Questions related to Mammalogy
There are two extant species of Hydrochoerus: Hydrochoerus isthmius, the lesser or Panamanian capybara, and the genotype species H. hydrochaeris. The latter is the more common species of capybara, found throughout most of South America, whereas the other is restricted to the northwestern side of the Andes ranging into Panama. However, H. hydrochaeris doesn't seem to have a useful common name to distinguish it from H. isthmius. It's referred to as the "capybara", but both species are capybaras, and it's never referred to as the "common capybara", "greater capybara", or "southern capybara". H. hydrochaeris is also much larger than H. isthmius (nearly twice the size of the latter species).
I am making a figure I intend to use to show to an educated layman audience, and am using capybara bones as an extant scale. I am trying to use the common name to not confuse my audience, but at the same time I want to make it clear I am referring to H. hydrochaeris and not H. isthmius so there is no confusion for people who are more familiar with scientific names. Given this, what would be the correct common name to refer to H. hydrochaeris such that I do not confuse my audience?
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?
Our Group (Laura Jaramillo and me) have two other parasite eggs found in faeces of the river Otter, that are different of the previously discussed
Is there any way in Europe to get a small amount of frozen citrate plasma (several microliters should be enough) of platypus and echidna from Australia?
The research would involve cross-check reaction of some monotreme plasma proteins on antibodies against human plasma proteins by western-blotting.
If anyone has an access to the animals and knows how to organize such matter I would be grateful for personal contact.
I am currently working in Cameroon on wildlife assessment with camera-traps, aim to compare faunal biodiversity between 4 different land tenures regimes. The objective is to quantify afterwards diversities indices between 4 sites. After discussions with specialists, I would like to have your opinion. I plan to divide my 44 cameras into 4 grids, corresponding to my 4 land tenures regimes, during 3 months. Cameras will be placed with a distance of 1.4 km between each other (TEAM protocol). I have 11 cameras available for each grid, but I have an important question about one of them: one of our land tenures regimes is composed of 3 distinct community forests of 5,000 hectares each.
We have three possibilities concerning the distribution of cameras for this land tenure: (i) we can use 3-4 cameras in each community forest during 3 months, (ii) we can set up 11 cameras in each community forest one by one and each during 1 month (but it will not be possible to keep the 1.4km distance between cameras, considering the small size of community forests), or (iii) we can put the all 11 cameras in only one community forest during 3 months, but taking maybe then less heterogeneity of landscape patterns.
Would it be better to maximize the sampled zone with a maximum of time and a minimum of cameras, or to reduce the time and put more cameras in each sub-zone? Are 700-800 camera-trap days ( < 1000 camera-trap days) enough to obtain valuable data ?
Thanks a lot!
I am interested to get ChR2 mammalian expression vectors. Please tell what are the sources I can get hese from. One I know is Addgene. Kindly inform if there are any other too...
Last monday the CONANP, a gubernamental institution in México, have released a familiy of (Canis Lupus baileyi) integrated by a couple of wolves and their five puppies. According with CONABIO another gubernamental institution in México, this specie has probably extincted in wildlife.
¿Is there is a number that indicates the minimum number of individuals in order to preserve genetic variability? According with the population of Canis Lupus bailey, aproximately 28 individuals, I understand that this poblation is not enough to preserve the variability genetic., Is there a research that support that?
Hello. I am deeply puzzled by the breaking-news study by Fennessy et al. (2016) "Multi-locus Analyses Reveal Four Giraffe Species Instead of One" (dx.doi.org/10.1016/j.cub.2016.07.036). The nuclear analysis seems to be solely based on 7 introns sequences, and contradicts a previous study, by Brown et al. (2007) "Extensive population genetic structure in the giraffe" (dx.doi.org/10.1186/1741-7007-5-57), which included over 3 times more individuals for 14 microsatellites loci, with samples from contact zones, and found up to eleven subpopulations clearly differentiated from a nuclear point of view (with possible hybrids)... without suggesting any taxonomic emendation though! As results of both studies are not clearly confronted in the 2016 paper (I barely found a terse "the statistical support is not clear" when mentioning the 2007 results), could any one explain me where are the new insights brought to the Giraffe's case by the latter? And who to follow?
We did a study on a free-ranging population of this species, in which we tested whether spatial associations (2 adult deer within 25 m of each other) varied with several factors such as season, home range overlap, genetic relatedness, and sex, age and disease status of pairs of deer.
Relatedness was not a good predictor of spatial associations.
Pairwise relatedness measures were estimated in SPAGeDi version 1.4 (Hardy & Vekemans 2002) using the estimator by Queller & Goodnight (1989).
Relatedness was on average -0.003 ± 0.005 (±SE, SD = 0.15, range -0.4 to 0.6).
I would like to compare these values with others generated in other mule deer populations, but I cannot find any reference!
Any guidance will be appreciated.
I found an injured bat in our college premises ( Veterinary College and Research Institute, Orathanadu, Thanjavur, Tamil Nadu, India.) with ectoparasites. The photographs of bat and the bat fly are enclosed for identification
It is possible that these molars have malformations? Or they were just growing in a wrong way ?
I identify them as Mammuthus primigenius? and the black spots represents burning traces. Probably found near a paleolithic site.
I am interested in learning the status of the following species with regards to the hairiness of the bottom side of their hind feet. The references I cite below are silent on this issue (for these species) which leads me to believe that they are all naked-footed. However, I would greatly appreciate it if someone who is familiar with these species could confirm this for me/correct me on this issue.
Mammals of Africa, Kingdon et al, 2013; Mammals of Sub-Saharan Africa, Monadjem et al, 2015; Kingdon Field Guide to African mammals, Kingdon 2015; Walker’s Mammals of the World, Nowak 1999; The Mammals of the Southern African Subregion, Skinner and Chimimba, 2005.
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?
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.
I have often heard in the "gray" literature and from several informal statements by researchers that there has been some suggestion that hyaenodontid "creodonts" may eventually be found to be afrotherians, given that some of the oldest known hyaenodontids are from Africa and that this continent seems to have been the center of the group's diversity. However, searching through the literature I have not been able to track down any paper that suggests this. Does anyone know of any paper that has suggested a afrotherian placement for Hyaenodontidae?
I'm working on a small project exploring the potential for coordinating the management and research of wolves in the Southern Caucasus (Georgia, Armenia and Azerbaijan) and Central Asia (Kyrgyzstan, Tajikistan, Kazakhstan and Uzbekistan) and want, first to get an idea of the current situation. I'm looking for paper/articles/chapters on the subject spanning the past 30 years or so (both Soviet and post-Soviet eras). Also, if you are currently working on wolf management in any of these countries, I'd like to hear from you.
I am wondering if there is a proper term for mammal species that are browsers, but primarily feed on the woody tissues of a plant as opposed to the leaves and shoots. Examples that come to mind are species like beavers (Castor spp.) and North American porcupines (Erethizon dorsatum). Would xylophagy be the correct term?
The commitee of Taxonomy of the Society of marine Mammalogy has recognized as Arctocephalus australis un-named subspecies to the "Peruvian fur seal" for differentiate it from the specimens of the eastern coast of Southamerica. (Committee on Taxonomy. 2014. List of marine mammal species and subspecies. Society for Marine Mammalogy, www.marinemammalscience.org).
I need know if this subspecies is studied by some researcher for a definitive classification.
A colleague of mine at a Cdn university is looking for a service provider to ID mammals from fecal pellet DNA. Any suggestions?
My aim is to store hair samples (bunches of hair collected from a small mammal during multiple years) at the conditions that would be least harmful for the DNA contained in hair follicles. With respect to the conditions I refer to temperature, humidity, container type, etc.
What is your personal experience with this kind of samples? How long one can store such samples at a cool and dry place e.g. in paper envelops?
Thank you for your advice.
I'm curious if anyone has looked at the difference in enamel band thickness between the upper and lower teeth of any mammal, but more specifically ungulates.
I would like to know if there has been a comparison of the differences between dogs, wolves, monkeys, or any mammals in the specific brain areas, primarily the pre-frontal cortex.
Does anyone know where to find and download information on genome size and number of protein-coding genes for as many eukaryotes as possible? I found small sets in Ensembl! and JGI but I suspect there must be more sequenced species. So far, I have the full set of fungi, about 40 mammals and some inverterbartes.
Taxa respond differently to anthropogenic challenges. Therefore, it is important to identify the most severe threat for a specific taxon. Bats are special among mammals owing to their ability of powered flight, their close association with humans, and many other things. Many bats are endangered, yet we are missing a global perspective on the specific causes. I am asking for your educated guess regarding what factors are most responsible for the decline in bat diversity? I suggest four causative factors, and I leave it up to you to rank those according to decreasing importance! (Plus, you may also add a factor in case I missed one)
I am looking for data on bite forces of extant Artidactyla and Perissodactyla. Anything is welcome, like experimental data or FEM calculations.
I am interested in doing a geometric morphometrics study on bat wings and am considering using specimens fixed in formalin. To do this, I would like to have the bat wing stretched out to its fullest extent, take standardized pictures of the outstretched wings, and place landmarks on representative features. This method has been used commonly with live specimens (e.g. in this great paper by de Camargo and de Oliveira 2012 -- http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0049734 ). However, the specimens I am considering using are fixed in formalin and vary somewhat in the amount of rigidity/flexibility. This makes me question how able I will be to stretch the specimens' wings out in a standardized manner.
Is there a way to restore flexibility to bat wings fixed in formalin, so that I could achieve a standardized degree of flexibility and stretch the wing to its fullest extent? Ideally, it would be great to restore the rigidity and when I am done. Anyone have any thoughts or experience changing flexibility of fixed specimens of other taxa, or literature to point me in the right direction?
I study white-footed mouse (Peromyscus leucopus) populations of Northern Michigan, and my research requires me to sacrifice individuals for the purpose of geometric shape analysis. However, this is confounded by the presence of deer mice (P. maniculatus) in some populations. These two congeners have been established as good species (i.e. no hybridization), but there are usually cryptic individuals in our trapping sites that are difficult to identify until we take a saliva and genetic sample back to the lab. In the field we use standard measurements such as ear length to determine one species from the other, but this is not fool proof as identifications made in the field are occasionally contradicted in the lab.
Is there a non-lethal way to determine species in small mammals in the field so that I do not mistakenly sacrifice deer mice and/or bias my P. leucopus samples by excluding cryptic individuals? Thanks for any and all help, and if you need further context just ask.
I am trying to connect locomotor and postural repertoire with ecological adaptations to investigate the adaptive significance in small-bodied bark gleaning squirrels