CATALOGUE OF THE PRIVATE BIRD-SKIN AND SKELETON COLLECTION

JOHANNES ERRITZOE

(Open for scientific research)

Copyright 1993 Johannes Erritzoe, latest revised 2006

This publication may be reproduced only if the original source is mentioned.

Document overview:

Introduction
Short history of the collection
Ethical guidelines
Sex determination
Age indication
Dates, locality and other measurements
Body mass indications
Colour disignations
Moulting and supplementary comments and methods
The osteological collection
Literature cited
Abbreviations
List of all papers based on this collection
Collection management guidelines
Preparation techniques
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
Reference list

Introduction

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. (6.000 skins, c. 4.000 part skeletons and 1000 skeletons). 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 the 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, p. 12-13).

SHORT HISTORY OF THE COLLECTION

The collection was started in 1947 under the supervision of the late Erik Petersen, Zoologisk Museum in Copenhagen. From the very beginning the following pieces of information have been recorded: Date and locality, weight and a drawing indicating the size of the sex organs, the measures taken with callipers. 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 measurement (which is also nowadays more computer adaptable). At first weight of the body mass was taken with an ordinary letter balance, but from 1964 a pair of scales was used - also on later expeditions. From 1995 a Sartorius 1608 MP have been used.

The number of items of information grew gradually year after year, some because I saw others use them, others were my own ideas. In 1965 in the museum in Copenhagen I showed Dr Finn Salomonsen some of my skins, 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 the legs crossed. The many museum-skins I saw with legs, tails or heads broken off gave me the idea of using excelsior (wood wool) instead of cotton wool, because with this material it is possible to use wire through the legs, neck and tail and thus obtain more stability. During the last few years wire has also been used for spread wings displaying important plumage features that folded wings on skins cannot show. In a series I try to make the degree of extension different. From about 2001 I have stopped to made skins with one wing spread and instead started a feather collection, mentioned latter.

Skeletons have been collected from 1986 onwards.

Ethical guidelines

In a catalogue like this only to be accessible to the scientific community it should be a waste of space and time to try to defend the legitimacy of a bird-skin, feather and skeleton collection.

The understanding of bird conservation has changed during the last decades. Most bird species which it was legal to shoot only 20 years ago are now protected by law. Also the attitude to conservation has changed concurrently due to the disastrous decline of many bird species.

All bird-skins or 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 one thousand House Sparrows speak for itself.-

Throughout the last two 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, concentrating on making larger series of the common species. All protected species are registered by the Danish authorities.

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).

Nomenclature

The following check-lists have been used:

Morony, J. J., W. J. Bock, & J. Farrand, 1975 & 1978: Reference-list of the Birds of the World. AMNH, New York.

For subspecies the following have been consulted:

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.

Sex determination

Sexing by collectors was formerly often based on some preconception on sexual differences rather than on anatomical confirmation. I find it important not only by dissection to determine the sex but also to show a latter researcher that the bird has been opened. Naturally the condition of the gonads also tell 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, possibly also hand-held magnifying glasses with factors of 10 and 20 respectively. For measuring 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.

Testes

Each testis is measured length x breadth in millimetres so testis volume can be roughly estimated. The left testis is the right when viewed from ventral and since 1995 I have used also to weight both testes.

If testes are discoloured, the abnormal colour of left or right or both is recorded (see p. 22-30). When it is difficult to measure the testes in the body, they are measured outside, which is often necessary, if you want to preserve the trunk skeleton. If large they are always weighted.

When birds have been sex-determined by plumage, this is invariably recorded. In most 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.

Spermatic ducts

As certainable cases of sinuous spermatic ducts are always noted. On the other hand the fact that there is no note on this - on small birds a very difficult matter - does not warrant the assumption that the spermatic ducts are straight.

Ovary

The Ovary is measured from the largest length to the largest breadth in millimetres.If two ovaries, both are measured. By 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. I believe that many such young females by former field workers by mistake are determined as males, confusing the adrenal glans for testes. If the ovary is granulated with ova larger than 2-3mm, the size of the biggest ovum is recorded. If ova are large, all eggs of more than 3.0 x 2.5 mm are removed, and their largest diameter given. In such cases there will be a note of the fact against the ovary measurement, e.g. Ovary - 4 ova .... If calies are found, this is recorded too.

Oviduct

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 measurements as above. If oviduct is greatly distended, the mass is recorded.

Anomalously shaped testes or ovaries are recorded, as is any other abnormality worth mentioning. If the sex determination or any other thing observed is astonishing, e. g. because of the 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 student: Do not leave that specimen out of the calculation, it is OK. If any doubt is left a ? is written or/and the organ in question is kept in alcohol.

Age indication

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." bird. (Only for members of Passeriformes, regarding skull ossification). After the complete moulting into the second plumage, it is set down as a 1st-calender year bird, "1 year", 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) or a "2 year" bird after that date, and in most birds the age indication changes into "ad" after the completion of the second moulting. For birds with a different plumage in their third calendar year compared with that of the adult bird, the age indication "3 year" is also used, but more often 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.

In most skins 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 named in: Svensson, L. 1970 (and later rev. ed.): Identification Guide to European Passerines. Lars Svensson, Stockholm. From catalog number 5509 the c. per cent is given as described by Kewin Winker (2000). Bursa Fabricii is measured length x breadth. Where b.F. is expected but not seen, owing to obesity and other factors, it is noted that b. F. was not found.

For more information about sex determination and age indication see:

Erritzöe, J. 1985: Geschlechts- und Altersbestimmung bei Vögeln. Der Präparator 31 (2):81-93 and page 21-33 in this catalogue.

Dates

In the absence of particular comments to the contrary, dates are perfectly exact. The months are given by at least a three-letter abbreviation to avoid any mistake with the American way of month numeral, e. g. means 5/11/1990 in Europe 5 November 1990 but in USA 11 May 1990! On older specimens the european method is used.

Locality

The coordinates are given for all Danish localities using the CD ROM Topografisk Atlas for Denmark. For all other countries the coordinates are taken from Times Worldatlas.

Other Measurements

All measurements are taken in millimetres but I have omit written mm, opposite mass where I always write g.

If a incubation patch is found, this is indicated on the label with length and breadth.

To now the index of size e. g. in connection to Bergmann's rule have always been the wing length and to a less degree the body mass (Zink & Remsen 1986). The following measurements can maybe give surprising new informations:

The length of femur and humerus are recorded.From 1998 are the average measurements (n = 2) given for both left and right femur and humerus. This is a great advantage because if there is great difference between the measurements you know that at least one of them is wrong, so you have to repeat the measurements until you are sure. In future tarsus will also be measured because with wire through the legs this measurement is difficult to measure in a latter stage. Wing- (max.), bill- (culmen to skull), and tail- lengths are also taken from the bird before skinning. From the dead body the following measurements are taken: Length of body from the front to the tip of pubis, the breadth and the height of the body. The length of the neck from a line between the dorsal end of furcula to the cranium. The diameter of both iris and the whole eye-ball. Finally the measurement "wingtip to tailtip" is taken on the dead bird with the wings and tail in a natural position, the two wingtips touching each other but not crossing, the measurement to the longest tail feather is taken. If the wingtip is outside the tailtip it is signed +....This measurement is mostly taken as a help for bird illustrators.

The spleen, bursa of Fabricius and the gall bladder are measured length x breadth; thymus are are now only weighted, first all on the left side of neck and then the right side. Caecum length and distance from the caudal end of caecum to cloaca.The two measurements are given with a dash between the two figures. If the two caeca have different lengths but the length to the cloaca from the end of both caeca are the same it is written e. g.: 10.0 + 9.6 - 14.4. If the same two caeca have different distances to the cloaca it is written: 10.0 + 9.6 - 14.4 + 13.8.

From CN 5133 and c. CNS 206 are wing-, bill, tail-, and tarsus length 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).

Body mass indications

The weight is given in grammes.

Such birds as have been weighed by myself (and this goes for nearly all) have been weighed with a pair of scales with an inaccuracy margin of ± 0.1 g, provided, 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 the fact is appended. After I have started to take DNA this is not neccessary anymore, because the DNA number is the date the bird is prepared.

In a experiments 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 was removed from them (personal obs.). (cf. Banks 1965).

Note, that a bird frosen for a short time, say one to twenty days, will often have a higher weight than the first recorded.

From Feruary 1995 all birds are weighed with a Sartorius 1608 MP.

If the 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.

Food and possible grit are weighed together, but a note is made if grit is found in the gizzard and since approx. 2003 grit ia always kept except from captive birds. Ithe inner layer of the gizzard (cuticula gastrica) was already by Aristotle recognised to be easily removed "which can be pulled away from the fleshy part",so it is alså weighted. The contents of the intestine is not weighed with the food except in the case of the nearly thousand House Sparrows collected between 1965 and 1969. Since 2005 the intestine with content is also 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 later is excise by cutting all vessels as near as possible to their origin on the heart and residual blood is forced from the heart by compression. Liver and spleen are also weighed, the external blood on the liver, connective tissues and blood vessels near their connection with the organ are removed before it is weighed. Finally, bursa of Fabricius, thymus and brain are weighed. In cases where the skeleton is taken brain is weightet after the following procedure: First is a piece of paper + some cotton wool weighted, the brain is then removed by putting wool inside the brain case and moved around. This procedure is repeated until no more brain is visible on the wool. The brain with paper, used and unused wool is now weightet and the weight before starting is subtracted.

The condition of the body mass is described as follows:

Emaciated: dead of starvation

Lean: breast muscle shrunken,

A little lean: breast muscle a little shrunken

Fat-free: breast muscle normal, no fat at all

Normal:  breast muscle normal, a very 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 much fat on skin and viscera.  

Colour designations

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 the 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 receive a bird just after its death, a colour-guide is used to get an exact colour. For a long time I used a Danish guide:

Kornerup, A. & J. H. Wanscher, 1962: Farver i Farver.

Politikens Forlag, Copenhagen.

Abbreviated FF

Later I used a German guide:

Küppers, H. 1978: DuMont's Farben Atlas

DuMont Buchverlag, Köln.

Abbreviated: FA

For the last few years I have used:

Smithe, F. B. 1975: Naturalist's Color Guide

AMNH, New York.

Abbreviated: NCG

I find the Danish one the best one, but nobody except the Danes know about it. It is translated into English, 3rd. edition 1983: Methuen Handbook of Colour. Methuen, London, but is now impossible to find on the markt. The German guide is also fine, but most English speaking people do not know it, therefore I use, the poorest of them all, the American Guide which all known, but only if I can find the exact colour, otherwise I explanin the difference, e.g. darker than ... or I use the Danish one.

Moulting

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 during the last few years (modified from Snow, D. W. 1967: A Guide to Moult in British Birds. BTO Field Guide No. 11 p. 3). One asterisk on the label against "Moult Card" signifies that the moulting is abnormal, e. g. asymmetric or fear-induced. Now I always spread one wing, when the bird is moulting, because it is very hard on 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 caharacteristic body feathers. If the moulting is finished but there are still black feather follicles on skin inside this is stated.

Supplementary comments and methods

If some albinistic feathers are 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 is notes. The tracheal bulla by ducks, cast still in the body, endo- and ecto parasites or food which is suitable for being dried. To facilitate later determination of seed a collection of seed from many plants in my region have been collected and sorted after size. When a bird is skinned, all loose feathers are kept separate and a note about that is written on the label.

If a bird is crested it is usually prepare with its head turned to one side so that the crest is conspicuous.

If the 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 the fat from the uropygial glands by washing is removed and thereby gives help to feather-degrading bacterias (Burtt 2009). If the bird is bloody or dirty a shampoo detergent was used in many years, now I prefer a common washing-up liquid in hand warm water. 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 int the skin I use much water to keep the skin smooth under this work. Is a bone in wing or leg broken all marrow is removed and the broken bone is repaired with a piece of wire. If the bird is very fat, 50% benzine and 50% alcohol are mixed together and the bird stay in this solution for about one hour.

On expeditions I 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.

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.

The skins have been treated inside with arsenic 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. I only use Eulane in rare cases because nobody knows how it affects the feathers and skins over a longer period.

Numbering

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. 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 the six-figure number is also written. This is the register number from the Danish Ministry of the Environment.

Minimum information

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 the breastbone with furcula, coracoid and scapula usually follow the skin.

From catalogue number 4080 and onwards the trunk skeleton is kept.

The last years the following informations are reported:

Scientific name, sex, age, date locality. masst, statement about the condition of the bird, weight of food and grit, weight of food in crop, brain, gizzard, heart, liver, bursa of Fabricius, thymus, and spleen.

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 were prepared. The method used is that described in Lars Svensson 1984: Identification Guide to European Passerines. Wing maximum length is used. As all measurements are made by the same person (JE), no correction factors for multiple measurers is necessary. Measurements of gonads, oviduct, straight or sinuous, breadth in both ends, femur, humerus, length between wingtip and tailtip, length of body from the front to the end of pubis, width and height of the body, length of the neck, diameter of iris and the whole eye-ball. On the label is space for the wing-, bill-, tail,- and tarsus length.

If a bursa of Fabricius, a gall bladder or a caecum exist it is also measured, the first two mentioned with length x breadth, caecum length and length from the caudal end of caecum and to anus. Spleen is also measured length x breadth. Often the length of intestine is recorded.

A statement is given of the skull ossification and a drawing of the "windows" are if any given. Notes about grit, food contents, fault bars and parasites, known age, captivity and if the bird is washed.

Colour of iris, bill, inside upper mandible, palate, mouth, tongue, legs and feet, sole and claws and if any bare parts on the skin. It is noted if the colours are identified short after death or later.

When tongues or ectoparasites are kept, they are only dried. Eyes are first prepared in alcohol and then dried. I have now stopped to keep the dried eye because they are often easten by pest.

From cataligue number CN 5509 and CNS 779 DNA tissue is taken.

THE OSTEOLOGICAL COLLECTION

After I had read Wood, D.S., R. L. Zusi, & M. A. Jenkinson, 1982: World Inventory of Avian Skeletal Specimens. AOU and Oklahoma Biological Survey, Normal, Oklahoma. and realized that at a rough estimate one-sixth of all bird species of the world are not represented in any museum, I immediately started collecting bird skeletons given the skeletons so high a priority that already the second species of a given species - whether beautiful or not - is skeletonized! And not without success:

So far, 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 the study of age changes (The ages of more than 200 are known). 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 third of the skeletons has been cleaned by dermestid beetles outside my house. Some have been cleaned by mealworm 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 become fat-free. To avoid the stench i have them in plastic boxed which are chosed tight as soon as the start to smell.

The data taken is the same as for the skins.

"There isn't one of us who hasn't at some time or another, misidentified a bird. If a bird is misidentified and then skinned, it is only a matter of time before someone corrects the error. If a bird has been misidentified 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 misidentified whole are precisely the ones whose misidentification is least likely to be detected as skeletons" (Matthiesen 1989).

In order to avoid later researchers failing to check the species determination, the primaries and tail feathers and some of the more distinguished smaller contour feathers are kept in archival plastic sheets. The last years one spread wing,the whole tail and some distinctive body feathers have been kept as a voucher to each skeleton.

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 this 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.

Literature cited

Anderson, S. 1965: Systematic Zoology 14:344-346.

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.

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.

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.

Svensson, L. 1984: Identification Guide to European Passerine. Lars Svensson, Stockholm.

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. (Also translated to Chinese)

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. 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. & H. Svenningsen, 1996: Dark-eyed Junco in Denmark 1980 and review of records from Europe and Greenland. Dutch Birding 18 (1):1-5.

Garamszegi, L. Z., A. P. Møller & J. Erritzøe. 2002: Coevolving avian eye size and brain size in relation to prey capture and nocturnality. Proc. Royal Soc. London B 269: 961-967.

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 (in press.): Sexually size dimorphic brains and song complexity in passerine birds. Behav. Ecol

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. Erritzoe, (in press): Thymus mass and T-cell mediated immune response in birds. Functional Ecology.

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. Erritzøe, & N. Saino, 2003: Seasonal changes in immune response and parasite impact on hosts. American Naturalist 161 (4): 657-671

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., 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.

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.

Storage facilities

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.

The gradual incorporation of new specimens in the collection

The time between the mounting of 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 hangs by its head wire in the laboratory for only 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 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 will be put in a 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.

Plastic bags

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 use now 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 - (6.000) of bird-skins, c. (4,000) breastbones, c.(2.000) trunk skeletons) and (1000) skeletons - in my collection (Anno 2006) every skin and skeleton is monitored twice a 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.

All cabinet door frames are painted twice 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 except a little Vapona.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 in the house. 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

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 use 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. 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 also makes it 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 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; in a few cases some feathers have been bent a little 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 thousands 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 student does not have the time to read this catalogue, forgets it, does not know about it, or cannot find the right page in a hurry? Attached labels are therefore 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.

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 in the top right-hand corner. The reasons for this procedure are several: I prefer to finish the skins, study books and papers and take the measurements of femur and humerus 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 the text in Danish, is rewritten, the old one always remains on the skin.

As twine a black nylon thread is now used, 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.

The trunk skeletons are kept in plastic bags in boxes in another place in the house with the catalogue numbers written on the outside of the box. On the label of the bird-skin in question it is mentioned that 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, tongue, food, grit, parasites, loose feathers.

The skeletons

The skeletons are also kept in plastic bags in boxes after families, with the catalogue numbers written on the skeletons and the labels. For the quick check I have a copy of Gruson s "Checklist of the Birds of the World" at hand, where the catalogue number of every skeleton under the bird in question is written. On the label of the skeleton an "F." is written if the feathers are kept, which is the standard procedure for all birds except for very dirty ones. The 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 and catalogue number.

Specimen in alcohol or its derivatives

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 are kept. The number of the alcohol preparation is stated on the label of the bird-skin or skeleton in question, and a number outside the plastic container with the organs kept in alcohol gives all the relevant information.

Preparation techniques

In the following methods and techniques not found in the literature I know about are reported. 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.

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 one pressed a finger on the body, it made a crackling sound. I suppose in a hundred years or less these skins would have broken into pieces like many old skins I have found in museum collections.

I restored them 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, fifteen years later, it has caused no disadvantages, and the skins are still soft.

I think the skin of a bird is comparable with the mammal-skin used, e.g. for book-binding. If you do not give the leather of a book someleather 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, both when skinning to avoid e.g. blood dirtying the feathers, and afterwards when the washed skin is to be dried.

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 very 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 maybe only five minutes more work for a small bird.

Wood flour is difficult to get out of the down again, and I have also tried to use magnesium carbonate but prefer potato flour. When I make black birds and birds with glossy colours I use blotting paper instead of flour. It works well.

Fat cleaning

Nearly all birds in temperate zones have some fat on the skin. If all is not 100% removed, the fat will 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 with a rounded and not too sharp scalpel. If water is used on the place of the skin where you work, it is easier to prevent to make holes in the skin. Potato flour is also very useful. The fat on the knee is ignored by some, but must also be removed. Are some bone with marrow broken it is very important to emty these, because the marrow otherwise will move to the feathers like the 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. Try to scrape using a rounded and blunt scalpel and lots of water!

Dirty skins

If one take a look in museum skin collections, the drawers are nearly always dirty from the skins. This make a effective pest control more difficult and make often the study of the skins rather grubby. To prevent this I always use a hair dryer. It gives the feather more life and remove most dirt and potato flour.

Bill closing

In order not to ruin the nostrils and damage the operculum 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 the clay dries before the bill begins to work and is easy to remove later when the 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 expanded polystyrene, 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 humerus 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 and label.

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 thrust 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.

My experience with sex determination

MALE

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.

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:

NORMAL WHITE AND DARK TESTES

STRUTHIONIDAE  

Struthio camelus 1n

RHEIDAE

Rhea americana 1gg, 1 b

TINAMIDAE

Crypturellus tataupa 1n

GAVIDAE

Gavia arctica 1 rb, G. immer 2 b, G. stallata 3 n

PODICIPEDIDAE

Podiceps cristatus 1n, Tachybaotus ruficollis 1: l.b, r. n.

PROCELLARIIDAE

Puffinus gravis 1grb

HYDROBATIDAE

Hydrobates pelagicus 2n, Oceanodroma leucorrhoa 2n

PHALACROCORACIDAE

Phalacrocorax carbo 4n

ARDEIDAE

Ardea cinerea 3n, 3b. Butorides striatus 1n, Ixobrychus sinensis 1n.

SCOPIDAE

Scopus umbretta 1n

Ciconiidae

Ciconia ciconia 1gg

PHOENICOPTERIDAE

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

FALCONIDAE

Falco tinnunculus 13n, F. ardosiaceus 1n

TETRAONIDAE

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

NUMIDIDAE

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

JACANIDAE

Actophilornis africana 1n

ROSTRATULIDAE

Rostratula benghalensis 2n

HAEMATOPEDIDAE

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,

RECURVIROSTRIDAE

Himantopus himantopus 1n, Recurvirostra avosetta 2n

PHALAROPODIDA

Phalaropus lobatus 1n

GLAREOLIDAE

Cursorius cursor 1n, Glareola pratincola 1n, G. maldivarum 1n

STERCORARIIDAE

Stercorarius parasiticus 1n, S. pomarinus 1n

LARIDAE (9 normal, 11 dark)

Larus argentanus 1n, 1b, L.canus 2n, L. glaucoides 1n,1b, L. marinus 1:l.b,r.n, L. ridibundus 4n, 4b, 2:l.n, r.b, Rissa tridactyla 1b, Sterna hirundo 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, Eolophus roseicapillus 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,

TYTONIDAE

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

CAPRIMULGIDAE

Caprimulgus europaeus 2n, C. ruficollis 1n

APODIDAE (21normal, 1 dark)

Apus apus 17n, A. melba 1n, A. pallidus 2n, Collocalia esculenta 1n, 1b

TROCHILIDAE

Amazilia amazilla 1n

COLIIDAE

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

CORACIIDAE

Coracias garrulus 1n, Eurystomus orientalis 2n

UPUPIDAE

Upupa epops 3n

PHOENICULIDAE

Phoeniculus senegalensis 1n

BUCEROTIIDAE

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

RAMPHASTIDAE

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,

EURYLAIMIDAE

Psarisomus dalhousiae 2n.

FURNARIIDAE

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,

CAMPEPHAGIDAE

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

CINCLIDAE

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.

PANURIDAE

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.

PLATYSTEIRIDAE

Batis senegalensis 1n, Platysteira cyanea 4n

MONARCHIDAE

Hypothymis azurea 1n,Rhipidura cyaniceps 2n, R. javanica 1n,

AEGITHALIDAE

Aegithalus caudatus 4n, A.concinna 6n, 2b

REMIZIDAE

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

CERTHIIDAE

Certhia brachydactyla 2n, C. familiaris 5n

RHABDORNITHIDAE

Rhabdornis mysticalis 1n

DICAEIDAE

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,

ZOSTEROPIDAE

Zosterops atricapilla 1n, Z. erytropleurus 1n, Z. montana 2n

MELIPHAGIDAE

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,

COEREBIDAE

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

ORIOLIDAE

Oriolus chinensis 1n, 1brown, O. oriolus 3n

DICRURIDAE

Dicrurus adsimilis 1n

ARTAMIDAE

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

FEMALE

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. Many researchers must formerly have sex determined many young female birds as males because of the suprarenal glands (see above). 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 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 the larger.

Two ovaries

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, the visible eggs often 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 defective 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 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).

Gynandromorpism

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 13.4 x 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 the ribbons in the 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 eggs 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 the 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 keeps in mind that the suprarenal glands will be much more visible in that way. If it is difficult to find the gonads it often help to put the body in water for a few minutes to remove the blood. To find the tranparent ovary of a young female is difficult, often you have to turn the body in different light angles.

Overview of the various kinds of study skins

1. Soft skin stuffed with cotton wool and crossed legs.

 Advantages:

 Quickest method. Weight and volume smallest.

 Disadvantages:

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 the inside of the skull to well beyond the tail.

 Advantages:

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.

 Disadvantages:

 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.

4. Firm skin with a wire through neck and body. Life size body of wood shavings, expanded polystyrene, balsa wood or cork. Legs crossed

 Advantages:

 Steadier than 1. 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.

 Disadvantages:

Greater volume and weight. More time consuming to prepare. Fat from legs on feathers and label, and legs often cut off. 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) Birds larger than a Blackbird with wire in both wings.

Advantages:

 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 e.g. benzine if fatty. Easy to study legs and thighs. Handy to hold by the legs or by the head- and tail wire.

Disadvantages:

 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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