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Paleontology/mammalogy/taxonomy colleagues! Does anyone have a complete citation (or, in a perfect world, a PDF) for Nordmann, 1850, the paper in which the Tribe Camelini was named?
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Nicholas; Pardon me! I submitted an answer to a different question.
Sorry, Jim Des Lauriers
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I am quite confused because some sources distinguish between 5 types: folded, lamellar, villous, trabecular, labyrinthine (e.g. ; ); whereas some sources distinguish only 3 types: villous, trabecular, labyrinthine (e.g. ; ).
So, who is right? Why the folded and the lamellar types do not appear in some sources? In the sources where they appear, Suidae are said to have folded interdigitations, but trabecular in sources where they do not appear. Carnivora are said to have lamellar interdigitations, but their interdigitations are referred as labyrinthine in sources where the word "lamellar" is not even mentioned. Is the distinction between folded and trabecular spurious? As well as the distinction between lamellar and labyrinthine? This seems odd, because on textbook's diagrams these types are very different.
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Fetomaternal interdigitations in placentas refer to the intricate connections between fetal and maternal tissues that facilitate nutrient and gas exchange during pregnancy. There are several types:Villous interdigitation: This occurs at the microscopic level within the placental villi, where fetal chorionic villi project into maternal blood spaces, increasing surface area for exchange.Decidual interdigitation: Involves the maternal decidua, which is the lining of the uterus during pregnancy. Decidual cells extend between fetal villi, contributing to the structural integrity of the placenta.Placental barrier interdigitation: This refers to the physical and biochemical barriers between maternal and fetal circulations within the placenta, ensuring selective exchange of nutrients, gases, and waste products while preventing direct mixing of blood.
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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?
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The capybara, capybara, chigüiro or chigüire, which is a species of rodent of the Caviidae family typical of South America, being the largest rodent in the world
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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?
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I would check other values like head-body length to see if there is intrapopulational variation. If the discrepancy is really due to intraspecific variation you would expect linear dimensions to show a similar pattern to body mass, since you would be measuring bigger dimensions on bigger animals.
However, it's also possible that the reported body masses are just plain discrepancies. I've noticed a lot of second-hand databases like PanTheria can be extremely inaccurate when it comes to reporting body mass values because they often just collate them from the previously published literature. A lot of times studies will use ballpark values or the midpoint of guestimated ranges from sources like Walker's Mammals of the World, which aren't very accurate or precise. A good example of this is the South American rodent Dinomys branickii. Most studies using body mass for Dinomys will report a value of 10-15 kg or 13-14 kg, which seems to be drawn from a ballpark range of 10-15 kg mentioned in a paper from the early 20th century (maybe Allen 1942). This doesn't appear to be based on any measured specimens, and no studies list specimens from where these data were collected. However, actual weights of Dinomys based on first-hand measurements of collected wild individuals (e.g., Osbahr et al. 2009, Gottdenker et al. 2001) finds that the body mass of Dinomys is actually only about 9.5 +/- 1.1 kg, a much lower estimate. Larger individuals are known but are highly gravid, captive ones. Nevertheless, people have been reporting a 13-15 kg body mass for Dinomys as if it were gospel.
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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?
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Hello Russell: Tungsten carbide is used as an abrasive. It comes in grit sizes that vary from extremely fine powder to pretty coarse grit. If you can get permission from collection managers, a heat sterilized jar of the grit of whatever sizes you determine to be appropriate, non-toxic and pest-free. A graduated cylinder provides the volume directly. Were I a collections manager, I can't imagine what the problem would be.
Apparently specimen loans are out of the question. Hmmm.
Best regards, Jim Des Lauriers.
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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
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First two images may be Aelurostrongylus abstrusus,
third one is Capillaria aerophila
with all best wishes
Afkar
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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.
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I know this is now quite old, but you could try Frank Grutzner at the University of Adelaide, or Paul Waters at University of NSW (I think). Also, Kathy Belov.
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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!
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Many thanks for all of your feedbacks!
Hein, forest elephants are not our only studied species. We focus more broadly on forest large mammals, such as forest antelopes, small primates, gorillas, chimpanzees and forest elephants. We point our cameras to animal tracks.
Mary, your recommendations exactly correspond to our first sampling plan for the “community forests” land tenure! We are just still wondering if it is not a problem to use a different design of camera grid for this land tenure (split in 3 sub-grids), compared with the three others where we will use regular grids of 11 cameras.
Fructueux, oui chaque forêt communautaire fait entre 4000 et 5000 ha, mais elles ne sont pas contigües. L’idée de diviser l’échantillonnage des 11 caméras entre trois forêts communautaires distinctes relève d’une perspective d’échantillonnage relative au régime foncier étudié (en acceptant une certaine variabilité due aux différences intrinsèques entre ces trois forêts communautaires), et non une étude du terroir au sens propre. Effectivement, c’est un problème d’installer une grille de 11 caméras dans chaque forêt communautaire durant un mois chacune en alternance, et nous éliminons donc cette possibilité. Il reste donc à voir si l’utilisation de 11 caméras dans une seule grille dans une seule des forêts communautaires serait plus pertinent ou non que d’utiliser 4 caméras par forêt communautaire avec une logique « agrégation par régime foncier ».
Kalinga, thanks for this confirmation and those recommendations! I agree with your comment!
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Simulation model for population of Koalas?
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Try articles at http://www/panearth.org;
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Small vs Medium?
Normal vs Easy Set?
Or are Tomahawks or XL Sherman a better fit?
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I also prefer Tomahawk traps.  Just make sure to check the ends of the wires - sometimes they can be pretty sharp, and can injure little paws as they try to dig around the trap.  A flat file can be used to take off the sharp edges.  Too large a trap can be a problem if the mesh size is big enough to get their nose through.
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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?
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Dear Isaac, interesting point that sometime is difficult to answer.
In your commnet I dont understand if the population of this subspecies (28) are in captivity? or in nature?
Many species have been in neckle bottle during thier evolution and letter had recovered thier population with a reduced genetic variability.
However is not the same to talk about species variability than subspecies variability. The level of genetic variation is different.  Have been the subspecie variability studied?
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Does any one have an idea of the dimensions of a canine open field arena. I want to assess novelty-induced behaviours in canine pups and young adults.
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My experience is with rodents, in which you can study both thigmotaxis and novel object recognition in the arena. For dogs, you could study object recognition memory. Here is a protocol adapted for the dog (the size of the arena was 0.609m x 1.1.5m x 1.08m):
Development of a protocol for studying object recognition memory in the dog, Callahan et al., 2000 (http://www.sciencedirect.com/science/article/pii/S0278584600001020)
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Caught in the Western Region of Ghana
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I think so and agree with the above viewpoints.
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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?
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... And you will find some people loooking and looooooooooooking to all genes of their dataset until they find THAT marker (in the middle of a neutral global result) that will show that HIS/HER SPECIES is statistically "real" (for example, frequencies of an allele coding for color under a strongly selective environment for that color). And the worse part: they get published in more-than-good journals. Now with RADs, for example, you could "find" a species almost everywhere, if you look enough.
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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. 
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By spatial association, do you mean social association (existing in the same group at the same time) or general spatial proximity (existing in a similar place)? What exactly did you do for your analysis for for "association strength?" I can think of a number of different metrics that I could call "association strength" and some would be weaker than others. And there are some statistical approaches that make more sense than others, as well.
So, if I've done my math right, you have n = 32 adult deer (assuming the 982 is a typo for 992, because 992 would give a half matrix of 32 individuals)? What's this in context of your overall population density? If we sampled Toronto for 30 people, we'd get very few individuals that are related to each other by chance. However, in a village of 300, for the same average family size, obviously there's going to be more per-capita relatives. My first suspicion is that given a mere 6 individuals have high relatedness, your sampling density was low (unless mortality is high and you have a stationary or declining population density). However, this assumes you sampled your deer at random. If your sampling protocol involved exhaustively sampling all animals that had contact with a group of focal individuals, and you were comparing them to random non-contacted controls from the same population, then suddenly you're looking at a decent number. It all depends on your sampling design.
A word of caution on the contact rate stuff: 0 < r < 0.125 puts you in a foxy place. Since you're thinking about mechanistic action (the animals makes decision x based on characteristic y), you need to think how that factor is going to be assayed in nature. You can make some argument based on odour cues and imprinting for close relatives, but once you get past 2nd degree kindred you need to consider whether the animal might reliably distinguish anything further than a cousin. You might consider using "cousin level or greater" as a binary predictive factor. Also, I'm assuming that you've stripped known parent/offspring-at-heel pairs out, since it wouldn't be very surprising to see that mom pays a lot of attention to her still-young fawns. If you don't strip that, you run the risk that your effect is entirely does with fawns or yearlings at heel; while technically, that's relatedness predicting contact rate, it's a very specific type of relatedness.
The group membership thing is most interesting. There was a recent paper (Biosa et al 2015 in J. Zool.) that found that their spatial segregation had a temporal component to it, with opposite sex relatives negatively associated during breeding periods. Without knowing much more about your study and study system, I'd wonder if you considered what the deer are 'trying to do' life-history-wise the time of year you observed/sampled them.
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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
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Dear Jeyathilakan,
The bat is most likely a juvenile Taphozous sp. while the fly is a Nycteribiidae.
Best
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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.
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In particular, what size of river could a pine marten cross?
And would they utilise road bridges?
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Andrew,
We know that martens have predated black throated diver nests on the Loch Maree Islands and must have crossed ~150m of water at some point to get to these islands. See SNH commissioned report 379 http://www.snh.org.uk/pdfs/publications/commissioned_reports/379.pdf.
The report suggests that crossing 30m of water (between 2 of the islands) is not a problem. This is standing water of course, so a 30m wide fast-flowing river would be different matter entirely. The report also refers to an American study in which American martens crossed either 30 or 64m of standing water.
Would they utilize road bridges? - of course! Pine martens began to colonize the Isle of Skye soon after the bridge was built.
Hope this helps.
Rob
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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.
Lophuromys xena
Lophuromys flavopunctatus
Lophuromys brevicaudus
Lophuromys melanonyx
Lophuromys chrysopus
Lophuromys sikapusi
Lophuromys woosnami
Uranomys ruddi
Lophiomys imhausi
Refs:
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.
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Dear Haya Rashed,
Welcome. I am waiting to read the result of your study. 
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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?
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Dear all,
My Gauthier-lab at Laval University studies lemming populations in an Arctic island since ~15 years now. Lemmings are ~ 20-50g. They are using a trapping grid and live-trap with baits (piece of apple as bait and they put cotton wool for a cosier trap ^^). Here are 2 references (with pdf) of their published study. They marked lemming with PIT-tag and use readers for individual recognition.
You could try to contact Dominique Fauteaux (current PhD student on lemming - present in ResearchGate) for more details about the study design.
For what I've seen during fieldwork (I was working on goose but also help sometimes on lemming project), there were some death due to heart attack but none due to infection.
Cheers,
Guillaume
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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. 
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Tanvir -
I am glad you found our paper interesting, but I have to add that these are relative densities, which is not the same as absolute densities let alone population size. Relative densities and absence-presence data are important, but in order to get the absolute population size you have to use other methods as indicated in the previous discussion.
Cheers, Andreas
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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?
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For anyone who's still interested, I think I may have tracked down where this rumor got started. In their 2007 paper on the ptolemaiid Kelba, Cote et al. mention the fact that (at least according to their study) researchers have tended to look outside Africa for the sister taxa of various African groups rather than looking for possible relationships with other African taxa. They mention Afrotheria as the prime example of how the previous idea may not be the case, and also mention that hyaenodontids may have originated in Africa, and they word it in such a way that it might be ambiguously interpreted as hyaenodontids possibly being afrotherians.
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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.
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Dear Gareth,
As far as I know, there are not many things published on that topic in Central Asia, and of course only in Russian. 
you have that book chapter in English, a bit more recent than the 1970 one: Bibikov, D.I. (1982) ‘‘Wolf Ecology and Management in the USSR’’, in F.H. Harrington and P.C. Paquet (eds) Wolves of the World. Perspectives of Behavior, Ecology and Conservation, pp. 120–33. Park Ridge, NJ: Noyes.
I got a copy of Vyrypaev & Vorobjev book on wolves in Kyrgyzstan:
Vyrypajev VA, Vorobjev GG. 1983. Volk v Kirgizii. Frunze: Ilim. 94 p.
However I would be cautious in using it since I detected some mistakes in the tables they give inside.
I published two papers about human-wolves relationships in Kyrgyzstan (they are available on RG, and you may find something in the references...), as well as a PhD in French. If you have any question about that specific topic, you are welcome to contact me!
Nicolas
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Exactly what it says on the tin.
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Humans have a thickened band in the scalp skin, approximately following the hair line over the ears. Sweat from this region has a peculiar navel-like odor, but I know no reports of the presence of apocrine sweat glands in there. It may be worth investigating.
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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?
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Russell, perhaps not the best phrased question given the responses you are getting, but I understand what you mean. Beavers eat bark as well as the cambium layer beneath it, roots, as well as many other plants and plant parts. Xylophagy may be a better fit for something like woodworm species (of which there are quite a few).
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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.
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EsEstimado Jose, 
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Please describe me about 'recce' (reconnaissance) survey method. How we can use it to estimate nocturnal mammal counter rate??
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Hi, recce is simply a way of getting apriory knowledge about your study area before starting your study. It does not have a statistical apparatus for you to calculate abundance of any species since it is not among any kind of sampling techniques. Even though it is not very statistically sound way, what you could do is, map and mark the whole recce area through GPS and also mark GPS locations of your target animal. You could then calculate the encounter rate as no. of Ind. Per km. 
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A colleague of mine at a Cdn university is looking for a service provider to ID mammals from fecal pellet DNA. Any suggestions?
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Try one of these authors (ms available on ResearchGate):
Kartzinel, T.R., P.A. Chen, T.C. Coverdale, D.L. Erickson, W.J. Kress, M.L. Kuzmina, D.I. Rubenstein, W. Wang, R.M. Pringle. 2015. DNA metabarcoding illuminates dietary niche partitioning by African large herbivores. PNAS, online early.
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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.
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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.
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The enamel isn't uniformly thick in upper or lower teeth, so I doubt you'll find any studies that have measures of this from exactly homologous points that could be comparable. Plus, a worn tooth is showing a section of an enamel band often at oblique sections, not a real measure of enamel thickness. And, that thickness will change from crown tip to base, so the real answer would be in estimating enamel thickness from something like microCT. That has been done with some primates and crocs, but not really for much in the context of ungulates, as far as I know. would be great to see done.
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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. 
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Robert Barton has published several comparative works that looked at specific (usually sensory) brain structures:
1 Barton R, Purvis A, Harvey PH. 1995. Evolutionary radiation of visual and olfactory brain systems in primates, bats and insectivores. Philos Trans R Soc Lond B Biol Sci 348:381–392. 
2 Barton R. 2006. Olfactory evolution and behavioral ecology in primates. Am J Primatol 68:545–558.
3 Barton R, Harvey PH. 2000. Mosaic evolution of brain structure in mammals. Nature 405:1055–1058.
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Has there been much research investigating whether cats behave differently under a natural range of temperatures?
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Yes , especially the reproductive system related behavior .
Cats are seasonal , so  high temperatures changes endocrine and reproductive system. What reflects the general behavior .
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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.
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The NCBI has a summary site at http://www.ncbi.nlm.nih.gov/genome/browse/, and you could look at genomesonline.org (which I think is also run by the JGI, but certainly should have more than 40 species).
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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) 
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Hi,
I rank the threats in the following way:
1) habitat deterioration - very important threat for many species, especially forest species or those with highly specialised habitat requirements, globally important.
2) climate change - may led to shifting distributions and habitat loss.
3) wind energy - important threat to some open space foragers, however a limited number of species is affected.
4) WNS: although huge numbers of bats die, a limited number of species is affected, the disease is still geographically restricted and hopefully the species can recover.
I would like to add traffic (casualties, fragmentation), poisoning (direct and indirect) and pets (especially cats) as additional threats, all three rank for me higher than wind energy.
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I am looking for data on bite forces of extant Artidactyla and Perissodactyla. Anything is welcome, like experimental data or FEM calculations.
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While there may not be exactly the data you are looking for, the Feeding Experiments End-User Database (FEED) may have some information that would provide the basis for a first approximation "one-off" calculation.
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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?
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Wow, that article says "even dimensions of essentially
bony elements are affected by the preservation process (Lee, 1982)". (LCC. J. C. (1982). Accuracy and precision in anuran morphometrics: artifacts of preservation. Syst. Zool. 31: 266-281. )
That makes it sound like all bone measurements from preserved specimens have to be taken with a grain of salt.
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We found one boa predate on jaguarundi.
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Well its possible if Boa constrictor can catch jaguarundi and kill him. I suppose his body shape mach to gape size of Boa snakes. Maybe its possible that Boa find fresh dead body of jaguarundi and constricted them (sometimes constrictors do this) and then going to swallow it?
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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.
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Very interesting new tool to consider: Carrion fly sentinels of mammalian diversity...
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I am trying to connect locomotor and postural repertoire with ecological adaptations to investigate the adaptive significance in small-bodied bark gleaning squirrels
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Sorry Dionisios! if I find recent information about your search I'll share it with you.