Questions related to Small Mammals
Is the energy conversion efficiency of herbivores better than carnivores and why birds and small mammals have low net production efficiency?
Why does a small mammal lose heat faster than a larger mammal and energy is transferred when a carnivore eats an herbivore?
In regards to biodiversity (general) and monitoring.
If geographical regions are significant, feel free to mention.
I am working on a small mammal detection-non detection data using dynamic occupancy models. The parameters I have are probabilities of occupancy, detection, colonization and extinction. The same dataset has been used for estimating survival, recruitment and other demographic parameters of species using Capture Mark Recapture models which show a positive influence of rainfall on survival probability. My analysis also show positive influence of rainfall on colonization of the small mammal community which makes a lot of sense. Now I want to connect my results to the previous results but am struggling to find a way or a study that links survival and colonization that I can refer to? It could be confusing because survival analysis is done at individual level and colonization analysis is done at species level. Probably survival would benefit colonization but I'd require an empirical notation or a study that has done both and shows that indeed these both are directly related? Thank you!
I recently deployed some insect pitfall traps and in several of them I unfortunately caught mice. How can I avoid this? I have seen suggestions of using shallow traps or small ladders, what is the best way to obtain quality insect samples and minimize risk to small mammals?
Hello all, I am analyzing body mass between small mammals and testing if it varies between sexes, reproductive status (breeding/non-breeding), and season (wet/dry). Would it be alright to have one variable called reproductive class as a factor with 4 variables: reproductive male, reproductive female, non-reproductive male, non-reproductive female. Is it possible to then use this in a linear model like body mass ~ reproductive class * season ?
Does sex and reproductive status have to be separate terms?
Additionally if I have significant differences between males and females, can I group the two sexes together to see if body mass overall changes with season?
Say a researcher was interested in determining the number of adults vs. juveniles of species X trapped during a small mammal survey. Does there exist a relatively reliable way of doing this based on standard field measurements?
Let’s say a total of 200 individuals of species X were sampled, and the following data recorded: sex, total length, tail length, hind foot length, ear length, and weight. For the sake of this question imagine no additional data is available (e.g. additional observations recorded in the field, access to collected specimen material, etc.).
- Is there a way to ascertain a point or “threshold” from a range of data based on the distribution of values to distinguish between juvenile and adult individuals with a meaningful degree of accuracy? For example, male species X with weight > 142 g = adults; < 142 g = juveniles.
- If yes, which of these measurements would be most indicative? Or perhaps a combination/ratio of more than one (e.g. ratio of hind foot length to ear height > 1 = adult, etc.)?
Thanks, and looking forward to the feedback.
I am interested in products of less than 1 g in weight, for bats of about 10 grams. Thank you for your recomendations in advance.
(Im aware that telemetric transimtors of such weight exist, yet am currently looking into other options)
I have been looking at weight values for rodents in the family muridae, specifically subfamilies: gerbillinae and deomyinae. I found some considerable discrepancies in the values for the same species from different references. Generally, I get similar values from sources concerned with African mammals (Mammals of Africa, Kingdon et al, 2013; Mammals of Sub-Saharan Africa, Monadjem et al, 2015; The Complete Book of the Southern African Mammals, Mills and Hes, 1997; The Contemporary Land Mammals of Egypt, Osborn and Helmy, 1982). The values I get from other sources, namely PanTheria, AnAge and Alhajeri et al (2015) are mostly similar amongst themselves but can be very different from those reported in the first (“African”) set of references.
The similarity within set cannot be solely explained as repeated citations from the same old reference; so I was wondering if it can be explained by biogeographic trends within widely distributed species. In other words, the set of references concerned with Africa is reporting species values from African populations only; while the other references report values from the world-wide distribution of the species. The observation that species with African and extra-African populations have wider ranges of values reported in PanTheria, AnAge and al-Hajeri compared to those in “African” sources for the species is consistent with this hypothesis. Furthermore, whenever a species is endemic to Africa, the two sets of references seem to largely agree.
Could somebody please corroborate/debunk this idea of mine, or suggest other explanations for these puzzling discrepancies?
I am interested in collecting brain size (endocranial volume) data for several modern species of mouse-sized rodents. However, I am struggling to figure out the best methodological way to obtain this data.
The gold standard for measuring endocranial volume would probably be to take CT scans of the specimens and measure endocranial volume off of the virtual endocasts. However this would be prohibitely expensive as it would be nessary to scan hundreds of skulls to obtain decent sample sizes (N > 8-10) for each species. Sufficient sample sizes exist but getting the data from them is the hard part. Only getting data from one or two individuals per species would not be rigorous enough to produce trustworthy results. Even if I got a grant to do the scanning the specimens I am interested in are housed in distant institutions that I can plan collections visits to but are too far away to visit regularly. I cannot drag a CT scanner to these institutions to get the neceasary data nor take out loans for hundreds of specimens.
The other major way I know that people have measured brain size is by filling up the endocranial cavity with glass beads or lead shot and estimating the volume from the density. However, at smaller and smaller body sizes lead shot or other spherical globules are going to increasingly poorly correlate with brain size, for the simple reason that you can pack fewer granules inside a spherical chamber. At large sizes the relative error is negligable, but at small sizes spherical pellets will poorly model the volume of the cavity and accuracy will become increasingly worse. I've even seen this in the literature with some small rodents, in which reported cranial volumes measured with lead shot seem suspiciously large or small compared to other studies on the same species, with endocranial volumes sometimes differing as much ad 50-70%. Additionally many natural history museum will not let you bring pellets like lead shot into the collections for fear that it could hide pests or get scattered and cause problems (something like this has happened to me a couple of times: I used sand bags to prop up larger modern mammal skulls for photograph and nearly got thrown out of the museum collection for bringing outside sand in).
Given this, what would be the best way to measure brain size in museum collections of mouse-sized rodents?
Happy new year, everyone!
I am looking for publications dealing with the influence of pesticides that are used to protect crops on small mammals (rodents/ shrews). My interest is focused on lethal and sublethal effects, bioaccumulation, and biodiversity,.
I am grateful for any hints!
I am doing a masters in Nature Management and am currently doing my thesis, which is on wild boar management in Denmark and the errection of the wild boar fence at the Danish/German border.
I want to investigate the following:
- Does the wild boar fence together with the shooting campaign in southern Denmark function as intended in keeping out wild boar from the country (investigated with camera trap data positioned at 8 of the 20 large openings in the fence and with individual counts from remaining populations).
- What detterents, if any, will keep wild boar from using the openings on the fence (tested on wild populations in Germany/Holland)
- What other animals are using the openings in the fence (there are 680 - 20x20cm openings positioned every 100m for small mammals) and how can we utilise this information in terms of management?
- Are there any negative effects of the fence construction on non-target species, what are the consequences, and how can these be mitigated? (Investigated with camera trap data - what animals are we not seeing, and literature research on home ranges, migration patterns and genetic variation in potentially affected species).
The thesis is of 9 months duration with a limited budget and I started 1 month ago.
I seek information on the following:
- What methods can be used other than camera trap data positioned at fence openings to investigate the impacts of fences on native species? (given the time and financial restrictions). I am already including how many animals are recorded as entangled.
- How can I set up a field experiment to test detterents that best imitates the real life situation? I am not sure a setup with food, detterents and controls as previously done best mimics the situation, because fence openings with a frightening scent, sound or smell might influence the boar's behaviour differently (because there is no instant reward of food on the other side).
- How can I investigate whether wild boar swim to Denmark via Flensborg Fjord? The area is too big to monitor with cameras or look for tracks and it is very difficult to get permission to fly drones because of the proximity to the German border.
I am also very open to comments or ideas about other aspects of my project (positive and negative). Note however, that my research is not an investigation of whether the fence was a good or bad idea or whether it will work in keeping away African Swine Flu, so I would appreciate not starting a debate of that nature.
I am interested to see what the damage on smaller animal long bones look like when attempting to extract marrow from long bones using percussive techniques. Most experiments and archaeological interpretations involve individuals using their teeth to open the cavities, with the assumption that stone will crush the small bones. It is not an optimal method for extracting marrow from smaller mammals but I'm interested to see if there are any archaeological relevance. Has anyone tried this before? Or is this just an assumption in the literature?
I am thinking about using JSDM for small mammals communities in Europe following HMSC package. When performing these kind of analyses for birds, I obtained the phylogenetic correlation matrix from http://birdtree.org/. However, in http://vertlife.org/ it seems that mammal phylogenetic subsets are not available for now. Is there any other source where I can find this kind of information?
Can someone please help me with this question? Why is bacteria isolate obtained from small mammals sampled in human induced habitats likely to be more resistant to antibiotic drugs than isolates obtained from small mammals sampled in their natural habitats?
For example, small mammals can trigger a camera with an infrared sensor because these are warm blooded, but perhaps not so for a reptile that is cold blooded. Resolution could be a problem for small animals. Any good model for general purpose?
I performed comparative histological, histochemical, and morphometric analysis on the tongue of two phylogenetically different small mammals with different feeding habitat. The reviewer gives me a comment " The purpose of what to clarify by comparison of two kinds of animals is not clear. It ought to focus only on the main aims and intentions of the work and why this work is regarded important".
Could you please, give me suggestions for a convinced answer?
All the best wishes
1. social organization,
2. varied diet (omnivorous diet based on fruits, acorns, roots, tubers, bulbs, rhizomes, insects and small mammals AND increase in the availability of cultivated plants,, ...),
3. reproductive system (high fertility, early first reproductive age, large size, high potential life expectancy,gestation time which is very short ),
4. low hunting and catching pressure,
5. extinction and regression of its natural predators especially the panther and the jackal,
Which statistical test is appropriate to test the following hypothesis?
H01: There is no difference in soil compaction across all locations (e.g., Location A, B, C, D, and E). No zeros.
H02: There is no difference in the total number of small mammal burrows across all locations (e.g., Location A, B, C, D, and E). Many zeros.
Each location consists of 7 plots that are located apart from each others and each plot is divided into quads. burrows are counted within each quads. therefore, enormous number of quads have zero counts. Sample size are not equivalent and data are not normally distributed.
Since the variable is independent, I am thinking of using multiple Mann-Whitney U test to test my hypothesis:
burrows in A versus burrows in B
burrows in A versus burrows in C, etc.
I will really appreciate your help
I collected data on small terrestrial mammals in two protected areas (PAs). In each PA I used Sherman's live trap to trap small mammals from three different land-use types. In each land-use type 20 experimental plots were sampled once with 40 traps. I also collected some environmental and habitat data at every other trap station.
In occupancy analysis what is the number of sites and occasions?
I want to measure activity, exploration, boldness, aggresivenes and sociality of small mammals (rodents and/or small opossums) in the field.
For activity, exploration and boldness, I find the protocol of Dammhahn (2012), for grey mouse lemurs, feasible and trustable (it lasts about 20 minutes per individual, and she found a high individual repeatability). Any better suggestions for rodents or opossums?
I read that aggressiveness can be measured during capture/manipulation, how it is done? it is repeatability considered?
About sociality, any recommended protocol?
Thanks a lot!
If somebody is using or used trail cameras in small mammal research/monitoring can he/she share some experience with me. I would like to know how you position traps on-field and how they react on small mammal movement. If you have any advice what not to do also please share.
I am looking for the best assay and biomarkers to use when measuring oxidative stress after chronic contaminate stress. Small mammals (shrews/field mice) will be collected from a contaminated area and assessed. I am just wandering what has worked best for everyone and what kind of assay you are using (i.e. western blot etc)
We have been discussing within our group the question of what to do with individuals of invasive species caught during projects, which for us are in preserves and national parks. I am also curious if you think it makes a difference if it is in a preserve or not.
I am looking to use metabarcoding to identify prey species in the feces of black bears and wolves, and I am searching for a lab to send the samples to carry the analysis.
We are interested in analyzing the feces of desert locust adults that have fed on grass in the field. A cup of feces will allow us to analyze the feces in relation to locust behavior. We want to know the name of the grass the locusts had fed in the field to produce the feces.
I am studyind small mammls in Quirimbas National Park, North of Mozambique, so i need help in ID of specimes that i am collecting. I have one Field Guide, but does not help me so much. i am looking for ID key to rodents in general.
I am investigating the ecological stoichiometry of Snowshoe Hares (L. americanus) and I need to estimate the age of wild-caught hares before chemical analyses. I am looking for a relatively simple, cost efficient method that uses a small sample of each hare to produce an age estimate.
My literature search yielded only two papers, one describing a method to count growth lines in mandibular bone cross sections of European hares (Iason, 1988), while the other provides a couple of allometric equations linking together hind foot length or body weight to age (Keith et al, 1968).
However, as both these sources are quite old, I was hoping someone could point me to more recently-developed techniques, or provide evidence of recent use/testing of the two methods mentioned above.
Thank you all in advance for your help!
I am extracting DNA from small mammals faecal samples with the Qiagen Mini Kit Stool and despite I have loads of buffers and tubes I am running out of spin columns and InibiteX tablets. On the manufacturer website of course it is not possible to buy these items separately. The kits are very expensive!
Does anyone know if I can use equivalent products (e.g. leftovers from other kits)? Or does anyone has any Stool Mini Kit leftovers is willing to share (I would pay for the shipping fee)?
I am a bit hung up on how to organize my data to analyze them. I have large sets of data that do not necessarily correlate. What I am interested in doing specifically is to analyze the distributions of my data and determine whether there are any associations. For instance, I have many samples of vegetation variables from a plot that I have also collected small mammal abundance data in. The sample size for my small mammal data in each plot is 8 throughout the year of sampling. However, I have 48 samples for each vegetation variables in the same plot. I would like to see if there is some relationship between the distribution of abundance in small mammals and the distribution in any of various vegetation variables (e.g. litter height, basal cover of forbs, basal cover of grass). I do not know how to tabulate the data for analysis in software due to the differences in sample sizes. I also have environmental data that could serve as another variable to analyze. Is there some sort of statistical method that tests for associations between distributions of independent variable data with that of a dependent variable such as abundance?
I have collected data for each season (e.g. Fall, Winter, etc.) for a full year of mark-recapture data on small mammals in Northcentral Texas at a tallgrass prairie. I trap in 3-night intervals and have two samples per replicate, with four site replicates in total from a 1,400-acre preserve. I collected quadrat samples randomly within the constraints of 60x60 m trapping plots, and measured percent coverage of vegetative categores like litter, grasses, forbs, etc. I also measured coverage at various heights ranging from ground level to 1-m. I would like to compile all of my data and determine which variables most strongly correlate and test for significance. I also have point count data for breeding birds during the summer along with the same vegetative variables and line-intercept data that I would like to add to the list of correlates to test for.
I am having trouble determining the best approach to statistically analyze these data. Any pointers?
The attached photo show some tracks from a small mammal from an Early Eocene site in western Washington, USA. The radiometric age is 54 Ma. The setting is a low elevation riverbank in a semitropical environment. The footprints consist of 5-toed tracks with narrow digits terminating in slender sharp claws.The stride/gait indications suggest a narrow-bodied animal with fairly long legs, about the size of a modern possum or racoon. Tracks like this have not previously been reported. I welcoming speculations as to a possible track maker.
Can anyone recommend proximity loggers for small mammals (up to 3 Kg) to use to study social networks in areas where there is no satteline reception?
I'm trying to fit a dispersal kernel using CMR data from small mammals, to parameterize a Cellular Automaton (CA) designed to model range shifts driven by environmental change. This CA have annual steps, so I need a dispersal kernel standarized to 1 year dispersal events. Does anybody knows how to fit such dispersal kernel using data of dispersal events experienced in just a couple days (no more than one month)?
if someone can share a R script, it would be great,
Thank you so much!
We are preparing a conference on forests and unique nature of Costa Rica and would like to illustrate the diversity of plants and animals. Have a look at the movies in the attachment. It shows probably a female of the Common Gray Four-Eyed Opossum (Philander opossum, Didelphidae). The movies were taken in the province of Cartago at ca. 700 m a.s.l.
Any information related to experiences of using keys for identification of European small mammals? I am considering purchasing one of the two:
- Collins "Mammals of Britain and Europe"
- Aulagnier "Mammals of Europe, North Africa and the Middle East"
I need to code the hairy-footedness of a large number of rodents for a statistical analysis. In most cases it is simple enough to code hairy-footed species as 1 and naked footed ones as 0; however, the following five species are described as intermediate in hairy footedness in different references. Furthermore, they are intermediate in different ways and to different degrees (see the description below), so each has to be considered separately. If I had to do a binary coding for this character, what is the more reasonable value (0 or 1) to be assigned to each of the species?
Gerbillus nancillus: “Soles of hind feet partly naked behind, with short hairs at level of the metacarpal bones” (Mammals of Africa, Kingdon et al, 2013) — "It is a relatively small gerbil with partially furred soles (posterior portion naked)", "an almost naked hindfoot sole (it is only partially haired in G.nancillus" (Mammals of Sub-Saharan Africa, Monadjem et al, 2015) — "Intermediate" (Lay, 1983)
Gerbillus nigerae: “[S]oles of hindfoot covered with hairs of variable length” (Mammals of Africa, Kingdon et al, 2013) — "These last three-mentioned species [G. henleyi, G. nancillus, G. nanus] also have naked or semi-naked soles of the hindfeet, which are completely haired over along their length in G.nigeriae" (Mammals of Sub-Saharan Africa, Monadjem et al, 2015) — "Intermediate"(Lay, 1983)
Meriones shawi: "Soles of hindfeet partly hairy" (Mammals of Africa, Kingdon et al, 2013) — “The sole of the hind feet is half covered with very fine fur” (Darvish, 2011)
Meriones tristrami: “Soles Partially covered with hair” (Mammals of Jordan, Amr, 2012) —“The sole of the hind feet is mainly hairy; the heel is bare with an irregular black line of demarcation” (Darvish, 2011)
Gerbillurus vallinus*: "Hairy footed gerbils" "soles partially furred" (Mammals of Africa, Kingdon et al, 2013) — "the anterior soles of the hindfeet are furred" (The Complete Book of the Southern African Mammals, Mills and Hes, 1997) —“Soles of hind feet of G. vallinus are naked from heel to the middle of the sole” (Dempster et al, 1999)
*I know it is called the brush-tailed hairy-footed gerbil, but unlike the other congeners that I have looked at the back half of the foot is described as naked).
Could somebody help find weight values for the following species or general body size descriptions for the following species: Meriones chengi, Psammmomys vexillaris, Acomys nesiotes and Acomys cilicicus?
I have searched through the following references with no luck so far:
The Contemporary Land Mammals of Egypt, Osborn and Helmy, 1982; Mammals of Africa, Kingdon et al, 2013; Mammals of Sub-Saharan Africa, Monadjem et al, 2015; The Complete Book of the Southern African Mammals, Mills and Hes, 1997; Mammals of China, Smith and Xie, 2013; Mammals of China and Mongolia, Allen 1938-1940; Rodents in Desert Environments (Prakash and Ghosh, 1975); Rodent Societies, an Ecological and Evolutionary Perspective, Wolff and Sherman, 2007; Ndiaye et al, 2011; Freudenthal et al., 2013; al-Hajeri 2015; PanTheria; AnAge.
I am working on population dynamics and reproductive ecology of a small desert marsupial (20-30g). This species is a bit difficult to mark. At present I am using ear knotching but I am looking for a more accurate way to do it. Do you have experience using this technology for small mammals?
I am interested in reviewing the topic of small mammal mortality within discarded bottles, and information available is severily biased towards the northern hemisphere and in developed countries (USA and Western Europe). Does anyone has information from tropical areas? or from other continents?. This is surpising to me, because I know that discarded bottles is a aproblem worldwide (result of human development). There is information of discarded bottles as a potential source for some diseases like dengue or malaria (the rain water contained in bottles allow mosquitoes to lay their eggs). But nothing about small mammals dead inside bottles.
We survey Bengal Slow Loris in several fixed transects in a forest of Northeastern Bangladesh, while we regularly encounter Particloloured Flying Squirrel (Hylopetes alboniger) and noted down with GPS co-ordinate. Surveys in the transect are not equal. Hence, a one year (at least 4 nights in a month) effort to the opportunistic encounters can reveal the accurate population size of the squirrel in the area?
If possible let me know data analysis patterns in estimating total population from the direct observations.
Best option may be body, but is this methodologically ok? I can use body without skin and intestines, is this better?
I would like to research distribution and movement patterns. Any advice on small tracking techniques would be great. Many thanks, Ryan
Hi. I'm looking to add some photographs to a publication I am doing on the Rodrigues fruit bat (Pteropus rodricensis) for the journal Mammalian Species. I need a photo of a live animal, plus a photo/illlustration of the skull-mandible (see attached file for an example). The specifications for the skull plates are fairly strict (but see image):
SKULL PLATE. Illustrations of the skull (dorsal, ventral, and lateral views) and lateral view of the mandible must be included. These either can be good quality images or line drawings. Do not include a scale bar. In the figure legend, indicate age and sex of specimen, the collection locality, full name of museum where specimen is on deposit, catalog number of specimen, and greatest length of skull or a similar measure. A statement including the origin of the photographs or name of illustrator (if other than the authors) and permission for use must be included. All views must be exactly to the same scale.
Can anyone help? Thanks.
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)?
We seek to attach spools to small nocturnal rodents and track their movement patterns. I have been told that most people use cyanoacrylate-base glues, but am unsure about the specific product type. Suggestions would be highly welcomed.
I would specifically like any information/papers on predation on other small mammals, but infanticide on their own young would also be of help. I have observed them predating Muscardinus avellanarius but would like to know of any other instances, if they are any out there!
we hope to try spooling small nocturanl mammals in order to find their burrows and movement patterns but been unable to find a place that can supply spools
I would like to measure the energy consumption of the European hedgehog during its hibernation under natural conditions. Has anyone experience in calorimetry in the field (preferably in small mammals)? Or are there other clever methods?
I like to measure depth and duration of anesthesia in rats. Does anyone have experience using non-invasive surface electrodes like Sumiyoshi et al. (A mini-cap for simultaneous EEG and fMRI recoding in rodents, 2011). Or are surgical interventions and screws in the skull needed?
Any suggestions about recommended EEG hardware and software also welcome.
Does anyone know whether the correct binomen for the monito del monte is Dromiciops gliroides or Dromiciops australis? Looking at the taxonomic history of the species, it looks like australis was named a year before gliroides, and the holotype was never lost, so why do all publications seem to use the name D. gliroides? Was there some ruling from the ICZN or something that suppressed D. australis?
I would like to measure immnune response in a big sample of small mammals. I heard about LPS intradermal injections to measure inflammatory response.
Does anyone have protocols or other suggestions ?
I am exploring new field and there is so much things that I don't know!
I am screening literature for studies on weevil population dynamics (species that oviposit in oak acorns are of my main interest). It seems that their numbers most heavily depend on availability of oviposition sites (here - acorns).
If oaks do not mast their populations remain low etc
What other factors are know to control their populations? I found papers about predation by small mammals (mice and shrews) but this was not of big significance.
In Manu and Debouzie 1993 authors also mention millipedes and fungi.
So - do the acorns are most important? What papers would you recommend?
I am looking for methodology pertaining to trapping designs (e.g., transects, web, grid, etc.) of small mammals to measure the edge effect of an acute disturbance. For example, if a parking lot were to appear in the middle of a grassland landscape, how far away would the biological footprint be felt by the small mammal community? 10m? 100m? Something in-between? What deisign might I use to determine this? The three statistics I want to investigate include species diversity, species richness and abundance. Thanks in advance, and I look forward to any input.
I am analyzing a 16-year data set of small mammals in a tropical forest. This study started in 1997 with a fixed number of traps in the study area. In 2009 we added more traps in the study area with the purpose of capture some understory small mammals (some of them were also captured in the forest floor). I have no doubt that the increase in the sampling effort in the study area will have an effect on the number of individuals captured and consequently on the population abundance estimates. So, my question is: How can I estimate population abundance when the sampling effort increased during the study?
Hey hallo, I want to mark shrews (Neomys fodiens, N. anomalus, Sorex araneus and S. minutus) for recaptures. The field study shall last 12 month. In the past, I tattooed them with high effort at their tail.
Is it possible to use small microchips? These chips are already in use by vets for marking cats and dogs. But I think that these are too large for small mammals, especially for Sorex minutus.
Does a kind of a very small microchip exists, which I can use for marking these small mammals?
I need to administrate specific dose of risk element water solution right to the rat gastrointestinal tract.
Does anyone have experience with any drop off tools on small animals (e.g. hare, marten, or even smaller)? We want to collar hares with e-obs-GPS-collars but need an additional drop off mechanism so that they lose the collar after a certain period of time, and we can get it back. Any suggestions?
For the past 18 months I have been monitoring two albino sibling hedgehogs living free amongst other wild hedgehogs in York, UK. Despite all the dismal forecasts concerning their survival they have so far managed to get through one winter in the UK. I recently found the male who weighed 1122g, a good weight for an adult male hedgehog. Does anyone have any comments regarding the rate of survival in the wild of other albino members of any small mammal species?
How soon do rats recover from acute and/or repeated restraint stress? Do people always use restraint or other methods to induce stress. What are the benefits of using it over other methods? Considering using repeated measures and want to determine how to space trials out.
I am studying Indian Giant Flying Squirrel in southern Rajasthan. Predation, moon light and reproductive status were found most important factors in animals communication during the night. But except these factors which another factors are possibly affecting their behaviour?