used, technique, processing). One paper that caught my attention was by Worachananant et al. (2016), but their capture process was also vague despite the wide range of tools used in the study. Any suggestions or recommended literature are greatly appreciated. Thanks in advance!
I am looking to track within breeding season movements of a 25 g bird. The species is not philopatric, so trackers that require recapture are not an option. Trackers must be under 0.75 g per USGS regulations. I'm thinking I am going to be limited to radio transmitters but if you know of other options, I would be very interested. Do you have a favorite radio transmitter brand? Cost is a major consideration. Thank you!
We will photograph feedings of birds (Great tits and also Barn owls) by nest boxes with camera traps to know who, what and when they bring a prey in. A PIR sensor is not working well enough during tests (Bushnell Agressors and Reconyx HC500 are tested). They missed to much events of the well isolated birds. The birds has pit tags, but that system failed +-10%. And that could be the clue of the research...
By that I was thinking to connect a simple and less energy consuming IR beam on a trail cam like this one: http://eltima.de/jokie-179/product/jokie2.html?details=features-180
I can also connect DSLR's on the IR beam, but that is most times not so easy to install and to much disturbance for barn owls.
I know the Reconyx PC-series can work with an external sensor as trigger with a special cable, but that price is (a bit) far above budget. (http://images.reconyx.com/file/Professional%20Settings%20User%20Guide.pdf)
Now I tried to connect a IR port on a Bushnell E2. But how can I switch of the PIR sensor? And where to solder the port for IR beam on? An extra plug is easy to install, but the input of the PIR is horrible to find...
Are there maybe some others who have experience with connecting trail cams on a IR barrier? Or maybe are there other brands that have also this option?
I like to come in contact with them and like to share experiences in this.
I am a hardcore experimental biologist. In particular, I am well-versed in molecular and cell Biology techniques. However, the area of Bioinformatics and Computational Biology have revolutionised the field of Biology.
Can anyone suggest basic hands-on-trainings/courses of international standard where we can get a good exposure on the following for beginners?
2. Systems Biology
3. Synthetic Biology
We will start soon a 1 year (at least) camera trap project, where the main target will be mesocarnivores. Unfortunately, some animals like mongooses don´t have spots or truly individual marks and many pictures might be blurred, especially at night.
The idea is to use tags (e.g. hear tags, hair cuts, etc.) that will increase the chance of individual identification, even in worst conditions!
At the same time, we will be conducting box trapping sessions to capture several individuals.
Can someone suggest any easy and durable method to mark animals with the lowest impact possible?
Thanks a lot in advance
We want to monitor nest and ambient temperature on the tundra, but have just been informed that they have again delayed our order and it won't arrive until AFTER our field season.
I'm planning a transect study using plasticine caterpillars and would like to take pictures in the field to document signs of predation, prior to transporting back to the lab.
Are pan traps more effective if coated with ultraviolet reflective paint? If so, what brands do people use or recommend? We have found these:
Any advice or thoughts welcome.
We would like to use trail cameras to record the behaviour of small carnivores at night. No need to say that we have a limited budget. The latest entry-level Bushnell model (Essential E2) does not allow to take video-clips longer than 15 s at night with 3 s trigger interval, whereas the Essential allowed up to 1 min with 1 s interval (this is what we call progress...). We are therefore looking for cheap but reliable alternatives. I found good reviews on the Bestguarder DTC-880V/SG009:
N.B. Apparently this model (or a slightly modified version) is also marketed under other brand names such as TEC.BEAN (and maybe XIKEZAN).
I also read some positive feedbacks from buyers on Amazon but noticed that a few people mentioned that the nocturnal video-clips and pics were not necessarily of good quality (i.e. "grainy").
We were wondering whether any of you have used this trail camera and if so, what is your opinion on/assessment of the quality of nocturnal video-clips?
Also, do you have any feedback on the reliability of this brand(s)/model(s)?
Thanks in advance for your precious assistance.
At times I catch 2000+ mosquitoes in a CO2 baited trap. I would like to get an idea of the population without identifying and counting every mosquito. What sampling methods are appropriate? I have been quartering the pile and multiply the results by 4, but is there a better way? And is there a sample size (sample size formula) I can use to ensure I have a large enough sample to accurately represent the trap population?
I am interested in how to classify activity measurements from GPS collars for kangaroos.
Clip cages are used in laboratories for biology determination of insects. Can anyone please tell me the best way to find out the biology of small insects like wheat aphids in the field condition?
Seeking alternatives to a Cessna 172 aircraft capable of being fitted with VHF antennae to complete radio-tracking surveys in remote terrain. Ideally I am looking for examples of successful use of twin-engine, turbine aircraft.
I'm using GPS coordinates for a particular species and need to enter this data into a database. Unfortunately, some of the researchers collecting the data did not record the GPS error. I am wondering if it makes sense to estimate error in those cases for entry into a database or if I should leave the GPS error field blank? Are there any publications or sources touching on this issue that anyone knows about?
I'm working on Flying-foxes monitoring and I want to know how I can built my protocol of fly-out count.
I have seen that determination of age by teeth analysis is possible ...
A student in a course with me has data on noxiously abundant flies
near a mink farm. Fly abundance was obtained monthly
at 8 equally spaced trap locations along 3 transects,
each within a narrow strip with a different treatment
(control, manure, manure + offal).
Clearly treatment and experimental unit (strip) are confounded.
It occurred to me that useful information might be obtained by
blocking at right angles to the strips, depending on the spatial
autocorrelation. The strip separation is the same as the trap
separation along a strip.
The within strip acf crosses from positive to zero at a lag of
around 3. Which suggests blocking the trap data at either end
of the parallel strips (two blocks, conservative).
Or perhaps even 3 blocks (either end and middle).
Does anyone have a reference on acf to post-stratify data?
Several google searches, nothing relevant
I plan on using fecal samples to identify individuals within the study area. Within each grid cell I was planning on rather opportunistically hiking in good habitat, along animal paths, etc. As long as the effort is even across grid cells I am hoping that capture/recapture models will be effective? Concerning grid size, I know that probability of recapture is important, so is basing the cell size on the species' typical home range size appropriate? Thanks.
Edit/Update: Additional Info -
I am planning on conducting this research on giant pandas on a pretty small scale (focusing on the borders of Wolong Nature Reserve, perhaps with a 3km buffer on either side). Other research (telemetry studies) has resulted in an average home range size of about 7 square km in this area, using minimum convex polygon (MCP) methods. There is also pretty high spatial overlap in this species. So far I have been considering grid cells of 2.25 square km to ensure effective coverage (the terrain is extremely difficult), which would also result in a grid 2 cells deep on each side of the reserve, keeping with the 3km buffer.
On the other hand, perhaps this increases the chances of unneeded recaptures and the cell size could be increased to around 7 square km? This would also save greatly on time, which is a consideration. I am not sure which is more effective and what the trade-offs are when considering modelling/statistical analysis of the data later.
I'm plan a new study that will focus on Eliomys melanurus community. I'm aware for two sample methods. the first one is to use Sherman traps and the second is to use IR camera. Do you have other ideas how to sampling population size of this extremely rare species?
I'm planning an undergraduate student field project with mark recapture on wild mice and voles, and thinking about marking methods. As individuals only need to be marked for a week I don't want to use tags. I know fur clipping is often used, but what about temporary marks with hair dye or food dye? As this is for a student project with minimal previous experience handling small mammals I'm a bit cautious about using clipping. If you have used food dye / hair dye or know of something published which did, which brand was used (or a link to the reference)?
The species are:
1)Muscovy duck (Goose)
(Cairina moschata) ***
2)Black Bellied Whistling Duck
(Dendrocygna autumnalis) ***
3)White faced Whistling Duck
(Dendrocygna viduata) ***
4) Fulvous Whistling Duck
(Dendrocygna bicolor ) ***
5) Snow Goose
(Chen caerulescens )
6) Comb Duck
(Sarkidiornis melanotos )
7) American Widgeon
(Anas americana )
(Anas crecca carolinensis)
9) Mallard Duck
(Anas platyrhynchos )
10) Northern Pintail
(Anas acuta )
11) White Cheeked Pintail Duck
(Anas bahamensis )
12)Blue Winged Teal
(Anas discors )
13) Southern Pochard
(Netta erythrophthalma )
14) Ring Necked Duck
(Aythya collaris )
(Aythya affinis )
16) Masked Duck
17) The Northern Shoveler
I am looking for a small mobile unit that could be used to perform DNA extraction, PCR and gel or capillary electrophoresis in the field, i.e. Not in laboratory conditions. The purpose of the kit is to perform species or individual ID. Samples would range from muscle to hair and PCR would use mtDNA or STR primers.
Hi, i am working with a new a rare specie of primate in Colombia named Callicebus caquetensis recently described.
Its is located in a high fragmented area and there are estimations of poblational size which barely reach 500 viable individuals.
I must stand a methodology for the study of its poblational density and i am not sure about the usual line transect method.
THanks in advance.
I just want to know about any marking technique for water monitors to identify an individual from a distance.
I am looking for researchers which can collaborate in a paper. What I need (attached in the jpg) is plastral measurements (PL, GuL, HumL, PecL, AbdL, FemL, and AnL) of many turtles of any species, with some recaptures along time. This measurements must be taken using callipers, not measurements from photographs.
If you have them and you can be interested in the collaboration, please, contact me as soon as possible.
Is it possible to publish a new species using only one specimen?
We are exploring potential tagging of large mammals in Namibia using small, low-cost, 'expendable' devices and we would be interested to know of experiences, publications, reports etc. that show success (so we can perhaps adapt these to the specifics of the locality) or even failures (so we don't repeat them). In particular are there any methods which have proven effective in attaching these for periods of time without needing to capture the animal.
I am about to start a field season studying cinnamon teal population dynamics and we will be taking basic vegetation measurements at each nest we find. I want to create a set of randomized points throughout the wildlife refuge in areas of different habitat, but I do not know how to determine how many points to create. Should it be equal to the number of nests I find during each field season or should I use the same number of systematic random points each year? Should they be in the same spots every year or should I re-randomize them in future field seasons? It is a 14,000 acre wildlife refuge with impoundments, some of which will have water this year and some of which will not. The teal mainly congregate in a couple of the main wet impoundments, which will therefore have very different vegetation characteristics than the dry. Thank you for any help!
We would like to observe the development of single insects on plants in the field. The insects usually settle on the bottom side of the leaf. Leaves are quite large (about palm-size) and more or less waved with some strong veins. The experiment will run about one month during summer, and therefore possible physical damage by the cages, rains and wind are additional challenges (the standard setup with metal clips and sticks can be problematic, since it is unstable and susceptible to wind especially with upper leaves which are about 30-50 cm above ground).
Now we are unsure what type of caging to use and a few ideas we had so far are:
- Light-weight clip-cages without sticks (still fragile setup, mounting only possible at leaf edges in larger leaves bec of clips)
- Gauze nets (covers whole leaf; prone to getting wet during rain, to heavy then or insects might drown)
- Perforated plastic bags (too high humidity and temperatures within bag)
Do you have any better setup for such caging or another cage type or an idea how to mount a clip cage safely on such leaves in the field?
Thanks for any input.
Im writing a report about the Superb Lyrebird vocalisations, however am struggling to find some decent papers explaining the 'hows', and the 'whys', looking at their physical form and function, and how it facilitates their mimicking behaviour, and for what behavioural reason they do this? Any help would be greatly appreciated! Thank you
I am investigating the possibility of setting up den cameras for my bat-eared foxes, which would allow me to record their social interactions in the den itself. These are natural dens/ hollows, and I want to do this in the least invasive way possible. Ordinary camera traps work fine for comings and goings outside the den, but I'm curious about the interior...
What equipment and set-up could you recommend?
A camera trap data study was developed, installing cameras for 21 days, but we completed all camera station points in a long period (24 months). Individuals of target species were identified.
This long period may include the inclusion or elimination of individuals of the study area, violating the assumption of closed population for capture-recapture methodology. There is another methodology to estimate abundance and density?
I have more than 300 genotypes and I want to measure CC in the field on these genotypes which are many. I can I do this in two subsequent days.
I am working on several projects with scorpions and Madagascar hissing cockroaches. If the camera also worked for shooting into a fish tank, that would be great too (as I also work on electric fish, and I have had issues in the past with the IR light source on Sony Handycams reflecting back off the glass of fish tanks.)
We are trying to estimate the occupancy of snow leopards in Rolwaling valley, Nepal. How much area should we cover for this study? Do we need to cover atleast area equivalent to approximate home range or should we take more than that, or can we take less than that?
I need to catch some Asian Openbill Storks to track their movement. Does anyone know how to catch them. This birds like feeding in shallow water areas and roosting on trees.
Color-banding related question: I'm studying Hispaniolan Woodpeckers in mid-elevation (~550-700 m) tropical, fairly open habitat, and I have had some issues with Darvic color bands (specifically, a large number of my Darvic bands inexplicably opened by themselves in my banding kit- a plastic tackle box). I'm considering incorporating Acetal color-bands (http://www.avinet.com/avi_order.taf?_function=view&ct_id=101) to supplement the striped celluloid bands I'm already using.
Has anyone run into major issues using Acetal bands in tropical climates? Do they have to be sealed shut? Is there a method other than application of heat that can be used to seal the bands?
I am looking for a GPS unit helpful in field biology. A handheld unit, affordable and easy to carry.
I recently work on the diet analyses for woodland migrants. I found couple of spring-like and brown or pinkish structures from bird fecal samples. It feels like rock or iron when I try to break it. Does anybody know what they are? Thanks in advance!
I estimated survival in Mark. All survival probabilities was estimated per month. To get annual survival I just raise into power of 12 (12 month). but how to obtain SE for this annual rates? Probably not by the same way, as SE for annual survival should be greater then for monthly rates, I suppose.
Field conditions may scientifically never be truly standardized because of perception constraints, logistic constraints or continuous dynamics in nature. The need to standardize experimental environments may depend on the scientific problems addressed or model systems considered. As an example, between-population variation in timing of reproduction in blue tits (a small European nest-box breeding bird) is sometimes larger in scientifically more standardized aviary conditions than in scientifically less standardized free-ranging conditions. Responses of blue tits to ‘artificial’ versus ‘semi-artificial’ conditions may result from scientifically ‘uncontrolled’ organism-environment interactions. Not all study populations may be preadapted to captivity and scientists most often don’t know this when they initiate laboratory experiments. Veterinarians may advise to use sterilized test cages minimizing infection. The practical problem is that blue tits will not breed in these highly artificial environments or that some populations cope better with captivity than other populations. Outdoor aviaries with ‘semi-controlled’ natural vegetation might be more appropriate for captive wildlife breeders, simply because they better simulate wildlife conditions. How can a veterinarian or members of ethical commissions that never worked with blue tits provide constructive advice about both scientifically and ethically acceptable experiments? Should ‘environmental sterility’ or the ‘mental state of captives’ been used to make decisions about how to conduct experiments? Do sanitary recommendations proposed by people not familiar with model species complicate execution of scientifically acceptable experiments that take background knowledge from model species into account? Sanitary conditions are never truly controlled in wildlife conditions anyway.
Currently, my colleagues and I are in the first stage of preparing a article about gender determination of Long-eared Owls (Asio otus) based on plumage coloration and biometric measurements supported of DNA analyzes. For now, we found several close related papers which we have been able to found on the Internet databases (Scopus etc). Thank you in advance for your help.
As part of my MSc project I am planning on collecting hair from wild foxes (Vulpes vulpes), but I am trying to avoid using barbed wire for ethical reasons.
We are developing a framework for monitoring free roaming dog populations in the cities of Tierra del Fuego. The objective is to monitor the mid and long term success of the policies to reduce free roaming dogs in the streets of the cities, like neuter and responsible pet ownership. Cities are expanding in surface and adding new neigborhoods over the course of the years.
For that reason, and to avoid restrictions of design-based surveys, we are considering the use of Density Surface Models, also to be able to explain changes in dog presence by covariates.
Then, we should be able to tell the towns to follow a series of tracks (streets) that need to be surveyed over the years. Adding covariates, we should be able to develop a DSM to assess abundance based on these covariates.
Then, here comest the two questions:
ONE: The issue that comes to my mind and I want to share with you hast to do with modelling all over the space of the city, when free roaming dogs are distributed only along the streets. The, the streets are like a mesh of something like 21m wide, separated by the about 800 blocks of around 90m side.
Potential solutions I was thinkg were:
1. Model the data as if they were collected and build a prediction grid where blocks can´t hold dogs (as if they are "0") meanwhile streets has a value of "1". Then I wonder how much the smoothing all over the surface will affect the predicted results.
2. Model the data, but adding "dummy" transects located over some of the blocks, just to tell the model that inside the blocks there are no dogs... However the modelling may be affected by the spatial spread of these dummy transects.....
3. Use something like the soap film smoother but I can´t figure out how we can handle such amount of blocks...
About detectability of dogs. When travelling along the streets we need to record dogs that can walk freely at the sidewalks or at the streets. Streets has parked cars along the sidewalks, and many houses do not have fence, so dogs can sleep inside a parcel but leave the parcel boundarie as soon as they want to bark or whatever. This impose visibility issues that affect detectability.
To reduce that I was thinking in make dogs more visible:
1. Survey the streets going behind garbage collectors, as these guys atract dogs either by their behavior (running along the streets) or leaving garbage behind ocasionally).
2. Broadcasting dog barks from the car meanwhile we go travelling. I know that this affect the results of a survey, but if we want to see dogs in the streets I understand that we need to make them out.
Any idea or previous experience in that is welcome. Many thanks
Asociacion Armonia is protecting a 11,000 acre tropical savanna in the Beni, Bolivia. We are going to use a savanna patch-burn management technique, but we are not clear of the potential climax level of older successional savannas. We are thinking the best measurement could be arthropod diversity and abundance. We are interested in an comparison arthropod studies in tropical savannas.
I'll be observing grey reef sharks (3) within an aquarium and trying to find a way to randomly allocate time slots for each shark for a focal sample. Observations will take place over several months for 2/3 days per week. Each observation day is then split into 3 time periods AM, NOON, PM (1 slot for each shark). This leads me onto...
Is there a method which takes all the observation days into consideration so that at the end- each shark would equally cover ever day of the week and all three time periods for each day?
Thanks for reading, I hope I've explained it ok?
What is the sex ratio in carabids?
As you all know pitfall trapping gives a bias estimate of abundance due to various reasons. Therefore higher abundance of males or females in the trap does not have to relate to their abundance. Do you know sex ratio of ground beetle populations obtained with more reliable methods than Barbet traps? Do you have some unpublished data on this topic?
With kind regards
We are having data on Bluethroats from its breeding and wintering grounds and would like to study their spatio-temporal migratory connectivity.
I am involved in analysing a capture-mark-recapture survey dataset that covers in excess of 30 years. I am trying to establish whether this is in fact the longest running survey of its type in existence. I have yet to find another survey that has been running continuously for such a long period of time. I though this would be the place to ask!
For example, if a dead tissue (e.g. metazoan) was left in an actively fermenting medium (e.g. beer), how would this affect the quality of the DNA one could extract or amplify from this tissue?
Personal experiences or papers about this?
Long-tailed macaques are primarily arboreal (and are quite elusive in our site). Tracking them in the wild to observe defecation events requires A LOT of input. I'm fully aware of the huge difference between captive and wild populations but would it still be acceptable to do so?
What have you found that is an essential app for field work? Voice recorder, stop watch, clicker and the like are obvious ones but are there anything you have found that makes a big difference to your research?
This may be an odd question for ResearchGate, but although I can find much in the way of products I can find little in the way of reviews and opinions. I am looking for a good, reliable, accurate hygrometer and thermometer (combo?) to use in the field (Spanish countryside) to record the humidity and air temperature at night. Any recommendations or advice on models? We don't want to go too cheap and risk bad data but we also do not need top of the line as it is supplementary information. Anyone have any experience?
Thank you in advance.
I am getting ready for this summer's field season and looking to buy a good sturdy GPS unit with reasonable accuracy including elevation. Any recommendations?