COLLECTION OF THE PRIVATE BIRD-SKIN AND SKELETON COLLECTION
(Open for scientific research)
Copyright 1993 Johannes Erritzoe, latest revised 2016
Short history of the collection
Body mass indications
Supplementary comments, samples collected and methods
The osteological collection
List of all papers based on this collection
Collection management guidelines
My experience with sex determination, male (with list of abnormally coloured testes)
female (with list of abnormal ovaries)
How to make safe sex determination
Overview of the various kinds of study skins
In comparison with the huge accumulations of study skins that many large museums around the world can muster, the collection presented here is indeed a very small one. (7.800 skins, c. 4.300 part skeletons, 1.300 skeletons and a large feather collection). Nevertheless I think it may be of interest to scientist because of the many pieces of information and skeleton derivatives kept with most of the skins and skeletons.
In a letter to me dated 28 March 1989 the Senior Curator of Birds of the Carnegie Mus. Nat. Hist. in Pittsburgh, Dr. Kenneth C. Parkes wrote as follows:
“I am sure you realize that many of your preparation techniques are not observed in most museums. The use of wires in head and feet, for example, I had not seen before, and your recording of almost every possible kind of data with respect to the dead bird. I don’t think I have ever seen bird specimens so thoroughly documented”.
Professor Anders Pape Møller, now Paris, wrote me in a letter dated 2nd December 1994:
“I have personally benefitted tremendously from using this collection for several research projects, and I can easily imagine that many other scientists could benefit as well from using the collection.” (cf. list of all papers based on this collection)
SHORT HISTORY OF THE COLLECTION
The collection was started back in 1947 under supervision of the late Erik Petersen, Zoologisk Museum in Copenhagen when I was 13 years old. From the very beginning the following pieces of information have been recorded: Date and locality, weight and a drawing indicating size of sex organs using a pair of compasses, and the other the measures taken with a calliper. If testes were discoloured, it was mentioned too.
Shortly after the beginning the size and sex organs were measured with a slide gauge in order to get a more exact meassurement (which nowadays is more computer adaptable). Weight of body mass was taken with an ordinary letter balance from the beginning, but from 1964 a pair of scales was used – also on three later expeditions. From 1995 a Sartorius 1608 MP has been used. For birds larger than 500g a kitchen weight with an accuracy of ‡10g have been used since 2000.
The number of information grew gradually year after year, some because I saw others use them, others Anders Pape Møller’s or my own ideas. Around 1965 I showed Dr. Finn Salomonsen some of my skins in the museum in Copenhagen, and he found I wasted my time collecting so many particulars. “Erritzoe”, he said, “if we want to examine a problem, we just shoot 2-300 specimens of that bird”.-
I would like to see the person who would dare to say that today!
Also the method used in making the skins has undergone a large development. The early ones were made as soft cotton wool skins with legs crossed. The many museum-skins I saw with legs, tails or heads broken off, or where fat from legs had discoloured the hind part of the birds and labels, gave me the idea of using excelsior (wood wool) instead of cotton wool, because with that material it is possible to use wire through the legs, neck and tail and thus obtaining more stability. In some years wire has also been used for spread wings displaying important plumage features that folded wings on skins cannot show. In a series I tried to make the degree of extension different. From about 2001 I stopped to made skins with one wing spread and instead started a feather collection, mentioned latter.
Skeletons have been collected from 1986 and onwards after I got World Inventory of Avian Skeletal Specimens, 1982 by D. Scott Wood et al.
A more detailed report of the collection is given later.
In a catalogue like this only to be accessible to the scientific community it should be waste of space and time to try to defend the legitimacy of a bird-skin, skeleton and feather collection.
The understanding of bird conservation has changed radical during the period I have collected bird skins. Most bird species which were legal to shot only 30 – 40 years ago are now protected by law. The attitude to conservation has also changed concurrently due to the disastrous decline of many bird species.
All bird-skins and skeletons in this collection have been collected in accordance with the national law and CITES regulations valid at the time of collection and without the normal stamp-collector mentality to obtain complete series, which would surely not be possible in a collection like this without breaking the law. I hope my more than one thousand House Sparrows speak for itself.-
Throughout the last more than four decades this collection has been increased only by birds killed by window collisions, traffic accidents, taken by cats, and many other man-made causes, but also birds found dead, e.g. dead from disease, and exotic birds dead in captivity, but with a priority of making larger series of common species. All protected species are registered by the Danish authorities and since 2013 all new species, except a few Danish which still may be hunted, all numbered with a six-number aluminium sign.
To a very small extent bird species have been exchanged with scientific museums. In such cases only salvaged dead bird-skins were used.
“Collectors should realize that the important thing they are collecting is information about animals. The specimens themselves are a basic source of this information, but not more basic than accurate, durable, and complete labels and carefully written notes” (Anderson 1965).
The following check-lists have over the years been used to name the species:
Morony, J. J., W. J. Bock, & J. Farrand, 1975 & 1978: Reference-list of the Birds of the World. AMNH, New York.
Mayr, E. et al., 1979-1986 (second ed.): Peter’s Check-list of Birds of the World. Vol. I-XVI. Museum of Comparative Zoology, Cambridge.
Howard, R. & A. Moore, 1991: A Complete Checklist of the Birds of the World. Academic Press, London.
and since 2003:
Dickinson, E. C. 2003: The Howard & Moore Complete Checklist of the Birds of the World. 3rd Edition. Christopher Helm, London.
The last has been used for the nomenclature in a database of the collection mentioned later.
Sexing by collectors was formerly often based on some preconception on sexual differences rather than on anatomical confirmation. I find it important by dissection to determine the sex and meassure the gonads, not only to get a safe sex determination but also because the condition of the gonads tells a lot about the bird in question.
When nothing to the contrary is mentioned (e.g. Collector …), identifications have been made by myself. For the determination of sex a four-factor illuminated magnifier is used. For meassuring I use a vernier calliper (Metr Auer) and a digital (Jocal Manual) ditto. When gonads are difficult to find, I use to put the body cavity in water with some salt. After an hour or so it is often possible to sex determine the bird.
Each testis is measured length x breadth in millimetres so testis volume can be roughly estimated but since 1995 I have used also to weight both testes without epididymis, when they are active. The left testis is the right when viewed from ventral.
If testes are discoloured, the abnormal colour of left or right or both is recorded. When it is difficult to measure the testes in the body and when you want to preserve the trunk skeleton, they are measured outside, which is often necessary e.g. Alcidae. With very small testes which are visibly of the same size, only one is meassured.
When birds only have been sex-determined by plumage, this is invariably recorded. In many cases, however, a bird is rejected if the determination of its sex proves impossible and the bird in question is not sexually dichromatic. If there is room for doubt, e. g. on account of advanced putrefaction, the fact is indicated by a question mark.
Certainable cases of sinuous spermatic ducts are noted when visible. On the other hand the fact that there is no note on this – on most birds a very difficult matter – does not warrant the assumption that the spermatic ducts are straight.
The ovary is measured from the largest length to the largest breadth in millimetres. If two ovaries, both are measured. If large ova are removed and measured the measurement of ovary is taken without these large ova. In quite young birds the ovary is transparent, often very difficult to find and only by turning the body in different angles to the light. Transparent ovaries are also recorded, but it is mentioned that it is transparent. I believe that many such young females by former field workers mistakingly are determined as males, confusing the adrenal glans for testes. If the ovary is granulated this is recorded and the size of ova larger than 1 mm is recorded. If ova are large, all eggs of more than 2.0mm are removed, and their mass and largest diameter is given. If calies are found they are recorded too. Black ovaries which mostly are found in sick or old birds are recorded.
Oviduct – means that the oviduct is thin and straight. Often followed by two measurements which give the width of the oviduct cranial (magnum) and caudal (uterus). Oviduct followed by a wavy line means that the oviduct is sinuous and possibly distended, often followed by two meassurements as above. If oviduct is greatly distended, the length in situ (if poosible) and out and mass is recorded.
Anomalously shaped testes or ovaries are recorded, as is any other abnormality worth mentioning. If sex determination or any other thing observed is astonishing, e. g. because of plumage, a ! is written to tell a later researcher, that also I was surprised, but I had checked it once more, and I am so sure that I will say to the researcher: Do not leave that specimen out of calculation, it is OK. If any doubt is left a ? is written and the organ in question is often kept in alcohol and this is mentioned on the label.
When a bird wears its first regular plumage and possesses a bursa Fabricii and/or an unossified or partially unossified skull, it is classified as a “juv.”. (Only for members of Passeriformes, regarding to skull ossification). After the complete moulting into second plumage, it is set down as a juvenile if in some other respects it displays the marks of a bird in its first year (such as straight oviduct, partially unossified skull, remains of the plumage of the juvenile bird, or/and a bursa Fabricii). Even though a bird has undergone no visible change of the plumage by the first of January, it will be registered as a immnature or ad (depending of the species, short-lived species = ad). For birds with a different plumage in their second or third calendar year compared with that of the adult bird, the age indication 2 or 3 year is also used, but more often only immature is used.
A bird registered as “ad” is one that has no bursa Fabricii or any other sign of being in its first year, and which wears the plumage that is the final one for the species. Birds with worn feathers in summer are nearly always adult birds.
In most passerine the stage of skull ossification is noted in autumn (fall) and winter, but seldom in spring time. In exceptional cases non-Passeriformes are also recorded. For the individual bird this is often done both with a drawing on the label and a reference to the stages (A-E) of ossification showed in Svensson 1984. Bursa Fabricii (b. F.) is measured length x breadth and mass taken. Where b.F. is expected but not seen, owing to obesity and other factors, it is noted that b. F. was not found. Thymus is also recorded, but I do not use it in age determination, because in spite it always in the literature is mentioned as only occuring by young birds, I have many examples, where adult birds also have thymus, especially females in the breeding conditions (Erritzøe 1985). Besides of this, thymus is difficult to find if the bird has been frozen more than a few months before skinning.
In absence of particular comments to the contrary, dates are perfectly exact. The months are given by a three-letter abbreviation to avoid any mistake with the American way of month nummerals, e. g. means 5/11/1990 in Europe 5 November 1990 but in USA 11 May 1990! However, on older specimens, where the labels were written in Danish, the European method has been used. For DNA tissues date of skinning is used, and here the European practice is followed, always with JER at the beginning followed by a number indicating if one, or more birds are maded the same day, e.g. JER-2 14.04.2006 means that it is the second bird made on 14 April 2006. If time of the day or night for death is known, e.g. by window accidents, this is also recorded. Number of days gone between death and preparation is given in later registrated birds, but it is easy for all specimens where DNA tissues are taken to work out this piece of information.
Locality where the bird is collected is always given as accurate as possible. The coordinates are given for all Danish localities using the CD ROM Topografisk Atlas for Denmark. For all other countries coordinates are taken from The Times Worldatlas and when not found there, from Google.
All measurements are taken in millimetres but I have always omitted written mm, opposite weight where g for gram always is written. All average are rounded off up.
Long ago I realized how destruction it is when measurements are taken on old skins, especially max wing length. Therefore I began to measure the dead bird before skinning. These measurements are comparable with these bird ringers take on alive birds.
To now the index of size, e. g. in connection to Bergmann’s rule, have always been the wing length and to a lesser degree body mass (Zink & Remsen 1986). The following measurements can maybe give surprising new information because it is possible to take therse with great accuracy:
The length of femur and humerus are recorded on the clean and dried bones, except for these not in the collection but with a six-number, where it is taken on the raw skeleton. From 1998 the average measurements (n = 2) are given for both left and right femur and humerus on the label, on my database all four. It is a great advantage to take two measurements of each item because if there here is a great difference between measurements you know that at least one of them is wrong, so you have to repeat taking the measurements until you are sure. In future tarsus will also be measured because with wire through the legs this measurement is difficult to take at a latter stage. Wing- (max.), bill- (culmen to skull), bill high and width (in this order) and tail- lengths are also taken from the bird before skinning. Wingspan and wing area are taken the last years. From the dead body the following measurements were earlier taken: Length of body from front of body to tip of pubis, the largest breadth and height of the body. The length of the neck from a straight line between the dorsal end of furcula to the cranium. Instead of these measurements I now only take the length of the keel. The diameter of both iris and the whole eyeball; because eyeball rarely is quite round three measures of diameters are taken, horizontal, vertical and at a tilt and an average of these three. Iris measurement is also taken three times, but because iris is normally quite round only taken from one point, if oval this is noted and measurement taken from both the longest and shortest place. Finally the measurement “wingtip to tailtip” is often taken on the dead bird with the wings and tail in a natural position. If wing tip is outside tail tip it is signed +….This measurement is mostly taken as a help for bird illustrators, but in a few bird species it can also be a help to indentify the bird, e. g. in shorebirds. A short time the lenght between the two pubis was also taken, because an aviculturist told me that he could sex all passerines in this way. I didn’t find any support for this.
The spleen, bursa of Fabricius (b.F.) and gall bladder are measured length x breadth, the two first mentioned are also weighted; thymus is now of time-consuming reasons only weighted, first all on left side of neck and then right side; however, birds frozen for a longer period are not possible to record because thymus glands are the first to dry up. The b.F. is often difficult to meassure, either because body is cut wrong at tail end, or there is too much fat of nearly same colour as b.F. (The three glands are removed with a curved tweezers Both caecae length and rectum (distance from caudal end of caecum to cloaca). The three measurements are given with a dash or comma between the numbers. Note, rectum is often only approximatly. If only one caecum, this is also mentioned, on the database with a 0 in second column.
To a study of cateract in wild birds, the diameter and thickness of both lenses are measured and the mass is taken after the yellowfish-like substans which often attached the lens is removed. In lenses with cataract the diameter of the grey area is recorded. Rarely the whole lens is milky, and collisions accidents and predator victims are mostly pale red or red. This is also recorded.
From CN 5133 and c. CNS 206 wing-, bill, tail-, and tarsus length are taken on the dead bird before preparation, wing length max. all measurements after the method described in Svensson (1984). About the degree of shrinkage in size after drying, see e.g. (Halftorn 1982). It should be banned to take max wing length on finished skins, and moult and fault bars in wings should also be prohibited to study on a dried skin because it disturb skins.
If an incubation patch is found, this is indicated on the label with length and breadth, but most birds I made have been frozen and then it is difficult to recognise an incobation patch.
The uropygial gland is measured length x breadth. If they are divided in two, only one is measured but both weighted, on the label this is recorded by adding a s to gland.
The weight is always given in grammes (g).
From about catalogue number CN 5100 and CNS 650 I have started to measure wing, bill, tail and tarsus length on the completely thawed bird before the bird was prepared. The method used is that described in Svensson (1984). Wing maximum length is used. Tarsus length are in most cases not taken on owls because it is so difficult on the fresh bird. As all measurements are made by the same person (JE), no correction factors for multiple measurers is therefore necessary.
Such birds as have been weighed by myself (and this goes for nearly all) have either been weighed with a pair of scales with an inaccuracy margin of ± 0.1 g, or with a electronic Sartorius with an accuracy of one miligram, that is, the bird was weighed before the freezing, if any took place. If the bird was not weighed until after having been frozen down in an airtight plastic bag for a considerable span of time, a note of this is added. After I have started to take DNA this is not neccessary anymore, because the DNA number is always the date the bird is prepared.
In a research with a small selection of birds (n = 9), all had been weighed before frozen and remained in cold storage (-20 ) for 5 to 351 months. All polythene bags had remained airtight and as much air as possible removed from the bags before closed. The plastic bags were, however, not of the same sort, which surely can explain some of the different results, because birds in thicker plastic naturally loose less weight and therefore this little study is only a hint without scientific importance (cf. Banks 1965).
|months||loss per month:|
|1 Hirundo rustica||19.000g/18.800g||5||0.0400g|
|1 Turdus merula||107.122g/106.886g||12||0.0197g|
|1 Accipiter nisus||141.300g/141.000g||20||0.0150g|
|1 Accipiter nisus||267.000g/265.413g||52||0.0305g|
|1 Turdus iliacus||48.300g/47.200g||52||0.0212g|
|1 Lanius collurio||28.600g/26.300g||95||0.0242g|
|1 Acrocephalus schoenobaenus||10.150g/7.900g||194||0.0116g|
|1 Oenanthe oenanthe||26.900g/23.650g||231||0.0141g|
|1 Sylvia atricapilla||13.600g/8.461g||351||0.0146g|
Note, that a bird frosen for a short time, say one to twenty days, will nearly always have a higher weight than the first recorded (own obs.).
From February 1995 all birds are weighed with a Sartorius 1608 MP. Time after time I calibrate it, and results with date are kept.
If collector is mentioned, it means that the bird has not been weighed by me. If I know the bird has been weighed with a pesola spring balance, this will be stated on the label. The finder or supplier of dead bird is not mentioned on the label, but via the six-numbered aluminium sign this is always possible to reconstruct, the same for the date of receipt. I use to put the fresh body in a plastic bag at the time it is removed from the bird, in order to avoid the breast muscles to shrink and first weighting it when the skin is finish.
Food in gizzard and possible grit are weighed together, but a note is made if grit is found, and since approx. year 2003 grit is always weighted and kept except from most captive birds. Grit is sorted out from food using water. Food in crop, esophagus and proventriculus are weighted separate The inner layer of gizzard (cuticula gastrica), which was already by Aristotle recognised to be easily removed, is also weighted. By some families, e. g. ducks it is difficult and time-consuming to remove and therefore no mass is here given. Intestine is weighed without the content in the case of the nearly thousand House Sparrows collected between 1965 and 1969. Since 2005 intestine with content but without the surrounding fat is weighted and the length of the intestine is taken when possible.
Before the gizzard and the heart are weighed, all external fat is removed. The gizzard i first weighted with the cuticula gastrica or koilin layer, and afterworth cuticula is weighted alone. The heart is excised by cutting all vessels as near as possible to their origin of heart muscle and residual blood is forced from heart by compression. Before liver, b.F. and spleen are weighed, the external blood on the liver, connective tissues and blood vessels near their connection with the organ of all are removed. Salt glans, lungs and kidney are also weighted, lungs only when not with dark blood, and then after “most” blod is removed by removing it ten times on both sides on a sucking newspaper, each time 2-3 seconds. Kidney and lungs only when easy to remove. The brain is weighed after removing it from underside of head and placing it on an already weighted absorbing paper and remaining part of the brain is sucked up with small pieces of same paper. If brain of a specimen has began to dry, it is not weighed. In cases where whole skeleton is kept for the collection, the brain is weighed using the following procedure: First a piece of absorbing paper and some cotton wool are weighed, the brain is then removed by putting wool inside the brain case through foramen with a pair of pincers, and this wool is moved around before being removed. This procedure is repeated until no more brain tissue is visible on the wool. The brain on the paper and the used and unused wool is finally weighed, and the weight of the paper and the wool before starting the extraction procedure is subtracted. This procedure may take up to five minutes.
The breastmuscles, (M. pectoralis and M. supracoracoideus) on both breast sides are weighted after they are removed from breastbone by cutting and stretching the muscles. Here an inexactitude of approx 5-10% must be incalculated, it is especially often difficult to get all muscle from furcula.
The condition of the body mass is described as follows:
Very lean or emaciated: dead or nearly dead of starvation
Lean: breast muscle shrunken,
A little lean: breast muscle a little shrunken
Normal: breast muscle normal, little fat on the skin and viscera
A little fat: a little fat covers muscle and skin but furcular area concave
Fat: fat fills furcular area, which is level or nearly so
Very fat: furcular area bulges with fat and or much fat on skin and viscera.
Tail and wings are examined for fault bars and breaks in feathers due to fault bars. Because there are great difference in fault bars they are from January 2017 divided in light, moderate and severe according to how much the feather is weakened by the fault bar: Light fault bars resemble thin lines crossing the feather vein and appear, on close inspection, as slight notch on the feather surface, due to feather malformation. Moderate fault bars consist of a conspicuous lack of keratin deposition in barbs and barbules, making the feather translucent at the fault bar. In this case the rachis often shows a clear malformation that is easy to see. Severe fault bars are seen as sections of feather vein a few mm wide or more that are free of barbules. Feather barbs often break at these fault bars, even the rachis. (Jovani & Rohwer 2016). Fault spots are transparent holes, often wide and maybe related to fault bars; they are also registered. The same with feather holes, small holes in barbs 05-1mm in diameter, which presumably are attributed to chewing lice. Pale bands on wings and tail are also noted.
If sign of sickness or other abnormalities are seen it is described as good as possible. Tumours are identified when they are clearly visible lumps of hard tissue with a diameter of more than one mm, meassured, mass and colour taken and kept in etanol. Tumours within tissues are not examined.
The time from the death to the time when colours are identified is noted as short after death or later. All descriptions of colours made more than a couple of days postmortem must be treated with some reserve, especially the colour of iris which may change postmortem.
The following colours are normally recorded: Iris, eye-ring, bill, inside upper mandible, palate, mouth, tongue, legs, (feet are only mentioned if they differ in colour), sole, claws and bare skin elsewhere.
The principle invariably adhered to is that e. g. “brown-black” is closer to black than “black-brown”; “brownish-black” is still closer to black than “brown-black”.
On expeditions and to some extent otherwise, when I rarely receive a bird just or soon after its death, a colour-guide is used to get an accurate colour. For a long time I used a Danish guide:
Kornerup, A. & J. H. Wanscher, 1962: Farver i Farver. Politikens Forlag, Copenhagen.
Later I used a German guide:
Küppers, H. 1978: DuMont’s Farben Atlas. DuMont Buchverlag, Köln.
For the last few years I have mostly used:
Smithe, F. B. 1975: Naturalist’s Color Guide. AMNH, New York.
I find the Danish one the best one, but nobody except Danes know it. It is even translated into English, 3rd. edition 1983: Methuen Handbook of Colour. Methuen, London, but is now impossible to find on the markt (I have heard due to stamp collectors). The German guide is also fine, but most English speaking people do not know it, therefore I mostly use, the poorest of them all, the American Guide which all know, but only if I can find the exact colour, otherwise I explan the difference, e.g. darker than … or I use the Danish one where all shades are shown.
Moulting study skins are frustratingly rare in most collections and mine is no exception. Where it is not possible to describe the moult in a few words on the label, a moult card has been used in some years (modified from Snow 1967: p. 3). One asterisk on the label against “Moult Card” signifies that the moulting is abnormal, e. g. asymmetric or fear-induced. Supposed right moult is now noted on a special column on the database. Primaries are counted descendent and secondaries ascendent. A time long I spread one wing, when the bird is moulting, because it is very hard to a bird-skin when a closed wing is examined. From CN 5000 nearly all birds in moult have been skeletonized, and one spread wing and tail have been kept + more characteristic body feathers. If the moulting is finished but there are still black feather follicles on skin inside this is stated, eventually also where on the skin it is found. Contour feathers on my database means all body feathers, also neck and head, but except wing- and tail feathers.
SUPPLEMENTARY COMMENTS, SAMPLES COLLECTED AND METHODS
If one or more white (albinistic) feathers is found it is always mentioned, otherwise they can easily latter be neglected, especially small feathers. If part of the skeleton is ossified due to old wounds or senescense and every sign of sickness or abnormallity are described. In a few cases it have been possible for me to make a diagnostic of the sickness using Tully et al. 2009 and Arnall & Keymer 1975). The tracheal bulla by ducks, cast still in the body, endo- and ecto parasites or food which is suitable for being dried are kept and it is mentioned on the label and database. To facilitate later determination of seed a collection of seeds from many plants in my region have been collected and sorted after size. When a bird is skinned, all loose, NOT washed feathers are kept separate, e. g. for stable isotope analyses, and a note about this is written on the label and data base.
When tongues or ectoparasites are kept, they are only dried, tongues are normally kept together with trunk skeletons or skeletons, and ectoparasites are kept separate in small plastic bags or in etanol. Eyes are first prepared in etanol and then dried. I have long ago stopped to keep the dried eye because they often are easten by pest, instead scleral ossicles is kept when not rushed away by water.
If the whole bird is washed, this is always noted with the abbreviation “Wa” but spot cleaning is not noted. I think this is an important piece of information, because presumably e. g. the fat from the uropygial glands by washing is removed and thereby gives help to feather-degrading bacterias (Burtt 2009) and I suppose it disturb any stable isotope analyses. If the bird earlier was washed a shampoo detergent was used in many years, now I prefer a common washing-up liquid in hand warm water and afterwards a woll wash liquid. I use much time to clean fat birds, e.g. 5-6 hours for a fatty duck. The fat is removed with a not sharf knife with a rounded end, and to prevent hules in the skin I use much water to keep the skin smooth under this work. Fatty birds where the end of the feather shaft is visible on skin inside (like ducks) are cut with a pair of scissors. Is a bone in wing or leg broken all marrow is removed with a wire and the broken bone is repaired with a piece of wire inside the bone.
On expeditions I have got good experience using salt on the inside of skin and bones, and letting the skin be half dried before it was placed in a tight plastic bag. On bigger birds I injected some saturated solution of salt in the legs. On places where the skin already was dried I also used this solution before salting it. This method has the disadvantage that the electroplated wire in the legs later became rusty.
About the effect of chemical on plumage colour and physical changes see: Rogers & Daley (1988).
If the ulna is stripped I have mentioned this on the label as an appeal to researcher to take extra care. Only the skins from Peru have been treated in this way by another collector in my collection.
Birds from captivity have also been collected, but only on a limited scale, because origin and date of death are rarely found. I have, however, avoided species, like e.g. bulgerigars and canaries, which have been breeding in so many generations that their origin characters are lost. Hybrids or crossings are not included in the collection with one exception, where I hoped it was a new species.
Wallace wrote about domesticated animals in 1858: “The domesticated animal… has food provided for it, is sheltered, and often confined, to guard it against the vicissitudes of the season, is carefully secured from the attacks of its natural enemies, and seldom even rears its young without human assistance. Half of its senses and faculties are quite useless, and the other half are but occasionally called into feeble exercise, while even its muscular system is only irregularly called into action.”
The skins have been treated inside with arsenic soap with c. 5-6% admixture of glycerine in order to prevent them from crumbling later on. Only a few skins have been treated with Eulane (U33), and if so it is always mentioned. In these cases I always treat the skin inside with glyserine. I only use Eulane in rare cases because nobody knows how it affects the feathers and skins over a longer period.
The catalogue number is prefixed either CN, CNS or CNF, the former for the bird-skins, the second for the skeletons, breastbones and craniums and the latter for the feather collection which normally contain one wing and eventually tail and some distinctive contour feathers. As nearly all items from the skeleton collection have one spread wing, the loose tail feathers and some body feathers kept in the feather collection, they have only a CNS number also in the feather collection. Attached the labels is for the protected Danish birds and all those listed on CITES a small aluminium sign with consecutive six-figure numbers, and in the catalogue and data base the six-figure number also figure. This is the register number from the Danish Ministry of the Environment. From 2013 all birds are rigistered with an aluminium number except a few Danish game birds.
Old skins, i.e. before catalogue number 1300, have as a minimum the following information: sex, date, locality, weight, size of the gonads. Very often also the weight of the food is given.
From catalog number 1300 and onwards to 4080 the breastbone with furcula, coracoid and scapula usually follow the skin.
From catalogue number 4080 and onwards the trunk skeleton is usually kept with the catalogue number to the bird in question attached.
The last years the following minimum informations are reported:
Catalogue number, scientific name, sex, age, date locality. mass, statement about the condition of the bird, cause of death, gonads size and mass, weight of food and grit, weight of food in crop, brain, gizzard, heart, lungs liver, kidney, bursa of Fabricius, thymus, spleen, intestine, M. pectoralis, M. supracoracoideus, uropygial gland. Endo- and ectoparasites are kept, moulting state, wingspan, wing area, wing length, bill length, hight, wide, tail length and tarsus + colours when not too old.
A statement is given for the skull ossification and a drawing of the “windows” are often given. Notes about if grit is kept and the mass, food contents and if kept, fault bars and parasites and if kept, known age, captivity and if the bird is washed.
Uropygial gland is measured and mass taken. If the gland is divided into two glands, it is written glands and only one is measured, but both are weighted.
Ringing recoveries are mentioned with all information obtained. Also if the age of captive birds is known.
From catalogue number CN 5509 and CNS 779 DNA tissue is taken, always from breast muscle.
THE OSTEOLOGICAL COLLECTION
After I had read (Wood et al. 1982). and realized that at a rough estimate one-sixth of all bird species of the world were not represented in any museum by a skeleton, I immediately started collecting bird skeletons given skeletons so high a priority that already the second species of a given species – whether perfect and beautiful or not – is skeletonized! And not without success:
So far (1995) a total of 40 skeletal specimens or sections of specimens are not represented in any other museum collection according to Wood, D.S. et al. They are : 12 skeletons in 8 species. 18 trunk skeletons from 14 species and 20 breastbone (with coracoid, scapula and furcula) from 15 species. Skeletons or derivative thereof of which only a total of one to five comprising until now 24 skeletons, 64 trunk skeletons and 110 breastbones; those of which six to ten are found in other museums, viz. 21 skeletons, 47 trunk skeletons and 78 breastbones. This listing is stopped from 1995.
Many skeletons in my collection are those of captive pet birds, which very often represent species that are difficult to obtain in the wild and are accordingly useful for many purposes e.g. identification and study of age changes (The ages of more than 300 skinds and skeletons are known in my collection). Raikov (1985) wrote: ” Museums often obtain specimens from zoos, aviaries, or private menageries. These generally lack age or locality data and may therefore be worthless for geographic or alpha-taxonomic work, yet they are perfectly good for many anatomical and phylogenetic studies”.
More than one fifth of the skeletons have been cleaned by dermestid beetles outside my house. Some have been cleaned by mealworms which at a small stage does the work nearly as well as the dermestid beetles, but there is a lot of work finding all the small bones, if you donot come just at the right moment, and the mealworms are only small for at short time. Therefore I now use maceration which have the advantage that the bones mostly become fat-free. To avoid the stench I have them in plastic boxes which are chosed tight as soon as they start to smell. When they are ripe I remove the rotten items with water. Afterwards I give the skeleton a short partboiling with some hydrogen peroxide added to the water, and finish with washing the skeleton in clean water.
The data taken are the same as for the skins.
“There isn’t one of us who hasn’t at some time or another, missidentified a bird. If a bird is wrong identified and then skinned, it is only a matter of time before someone corrects the error. If a bird has been missidentified and then prepared as a skeleton, it is entirely possible that the mistake may never be realized, the consequences of which are nightmarish. Bear in mind that, in general, the bird that are most likely to be missidentified whole are precisely the ones whose missidentification is least likedly to be detected as skeletons” (Matthiesen 1989).
In order to avoid later researchers failing to check the species determination, wing- and tail feathers and some of the more distinguished smaller contour feathers are kept in archival plastic sheets. The last many years one spread wing, the whole tail and some distinctive body feathers have been kept as a voucher to each skeleton.
THE FEATHER COLLECTION
Very often a salvaged birds is found which is too bad for skin preparation, and the skeleton damaged. Instead of discart such a cadaver I often use it for the feather collection if I have date and locality and at least one wing and tail are undamaged and I can get some other information. As already written, the feather collection contain also feathers from nearly all in the skeletal collection. This collection is not thought as an ornamental collection, though many feathers are very beautiful, first of all it is thought as a tool to check the species and subspecies determination of the skeletons, and second for many research, where the skins can be spared. They are all kept in thick plastic bags.
The data taken are the same as for the skins and skeletons.
The collection await imaginative approaches to the study using current technologies as well as the technologies still to be discovered. New insight may be gleaned when thise data are examined with fresh eyes and minds.
Due to the complicated rules nowadays I donot lend any bird skin or skeleton anymore, but researchers are wellcome to come and study the collection.
Anderson, S. 1965: Systematic Zoology 14:344-346.
Arnall, L. & I. F. Keymer. 1975: Bird Diseases. T. F. H. Publications, Nepture City, N. J.
Bangs, R. C. 1965: Weight change in frozen specimens. Journal of Mammalogy 46 (1):110.
Burtt, E. H. Jr. 2009: A future with feather-degrading bacteria. Avian Biol. 40: 349-351.
Erritzöe, J. 1985: Geschlechts- und Altersbestimmung bei Vögeln. Der Präparator 31 (2):81-93.
Halftorn, S. 1982: Variation in body measurements of the Willow Tit Parus montanus, together with a method for sexing live birds and data on the degree of shrinkage in size after skinning. Fauna norv. C. Cinclus 5:16-26.
Jovani, R. & S. Rohwer. 2016: Fault bars in bird feathers: Mechanisms, and ecological and evolutionary causes and consequences. Biological Reviews: April 2016.
Matthiesen, D. G. 1989: The curation of Avian Osteological Collections, p. 71-110. in:Rogers, S. P. & D. S. Wood: Notes from a Workshop on Bird Specimen Preparation held at The Carnegie Museum of Natural History in Conjunction with the 107th Stated Meeting of the American Ornithologist’s Union. Carnegie Mus. Nat. Hist., Pittsburgh.
Raikov, R. J. 1985: Museum collections, comparative anatomy and the study of phylogeny. in: E. H. Miller (ed.) Museum collections: Their roles and future in biological research. Occ. Papers, Brit. Columbia Provincial Museum No. 25.
Rogers, S. P. & K. Daley, 1988: The effect of preparation and preservation chemicals on plumage color and condition. Carnegie Museum Nat. Hist., Pittsburgh.
Snow, D. W. 1967: A Guide to Moult in British Birds. BTO Field Guide No. 11.
Svensson, L. 1984: Identification Guide to European Passerine. Lars Svensson, Stockholm.
Tully, T. N. Jr., G. M. Dorrestein & A. K. Jones. 2009: Avian Medicine, Second Edition. Saunders & Elsevier, Edinburgh.
Wood, D. S., R. L. Zusi, & M. A. Jenkinson. 1982: Inventory of avian skeletal specimens. Amer. Ornithol.s’ Union and Oklahoma Biol. Survey, Norman,Oklahoma.
Zink, R. M. & J. V. Remsen Jr. 1986: Evolutionary processes and patterns of geographic variation in birds. Current Orn. 4:1-69
LIST OF ALL PAPERS BASED ON THIS COLLECTION
Erritzöe, J. 1985: Geschlechts- und Altersbestimmung bei Vögeln. Der Präparator 31:81-93.
Erritzøe, J. 1990: Einflügelige und flügellose Vögel. Natur und Museum, Frankfurt a M. 120 (1):10-15.
Erritzoe, J. 1993: The Birds of CITES and How to Identify Them. Lutterworth, Cambridge.
Erritzoe, J. 1994: First record of a Dunlin from the Philippines. Bull. Brit. Orn. Club. 114 (2):128-129.
Erritzoe, J. 1995:A small collection of birds from the Philippines with notes on body mass, distribution, and habitat. Nemouria, 40 Delaware Mus. Nat. Hist., Delaware.
Erritzoe, J. 1996: Den almindelige gråspurv er minsandten også spændende.
Hjejlen 14 (4): 5-8 & Panurus 31 (2): 6-9. (In Danish).
Erritzoe, J.1998: The Birds og CITES and How to Identify Them.
Chinese Wild Bird Federation, Taiwan. (In Chinese).
Erritzoe, J. 1998: Body masses(weights) of parrots. Avicultural Magazine 104 (1):27-33.
Erritzoe, J. 1999: Causes of Mortality in the Long-eared Owl. Dansk Ornitol.Forenings Tidskrift 93 1999: 162-164.
Erritzoe. J. 2002: Mauersegler Apus apus mit asymmetrisch wachsenden Flügel- und Schwanzfedern. Ornithol. Mitt. 54 (5): 159-160.
Erritzoe, J. 2003: Family Pittidae (Pittas). Pp. 106-160 in: del Hoyo, J., A. Elliott & D. A. Christie (eds.): Handbook of the Birds of the World. Vol. 8. Broadbills to Tapaculos. Lynx Edicions, Barcelona.
Erritzoe, J. & H. B. Erritzoe. 1998: Pittas of the World. Lutterworth, Cambridge.
Erritzoe, J. & R. Fuller. 1999: Sex differences in winter distribution of Long-eared Owls Asio otus in Denmark and neighbouring countries. Die Vogelwarte 40 1999: 80-87.
Erritzøe, J., C. F. Mann, F. P. Brammer & R. A. Fuller. 2012: Cuckoos of the World. Christopher Helm, London
Erritzøe, J. & H. Svenningsen, 1996: Dark-eyed Junco in Denmark 1980 and review of records from Europe and Greenland. Dutch Birding 18 (1):1-5.
Galvan, I., A. P. Møller & J. Erritzøe. 2011: Testicular melanization has evolved in birds with high mtDNA mutation rates. J. Evol. Biol.: 1-11. (pdf)
Galván, I., J. Erritzøe, K. Wakamatsu & A. P. Møller. 2012: High prevalence of cataract in birds with pheomelanin-based coloration. Comp. Biocehm. Physiol. A 162: 259-264.
Galván I., J. Erritzøe, F. Karadas & A. P. Møller. 2012:
High levels of liver antioxidants are associated with life-history strategies characteristic of slow growth and high survival rates in birds J. Comp. Physiol. B 182: 947-959.
Galván, I., J. Erritzøe, K. Wakamatsu & A. P. Møller. 2012: High prevalence of cataract in birds with pheomelanin-based coloration. Comp. Biocehm. Physiol. A 162: 259-264.
Galván, I. A. Naudi, J. Erritzøe, A. P. Møller, G. Barja & R. Pamplona. 2015: Long lifespan have evolved with long and monounsaturated fatty acids in birds. Evolution 69-10: 2776-2784. (Article in PDF)
Garamszegi, L. Z., A. P. Møller, & J. Erritzøe, 2003:The Evolution of immune defense and song complexity in birds. Evolution 57 (4): 905-912. pdf (92 KB)
Garamszegi, L. Z., M. Eens, J. Erritzoe, & A. P. Møller. 2005: Sperm competition and sexually size dimorphism in birds. Proc. R. Soc. B. 272: 159-166.
Garamszegi, L. Z., M. Eens, J. Erritzoe, & A. P. Møller. 2005: Sexually size dimorphic brains and song complexity in passerine birds. Behav. Ecol. 16:335-345.
Garamszegi, L. Z., J. Erritzøe & A. P. Møller. 2007: Feeding innovations and parasitism in birds. Biol. J. Linn. Soc. 90: 441-455. (Article in pdf)
Garamszegi, L. Z., A. P. Møller, & J. Erritzoe. 2012: Coevolving avian eye size and brain size in relation to prey capture and nocturnality Proc. Royal Soc. London B. 269: 961-967.
Møller, A. P., Ph. Christe, J. Erritzøe, & J. Mavarez, 1998: Condition, disease and immune defence. Oikos 83:301-306.
Møller, A. P., R. Dufva, & J. Erritzøe, 1998: Host immune function and sexual selection in birds. J. Evol. Biol. 11: 703-719.
Møller, A. P. & J. Erritzoe 1988: Badge, body and testes size in House Sparrows, Passer domesticus. Ornis Scandinavia 19 (1): 72-73
Møller, A. P. & J. Erritzoe 1992: Acquisition of breeding coloration depends on badge size in male house sparrows Passer domesticus. Behav. Ecol. Sociobiol. 31: 271-277.
Møller, A. P. & J. Erritzøe, 1996: Parasite virulence and host immune defence: Host immune response is related to nest reuse in birds. Evolution 50 (5):2066-2072.
Møller, A. P. & J. Erritzøe. 1998: Host immune defence and migrating in birds.
Evolutionary Ecology 12: 945-953.
Møller, A. P. & J. Erritzøe. 2000: Predation against birds with low immunocompetence. Oecologia 122: 500-504.
Møller, A. P. & J. Erritzoe 2001: Dispersal, vaccination and regression of immune defence organs. Ecological Letters 4: 484-490.
Møller, A. P. & J. Erritzoe 2002: Coevolution of host immune defence and parasite-mediated mortality: Relative spleen size and mortality in altricial birds. Oikos 99: 95-100.
Møller, A. P. & J. Erritzoe, 2003: Climate, body condition and spleen size in birds. Oecologia 137: 621-626.
Møller, A. P. & J. Erritzøe. 2009: Why birds eat colourful grit: colour preferences revealed by the colour of gizzard stones. J. Evol. Biol. 23 (2010): 509-517.
Møller, A. P. & J. Erritzøe. 2010: Flight distance and eye size in birds. Ethology 116: 458-465.
Møller; A. P. & J. Erritzøe. 2014:
Predator–prey interactions, flight initiation distance and brain size J. Evol. Biol. 27: 34-42.
Møller, A. P. & J. Erritzøe. 2015:
Brain size and urbanization in birds. Avian Research 6:8-14.
Møller, A. P. & J. Erritzøe. 2016:
Brain size and the risk of getting shot. Biol. Lett. 12: 20160647
Møller, A. P., H. Erritzøe & J. Erritzøe. 2011: A behavioral ecology approach to traffic accidents: Interspecific variation in causes of traffic casualties among birds. Zool. Research 32 (2): 1-13.
Møller, A. P., J. Erritzøe, & L. Z. Garamszegi, 2005: Covariation between brain size and immunity in birds: Implications for brain size evolution. J. Evol. Biol. 18:223-237
Møller, A. P., J. Erritzøe & F. Karadas. 2010: Levels of antioxidants in rural and urban birds and their consequences. Oecologia 163: 35-45. pdf
Møller, A. P., J. Erritzøe, F. Karadas & T. A. Mousseau. 2010: Historical mutation rates predict susceptibility to radiation in Chernobyl birds. J. Evol. Biol. 23: 2132-2142.
Møller, A. P., J. Erritzøe & J. T. Nielsen. 2009: Frequency of fault bars in feathers of birds and susceptibility to predation. Biol. J. Linnean Soc. 97:334-345. pdf
Møller, A. P., J. Erritzøe & J. T. Nielsen. 2010: Causes of interspecific variation in susceptibility to cat predation on birds. Chinese Birds 1 (2): 97-111.
Møller, A. P., J. Erritzøe & J. T. Nielsen 2010: Predators and microorganisms of prey: Goshawks prefer prey with small uropygial glands. Functional Ecology 24: 608-613. pdf.
Møller, A. P., J. Erritzøe & L. Rózsa. 2010:. O Ectoparasites, uropygial glands and hatching success in birds Oecologia 163: 303-311.
Møller, A. P., J. Erritzøe, & N. Saino, 2003: Seasonal changes in immune response and parasite impact on hosts. American Naturalist 161 (4): 657-671
Møller, A. P., P-Y, Henry & J. Erritzøe. 2000: The evolution of song repertoires and immune defence in birds. Proc. R. Soc. London B 267: 165-169
Møller, A. P., R. T. Kimball, & J. Erritzøe, 1996: Sexual ornamentation, condition, and immune defence in the house sparrow Passer domesticus. Behav. Ecol. Sociobiol. 39:317-322.
Møller, A. P., J. T. Nielsen & J. Erritzøe. 2006: Losing the last feather: feather loss as an antipredator adaptation in birds. Behav. Ecol. Sept.2006:1-11
Møller, A. P., G. Sorci, & J. Erritzøe, 1998: Sexual dimorphism in immune defence. American Naturalist 152 (4): 605-619.
Møller; A. P. & J. Erritzøe. 2014:
Predator–prey interactions, flight initiation distance and brain size J. Evol. Biol. 27: 34-42.
Møller, A. P. & J. Erritzøe. 2015:
Brain size and urbanization in birds. Avian Research 6:8-14.
Vágási, C. I., P. L. Pap, O. Vincze, G. Osváth, J. Erritzøe & A. P. Møller. 2016: Morphological adaptations to migration in birds. Evol. Biol. 43: 48-59. (Read article in PDF)
COLLECTION MANAGEMENT GUIDELINES
The computerization of the collection
From the very beginning in 1947 all data concerning the bird-skin collection were written into catalogues. As the collection grew it became obvious that this system alone was not satisfactory for many reasons, e.g. the storage of data relevant to natural history museum collections using computer aids, the task of collection management, and enhanced research uses of the materials.
In 1993 I decided to carry out this computer cataloguing before the collection had grown to the point where this work would have been a major undertaking. It took four months of data entry before computerization was completed. I had every skin, breastbone, trunk skeleton or skeleton in my hand to obviate any mistake. However, in this database only the following data were recorded: Number, scientific name, month, country where collected, and at the end different codes for DNA kept, captivity, feathers kept, trunk skeleton kept etc.
However, I soon realised that his database only gave a list of the birds and the other items stored, just like the catalogues, but here sorted after families and not after the date of preparation, and therefore of less use for researchers, who wanted to use all the information laid down in the collection. Therefore I soon started a new database in Excel, where all the about 200 information now taken each became a column. This work was finished in August 2013 with 10.800 entries and will now be updated each time new specimens are added to the collection.
The temperature where the collection is kept in our house is not the best: in winters about 5-10 (42-50 F.) and in summers up to 30 (87 F.), but the relative humidity is rather constant between 70 and 80.
The light is normal, but most of the skins are stored in wooden cabinets where no light and very little dust can get in. The rest is placed in boxes. Each bird is stored in a closed plastic bag.
The gradual incorporation of new specimens in the collection
The time between the skinning the bird and until it is quite dry is the most dangerous period as to insect pest attacks. I try to cope with that problem in the following way:
The finished skin lay on its bag on a expanded polystyrene a few days, fastened with pins crossed to the head- and tail wire; thereafter hung by its head wire in the laboratory for other three to five days according to size. It is enough to dry the feathers in the wanted position so the skin can be laid down on its back without disturbing the pattern of the feathers. Then – not only to protect it from the attack of pests but also to safeguard it against dust – it is put in a cardboard box in my office to finish its drying there. Such a box is not airtight, but dust, moths and beetles do not so easily find their way to the skins. (If I had a large cold store I would dry the skins there).
When the skin is quite dry, (with the temperature and humidity in our house it takes c. 3 weeks for a passerine and c. 5 weeks for a limicoline), and the label has been written, the skin is put in a closed plastic bag and placed in a deep freezer (-20 ) for at least 3 days to kill possible unwanted live animals. Afterwards the skin is placed in isolation in a cardboard box again for the next month, still in its plastic bag, to give possible eggs (which will not be killed at -20 ) the time to develop. Then the procedure in the deep freezer is repeated before the skin at last is placed in the collection.
All the bird-skins (and skeletons) are kept in plastic bags not thinner than 0.003mm. I have used plastic bags from about 1965 and onwards without any problems except that in late years some of the earlier plastic bags have got a slightly sticky surfaces without any visible effect on the skins or labels. From time to time such plastic bags are found and replaced. I can therefore heartily recommend the use of plastic bags in a collection like mine not kept in expensive airtight metal skin cabinets but only in wooden cabinets. By the way, BMNH now use vacuum closed plastic bags for their types.
Insect pest control
The most acceptable level of pests in a bird-skin collection is zero, but I know no curator who has achieved this goal, and I cannot put forward the solution of this severe problem either, but only state what method I use with a reasonable good result:
Because of the small number – (7.000) of bird-skins, c. (4,000) breastbones, c. (3.000) trunk skeletons) and (1200) skeletons – in my collection (Anno 2013) every skin and skeleton is monitored each year. Owing to the plastic bags the affected part is very easy to find and I have never found more than one skin attacked in the same drawer. The pest problem has not until now been serious in the collection. On average two or three skins are found every year attacked by dermestid beetles Dermestes lardarius, never clothes moths Tineola bisselliella. I think moths cannot bite through a plastic bag. Once another beetle Ptinus tectus was found in the collection without having done any harm to the skins yet. They were never found again. The last two years I have not found any pest attack.
All cabinet door frames are painted once every year with cockroach poison, “Mortalin” a Danish product, containing e.g. 2.5% chlorpyriphos and 1.5% xylene, and it seems to be a helpful agent, because dead beetles are found nearly every year just inside the doors. (If you touch the poisoned door frame a disagreeable prickling starts immediately at the point of contact).
No other chemical control is used. I have never used paradichlorobenzene and naphthalene, because I once saw a drawer full of naphthalene flakes around the skins among hundreds of live moths!
The dermestid beetles are commonly found every summer in our garden or even seen entering the house. The Museum beetle Anthrenus musaeorum has never been found in the house. We also sometimes find clothes moths. I try to remove dead young birds and mice from under the roof, because their smell attracts the beetles, and in summer doors and windows are only open to a very small extent.
The label I use is made of a fine quality glazed and thick (0.24 mm) cardboard. In many collections you can see how old thin labels bend and make it difficult to place the skins well. The paper of the labels is in two colours: pale blue-grey for the male and pale grey for the female. If you have to sort a large series of a species without sexual dimorphism, the colour difference on the labels makes it easier to sort the sexes quickly. For the text I used permanent, waterproof black Indian ink (Faber-Castell, Germany). Due to problems with continuously constipation of the pen I have the last year used “Artline 853 with permanent and water-resistant ink. However, it make too thick a line. Now I use Micron 005 with good result. If there is any doubt about the identification of the bird in question, the scientific name is written provisionally with a pencil.
Most labels I have seen in other collections have only a few items of information printed: the name of the museum or private collector, number. or catalogue number and even date of registration! Because of human forgetfulness I have left space for many particulars, each printed on a separate line or box. This makes it also easier to sort out a special class of information if you can always find it in the same place on the label.
From the very beginning of the history of bird-skin collecting it was usual to make the labels as small as possible, e.g. for a hummingbird smaller than the body of the bird and otherwise the normal size 5.5 x 1.3 cm to 6.0 x 1.8 cm. The reason was said to be that a larger label would be likely to cause serious damage to the skin and make the weight heavier. Owing to the many particulars, my label is 10.0 x 4.3 cm and for each bird I use two different labels, sometimes three, the last is blank and in spite of this I have never observed serious damage to a skin caused by a label except cases where some feathers have been bent a little on hind neck but easily repaired over hot water-steam. The weight of two of my labels is 1.578 g, compared with the 0.3 to 0.8 g of a normal museum label. I think this difference will only affect the freight rate if many hundreds skins have to be sent.
Several curators have told me that if there is more information available than a label can contain, the researcher only has to study the field notebook. How much information will never be used because the researcher on a museum visit does not have the time to read this field notebook, forgets it, does not know about it, or cannot find the right page in a hurry? Attached labels are therefore after my opineon the most important and most used documents for data retrieval. For me it is all the same whether a skin has one, two, three or ten labels. All information must only be found by the skin.
In 1940 Alden Miller wrote in Museum News 17 (17):6: “The original label written when the animal is taken and prepared is a scientific document. It must never be destroyed or replaced and the essential data it is to bear must be entered at the time, not later. The practice of writing temporary labels is pernicious in the extreme”.
In spite of these very clear words I must confess that I write temporary labels on normal papers which I afterwards keep in files with the catalogue number of the bird on the top. The reasons for this procedure are several: I prefer to finish the skins, study books and papers and take the measurements of femur, humerus and keel on the dried trunk skeleton in my office where it is cleaner and safer for such work. But of course, if an old label, e.g. with text in Danish, is rewritten, the old one always remains on the skin.
As twine I formally used a normal sewing thread but now I use a black nylon thread, (note, here 4-5 knots must be made to fasten the label) and because the label paper is 0.24 mm thick I have not found it necessary to make an expensive metal eye-hole to secure the attachment to the skin.
The trunk skeleton
The trunk skeleton receives the catalogue number corresponding to that of the skin. It is written on one side of the breastbone with the same waterproof black Indian Ink as used on the label. I take care not to write in a place where it will disturb interesting features. In later years I have only put a number on cardboard with the trunk skeleton into the plastic bag.
The trunk skeletons are kept in plastic bags in boxes in another place in the house as the skin collection, with the catalogue numbers written on the outside of the box. On the label of the bird-skin in question it is mentioned if the trunk skeleton is kept, but the trunk skeleton has no separate label. Together with the trunk skeleton other availably remains may be kept, e.g. eyes and tongue. Food, grit, parasites, cast and loose feathers are, however, kept separate.
The skeletons are also kept in plastic bags in boxes after families, with the catalogue numbers beginning with CNS written on the labels. For the quick check I have a copy of Grusons Checklist of the Birds of the World at hand, where the catalogue number of every skeleton for a quick overwiew is written. On the label of the skeleton an “F.” is written if the feathers and one wing are kept on the early skeletons, but now it is the standard procedure for all birds except for very dirty ones. The one spread wing and loose feathers are kept in plastic folders Din A4 for the smaller ones, which are closed airtight and kept in files in systematic order, but within the family alphabetically. A label in each plastic folder displays the scientific name, sex, catalogue number and all other information.
Items in alcohol
In a private house without extra fire prevention it is not wise to store more alcohol specimens than necessary. Therefore only abnormal organs or organs for which I am not quite sure of my determination + especially endoparasates are kept. The number on the alcohol preparation is the same as on the label of the bird-skin or skeleton in question.
In the following methods and techniques not found in the litterature I know about is reportet. A reference list of some of the most important books and papers consulted is given at the end.
Preventing old skins from crumbling
It is a known fact that old museum-skins often crumble. To prevent this I use 3-5% clean glycerine in the arsenic for the following reason: Many years ago I received some very old and badly made bird-skins from Italy. I tried to prepare them once more and was very surprised to find how easy it was to get these skins soft and flexible again after a few days in salt water. Otherwise it is well-known to be very difficult to restore old bird-skins, because the skins are so crumbly and inflexible. When asked my Italian supplier explained that taxidermists in Italy had always mixed some glycerine in the arsenic.
In my collection I have some old skins from C. J. Aagaard’s collection:
Halcyon smyrnensis 20 Nov. 1930. CN 3903
Coracias benghalensis 18 Nov. 1930. CN 3902
Irena puella 20 Feb. 1929. CN 3906
Padda oryzivora 15 Oct. 1930 CN 3907
They were all crumbling and very dry. If you pressed a finger on the skin, it made a crackling sound. I suppose in a hundred years or less these skins would have broken into small pieces like many old skins I have found in museum collections.
I restored them all in a simple way: Some clean glycerine was injected in the head and body. To secure the feathers the skin was afterwards wrapped up in blotting paper for 2-3 months. Now, 25 years later, it has caused no disadvantages, and the skins are still soft.
I think a skin of a bird is comparable with mammal-skin, used, e.g. for book-binding. If you do not give tanned leather of a book some leather oil or shoe polish from time to time (I do it every ten years) after some years you will find that the leather starts crumpling.
The use of potato flour
Potato flour is in common use among taxidermists in Europe when skinning birds to avoid e.g. blood dirtying the feathers, and to absorb fat.
Putting the flour on the wet feathers in connection with a hair dryer is a quick method to give the feathers their “life” back again. But potato flour has one disadvantage: some flour always remains in the feathers afterwards as will appear through a microscope. On black birds it is even easy to see without any magnification. That is the reason why I only use potato flour sparingly when skinning and NEVER to dry a washed bird. It is a little more time consuming to use a hair drier only but using the warm air with much care it is only a few minutes more work for a small bird.
Wood flour is difficult to get out of the down again, especially on owls and ducks. I have also tried to use magnesium carbonate, but prefer potato flour. However, when I make black birds and birds with glossy colours I use blotting paper instead of flour. It works well.
Nearly all birds in temperate zones have some fat on the skin. If all is not 100% removed, the fat will – as every curator knows- sooner or later move outside to the feathers and give the skin sticky yellow feathers. I have tried lots of different methods to remove or neutralize this fat, but all in wain. The only way which I can recommend is the labour-intensive way to remove it with a rounded and not sharp scalpel. If water is used on the place of the skin where you work, it is easier to prevent to make holes. Potato flour and a wash in hot wateris also very useful to take the last fat. In bird familes where the feather shafts are visible on the skin inside I use a pair of scisors to cut the feather follicles of, but never if they are black. Are some bone with marrow broken it is very important to emty these, because the marrow otherwise will move to the feathers , with the same result as fat. Of this reason I prefer always to cut the bones in their joints. Feather tracts very often hide fat not visible for the human eye. Even a starving bird can retain a little fat there. In the last years I have in cases with black feather folliches often used to unwind the artificial body and neck with strong sucking toilet paper, in the hope that fat will choice to move inside instead of outside. The most difficult place on very fat birds are the wing joints. However, unimportant how careful I have removed the fat, I often later recognise fat on the feathers, so I can not claim to have solved the problem.-
I prefer not to wash skins, because it makes state of feathers impossible and when washes it is noted on the label, however not if only partiel. If one takes a look in museum skin collections, the drawers are nearly always dirty from the skins. This make a effective pest control more difficult and makes often the study of skins rather grubby. To prevent this I always use a hair dryer before I arrange the plumage. It gives the feather more life and remove most dirt and potato flour.
In order not to ruin the nostrils and damage the operculum a thread through the nostrils has never been used, but the bill has been closed with a pin from between the gonys to inside the upper mandible. With seed-eating passerine this is often difficult, and in such cases I use a piece of clay on the tip of the closed bill, . It works, because clay dries before the bill begins to work and is easy to remove later when bill is fixed. Another thing is that it is sometimes necessary to stop the blood in the nostrils with a little piece of cotton wool. In these cases the operculum will naturally be spoiled.
Using wire in bird-skins
Using wire in study-skins is not a brilliant new idea. It has already been used in mounting birds and mammals for the last two hundreds years, as you surely know. So I will not give a long explanation of this method, which can be found in every taxidermy-book, but only describe the difference between the normal procedure of mounting a bird and my way to make a skin:
The first condition for using wire in stuffed or skinned birds is the use of a material for the artificial body that is firm yet capable of being pierced. Cotton wool does not have this quality. I use fine wood shavings (excelsior), but balsa wood or cork can also be used and has the advantage that you can carve it. I prefer wood shavings, because it is the cleanest material when you soray it with water from an atomizer. It is also the cheapest. Even with wood shavings it is possible to form the moist wood shavings in such a way that, e.g. the notch on the front of the bird-body is perfect! (Important in owls and bird of prey).
For bird-skins I use a thinner wire than if the bird is stuffed, on average one number thinner. At both ends of the bird the wire ends in an eye to prevent any damage and to facilitate hanging the bird to dry and later handling the skin.
As I now keep both femurs and humeri on the trunk skeleton I usually insert the leg-wire in the artificial body in the same place as the proximal end of femur, i.e. on the side of pelvis, and let the wire have about the same length as the femur before I bend it. That is a very easy and fast method and one of the most important secrets if one wants to make a nice skin where all the feathers are placed correctly. This method allows you also to give the legs just the position you want and you can be sure the legs are in an anatomically correct position! Another advantage: with the tarsus and toes free of the feathers no fat from the legs will later soil under tail-coverts, tail and label. On large birds I use to tie the artificial femur with a thinner wire through the body to give the legs more stability.
For a skin with one wing spread, and for closed wings of birds from the size of a jay and larger, a thin wire is used. It is pressed as far out between the finger bones as possible without splitting the outermost primaries. The other end is wired around the humerus before the rest of the wire is thrusted through the artificial body. If both humeri are kept on the trunk skeleton it is easy to make an artificial humerus only using the wire: after having pushed the wire out into the finger bones, make the innermost secondaries free and wind the wire around the end of ulna and radius. Take the length of the humerus and bend the wire back again to the end of radius and ulna, bend it then once again, so that the artificial humerus is now three lengths of wire. Then wrap a little cotton around the humerus. A small but important tip if one or both wings are spread: the skin of the front wing (prepatagium) must be secured to the body with a pin, otherwise it is not possible to place the feathers in a correct position. Now I fasten the wings of smaller birds using the american method tieing the two ulna or radius together inside the body.
If a bird is crested it is usually prepare with its head turned to one side so that the crest is conspicuous.
All birds collected before they were protected, were, when wounded or winged killed by thoracic compression.
My experience with sex determination
The paired testes are found in the body cavity just ventral to the anterior end of the kidneys. In breeding they are large, most often asymmetrical. The paired suprarenal glands are visible at the end of the kidney, they are mostly of the same whitish colour as testes but where testes stand distinctly separate from the kidney, the suprarenal glands are more ingrown and symmetrical. In the full grown testes the vascular system is visible on the surface, and under both testes is a sprongy mass called the epididymis. The form of the testes is ellipsoid, only rarely are other shapes such as triangular, seen, but such abnormal forms are mostly found in very old birds. The right testis is often less than left, and very often more rounded.
One testis (or the second testis so small that it is not visible) is rare. I have found it in the following species:
Gavia immer, 2 cases (n. n.), Phoenicopterus minor (n. n.), Buteo buteo
(Alcohol No. 23), Scolopax rusticola (n. n.), Platycercus venustus (CN 4276), Trichoglossus haematodus (CNS 339), Platycercus venustus (CN 4276). Upupa epops (CN 4577), Pitta guajana (CN 4394), Turdus merula (n. n.), Pseudoleistes virescens (CN 4675), Coccothraustes coccothraustes (216175). In all cases (n = 13) except 3 the right testis was missing or microscopic. n. n. = no number and not in the collection.
Three testes is very rare: Forpus passerinus (Alcohol No. 109) and Emblema guttata (CNS 422).
The colour of the testes is NORMALLY creamy-white, pale pinkish white to more rarely pale buffish or pale grey-brown. In breeding the colour often changes into yellow or orange, but that is also seen outside the breeding season. Dark grey-brown, dark grey-green, dark red-brown, olive and black testes can be found, most commenly outside the breeding season, but also in half- or full grown testes, e.g. Alisterus chloropterus (Alcohol no. 6), which is contrary to King & McLelland (1981, p.8). Dark testes are very common in some genera or families, e.g. Turdidae and Sturnidae, in others rare or absent, e.g. Paridae and Ploceidae. In some families discoloured testes are common among some species, rare or absent in other species e.g. in Psittacidae, Strigidae and Sylvidae. In the following overview all dark grey-brown, dark red-brown, dark grey-green to dusky, blackish-brown and totally black testes figure as dark . Testes only dark in one end are also for space-saving reasons classified as dark.
ABBREVIATION USED FOR THE NORMAL WHITE AND DARK TESTES
b = black, bb = blackish-brown, br = brown, d = dull, g = dark grey, gb = dark grey-brown, gg = dark grey-green, grb = dark grey red-brown, l. = left, n = normal white or pale coloured, r. = right, rb = dark red-brown (i. e.
l.b r.n means: left testis black and right normal).
The following has not been updated since 2000:
NORMAL WHITE AND DARK TESTES
Struthio camelus 1n
Rhea americana 1gg, 1 b
Crypturellus tataupa 1n
Gavia arctica 1 rb, G. immer 2 b, G. stallata 3 n
Podiceps cristatus 1n, Tachybaotus ruficollis 1: l.b, r. n.
Puffinus gravis 1grb
Hydrobates pelagicus 2n, Oceanodroma leucorrhoa 2n
Phalacrocorax carbo 4n
Ardea cinerea 3n, 3b. Butorides striatus 1n, Ixobrychus sinensis 1n.
Scopus umbretta 1n
Ciconia ciconia 1gg
Phoenicopterus ruber 3n, P. minor 1n
ANATIDAE (55 normal, 5 dark)
Anas acuta 1n, A. clypeata 2n, A. crecca 5n, A. penelope 2n, A. platyrhynchos 7n, Anser anser 1n, Aythya nyroca 1n, A. ferina 1n, Branta bernicla 1n, B. leucopsis 1n, Bucephala clangula 3n, 1rb, Clangula hyemalis 7n, 1rb, Cygnus olor 2n, Melanitta nigra 2n, 1rb, Mergus _ain merganser 2n, M. serrator 2n, Mergus cucullatus 1n, Oryura leucocephala 1n, Somateria mollissima 10n, 1 gb, S. spectabilis 2n, 1:r.rb,l.n
ACCIPITRIDAE (114 normal, 4 dark)
Milvus milvus 1n, Accipiter gentilis 8n, A. nisus 50n, 2 gb, Buteo buteo 53n, 1gb, 1gg, B. lagopus 1n, Circus aeruginosus 1n, C. cyaneus 1n, Pernis apivorus 2n
Falco tinnunculus 13n, F. ardosiaceus 1n
Lagopus lagopus 2n, L. mutus 1b, Lyrurus tetrix 1n, Tetrao urogallus 1b, Tetrastes bonasia 4n, 2d, 2b
PHASIANIDAE (44 normal, 39 dark)
Alectoris barbara 1n, A. philbyi 1n, A. rufa 1n, Arborophila brunneopectus 1gb, Bambusicolo thoracica 1n, Callipepla squamata 1:l.gb r.n, 1 l.n., r.d. Colinus virginianus 2gb, 1n, Chrysolophus amherstiae 1gb, 1b, Gallus gallus 2n, G. sonneratii 2n, Lophortyx californicus 1n, 1b, L. gambellii 1b, Lophura edwardsii 1n, 1b, argaroperdix madagarensis 2n, Pavo cristatus 2n, 1:l.g,r.n, 1 gb, Perdix perdix 1b,1: r.b l.n, 1rb, 1gb, 1n, Phasianus colchicus 27n, 9gb, 1:l.b r.gb, 1gg, 1rb, Polyplectron bicalcarata 1gg, P. emphanum 1b, Pucrasia macrolopha 1gb,1b, Rollulus rouloul 2b,1 gb, Syrmaticus soemmerringi 1n,1b, Tetraogallus himalayensis 1b, Tragopan satyra 2b, T. temmincki 1b
Acryllium vulturinum 1n
RALLIDAE (21 normal)
Amaurornis phoenicurus 1n, Crex crex 1n, Fulica atra 2n, Gallicrex cinerea 1n, Gallinula alleni 1n, G. chloropus 3n, Laterallus leucopyrrhus 1n, Poliolimnas cinereus 1n, Porzana fusca 2n, P. tabuensis 1n, Rallus aquaticus 4n, R. striatus 2n, R. torquatus 1n
Actophilornis africana 1n
Rostratula benghalensis 2n
Haematopus ostralegus 2n
CHARADRIIDAE (29 normal, 4 dark)
Charadrius alexandrinus 1n,1:l.n, r. d, C. dubius 3n, C. hiaticula 2n, C. leschenaultii 7n, 1g, 1: l.b, r.br, 1:l.b, r.n, C. mongolus 1n, C. peronii 1n, Eudromias morinellus 1n, Pluvialis apricaria 4n, P. dominica 1n, P. squatarola 1n, Vanellus coronatus 1n, V. gregarius 1n, V. tectus 1n, V. vanellus 4n
SCOLOPACIDAE (115 normal, 18 dark)
Arenaria interpres 3n, Calidris acumulata 3n, C. alba 6n, C. alpina 33n, 8b, 5:l.b, r.n,2: l.n, r.b, 1: l.g, r.n, C. maritima 17n, 1:l.b, r.n, Gallinago gallinago 5n, G. media 1n, G. megala 1n, Lymnocryptes minima 6n, Scolopax rusticola 26n, Tringa erythropus 1n, T. glareola 3n, T. hypoleucos 1b, T. nebularia 3n, T. stagnatilis 2n, T. totanus 5n,
Himantopus himantopus 1n, Recurvirostra avosetta2n
Phalaropus lobatus 1n
Cursorius cursor 1n, Glareola pratincola 1n, G. maldivarum 1n
Stercorarius parasiticus 1n, S. pomarinus 1n
LARIDAE (9 normal, 11 dark)
Larus argentanus 1n, 1b, L.canus2n, L. glaucoides 1n,1b,L. marinus 1:l.b,r.n, L. ridibundus 4n, 4b, 2:l.n, r.b, Rissa tridactyla 1b, Sternahirundo 1n, S. paradisaea 1b
ALCIDAE (14 normal, 2 dark)
Alle alle 3n, 1:l.d, r.n, Cepphus grylle 4n, Fratercula arctica 1n, Uria aalge 4n, 1: l.b, r. n., U. lomvia 2n
COLUMBIDAE (24 normal, 2 dark)
Chalcophaps indica 2n,1b, Claravis pretiosa 1n, Columba oenas 1n, C. palumbus 3n,Columbina minuta 1n, C. passerina 1n, C. talpacoti 1n,Geopelia striata 4n,Geotrygon montana 1n, Oena capensis 1n, Phaps chalcoptera 1b,Streptopelia decaocta 2n, S. turtur 1n, Turtur afer 1n, T. chalcospilos 1n, T. tympanistria 2n, Zenaida macroura 1n
PSITTACIFORMES (following the systematic in Howard & Moore 1991)
(176 normal, 45 dark)
Chalcopsitta atra 1n, C.duivenbodei 3n,Eos squamata 2n, Trichoglossus haematodus 6n, T. euteles 1n, Lorius lory 1n, L. garrulus 1n, Eolophusroseicapillus 1b, Opopsitta diophthalma 1n, Psittaculirostris desmarestii 2n, Psittinus cyanurus 1n, Prioniturus discurus 1n, Tanygnathus lucionensis 4n, Eclectus roratus 1n, Alisterus scapularis 10b, A. chloropterus 1b. A. amboinensis 1n, 1b, Aprosmictus erythropterus 6b, Polytelis swainsonii 2b, P. anthopeplus 2n, 1bb, P. alexandrae 4b,1g, Purpureicephalus spurius 4n, Barnardius barnardi 5n, 1b, 1:l.n, r.b, B. zonarius 3n,1b,1g, Platycercus elegans 2n,1b,2g, P. flaveolus 1b, 1: l.n, r.b, P. adelaidae 2n, 1:l.b,r.n, P. eximius 1b, P. adscitus 1b, 1: l.bg, r.n, P. venustus 1b, 1gb, P. icterotis 2b, 1: l.b, r.n, Psephotus haematonotus 2n, 1: l.n, r.b, P. varius 5n, P. haematogaster 2n, P. chrysopterygius 3n, Cyanoramphus novaezelandiae 2n, C. auriceps 1n, Neophema bourkii 3n, N. chrysostoma 4n, N. elegans 1n, N. pulchella 2n, N. splendida 7n, Lathamus discolor 3n, Coracopsis nigra 1n, Psittacus erithacus 7n, Poicephalus rufiventris 2n, P. meyeri 5n, P. senegalus 3n, Agapornis cana 3n, A. pullaria 2n, A. taranta 2n 1: l.g, r.n, A. roseicollis 1n, A.lilianae 1n, A. nigrigenis 1n, Loriculus vernalis 3n, L. galgulus 1n, Psittaculas krameri 2n, P. cyanocephala 4n, P. derbiana 1n, P. alexandri 2n, Ara ararauna 1n, Aratinga erythrogenys 2n, A. auricapilla 1n, A. pertinax 1n, Pyrrhura melanura 1n, Enicognathus ferrugineus 1n, E. leptorhynchus 1n, Bolborhynchus aymara 1n, B.lineola 2n, Forpus cyanopygius 1n, F. passerinus 4n, F. xanthopterygius 4n, F. conspicillatus 1n, F. xanthops 4n, Brotogeris chrysopterus 1n, Pionites leucogaster 1n, P. melanocephala 2n, Pionopsitta pileata 1n, Pionus chalcopterus 1n, Amazona albifrons 1n, A. viridigenalis 2n, A. finschi 2n, A, autumnalis 4n, A. aestiva 5n, A. ochrocephala 2n, A. amazonica 7n, A. farinosa 1n, Deroptyus accipitrinus 1n.
MUSOPHAGIDAE (4 normal, 6 dark)
Crinifer piscator 1n, Criniferoides leucogaster 1n, Musophaga violacea 1n, 1: l.b, r.n, Tauraco persa 1n, 1gb, T. hartlaubi 3b,1gg,
CUCULIDAE (9 normal, 1 dark)
Centropus senegalensis 1n, 1: l.gg,r.n, Cuculus canorus 4n, Dasylophus superciliosus 2n, Guira guira 1n, Rhamphococcyx curvirostris 1n,
Tyto alba 21n, 2gb,1b
STRIGIDAE (25 normal, 18 dark)
Asio flammeus 1:l.b r.n, A. otus 21n, 4:l.n, r.gb, 1: l.n, r.g, Bubo bubo 1n, Strix aluco 3n, 1 l. n. r. b., 10b, 3gb
Caprimulgus europaeus 2n, C. ruficollis 1n
APODIDAE (21normal, 1 dark)
Apus apus 17n, A. melba 1n, A. pallidus 2n, Collocalia esculenta 1n, 1b
Amazilia amazilla 1n
Colius indicus 3n, C. striatus 1n
ALCEDINIDAE (18 normal, 1 dark)
Alcedo atthis 8n, Ceryle rudis 1n, Corythornis cristata 1n, Dacelo novaeguineae 1n, Halcyon coromanda 1b, H. chelicuti 1n, H. chloris 1n, H. leucocephala 1n, H. pileata 1n, H. senegalensis 1n, H. smyrnensis 1n, Ispidina picta 1n
MEROPIDAE (11 normal)
Merops albicollis 1n, M. apiaster 1n, M. bullockoides 1n, M. orientalis 2n, M. philippinus 2n, M. pusillus 3n, M. viridis 1n
Coracias garrulus 1n, Eurystomus orientalis 2n
Upupa epops 3n
Phoeniculus senegalensis 1n
Lophoceros nasutus 1n, Tockus erythrorhynchus 2n
CAPITONIDAE (17 normal, 5 dark)
Eubucca bourcierii 1bg, Lybius dubius 4b, L. torquatus 1n, Megalaima asiatica 2n, M. haemacephala 2n, M. henrici 1n, M. lineata 1n, M. mystacophanos 1n, Psilopogon pyrolophus 7n, Semnornis ramphastinus 1n, Trachyphonus vaillantii 1n
Andigena bailloni 4n, Pteroglossus acacari 2n, P. torquatus 1n, Ramphastes toco 2n
PICIDAE (34 normal)
Chrysoptilus melanolaimus 1n, Colaptes campestris 1n, Dendrocopus fuscescens 1n, D. leucotos 1n, D. major 20n, Jynx torquilla 6n, Picus viridis 4n,
Psarisomus dalhousiae 2n.
Furnarius rufus 1n, 1b, Cinclodes fuscus 1n
PITTIDAE (8 normal, 3 dark)
Pitta erythrogaster 1b, P. guajana 4n, P. nympha 1n, P. moluccensis 1n, P. sordida 2n, 1gg, 1 l.gb r.b.
TYRANNIDAE (7 normal)
Hymenops perspicillata 1n, Lessonia rufa 3n, Myiopagis gaimardii 1n, Pitangus sulphuratus 1n, Serpophaga subcristafa 1n
ALAUDIDAE (86 normal, 2 dark)
Alauda arvensis 31n, A. gulgula 2n, Alaemon alaudipes 3n, Ammomanes cincturus 1n, A. deserti 3n, Calandrella cinerea 6n, C. rufescens 4n, 1 l.b r.n, Chersophilus duponti 1n, Eremophila alpestris 2n, 1 l.b, r.n, E. bilopha 1n, Galerida cristata 14n, G. theklae 2n, Lulula arborea 3n, Melanocorypha calandra 4n, M. leucoptera 1n, Mirafra assamica 1n, M.javanica 6n, Ramphocorys clot-bey 1n
HIRUNDINIDAE (36 normal, 5 dark)
Delichon urbica 3n, Hirundo rustica 25n, 4 l.n r.b,1 l.b.,r.n., H. rupestris 2n, H. striolata 1n, Riparia riparia 5n
MOTACILLIDAE (63 normal, 6 dark)
Anthus berthelotii 5n, A. campestris 3n, A. cervinus 1b, A. novaeseelandiae 8n, A. pratensis 8n, 3b, A. spinoletta 1n, 2b, A. trivialis 5n, Motacilla alba 18n, M. cinerea 1n, M. flava 14n,
Pericrocotus flammeus 1n, P. roseus 2n, P. trevirostris 1n
PYCNONOTIDAE (6 normal, 4 dark)
Hypsipetes madagascariensis 2n, H. philippinus 1: l.n r.b. Pycnonotus barbatus 2n, 1:l.b r.n, P. leucagenys 1:l.n r.d, P. melanicterus 1n, P. urostictus 1n, Spizixos semitorques 1b
IRENIDAE (8 normal)
Aegithina tiphia 1n, Chloropsis cyanopogon 1n, C. hardwickii 1n, Irena puella 5n
LANIIDAE (28 normal)
Corvinella corvina 1n, Dryoscopus cubla 1n, Laniarius barbarus 2n, Lanius collurio 7n, L. cristatus 3n, L. excubitor 7n, L. minor 2n, L. schack 1n, L. senator 3n, Tchagra senegala 1n
BOMBYCILLIDAE (24 normal)
Bombycilla garrulus 21n, B. japonica 3n
Cinclus cinclus 1n, 1b
TROGLODYTIDAE (13 normal, 1 dark)
Troglodytes aedon 1n, T. troglodytes 12n, 1: l.n r.gb
PRUNELLIDAE (23 normal, 4 dark)
Prunella collaris 3n, P. modularis 19n, 1b, 1gb, 1:l.b r.n, 1: l.n r.rb, P. montanella 1n
TURDIDAE (164 normal, 147 dark [Turdinae 62n, 107 d.])
Brachypteryx montana 1n, Cercotrichas galactotes 1:l.n r.b, Copsychus luzoniensis 1b, C. niger 1n, Cossypha albicapilla 1n, C. malabaricus 2n, 1b, C. niveicapilla 2b, Erithacus rubecula 3n, 20b, 1gb, E. cyane 1n, 2b Irania gutturalis 4n, Luscinia cyane 2n 1gg, L. luscinia 1:1n r.b, 1:ln r.g, L. megarhynchos 1n, Monticola gularis 1n, M. solitarius 2b, 1:l.n r.b, Myrmecocichla cinnamomeiventris 1n, Oenanthe deserti 4n, O. hispanica 5n, 1gg, O. isabellina 6n, O. leucopyga 1n, 1gg, O. leucura 2b, O. lugens 5n, 1:l.g r.n, 1:l.n r.gg, O. moesta 2n, O.oenanthe 16n, Phoenicurus auroreus 4n, P. moussieri 2n, P. phoenicurus 12n, 3b, P. ochruros 6n, Saxicola caprata 6n, S. rubetra 5n, S. torquata 4n, 1: l.n r.b, Stiphrornis erythrothorax 1n, Turdus chrysolaus 1b, T. iliacus 4n, 7b, 1 other discoloured, T. merula 41n, 56b, 15 other discoloured, T. olivaceous 1n, T. philomelos 19n, 14b, 4 other discoloured, T. pilaris 4n, 4b, T. torquatus 1b, T. viscivorus 2b, 1:l.n r.b
TIMALIIDAE (10 normal, 21 dark)
Actinodura ramsayi ln, rb, Garrulax leucolophus 1:l.b r.n, G.sp. 1n, Heterophasia capistrata 1n, Leiothrix argentauris 9b,1:l.b r.gb, 1:l.g r.gg, L. lutea 1n, 2b, 1:l.n r.b, 2:l.b r.n, Minla ignotincta 1b,1n, 1:l.n r.b, Trichastoma sepiaria 1n, Turdoides fulvus 3n, Yuhima brunneiceps 1n, Y. nigrimenta 1n, Y. zantholeuca l.b, r.n.
Panurus biarmicus 4n, Paradoxornis webbianus. 1 rb,ln
SYLVIIDAE (114 normal, 27 dark)
Acrocephalus arundinaceus 1n, A. orientalis 3n, 1:l.n r.b, A. palustris 2n, A. schoenobaenus 5n, A. scirpaceus 8n, 1:l.n r.b, A. stentoreus 1n, Cisticola juncidis 3n, Hippolais icterina 3n, H. pallida 1n, Locustella certhiola 1n, L. lanceolata 2n, L. naevia 2n, Magalurus palustris 2n, Orthotomus sp. 1n, Phylloscopus bonelli 1n, P. collybita 8n, P. sibilatrix 3n, P. trochilus 13n, 1: r.d, Polioptila dumicola 1n, Regulus ignicapillus 3n, R. regulus 5n, 1b, Scotocerca inquieta 2n, Sylvia atricapilla 1n, 5b, 7gb, 2gg, 2rb, S. borin 2n, 4b, 2gb, 1gg, S. cantillans 2n, S. communis 11n, S. conspicillata 1n, S. curruca 15n, 1:lgb r.n, S. deserticola 1n, S. hortensis 1n, S. melanocephala 5n, S. nana 1n, S. rüppelli 2n, S. sarda 1n,
MUSCICAPIDAE (12 normal,11 dark)
Ficedula albicollis 2n, F. hypoleuca 6n, F. zanthopygia 2b, Muscicapa striata 2n, 1b, 1: l.n r.b, M.thalassina 1b, Niltava hainana 1gb, N. rufigaster 1n, 3b, N. sundara 1n, 1b, N. grandis 1b.
Batis senegalensis 1n, Platysteira cyanea 4n
Hypothymis azurea 1n,Rhipidura cyaniceps 2n, R. javanica 1n,
Aegithalus caudatus 4n, A.concinna 6n, 2b
Remiz pendulinus 2n
PARIDAE (68 normal)
Parus ater 4n, P. caeruleus 16n, P. cristatus 3n, P. cyanus 1n, P. elegans 3n, P. major 29n, P. montanus 2n, P. palustris 7n, P. venustulus 3n
SITTIDAE (5 normal, 10 dark)
Sitta carolinensis 1n, S. europae 1n, 6b, 1g, S.frontalis 1n, S. krüperi 2n, S. neumayer 1gg, S. tephronota 1g, Tichodroma muraria 1b
Certhia brachydactyla 2n, C. familiaris 5n
Rhabdornis mysticalis 1n
Dicaeum australa 2n, 1: l.n r.g, D. trigonostigma 1n
NECTARINIIDAE (24 normal, 2 dark)
Anthreptes neglectus 2n, A. platura 1n, Cinnyris coccinigaster 1n, Nectarina jugularis 2n, N.pulchella 7n, N. senegalensis 2n, 1g, N.venusta 9n, 1g,
Zosterops atricapilla 1n, Z. erytropleurus 1n, Z. montana 2n
Philemon buceroides 1b, 1:l.n r.b Melitreptus albogularis 1b
EMBERIZIDAE Emberizinae (169 normal, 7 dark)
Atlapetes citrinellus 1n, Calcarius lapponicus 10n, Catamenia analis 1n, Coryphospingus cucullatus 2n, C. pileatus 1n, Emberiza aureola 1n, E. bruniceps 5n, E. caesia 3n, E. calandra 7n, E. cia 1n, E. cioides 2n, E. cirlus 2n, E. citrinella 43n,21:l.b r.n, 1:l.n r.b, E. elegans 4n, E. flaviventris 1n, E. hortulana 4n, E. melanocephala 5n, E. rustica 1n, E. rutila 2n, E. schoeniclus 7n, E. striolata 2n, E. tahapisi 1n, Embernagra platensis 1n, Junco hyemalis 1n, Lophospingus pusillus 1n, 1gg, Plectrophenax nivalis 41n, 1:l.n r.g, Phrygilus alaudinus 1n, 1l.b r.n, P. fructiceti 2n, P.patagonicus 1n, Poospiza melanoleuca 1n, P. nigrorufa 1n, P. torquata 1n, Ramphocelus carbi 1n, Rhodospingus cruentus 2n, Saltatricula multicolor 1n, Sicalis olivaceous 1b, S. taczanowskii 1n, Sporophila americana 1n, S. caerulescens 1n, S. castaneiventris 1n, S. collaris 1n, S. lineola 1n, S. torqueola 1n, Zonotrichia capensis 1n,
EMBERIZIDAE Cardinalinae (11 normal, 3 dark)
Cardinalis cardinalis 2n, C. sinuatus 1b, Gubernatrix cristata 2n, 1: l.b r.n, Paroaria capitata 1n, P. coronata 2n, Passerina ciris 1n, 1:l.n r.gb, P.cyanea 1n, P. leclancherii 1n, Saltator aurantiirostris 1n
EMBERIZIDAE Traupinae (23 normal, 3 dark)
Anisognathus flavinucha 1n, Buthraupis montana 1:l.d r.n, Calyptomena viridis 1n, Cyanerpes caeruleus 1b, 1gb, C. cyaneus 1n, Dacnis cyane 1n, Ramphocelus carbo 1n, Stephanophorus diadematus 1n, Tachyphonus rufus 1n, Tangara arthus 1n, T. chilensis 3n, T. cyanicollis 1n, T. gyrola 1n, T. icterocephala 4n, T. nigroviridis 2n, T. schrankii 1n, Traupis bonariensis 2n, T. palmarum 1n,
Coereba flaveola 7n
ICTERIDAE (7 normal)
Cacicus leucoramphus 1n, Icterus cayanensis 1n, Molothrus bonariensis 1n, Pseudoleistes virescens 1n, Sturnella loyca 3n
FRINGILLIDAE (346 normal, 22 dark)
Acanthis cannabina 31n, 1rb, A.flammea 23n, A. flavirostris 21n, Carduelis ambigua 1n, C.atratus 1n, 1gg, C.carduelis 32n, C.chloris 38n, 1b, 1gg, C. cucullata 1n, C. magellanicus 2n, C. psaltria 1n, C. sinica 2n, C. spinus 11n, Carpodacus sp. 1:l.b r.n, C. nipalensis 1n, C. roseus 3n, C. trifasciatus 1n. Coccothraustes affinis 1b, C. carnipes 4n, C. coccothraustes 18n, Fringilla coelebs 44n, 3b, 2:l.b r.n, 2:l.n r.b, F. montifringilla 19n, 1:l.n r.b, Leucosticte arctoa 1n, Loxia curvirostra 13n, L. pytyopsittacus 3n, Pyrrhula pyrrhula 26n, 2b, 2:l.br.n, 1:l.n r.b, 1:l.b,r.n, P. erythaca 5n, 1b, Rhodopechys githaginea 3n, R. mongolica 1n, Serinus alario 1n, S. albogularis 1n, S. burtoni 2n, S. canaria 2n, S. canicollis 1n, S. citrinelloides 1n, S. flaviventris 2n, S. gularis 3n, S. mozambicus 2n, S. serinus 2n, S. sulphuratus 1n, S. pusillus 23n, Uragus sibiricus 1n, 1:l.b r.n,
ESTRILDIDAE (129 normal, 6 dark)
Aegintha temporalis 1n, Amandava amandava 2n, 1:l.b r,n, A. subflava 1n, Chloebia gouliae 4n, Cryptospiza reichenovii 1n, Euschistopspiza dybowskii 1n, 30 Emblema guttata 1n, Erythrura coloria 1n, E. hyperythra 1n, E. prasina 4n, E. psittacea 1n, E. trichroa 1n, E. tricolor 1n, Estrilda melanotis 1b, E. troglodytes 1n, Euschystospiza dybowskii 1n, 1:l.n r.d, Hyparges niveoguttatus 41n, Lagonosticta senegala 2n, Lonchura atricacapilla 1n, L. bicolor 3n, L. cantans 1n, L. castaneothorax 6n, L. cucullata 1n, L. flaviprymna 1n, L. fringilloides 1n, L. fuscans 2n, L. grandis 1n, L. griseicapilla 1n, L. kelaarti 1n, L. leucogastra 1n, L. leucosticta 1n, L. maja 1n, L. malacca 1n, L. malabarica 1n, L. molucca 1n, 1:l.b r.n, L. nana 1n, L. nevermanni 4n, L. pallida 1n, L. punctulata 1:l.b r.n, L. quinticolor 3n, L. spectabilis 2n, L. tristissima 3n, Neochmia phaeton 1n, N. ruficauda 1n, Nigrita bicolor 1n, Ortygospiza atricollis 1n, Padda fuscata 1n, P. oryzivora 1n, Pyrenestes sanquineus 4n, Poephila acuticauda 1n, P. personata 1n, Pytilia hypogrammica 4n, P. melba 1n, P. phoenicoptera 1n, Spermophaga haematina 3n, 1g, Uraeginthus bengalus 1n, U. cyanocephala 2n, U. granatina 1n, U. ianthinogaster 1n
Ploceidae Viduinae (18 normal)
Vidua chalybeata 8n, V. fischeri 2n, V. macroura 2n, V. orientalis 4n, V. paradisaea 1n, V.regia 1n
PLOCEIDAE Passerinae (615 normal, 10 dark)
Auripasser luteus 2n, Dinemelia dinemelia 1n, Euplectes afer 3n, 1:l.n r.g, E. macroarus 2n, E. orix 11n, Montifringilla nivalis 2n, 2b, Passer domesticus 525 n, 1b, 2:l.b r.n, 1:l.gb r.n, P. griseus 3n, P. hispaniolensis 4n, P. moabiticus 1n, P.montanus 55 n, 1rb, 1: l.n r.b, P. rutilans 1n, Petronia petronia 2n, Ploceus cucullatus 1n, 1b, P. velatus 2n, Quelea erythrops 1n.
STURNIDAE (29 normal, 48 dark)
Acridotherus cristatellus 1n, A. tristis 1n, Basilornis galeatus 1b, Gracula religiosa 2n, 1b, 1gb, 2: l.n r.b, Lamprotornis purpureus 3b, L. chalybaeus 1gg, L. iris 1b, Cinnyricinclus leucogaster 1b, Mino anais 1n, M. dumontii 2n, Scissirostrum dubium 1n, Spreo pulcher 1gg, S. superbus 7n, 1b, 2gg, 3gb, Sturnus vulgaris 12n, 26b, 1gg, 2gb, S. malabaricus 1n, S. unicolor 1n,1b
Oriolus chinensis 1n, 1brown, O. oriolus 3n
Dicrurus adsimilis 1n
Artamus leucorhynchus 1n
CORVIDAE (59 normal, 4 dark)
Cissa chinensis 1n, Corvus corax 6n, C. frugilegus 3n, C. corone ln, r.d, C. monedula 2n, Cyanocorax chrysops 1n, C. yncas 1n, Garrulus glandarius 15n, 1: 2.d r.n, Nucifraga caryocatactes 4n, Pica pica 23n, Platylophus galericulatus 1n, Ptilostomus afer 1n, Pyrrhocorax graculus 1b, Urocissa erythrorhyncha 1n
The left ovary is found in the same place as the left testis. Most often the right ovary is missing. The ovaries of nestlings and juveniles in their first months lack the small follicles that produce the granular appearance found in older females, but are distinguishable by being flattened against the kidney like a small piece of transparent plastic. Such an ovary can be very difficult to find, particularly if the bird has not died recently. The shape of the ovary is normally triangular with the caudal end pointed, or oval, but all shapes can be found. The ovary undergoes the same great variation in size as the testes, depending on the time of the year.
The normal occurence is one ovary, but I have found females with two ovaries in many families; in most cases the left ovary is then the larger.
Two ovaries are very common in some families but rare or absent in others. Cf. the following overview from the collection:
PHALACROCORACIDAE: Phalacrocorax carbo
ACCIPITRIDAE: Accipiter nisus (always and sometimes also 2 oviducts Alcohol No.25 & 103), Buteo buteo (rare), Melierax metabates
FALCONIDAE: Falco tinnunculus (egg in development in both ovaries found)
PHASIANIDAE: Oreortyx pictus
CHARADRIIDAE: Charadrius leschenaultii
SCOLOPACIDAE: Phalaropus lobatus, Philomachus pugnax, Tringa ochropus,
PSITTACIDAE: Alisterus scapularis Amazona farinosa, A. finschi, A. tucumana, Ara maracana, Aratinga jandaya, Barnardius barnardi, B. zonarius, Brotogeris pyrrhopterus, Cyanoliseus patagonus, Eclectus roratus, Forpus cyanopygius, F. xanthops, Neophena chrysostoma, Pionopsitta pileata, Platycercus caledonicus, Poicephalus meyeri, Polytelis anthopeplus, P. swainzonii, Psephotus haematogaster, Psittinus cyanurus, Psittacula krameri, Psittaculirostris edwardsii, Purpureicephalus spurius
MUSOPHAGIDAE: Tauraco hartlaubi
TYTONIDAE: Tyto alba
STRIGIDAE: Bubo bubo
RAMPHASTIDAE: Baillonius bailloni, Ramphastos ambiguus, Pteroglossus aracari
PITTIDAE: Pitta guajana
BOMBYCILLIDAE: Bombycilla garrulus
TURDIDAE: Turdus philomelos, T. viscivorus
EMBERIZIDAE: Emberiza citrinella (left grey-green, right pink), Plectrophenax nivalis
ICTERIDAE: Cacicus cela
FRINGILLIDAE: Coccothraustes, coccothraustes, Loxia curvirostra, Serinus mennelli, Uragus sibiricus
ESTRILDIDAE: Hypargos niveoguttatus (right ovary the larger), Lonchura flaviprymna, Neochmia phaeton
STURNIDAE: Lamprotornis chalybaeus, Mino dumontii, Spreo superbus, Sturnus vulgaris
CORVIDAE: Corvus corone, Garrulus glandarius (eggs in development in both ovaries, which were both the same size)
Three ovaries have been found in an Accipiter nisus alcohol no. 25, Amazona farinosa, CN 4226 and Amazona tucumana CN 4574.
Colour of Ovary
When after the breeding the ovary shrinks, visible eggs rarely become black, e.g. a 15-year-old Paroaria coronata (alcohol No. 8), an Ara ararauna with cancer in the whole body cavity (Alcohol No.6c), or an Alisterus scapularis which had never laid fertile eggs and with two defect bones (CN 4977), a Cacatua alba (CN 5331 & CNS 622), or a very old Anodorhynchus hyacinthinus in which only the cranical end of ovary was black (235369), or an 8 years old Lophura swinhoii (CN 5324), but a black ovary (8.0 x 4.0) has also been found in a visibly normal immature Alle alle with straight oviduct (171869). Ovaries where some eggs are one colour and others another colour are often found, e.g. a Fringilla coelebs with pink and grey-green eggs (235013).
Calies also called post-ovulatory follicles, disappear soon as a rule and only one is normally found, but once I found seven in a Strix aluco (299507), three in a Turdus merula (n.n.), and four in a Gavia stellata (n.n.).
Oviduct: two oviducts have been recorded in several Accipiter nisus and one Buteo buteo. (A Pitta iris is reported with 2 eggs in its oviduct (Frith & Hitchcock 1974). I have never observed such a case and believe it is extremely rare).
A case of gynandromorpism or hermaphroditism has been found in the collection, a Rollulus rollouides with one yellowish right testis 4.3 x 3.3 and a left ovary 13.4x 7.0, the ovary high and granular and black. (Alcohol No. 105).
In the collection there is also a Turdus merula which has one side of the ventral body feathers male, the other side female (CN 1779).
HOW TO MAKE SAFE SEX DETERMINATION
Many researchers habitually only cut a little hole in ribbons on left body side and take a look at the gonads through this hole. I cannot seriously enough warn against using this method. Once I saw a researcher sexing a bird as an old male with two large testes in such a way. Checking his statement I found that it was an old female with two large ova in development, which looked like testes! I always open the abdomen, take out the whole intestine without damaging the oviduct, remove the various membranes which hide the gonads, and only then look at the gonads. Many usually break the bones in the middle of body where the pelvis ends to get a better view. If the skeleton is not to be used, this is a good method of determining sex if only one keep in mind that the suprarenal glands will be much more open and visible in that way.
OVERWIEW OF THE VARIOUS KINDS OF STUDY SKINS
1. Soft skin stuffed with cotton wool and crossed legs.
Quickest method. Weight and volume smallest.
Fragile skins, very often skins with broken neck, detached wings or tail and legs. Sometimes I have found several cut-off legs in the same drawer with the labels attached to the legs, so that it was no more possible to determine to which bird the label belonged. Fat from the legs on feathers and label. Difficult to study legs. Feathers impossible to place in the correct patterns, e.g. I have found a skin where a diagnostic collar was entirely hidden etc.!Very often the carpal joints of the wings are put under the skin of the shoulder, (e.g. many famous earlier ornithologists and Philippine skinners nowadays practise this method), which makes wing measurement impossible. The same goes for the tail.
2. Soft skin stuffed with cotton wool and crossed legs with a stick anchored inside the skull to well beyond the tail.
Quick method. Not so fragile as number 1. You can spare the skin by holding the stick instead of handling the feathers. Weight and volume small.
Fat from the legs on feathers and label. The legs and thighs very difficult to study. Feathers impossible to place in the correct patterns. Sometimes seen skins, where the legs were cut off but still attached to the stick.
3. Flat skin
I have not enough experience with them but I do not like them.
They are no aesthetic pleasure and no use for many purposes, e.g. drawings and study of plumage patterns, but good for stydying pterylography.
4. Firm skin with a wire through neck and body. Life size body of wood shavings, balsa wood or cork. Legs crossed
Steadier than 1-3. Easier to place the feathers correctly, because the body and neck shape are like the natural ones. Wings can be placed correctly and sticks between ulna and radius can secure them to the body. Handy to hold by the head- and tail wire.
Greater volume and weight. More time consuming to prepare. Fat from legs on feathers and label, and legs often loose. Legs difficult to study.
5. Firm skin with wire through head, neck and body, legs and tail (possible with the wire ending outside head and tail in an eye). To give the head more stability I put glue (wood glue) in the brain case. Birds larger than a Jay with wire in both wings, in smaller the two ulna or radius are tied together.
Steadiest. Easy to place wings and feathers in a correct position. Fat from the legs will not make feathers and label dirty, and it is easy to clean the legs later with benzine if fatty. Easy to study legs and thighs. Handy to hold by the legs or by the head- and tail wire.
Greatest volume and weight. Most time consuming to prepare. The tarsus measurement often difficult to take (take it during the skinning).
6. ROM skin (half skin, half skeleton)
I have no experience because I do not like them, but unquestionably they allow maximal use of the skin.
7. Shmoos (muppets), skins where the bill follow the skeleton
I have no experience with this method too but described in detail by Winker (2000).
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