Jeże anatomia histologia i fizjologia (2)

www.naturesweb.ie

Spring 2010

Animal Life

TTTT

he hedgehog is easy

to recognise. Its

head and back are

covered with sharp spines,

each 2-3 cms long. These

spines are actually

modified hair (much

harder than normal hair).

If frightened or attacked,

it will curl up into a ball,

and uses the spines to

protect its body. An

adult hedgehog has

approximately 5,000-

7,000 spines.

A hedgehog is about 25

cm long. It has a sharp

snout, relatively long legs

(for its size), and a short

tail. Its small bright eyes

cannot see very well but

it has a very good sense

of smell and great

hearing.

For food, hedgehogs eat

caterpillars, earthworms,

slugs, beetles, snails and

insects – and are very

noisy eaters! They also

make pig-like squeals

when distressed and

grunt when courting.

Hedgehogs generally live

alone and only look for

company when they are

mating. They live in

woodlands and gardens.

In the wild, a hedgehog

can live for about three

to five years, but some

can live up to 10 years.

Hedgehogs are nocturnal

and wander about at

night, travelling about 3

km in search of food. If

you see one during the

day, it is possible that it

could be ill.

Hedgehogs hibernate in

winter when food

becomes scarce. They

wake up now and then

and feed, often when the

weather is mild. They

build nests out of leaves,

grass and other

vegetation, often under

hedges, in compost heaps

and beneath piles of

wood.

Few animals will eat a

hedgehog because of its

spines, but badgers will.

The biggest killer of

hedgehogs are cars and in

the garden, slug pellets.

Hedgehogs are a

gardener's friend, eating

slugs and caterpillars and

not doing any damage.

The Hedgehog

Are hedgehogs and

porcupines related?

As both hedgehogs and porcupines

have spines, you might think that they

are related, but they are not.

Hedgehogs belong to a group of

animals known as "insectivores", small

mammals that feed mainly on insects

and similar small creatures.

Porcupines are rodents and, being

herbivores, mostly eat plant food such

as bark and leaves.

The hedgehog doesn't really have any

close relatives. However, it is thought

that there is some distant link to moles

and shrews.

Hedgehogs in

Ireland

Though hedgehogs

are common

throughout Ireland, they are not

native to this country. It is thought

they were introduced by humans,

possibly the Normans in the 13th

century.

The hedgehog in Ireland is the same

species as that found in the rest of

Europe. There are about a dozen

other species of hedgehog and these

are found in South East Asia, China

and Africa.

Hedgehogs are protected in Ireland. If

you want to keep one captive

(perhaps because it is ill)

you need to apply to

the National Parks and

Wildlife Service for a

license. Hedgehogs

cannot be sold.

Latin: Erinaceus europaeus

(“Erinaceus” means “spiky wall”)

Irish: Gráinneog

(meaning “horrible one”)

Hedgehogs and their young

The hedgehog will have four or five young at a time, one

litter between May and July and often another in August

or September. The young are born with soft spines,

which will soon harden.

A baby hedgehog is called a hoglet and is also known as a

pup, kit or piglet.

Porcupines (above) are not related to hedgehogs.

Cou

rtes

y Sa

uel P

alm

er CC-

-SA

-2

y of

www.

Introduction

No other exotic animal has caught the attention of

the public quite like the hedgehog has. Their spines,
friendly nature, and an ever-smiling expression have
endeared them to millions of confessed hedgehog
lovers around the globe.

In the evolutionary development of the vertebrate

heart, the specialized atrioventricular conduction
system appears as a phylogenetically new structural
entity, which, to date, has been documented only in
mammals and birds (Szabo et al., 1986). Moreover, in
considering its development, it is very important to
compare the cardiac conducting system in different
species.

Cardiovascular diseases are an important cause of

human mortality worldwide, especially in developing
countries. In addition to high rates of mortality, the
costs associated with treating these diseases are high.
The negative economic, social, industrial, and
psychological effects of these diseases are significant. In
order to understand cardiac function, research on the
histological structure of the cardiac conduction system,
especially the atrioventricular system, is necessary. Two
principal components of the atrioventricular
conducting system are the atrioventricular node (AVN)
and atrioventricular bundle (AVB). For example, some
cardiac arrhythmias are due to pathological lesions and
anatomical defects in the AVN and AVB, or in their
blood supply.

237

Research Article

Turk J Zool
34 (2010) 237-242
© TÜBİTAK
doi:10.3906/zoo-0810-17

The anatomy and histology of the atrioventricular conducting

system in the hedgehog (

Hemiechinus auritus) heart

Abolghasem NABIPOUR*

Department of Anatomical Sciences, School of Veterinary Medicine, Ferdowsi University of Mashhad, Mashhad,

P. O. Box: 91775-1793, IRAN

Received: 30.10.2008

Abstract:

This study examined the atrioventricular conducting system in 4 adult male hedgehogs (Hemiechinus auritus).

The histological structure of these components was studied using routine histological methods. The AVN was located at
the lower and anterior part of the interatrial septum, near the root of the aorta. It was almost oval and consisted of twisted
cells. Internodal pathways in the hedgehog heart were not observed, but there were numerous purkinje-like fibers within
the myocardium of the atrium. The AVB was a continuation of the AVN, as a compact structure, extended obliquely
through the fibrous ring toward the apex of the interventricular septum, and was composed of many purkinje cells.

Key words:

Hedgehog, heart, histology, atrioventricular node, atrioventricular bundle

* E-mail: nabipour@ferdowsi.um.ac.ir

The anatomy and histology of the AVN and AVB

have been studied in humans (Lev and Lerner, 1955;
Titus et al., 1963; James, 1970; Titus, 1973), dogs and
monkeys (Nonidez, 1943; James, 1964), hoofed
animals (Meyling and Terborg, 1957; Prasad and
Sinha, 1980), rabbits (James, 1967), birds (Szabo et al.,
1986), lizards (Prakash, 1990), camels (Ghazi and
Tadjalli, 1993, 2002), cats (Ghazi et al., 1998; Tadjalli
et al., 1999), cattle (James, 1965), horses (Bishop and
Cole, 1967), goats (Nabipour, 2002; Nabipour et al.,
2002), guinea pigs (Nabipour, 2004a, 2004b), and
recently in ovine fetuses (Nabipour and Shahabodini,
2007). However, precise data are not available on the
anatomy and histology of the AVN and AVB in
hedgehogs. The present study follows others on the
histological structure of the AVN and AVB in
different animal species. Histological knowledge of
the AVN and AVB provides the basis for
understanding their physiological function, and
delineation of their structure in the hedgehog will
provide additional insights into the significance of
their structure.

Materials and methods

The study included 4 adult male hedgehogs

(Hemiechinus auritus) with an average weight of 357
g (Figure 1). They were euthanized with an overdose
of sodium pentobarbital administrated
intraperitoneally. After removal of the pericardium,

the heart was flushed with warm (40 °C) normal
saline and for fixation was perfused with 10% neutral
buffered formalin solution. The lower part of the
interatrial septum (from the level of the upper part
with the coronary sinus) along with the upper part of
the interventricular septum were removed. The
samples were placed in the same fixative, and then
through a series of graded alcohols and xylene, and
eventually into paraffin wax. Serial sections 6-μm
thick were made longitudinally, starting from the
right side of the samples. The sections were preserved
and then selected by the interval of 3, stained with
Masson’s trichrome green and PAS-Alcian blue
(periodic acid Schiff-Alcian blue) (Luna, 1968). The
stained sections were studied under a light
microscope.

Results

The hedgehog AVN was located at the lower and

anterior section on the right side of the interatrial
septum, near the root of the aorta, and was almost
oval (Figure 2). Morphologically, the hedgehog AVB
was a continuation of the AVN. There was no
detectable border between the node and the AVB. The
AVB extended obliquely through the fibrous ring to
the apex of the interventricular septum, as a compact
structure (Figure 3).

Within the AVN there was a mass—an interlacing

bundle of fibers that were smaller than ordinary

The anatomy and histology of the atrioventricular conducting system in the hedgehog (Hemiechinus auritus) heart

238

Figure 1. The Hemiechinus auritus species of hedgehog studied

in this research.

Figure 2. Photomicrograph showing the location and shape of

the atrioventricular node (AVN); interatrial septum
(IAS); interventricular septum (IVS); fibrous ring (FR),
(green Masson’s trichrome staining, ×160).

myocardial fibers. The myofibrils of the nodal cells
were a little smaller than ordinary myocardial fibers.
As such, the difference in color between the node and
the surrounding myocardium was minimal (Figure
2). There was a framework of collagen fibers between
the AV nodal fibers. There were 2 types of cells in the
hedgehog AVN: P (pacemaker-like) cells and other
cells with darker cytoplasm. The cytoplasm of P cells
contained a perinuclear clear zone (Figure 4). There
were no detectable mucosubstances in the cells of the
atrial and ventricular myocardium, AVN, or AVB
(Figure 5).

The AVB was composed of many purkinje cells.

Myofibrils were located at the periphery of the cells
and a perinuclear clear zone was obvious, whereas the
other cells of the AVB had darker cytoplasm (Figure
6). Intercalated discs between the cells of the AVB
were present. Internodal pathways were not observed
in the hedgehog heart, but there were numerous
purkinje-like fibers within the myocardium of the
atrium and auricle (Figure 7). Several arterioles, nerve
fibers, and ganglions were present at the caudodorsal
section of the AVN and AVB to supply them.
Additionally, there was fibrous cartilage in the
hedgehog atrioventricular fibrous ring (Figures 8, 9).

A. NABIPOUR

239

Figure 3. Photomicrograph showing the AVB that is passing

through the fibrous ring. Atrioventricular bundle
(AVB); interventricular septum (IVS); fibrous ring
(FR), (green Masson’s trichrome staining, ×160).

Figure 4. Histological structure of the AVN in the heart

hedgehogs. Pacemaker like cells (P); darker cells (D);
collagen fibers (arrows), (green Masson’s trichrome
staining, ×640).

Figure 5. The photomicrograph does not show detectable

mucosubstances in the ventricular myocardium of
hedgehog, (Periodic Acid Schiff-Alcian blue staining
×640).

Figure 6. Showing the AVB in the heart of hedgehogs.

Atrioventricular bundle (AVB); interventricular
septum (IVS); fibrous ring (FR). Note the high number
of the purkinje fibers (arrows), (green Masson’s
trichrome staining, ×320).

Discussion

The anatomic location of the AVN in the

hedgehog heart was similar to that in rabbits (James,
1967), guinea pigs (Nabipour, 2004a), and ovine
fetuses (Nabipour and Shahabodini, 2007). Because
the ostium of the coronary sinus is so large in the
rabbit (James, 1967) and guinea pig (Nabipour,
2004a), the AVN is displaced anteriorly and occupies
the entire region, and the AVB is foreshortened. As
these animals normally have a left cranial vena cava,
the ostium of the coronary sinus (embryologically
derived from the terminal portion of the left cranial

vena cava in most mammals) is unusually large. This
effectively displaces the AVN and AVB anteriorly
toward the root of the aorta. However, in sheep
(Copenhaver and Truex, 1952), humans (Titus et al.,
1963), dogs (James, 1964), horses (Bishop and Cole,
1967), cattle (James, 1965), camels (Ghazi and
Tadjalli, 2002), cats (Tadjalli et al., 1999), and goats
(Nabipour, 2002) the AVN is located in the posterior
section of the interatrial septum, anterior to the
coronary sinus. The hedgehog AVN was oval, whereas
in ovine fetuses, as in adult sheep (Copenhaver and
Truex, 1952), the AVN is almost spherical. It is oval
or fan-shaped in humans (Titus et al., 1963), is like a
tiny spleen in dogs (James, 1964), has a flattened
oblong shape in horses (Bishop and Cole, 1967), is
ovoid in cattle (James, 1965), is an irregular elongated
oval in goats (Nabipour, 2002), is an irregular ellipse
in camels (Ghazi and Tadjalli, 2002), is an irregular
elongated oval in cats (Tadjalli et al., 1999), and is
almost spherical in guinea pigs (Nabipour, 2004a).
The AVN in avian hearts is not morphologically
definable (Szabo et al., 1986).

The hedgehog AVB was displaced anteriorly, near

the root of the aorta. This location is similar to that
in rabbits (James, 1967), guinea pigs (Nabipour,
2004b), and ovine fetuses (Nabipour and
Shahabodini, 2007). The shortness of the hedgehog
AVB is also similar to that in goats (Nabipour et al.,
2002), ovine fetuses (Nabipour and Shahabodini,
2007), and cattle and horses (Meyling and Terborg,

The anatomy and histology of the atrioventricular conducting system in the hedgehog (Hemiechinus auritus) heart

240

Figure 7. Showing numerous purkinje-like fibers within the atrial

myocardium in the heart of hedgehogs. Purkinje-like
fibers (arrows), (green Masson’s trichrome staining,
×640).

Figure 9. Fibrous cartilage in the right atrioventricular fibrous

ring of hedgehogs. Collagen fibers (CF); lacuna and
chondrocyte (arrow), (green Masson’s trichrome
staining, ×640).

Figure 8. Showing a parasympathetic ganglion near the AVN and

AVB of hedgehogs. Capsule (C); perikaryon (P); nerve
fibers (NF); amphicyte (arrow), (green Masson’s
trichrome staining, ×640).

1957). Due to the absence of the membranous part of
the interventricular septum in these animals, the AVB
is short; however, in animals in which the
membranous part is present, e.g. cats (Ghazi et al.,
1998), the AVB is long.

The AV node cells and their arrangement as a mass

of interlacing bundles interwoven with collagen fibers
in the hedgehog is similar to that in humans (Titus et
al., 1963), dogs (James, 1964), horses (Bishop and
Cole, 1967), cattle (James, 1965), camels (Ghazi and
Tadjalli, 2002), goats (Nabipour, 2002), cats (Tadjalli et
al., 1999), rabbits (James, 1967), guinea pigs
(Nabipour, 2004a), and ovine fetuses (Nabipour and
Shahabodini, 2007). The difference in color between
the node and ordinary myocardial fibers we observed
in the hedgehog is less than has been reported in other
animals, which is because there were more myofibrils
in the cytoplasm of the cells of the hedgehog AVN.
There is a small quantity of elastic fibers scattered
within the AVN in humans (Titus et al., 1963), dogs
(James, 1964), and cats (Tadjalli et al., 1999).

The number of P cells in the hedgehog AVN was

low, which is similar to other animals, while the AVN
of the guinea pig (Nabipour, 2004a) and ovine fetus
consisted of numerous P cells. The level of
carbohydrates in the AVN cells of ovine fetuses is high
(Nabipour and Shahabodini, 2007); however, there is
no glycogen in the AVN cells in goats (Nabipour,
2002), camels (Ghazi and Tadjalli, 2002), or guinea
pigs (Nabipour, 2004a). There is a small quantity of
nerve fibers within the hedgehog AVN; in this respect
it is similar to that in humans (Titus et al., 1963), dogs
(James, 1964), cats (Tadjalli et al., 1999), guinea pigs
(Nabipour, 2004a), and ovine fetuses (Nabipour and
Shahabodini, 2007). In contrast, in cattle (James,
1965), horses (Meyling and Treborg, 1957), and goats
(Nabipour, 2002) an abundance of nerve fibers are
present in the node. In the hedgehog heart, as in that
of humans (Titus et al., 1963), dogs (James, 1964),
horses (Bishop and Cole, 1967), cattle (James, 1965),
camels (Ghazi and Tadjalli, 2002), cats (Tadjalli et al.,

1999), rabbits (James, 1967), goats (Nabipour, 2002),
and ovine fetuses (Nabipour and Shahabodini, 2007),
ganglia are present in the posterior part of the AVN,
but not in the node. Additionally, there are no ganglia
at the periphery or within the node in the guinea pig.
In the present study internodal pathways in the
hedgehog heart were not observed, but there were
numerous purkinje-like fibers within the myocardium
of the atrium and auricle. The distribution of these
fibers suggests that they may be involved in the
interatrial spread of excitation. In humans (James,
1963), dogs (Glomset and Glomset, 1940), rabbits
(James, 1967), and guinea pigs (Nabipour, 2004a)
internodal pathways are connected to the margins of
the AVN.

Histologically, there were 2 types of cells in the

hedgehog AVB: purkinje cells and cells that did not
have the typical characteristics of purkinje cells. In
this respect it is similar to that in guinea pigs
(Nabipour, 2004b) and ovine fetuses (Nabipour and
Shahabodini, 2007). Typical purkinje cells
(Copenhaver and Truex, 1952), as seen in the AVB of
ungulates (James and Sherf, 1971), have a distinct
perinuclear light zone and a much larger diameter
than cardiac cells. Ungulate purkinje cells are almost
spherical or polyhedral, and make contact with other
cells along virtually their entire periphery, whereas the
cells in the AVB of canines and humans are elongated
and oblong, and make contact to some extent along
their lateral margins, but more often at their terminal
ends (James and Sherf, 1971). Partitioning of the AVB
was not observed in the hedgehog heart, which is in
contrast to the results reported for humans and other
animals.

Acknowledgements

The author wishes to express his appreciation to

the Ferdowsi University of Mashhad Research
Council for their financial support and to thank Mr.
Pouradibi for his technical assistance.

A. NABIPOUR

241

Bishop, S.P. and Cole, C.R. 1967. Morphology of the specialized

conducting tissue in the atria of the equine heart. Anat. Rec.
158: 401-416.

Copenhaver, W.M. and Truex, R.C. 1952. Histology of the atrial

portion of the cardiac conduction system in man and other
mammals. Anat. Rec. 114: 601-625.

References

The anatomy and histology of the atrioventricular conducting system in the hedgehog (Hemiechinus auritus) heart

242

Ghazi, S.R. and Tadjalli, M. 2002. Anatomy of the atrioventricular

node of camels (Camelus dromedarius). Iranian J. Vet. Res. 3:
93-99.

Ghazi, S.R. and Tadjalli, M. 1993. The anatomy of the atrioventricular

bundle in the heart of camels (Camelus dromedarius). Vet. Res.
Commun. 17: 411-416.

Ghazi, S.R., Tadjalli, M. and Baniabbas, A. 1998. The anatomy of the

atrioventricular bundle in the heart of domestic cats (Felis
catus). J. Fac. of Vet. Med., Univ. of Tehran, 53: 87-91.

Glomset, D.J. and Glomset, A.T.A. 1940. A morphologic study of the

cardiac conduction system in ungulates, dog and man. Am.
Heart J. 20: 389-398.

James, T.N. 1964. Anatomy of the AV node of the dog. Anat. Rec. 148:

15-27.

James, T.N. 1965. Anatomy of the sinus node, AV node and os cordis

of the beef heart. Anat. Rec. 153: 361-372.

James, T.N. 1967. Anatomy of the cardiac conduction system in the

rabbit. Circ. Res. 20: 638-648.

James, T.N. 1970. Cardiac conduction system: Fetal and postnatal

development. Am. J. Cardiol. 25: 213-225.

James, T.N. and Sherf, L. 1971. Fine structure of the His bundle. Circ.

44: 9-29.

Lev, M. and Lerner, R. 1955. A histology study of the normal

atrioventricular communications of the human heart. Circ. 12:
176-184.

Luna, L.G. 1968. Manual of histologic staining methods of the armed

forces institute of pathology, 3rd Ed., McGraw-Hill Book
Company, New York, PP: 94-95 and 168-169.

Meyling, H.A. and Terborg, H. 1957. The conducting system of the

heart in hoofed animals. Cornell Vet. J. 47: 419-447.

Nabipour, A., Khanzadi, S. and Banihassan, M. 2002. Anatomy and

histology of the atrioventricular bundle in the heart of goats
(Capra hircus). J. Appl. Anim. Res. 22: 155-160.

Nabipour, A. 2002. Anatomy and histology of the atrioventricular

node of goats (Capra hircus). J. Appl. Anim. Res. 22: 67-71.

Nabipour, A. 2004a. Anatomy and histology of the atrioventricular

node in the heart of guinea pig (Cavia percellus). Iranian J. Vet.
Res. 5: 204-209.

Nabipour, A. 2004b. Histology of the atrioventricular bundle in the

heart of guinea pig (Cavia percellus). Iranian J. Vet. Res. 5: 7-13.

Nabipour, A. and Shahabodini, M.R. 2007. Histological study of the

atrioventricular node and bundle in the heart of ovine fetus.
Iranian J. Vet. Res. 8: 64-70.

Nonidez, J.F. 1943. The structure and innervation of conductive

system of the heart of the dog and rhesus monkey as seen with
a silver impregnation technique. Am. Heart J. 26: 577-597.

Prakash, R. 1990. The heart and its conduction system in the lizard

Calotes versi color (Daudin). Anat. Rec. 136: 469-475.

Prasad, J. and Sinha, R.D. 1980. Histological and histochemical studies

on the right branch of atrioventricular bundle of Indian buffalo.
Indian Vet. J. 57: 373-376.

Szabo, E., Viragh, S. and Challice, C.E. 1986. The structure of the

atrioventricular system in the avian heart. Anat. Rec. 215: 1-9.

Tadjalli, M., Ghazi, S.R. and Baniabbas, A. 1999. Anatomy of the

atrioventricular node in the heart of cat. J. Appl. Anim. Res. 15:
35-40.

Titus, J.L., Daugherty, G.W. and Edwards, J.E. 1963. Anatomy of the

normal human atrioventricular conduction system. Am. J. Anat.
113: 407-415.

Titus, J.L. 1973. Normal anatomy of the human cardiac conduction

system. Mayo. Clin. Proc. 48: 24-30.

Lorraine A. Corriveau,DVM

Purdue University Veterinary Teaching Hospital

Small Animal Community Practice Clinician

Hedgehog Medicine

Description
The African pigmy hedgehog (Atelerix albiventris) has become a very popular pet in the early 1990’s.
They were a fad that hit the ‘get rich quick’ crowd initially but now has a small but very loyal following. At
first these pets sold for $120-200, but as more people bred them the prices dropped and everyone who
wanted them had them. It is not difficult to find them given up for adoption now. Now color variations
exist, hedgehog clubs have been formed, and shows are held. Clinical information is becoming more
abundant and easy to find.

Hedgehogs are members of the insectivore family. They possess 36-44 teeth with the first

incisors being notably longer than the rest and are spaced apart. The lower incisiors “fit” into this space
when the jaw is closed.

Dental formula: 2(3/2,1/1,3/2,3/3). Their teeth are brachyodontic like

carnivores. The most obvious feature of the hedgehog is the dermal spines. These are modified hairs
that provide protection from predators. Beneath the spines is a thick layer of subcutaneous fat.
Hedgehogs will roll up when alarmed into an impermeable ball of spines, making them clinically
challenging. They have sharply pointed snouts and typically small eyes. All possess a collarbone
(clavicle). They are nocturnally active, and like to hide during daytime hours. The olfactory and auditory
senses of all insectivores are highly developed, making them good hunters and foragers. Hedgehogs
have a plantigrade gait, meaning that they walk on the entire soles of the feet rather than the toes alone.
The most common pet species in the United States is the African pigmy hedgehog. Adults of this species
generally weigh 300-600 grams. Longevity is about 5-7 years in captivity. Some color varieties are
available now and if the pattern follows true, many genetic defects and a generally weakening of the
species may follow from the inbreeding practices used to produce these. Hedgehogs are easy to sex,
because males have a distinct prepuce like in dogs but testicles are abdominal. Females have a vulva.
Hedgehogs are induced ovulators year round. Litters of 3-6 young are produced following a gestation of
34-37 days. The offspring have a protective coating over the spines during parturition, which is lost in the
first 24 hours to expose the spines. Babies wean in 30-40 days and become sexually mature at about 2-3
months of age.
Diet
In the wild, hedgehogs consume insects, small vertebrates, and carrion. In captivity, hedgehogs fare
well on formulated hedgehog or low fat dog or feline diets. Some recommend soaking the dry food prior
to feeding. Reduced calorie formulations should be used, as obesity is the most commonly encountered
nutritional problem. Fruits and vegetables should be added to the diet to dilute out the high calorie foods
and to offer extra fiber. Cottage cheese, eggs, and other such protein sources can be given to breeding
hedgehogs, but should be avoided or limited in sedentary animals. Insects are relished and can be given
as a treat, but are calcium deficient and due to improper Ca:P ratio should not be the sole diet.
Recommend feeding once daily in the evening. Also encourage evening exercise to prevent obesity.
Housing
Usually caged alone but can be housed in groups if given enough space. Excellent climbers so cages
need to be smooth walled and high enough to prevent escape. It is recommended to avoid wire floors.
Solid floors should have bedding such as newspaper, recycled newspaper, or wood shaving (pine or
aspen). A hide/sleeping area should be provided. One can be creative in the type of hide/sleep area
provided.

Examination
Hedgehogs can be very difficult to work with as patients. Their ability to roll up into a ball makes them
impossible to examine if they do not want to be examined. Lightweight leather gloves should be used to
prevent the spines from pricking the handler’s hands. Sometimes, an aquarium with a shallow layer of
water can be used to allow for visual examination. When the hedgehog is placed in the water, it must
unroll so as to not breathe the water. This helps keep the hedgehog unrolled for examination. Some
hedgehogs uncurl with back stroking of the rump spines. Anesthesia is almost always required for a
complete examination. Isoflurane or Sevoflurane are the recommended gas anesthesia. A small mask,
fashioned from a syringe case can be slipped into the opening of the ball and over the snout.
Alternatively, the hedgehog can be placed in an anesthetic chamber for induction. Once anesthetized,
the mask can be positioned better or an endotracheal tube can be placed (very challenging). At this point
the hedgehog can be easily examined. The examination should be systematic. A visual of the eyes,
nose, ears, oral cavity, teeth, spines, anal and urogenital openings, palpation of lymph nodes and
abdominal cavity, and auscultation of the thorax and abdomen will detect most disorders. In many cases,
the problem will require further evaluation.
Clinical Pathology
Hematology and serum biochemistry are often the first stop in evaluating more vague illnesses in
hedgehogs. Blood collection can be a bit challenging but can be mastered with a bit of practice. The
jugular or cranial vena cave is generally used for venipuncture. The jugular is relatively short and runs
from the thoracic inlet to the ear. A large lymph node lies just lateral to the jugular. It usually cannot be
palpated, but can be blindly entered in the anesthetized hedgehog. With the hedgehog anesthetized and
in dorsal recumbency, the neck should be extended. While it cannot usually be visualized, the jugular
vein will generally course from the manubrium sterni to the ear. Large lymph nodes lie just lateral to the
vein and serve as a landmark. The jugulars should be entered blindly with slight suction on the syringe
until a flash is seen. Slow and steady pressure is used until the desired quantity of blood is obtained.
One percent of the body weight can be taken, but 1ml should be adequate for routing analysis. Pressure
should be applied briefly following venipuncture. Fecal examination, urinalysis, cytology are run and
interpreted in the same manner as the more familiar mammals. Clinical pathology data is limited but
empirically, most values are similar to canine and feline patients.
Radiographs
Radiographs are most easily taken in the anesthetized hedgehog. A ventrodorsal and lateral view can be
taken on a single small film. It may be beneficial on the lateral view to elevate the excess fat pad that
holds to dorsal spines. The can be accomplished by using a “chip clip”. Knowledge of normal anatomy is
necessary for interpretation of radiographs. A radiographic atlas of exotics is available for normal
anatomy.
Therapy
The diagnosis is not the only challenge in hedgehog medicine. The same defense that prevents
examination can also make treatment difficult. Once a diagnosis is made, or at least initial diagnostic
procedures are finished, therapy should be started. Early treatment is crucial to success. The small size
of hedgehogs makes them very susceptible to starvation and dehydration. If they are not eating or
drinking, they should be force-fed and administered fluids. If shocky or critically dehydrated, fluids should
be given intraosseously. A needle can be placed in the femur, in the same fashion as an intramedullary
pin, and fluids or drugs can be administered in this fashion. The fluids are taken up into circulation so
rapidly that this technique is equivalent to intravenous infusions, which is very difficult in hedgehogs.
Less severely ill pets can be given subcutaneous or oral fluids. Maintenance fluid requirements are 60-
100ml/kg/day. Therefore a 400gram hedgehog will need 24-40ml per day for maintenance plus the
deficit. Force-feeding energy requirements can be calculated with the following formula:

Basal energy requirement (BER) =

68*(body-weight in kg

.75

)

Maintenance energy requirement (MER) =

1.25*(BER)

Actual energy requirements will vary from 1-2 times maintenance energy, depending on the medical
condition. A 400gram hedgehog will require 45-90 kcal of energy per day for maintenance. Using the
higher calorie formula, this means about 23-45ml per day.

Normal room temperature should be used for housing hospitalized mammals unless they are

hypothermic or hyperthermic. Hedgehogs will go into a torpid state when cold which could be dangerous
if they are sick. Care should be taken, however, that they are not overheated in an avian or reptile
intensive care unit. Minimizing stress is critical and somewhat more difficult when treating ‘prey’ type

species. These secretive creatures will feel more comfortable when adequate hiding spaces are
provided. They preferably should be kept in a quieter location, away from barking dogs.

Information regarding drug dosing and efficacy is largely anecdotal at this point. There are no

label-approved drugs for hedgehogs. All medications should be used with caution and with informed
consent of the owner. Generally drugs and dosages published for ferrets are appropriate for hedgehogs.
When exotic animals are treated, follow up evaluation takes on a critical role since we are still feeling our
way along. The safest choice of available drugs should always be made. Often, injectable medications
are preferred since they can be given when the hedgehog is in “spiny ball” mode. Oral medications are
given in liquid suspensions or solution form. Very tame hedgehogs can be medicated easily by their
owners, but some will have to have the medications put on a favorite food item. Eye and ear medications
are very difficult to administer. Solution preparations are easier to apply but ointments will last longer and
require less frequent administration.

Dental care for hedgehogs is difficult. Few owners can brush the teeth and yet periodontal

disease, tooth root abscesses, and decay are all common problems. Recommendations include using
dry food formulated for dental care, using crisp vegetables, and if possible, the use of oral cleansing gels
(example: Maxiguard®).

Client education is critical to proper treatment. In addition to demonstrating the proper method of

treatment, owners should be informed of predisposing factors and how to correct them.
Anesthesia
Certain diagnostic and therapeutic procedures may require anesthesia for restraint and prevention of
pain. In most cases, isoflurane gas, administered by mask is the simplest, safest, and most rapid
method. Endotracheal intubation is difficult and requires that the hedgehog be deeply anesthetized, so
not recommended for novice clinicians. About 1.5-2 mm endotracheal tube is needed for an adult
hedgehog. Injectable anesthetics carry the inherent disadvantage of greater difficulty to control depth of
anesthesia. Hedgehogs loose much heat more rapidly, the tracheal lumen occludes more easily, the
patient is more difficult to monitor. A supplemental heat source is essential, and cotton tipped applicators
should be available to swab out the throat. A small endotracheal tube should be available for emergency
intubation. Clear adhesive drapes facilitate monitoring. Most importantly, a technician should be
dedicated to the constant monitoring of the patient. The tidal volume is generally too low to move the bag
on most systems so respiration cannot be monitored in this way. The low tidal volume also leads to a
large amount of dead space within the delivery system. Semi-open, non-rebreathing systems must be
used for hedgehogs. Special bags are available that adjust to the small tidal volume. Alternatively, a
balloon can be used as a bag on the non-rebreathing circuits. A respiratory monitor helps detect early
changes in respiratory rate or tidal volume. An ECG facilitates monitoring well.
Surgery
Preparation of the surgical site can be a challenge if surgery is performed on the dorsal part of the body.
The spines must be removed so a surgical field can be formed. They can be plucked or clipped at the
base. These spines are very difficult to pluck. Clipping is done with a pair of scissors just at the base,
leaving a little behind. The skin is then prepared as in other animals. Surgery of hedgehogs can be
enhanced by the use of several types of instrumentations not commonly used in traditional pets. Their
small size requires finer instruments, methods of controlling small amounts of hemorrhage, magnification
and directed source of light. Microsurgical or ophthalmic instruments are frequently used. Microsurgical
instruments should be counterbalanced and have rounded handles to allow them to be manipulated by
gently rolling them between the fingers. There should be no locking mechanism on needle holders as
releasing these causes considerable jarring. Delicate surgery should be performed while seated with the
wrists supported on the table to minimize motion. Hemorrhage can be controlled by the use of
electrocoagulation. Bipolar instrumentation is preferred. Vessels must be isolated and then coagulated.
If bipolar instruments are not available, the vessel can be isolated with forceps and the electrode is then
touched to the forceps. Care must be taken not to cause excessive time destruction. Ligatures will be
required on larger vessels (>2mm). Small sizes of suture (4-0 to 10-0) can be used for these when they
are in an accessible area. Vasculare clips (Hemoclips) are preferred when working in a restricted area
such as the body cavity or when speed is required. Occasionally hemorrhage will occur when none of the
above will be applicable. In these cases, the area can be packed off with absorbable foam sponges (Gel
Foam). Due to the small size of hedgehogs, magnification of the surgical field is advantageous. Optical
loupes can be used for many procedures and are reasonably priced. Operating microscopes will provide
greater magnification and also lighting and are very beneficial for many procedures. Working under
magnification is very different from standard surgery. Every movement is magnified and the field is very
restricted. It is sometimes difficult to even find instruments. Surgical procedures performed under a

microscope can be done with much greater precision, but they also take more time. Considerable
practice is required to master microsurgery. The most commonly performed surgeries are tumor removal
procedures and pyometra sugery.
Common Problems
Ectoparasites are the single most commonly presented problem of hedgehogs. Mites, especially
Chorioptes and Caparina sp. are very common in hedgehogs. These parasites are just barely visible to
the naked eye and the crusts can be seen to move in heavily infested hedgehogs. Loss of spines,
pruritus, and scaling of the skin are the primary clinical signs of this disease. A skin scraping should be
performed routinely on hedgehogs as these mites can take a long time to cause overt disease. When
diagnosed, hedgehogs should be treated with an ivermectin product. A dose of 0.5mg/kg given
subcutaneously once weekly for 3-6 weeks is generally effective in eliminating the parasite. Revolution



(Selamectin) used topically at 9mg/pound once and then repeated in 3 weeks has shown good success
also in the treatment of these mites.

Endoparasites were quite common in imported hedgehogs but appear to be uncommon in the

domestic raised ones commonly encountered in private practice now. Fecal examinations should be
routinely performed to determine if any are present and then they should be treated appropriately.

Periodontal disease occurs frequently in hedgehogs. This also frequently progresses to tooth rot

abscesses. Affected hedgehogs will lose weight, show abnormal feeding behavior, may have swelling of
the jaw, maxilla, or periorbital region or may become anorectic. A thorough examination of the mouth,
including probing the periodontal pockets will detect most problems. Skull or dental radiographs may be
needed in some cases. Minor cases may respond to supragingival and subgingival scaling, antibiotics,
and nutritional support. More often, extensive extractions must be performed. Hedgehogs appear to do
well with few or no teeth. If no teeth are present, softer foods may be needed.

Hedgehogs are emerging as rivals of budgerigars, ferrets, and boxers in the arena of tumor

production. Neoplasias of many different types have been reported in hedgehogs, despite the relatively
sparse literature on them in general. Squamous cell carcinoma of the oral cavity and lymphosarcoma of
any body part are over represented in the literature. Whenever a lump is encountered, it should be
aspirated or removed for histopathology. Vague clinical signs of illness often result in a final diagnosis of
cancer. Thorough palpation of the lymph nodes, neck, abdomen, and auscultation of the thorax should
be done on each exam. Radiographs and ultrasound are helpful for early detection or gaining further
information about a tumor. Currently, the treatment of choice for tumors is surgical resection. If done
early, this can achieve a cure in many cases.

Obesity is a common occurrence in pet hedgehogs. The combination of unlimited supply of high

calorie food and the lack of exercise results in a calorie excess (fat deposition). Some are so fat that they
cannot or have difficulty rolling into a ball. Treatment of obesity includes changing the diet to a low calorie
cat/dog food, limiting the amount given to 2/3 of the current ration, giving fresh fruits and vegetables for
fiber, and increasing exercise by giving a wheel and allowing more play-time outside of the cage.

Trauma can also occur to the limbs of hedgehogs. One must be careful to not have string-like

material on the floor or in the cage. There have even been reports of even human hair causing
constriction bands around the limbs/digits leading to avascular necrosis.

Enteritis has been sporadically encountered in hedgehogs. Often clostridial organisms are seen

on cytological examination of the feces. Amoxicillin and metronidazole are both affective in elimination of
the clostridium and the diarrhea in most cases.
Preventive Medicine
Client education is the most important key to maintaining a healthy hedgehog. A combination of written,
spoken, and visual aids are usually necessary for adequate retention of information. Information about
housing, nutrition, sanitation, behavior, and health care should be given to each client.

Nutrition is key to keeping any animal healthy and with exotic pets, where formulated diets are not

available; the task is much more difficult. Common sense is important. A diet that would not be balanced
for more familiar species would not be for hedgehogs as well.

Parasite control should be routinely applied/performed. Fecal samples should be checked once

or twice annually and any parasites treated. Skin scrapings are very important for detecting external
parasites.

Early detection and treatment of diseases is critical. Small “prey” species try to hide any signs of

illness or weakness as a mechanism to prevent predation. In captivity, this trait leads to presentation late
in the course of the disease. A hedgehog that looks sick is often very sick. A “wait and see what
happens” approach is very dangerous in exotic animals. Rapid diagnosis and treatment is essential.

Suggested reading
1. Gregory MW, Stocker L: Hedgehogs. In Beynon PH, Cooper JE (eds); Manual of exotic pets.
Gloustershire, England, British Small Animal Veterinary Association, 1991, pp 63-68.
2. Hoefer HL: Hedgehogs. In Vet Clin North Am Sm Animal Pract 24 (1)pp 113-120, 1004.
3. Reeve, N: Hedgehogs. T&AD Poyser Ltd, London, 1994.

• 435-644-2001 • www.bestfriends.org

Caring for Your Hedgehog

By Mark Burgess, DVM

Hedgehogs are small, shy, insect-eating mam-
mals with quills on top and fur on the face and
belly. The quills resemble those of porcupines,
but are not barbed and do not imbed in the skin
when touched. When frightened, hedgehogs roll
into a ball and emit an odd rattling hiss. They
may bounce to try to poke you with their quills,
but they rarely attempt to bite. They can make
decent pets if socialized and they typically live
two to four years.

There are many species of hedgehogs, including the large European hedgehog and
many African species. The African pygmy hedgehog is the species commonly acquired
as pets. It is illegal to have hedgehogs as pets in certain areas of the U.S., so please
check your state, city, and county ordinances before getting a hedgehog.

If you do decide to get one, please adopt from a rescue group rather than buying from a
pet store or breeder. There are many wonderful hedgehogs out there just waiting to be
adopted. To ﬁnd a hedgehog rescue, do a search for “hedgehog rescue” on the Internet
or visit the Hedgehog Welfare Society at

www.hedgehogwelfare.org

Housing

Since African hedgehogs are native to warm regions, they do not tolerate cold tempera-
tures very well. They should be kept above 70 degrees Fahrenheit.

They may be housed in wood or wire cages with solid ﬂoors; a minimum cage size is 24
x 24 inches; cage height is not critical. Avoid wire ﬂoors, since their feet may become
caught and bones broken as the pet tries to free himself. Wire ﬂoors also tend to cause
sores on the feet. Provide a thick layer of good bedding, such as recycled paper or
hardwood shavings (not cedar or pine). Soft, clean towels can be used, but should be
removed if your pet chews them or if they become frayed; the hedgehog may become
tangled in any loose threads.

Hedgehogs are quiet animals, but they can be very active, especially at night. Ideally,
give them daily exercise outside of the cage for at least 30 minutes. Hedgehogs occa-
sionally can be aggressive with each other if housed together, so they may need to be
kept separate to prevent ﬁghting, especially with males.

Feeding

The bulk of the hedgehog’s diet should be low-fat hedgehog pellets (not a seed and fruit
mix). Pellets may be fed free-choice, unless obesity occurs. Use a formula with no more
than 7 percent fat – Pretty Pets is a good brand of food. You can also use a very low-fat
cat food, such as Hill’s W/D (7 percent fat). The hedgehog’s natural diet is mostly bee-

• 435-644-2001 • www.bestfriends.org

tles. In captivity, occasional insects such as crickets or mealworms can be fed, but these
are not nutritionally balanced and should be used sparingly. Invertebrates with better
nutritional content would include slugs, earthworms, and silkworms.

Fresh water, of course, should always be available. Ball-bearing bottles are cleaner and
not as easily spilled as bowls. Remember to clean your hedgehog’s cage, food bowls
and water sources regularly.

Common Diseases

Obesity. The most common medical problem seen in captive hedgehogs is obesity. The
usual cause is feeding high-calorie foods, such as regular cat food, seeds or nuts, or
fatty insects such as mealworms and crickets. But hedgehogs can overeat even on a
low-fat hedgehog diet, so food intake often must be restricted to maintain proper body
weight. Healthy hedgehogs should appear twice as long as they are wide; they are not
round in shape, except when rolled up. Obesity increases the risk of tumor development
and may shorten your pet’s lifespan. Weight loss should be accomplished slowly, since
obese pets who lose weight too rapidly may develop liver disease.

Weight loss. Because hedgehogs are aggressive eaters and rarely lose weight unless
put on a strict diet, spontaneous weight loss is a cause for concern. It usually indicates
signiﬁcant illness, such as dental disease, cancer, heart disease, or uterine disease.
Seek immediate veterinary advice if you notice your pet losing weight for no apparent
reason.

Dental diseases. Hedgehogs develop gum disease and tooth infection with age. Signs
may include salivation, difﬁculty eating, and bad breath. Infected teeth may be loose and
painful. Treatment is cleaning and/or extraction of the affected teeth, and oral antibiot-
ics. Oral odor and salivation can also be a sign of oral cancer, which is common in older
hedgehogs.

Mites. These are common skin parasites, but they often produce no symptoms until the
mite population grows large. When numerous, the microscopic mites cause itching, ﬂak-
ing, and quill loss. A hedgehog with a severe case of mites may develop scabs or sores
from intense scratching and biting at the skin. Mites respond to treatment with ivermectin
(oral or injectable) given weekly for 6-8 weeks. Lyme sulfur dip may help when applied
once or twice weekly for 6-8 weeks, but it is more labor-intensive and must be applied
thoroughly.

Cleaning the cage weekly when treating the mites may help reduce re-infestation, but
long-term environmental treatment is unnecessary, since the parasites die if they are off
their host for long. These parasites are species-speciﬁc but highly contagious between
hedgehogs, so use caution when introducing new pets to an existing group.

Respiratory infection. Signs include sneezing, wheezing, lethargy, nasal or eye dis-
charge, or difﬁculty breathing. Various bacteria may cause this type of infection. Treat-
ment is with antibiotics. You can minimize your hedgehog’s risk of respiratory disease
by providing her with a warm, clean cage; avoiding the use of wood-chip beddings; and
feeding her a balanced diet.

• 435-644-2001 • www.bestfriends.org

Head tilt (wry neck, torticollis). This is usually due to internal ear infection, and is
sometimes secondary to a respiratory infection. The hedgehog usually tilts his head to
one side and loses his balance, often falling or circling when trying to walk. Bacteria are
the usual cause and treatment is with antibiotics.

Fight wounds. Hedgehogs can occasionally be aggressive with each other, and some-
times ﬁght if housed together. This may result in bite wounds. Any visible wounds require
immediate medical treatment; the risk of infection is great, and early antibiotic therapy to
prevent infection is the safest option.

Cancers. Older hedgehogs are highly prone to many types of cancer. Common types
include oral tumors, and mammary tumors (breast cancer) in females. Any visible lump
should be checked immediately by a veterinarian. Oral odor, drooling, or difﬁculty eat-
ing are also cause for concern. Many tumors are curable if caught early and removed.
Weight control may reduce the risk of some tumors. Spaying female hedgehogs most
likely reduces the risk of mammary tumors, and eliminates the risk of uterine cancer.

Heart disease (cardiomyopathy). This degenerative disease of the heart is seen in
many pet species, including dogs, cats and ferrets, and also in humans. The causes are
unknown, but some forms in dogs and cats have been linked to nutritional deﬁciencies.
Signs of heart failure include bloating, lethargy, and difﬁculty breathing. Treatment may
control symptoms for months, but isn’t likely to cure the disease. The risk of heart dis-
ease might be reduced by feeding a balanced diet and preventing obesity.

Progressive paralysis (degenerative myelopathy). This poorly understood disease
of unknown origin causes slow deterioration of the spinal cord. A gradual weakness and
paralysis begins in the rear legs and usually progresses to the front legs over time. Total
paralysis can result. There is no effective treatment in most cases, and the condition is
often fatal.

Hair or thread entanglements. Hedgehogs are prone to becoming entangled in long
pieces of thread (from bedding, such as frayed towels) or in strands of their people’s
hair. The strands wrap around a leg or foot (or occasionally the penis in males) and act
like a tourniquet, cutting into the skin and cutting off blood ﬂow. Infection and loss of the
foot may result. Minimize exposure to long hairs, strings or threads in the cage environ-
ment. If your pet is limping or has a swollen foot, seek immediate veterinary care.

Veterinary Care

No vaccines are given, but regular exams are recommended for early disease detection.
Have an exam done when you ﬁrst get your hedgehog, then at one year old, then every
six months after that. With good care, your hedgehog can be a happy and lovable pet!

Dr. Mark Burgess is owner of Southwest Animal Hospital/The Exotic Animal Practice in
Beaverton, Oregon. Ninety-ﬁve percent of his practice is small exotic pets, including fer-
rets, rabbits, rodents, reptiles, hedgehogs, marsupials, and some wildlife. He lectures at
conferences and has published articles on exotic pet disease in medical journals.

Introduction to the African Pygmy Hedgehog

Heidi L. Hoefer, DVM, ABVP

Introduction

Hedgehogs are small mammals that belong to the order Insectivora. There are

several species that are found throughout the British Isles, Europe, and Asia and
Africa. Hedgehogs are not native to North America. The African pygmy hedgehog

(Atelerix albiventris) originated in the African
savanna, and is now widely bred in North
America for the pet industry. The European
hedgehog (Erinaceus europaeus) is a larger
species, native to England and Europe and
protected by law in many areas.

The African pygmy hedgehog is small,

nocturnal, and spiny-coated. They vary in color

from brown to almost black with a white ventrum. The adult ranges in weight from
300-600 grams (1 lb = 450 grams). Life expectancy averages 3-4 years in the wild
but up to 10 years in captivity.

Anatomy & Physiology

Hedgehogs are characterized by the short, grooved white and brown spines

that cover the upper part of the body. The face and belly are covered with soft, light-
colored fur. The hedgehog can assume a defensive posture by rolling up and
erecting the spines to resemble a tight ball of sharp spines.

Gender (sex) is easy to identify in hedgehogs. The male has a penis and

prepuce located midway along the abdomen. The testes are usually intra-abdominal
and are not easily seen. The female has a vulva located close to the rectal opening.
African hedgehogs breed year-round. The gestation period is 34-37 days. Litter size
ranges from 1-7 with an average of 3-4 pups. The young are born blind with soft
white spines. New brown spines appear in 2-3 days and the eyes open in 2-3
weeks. Weaning occurs at 4-6 weeks.

In their native habitat, a hedgehog will dig its burrow under logs, leaves, among

rocks, or tree roots and sleep most of the day. They are solitary and nocturnal,
emerging at dusk to forage for insects. When undisturbed, it moves with an
unsteady, waddling gait but can run quickly. The hedgehog has a keen sense of
hearing and smelling, making it an adept hunter and forager. The European species
will hibernate in the winter and the African species will aestivate in the hot, dry
season. Hibernation is not essential and is not recommended for captive hedgehogs.

Housing & Diet

Hedgehogs are usually caged alone but can be housed in groups if given

enough space. Hedgehogs are excellent climbers, so cages should be smooth-
walled and high enough to prevent escapes. Wire flooring should be avoided due to
the potential for toe and limb injury. Newspaper or wood shavings (pine or aspen)
can be used as bedding, but it must be changed frequently. A sleeping area can be
made from cardboard boxes, hollowed logs, wooden boxes, or plastic flowerpots and
filled with hay or leaves.

The native diet consists of a variety of insects, occasional small vertebrates,

and carrion (dead animals). Captive pets can be fed a diet of soaked low-fat dog or
cat chow, smaller amounts of mealworms or earthworms, and a small amount of
chopped fruit and vegetables. Because of the calcium-phosphorus imbalance, a diet
solely of insects must be avoided. Hedgehogs should be fed once daily in the
evening. Captive hedgehogs have the tendency to become obese; early evening
exercise should be encouraged.

Physical Examination & Restraint

Hedgehogs typically roll-up during clinic visits making a full physical

examination impossible without sedation. Light leather gloves are often used to
protect against the sharp spines. Some hedgehogs can be scruffed behind the ears
if caught before they ball up, however, the majority will need to be sedated.
The preferred method of sedation is isoflurane gas. The hedgehog is initially placed
in a small plastic box and then switched to a face mask to allow inhalation of the gas.
Isoflurane is generally safe for most hedgehogs.

Common Conditions And Diseases

There is very little information in the literature regarding the diseases of pet

African pygmy hedgehogs. The following list of conditions represents a review of the
literature as well as the author's own clinical experience.

Cancer of almost any body part, but especially the mouth and skin

Dental disease

Heart disease

Kidney failure

Leg and foot injuries

Obesity

Overgrown nails

Quill loss (mites are a common cause)

Respiratory problems

References
1. Allen ME: The nutrition of insectivorous mammals. In Proceedings of the Annual
Meeting of the American Association of Zoo Veterinarians, Oakland, CA 1992, pp
113-115
2. Done LB, Dietze M, Crnafield M, et al: Necropsy lesions by body systems in
African hedgehogs: Clues to clinical diagnosis. In Proceedings of the Annual
Meeting of the American Association of Zoo Veterinarians, Oakland, CA 1992, pp
110-112
3. Gregory MW, Stocker L: Hedgehogs. In Beynon PH, Cooper JE (eds): Manual of
Exotic Pets. Gloustershire, England, British Small Animal Veterinary Association,
1991, pp 63-68
4. Hoefer HL: Hedgehogs. In Vet Clin North Am Sm Anim Pract 24 (1) pp 113-120,
1994
5. Isenbugel E, Baumgartner R: Diseases of the Hedgehog. In Fowler, ME (ed): Zoo
and Wild Animal Medicine, Current Therapy 3. WB Saunders, Philadelphia, 1993,
pp 294-302
6. Smith AJ: Husbandry and medicine of African hedgehogs. J Small Exotic Anim
Med 2: pp 21-28, 1992.

213

VETERINARSKI ARHIV 72 (4), 213-220, 2002

Contact address:

Dr. Zait Ender Özkan, Department of Anatomy, Faculty of Veterinary Medicine, Fýrat University, 23159, Elazýđ,
Turkey. Phone: +90 424 237 0000, Fax: +90 424 238 8173

ISSN 0372-5480
Printed in Croatia

Macro-anatomical investigations on the skeletons of hedgehog

(Erinaceus europaeus L.). II. ossa membri pelvini

Zait Ender Özkan*

Department of Anatomy, Faculty of Veterinary Medicine, Fýrat University, Elazý

g, Turkey

ÖZKAN, Z. E.: Macro-anatomical investigations on the skeletons of hedgehog

(Erinaceus europaeus L.). II. ossa membri pelvini. Vet. arhiv 72, 213-220, 2002.

ABSTRACT

In this study, three adult male hedgehogs (Erinaceus europaeus Linnaeus) were used to

investigate the bones of hind limb. The spina iliaca ventralis caudalis was absent. The symphysis
pelvis was formed by symphysis pubis and it was an interpubic ligamentous tissue in hedgehogs.
The average Vialleton angle was measured as (8º). Average distance between the midacetabulum
and tuber coxae was 39.2 mm and average distance between the midacetabulum and ischial
tuberosity (tuber ischiadicum) was 13.8 mm. There were three trochanters on the femur. The
tibia and fibula were fused almost in the distal half. There were 7 tarsal bones and the pedis was
complete with five digits.

Key words: hedgehog, Erinaceus europaeus L., ossa membri pelvini

Introduction

Hedgehogs (Erinaceus europaeus L.) belong to the Erinaceidae family,

order Insectivora (

VAUGHAN, 1972; DEMÝRSOY, 1996; 1997; 1998

). The most

important features of hedgehogs are the bristles transformed to spine form
on the dorsal and lateral sides of the body, forming a quite round body

214

Vet. arhiv 72 (4), 213-220, 2002

Z. E. Özkan: Macro-anatomical investigations on the skeletons of hedgehog (Erinaceus

europaeus L.). II. ossa membri pelvini

when threatened and in order to sleep in winter under a 4

°C environmental

temperature (

DEMÝRSOY, 1997; 1998

).The literature on the macro-anatomical

features of the skeletal system in hedgehogs is very meagre.

There have been some macro-anatomical investigations carried out on

the skeletal systems of wild animals such as lagomorphs (

DUBRUL, 1950

African rhinocerus (

GOMERČIĆ and HUBER, 1982

), hyena (

TECÝRLÝOĐLU,

1983

), wolf and fox (

GÝRGÝN et al., 1988

), mink (

DURSUN and TIPIRDAMAZ,

1989

), raccoon dogs and badgers (

HIDAKA et al.,1998

), porcupine (

YILMAZ et

al., 1998; 1999

), otter (

DÝNÇ et al., 1999

), but the skeletal systems of hedgehogs

of the order Insectivora have not been investigated in detail.

The aim of this study is to investigate the ossa membri pelvini part of

the skeletal system in hedgehogs and to contribute to the fund of information.

Materials and methods
The bones examined were obtained from three adult male hedgehogs

caught by the villagers in Elazýđ. Maceration of bones was carried out by
the method of

BARTELS and MEYER (1991)

and TAŢBAŢ and TECÝRLÝOĐLU

(1966)

For terminology, Nomina anatomica veterinaria (

ANONYMOUS, 1994

)

was used.

Vialleton angle (

LESSERTISSEUR and SABAN, 1967

) was measured as an

angle between the line lying from the midacetabulum to the midpoint of the
crista iliaca and the line lying from the midacetabulum to the centre of the
facies auricularis.

Results
Os coxae. Linae glutae were not prominent. Spina iliaca dorsalis

cranialis, spina iliaca dorsalis caudalis, spina iliaca ventralis cranialis were
prominent; spina iliaca ventralis caudalis was not present. The spina
ischiadica was not well developed. The great sciatic notch (incisura ischiadica
major) was deeper and wider than the lesser sciatic notch (incisura
ischiadica minor). The iliac tuberosity (tuberositas iliaca) on the sacropelvic

215

Vet. arhiv 72 (4), 213-220, 2002

Z. E. Özkan: Macro-anatomical investigations on the skeletons of hedgehog (Erinaceus

europaeus L.). II. ossa membri pelvini

surface (facies sacropelvina) was prominent, and facies auricularis was
formed as a small area.

Tuber ischiadicum was prominent and had a single process. There was

a small notch on the caudal side of the tabula ossis ischii. The symphysis
pelvis was formed by symphysis pubis and was in the form of a ligament
connecting the two caudal branch of the pubic bones (ramus caudalis ossis
pubis) (Fig. 1).

The Vialleton angle was measured as (8ș). Average distance between

the midacetabulum and tuber coxae was 39.2 mm, and the average distance
between the midacetabulum and ischial tuberosity (tuber ischiadicum) was
13.8 mm. The average sagittal length and width of the foramen obturatum
were 11.4 mm and 6.6 mm, respectively.

Fig. 1. Ventro-lateral aspect of os coxae of hedgehog (Erinaceus europaeus L.)

a) ala ossis ilii, b) crista iliaca, c) spina iliaca dorsalis cranialis, d) spina iliaca dorsalis
caudalis, e) spina iliaca ventralis cranialis, f) acetabulum, g) incisura acetabuli, h) for.
obturatum, ý) tabula osis ischii, i) a small notch on the caudal side of the tabula osis ischii,
j) incisura ischiadica major, k) incisura ischiadica minor, l) tuber ischiadicum, m) interpubic
ligamentous tissue.

216

Z. E. Özkan: Macro-anatomical investigations on the skeletons of hedgehog (Erinaceus

europaeus L.). II. ossa membri pelvini

Vet. arhiv 72 (4), 213-220, 2002

Femur. There were three trochanters on the femur: the greater

trochanter (trochanter major), the lesser trochanter (trochanter minor) and
the third trochanter (trochanter tertius). The trochanteric fossa (fossa
trochanterica) was wide and deep and the trochanteric ridge (crista
intertrochanterica) was present between the lesser and the greater
trochanters. Condylus lateralis, condylus medialis, epicondylus lateralis,
epicondylus medialis, linea and fossa intercondylaris were prominent (Fig.
2).

Fig. 2. The femur, ossa cruris and the patella in hedgehog (Erinaceus europaeus L.)

i- caudal aspect of the femur; ii- cranio-lateral aspect of the tibia and fibula

a) caput ossis femoris, b) trochanter major, c) trochanter minor, d) trochanter tertius, e)
crista intertrochanterica, f) fossa trochanterica, g) condylus lateralis, h) condylus medialis,
ý) epicondylus lateralis, j) epicondylus medialis, k) linea intercondylaris, l) fossa
intercondylaris, m) tibia, n) fibula, o) tuberositas tibiae, p) patella, r) facies cranialis, s)
basis patellae, t) apex patellae

Patella. The cranial surface of the patella was convex; apex patellae

was pointed.

Skeleton cruris. The tibia and fibula were fused almost in the distal

half and there was a wide spatium between the tibia and fibula in the

217

Vet. arhiv 72 (4), 213-220, 2002

Z. E. Özkan: Macro-anatomical investigations on the skeletons of hedgehog (Erinaceus

europaeus L.). II. ossa membri pelvini

Table 1. Specification of myomorphus mammals examined by renoculture and

microscopic agglutination acording to the trapping area with corresponding results

proximal half. Tuberositas tibiae was prominent. There was a prominence
near the lateral malleolus and the cochlea tibiae was sagittal (Fig. 2).

Ossa tarsi. There were 7 tarsal bones. The proximal row consisted of

the talus and calcaneus. Os tarsi centrale was in the distal of the talus.
Facies articularis navicularis of the talus was convex and there was a
small pit on the trochlea tali. The distal row bones from medial to lateral
were os tarsale I, os tarsale II, os tarsale III, and os tarsale IV. The
comparative sizes of the distal tarsal bones were: IV>I>III>II.

Ossa metatarsalia I-V. The pedis was complete with five digits and

there were five distinct metatarsal bones lying between the tarsal bones
and phalanges. The comparative lengths of the metatarsal bones were:
IV>III>II>V>I.

There were two plantar located sesamoid bones in pairs at each of the

metatarsophalangeal joints.

Ossa digitorum pedis. There were two phalanges in the first and fifth

digit and the other three digits comprised three phalanges. The distal

Fig. 3. Dorsal aspect of the tarsal and metatarsal bones of hedgehog (Erinaceus europaeus
L.). a) talus, b) calcaneus, c) os tarsi centrale, d) os tarsale I, e) os tarsale II, f) os tarsale III,
g) os tarsale IV, h) os metatarsale I, ý) os metatarsale II, i) os metatarsale III, j) os metatarsale
IV, k) os metatarsale V

218

Vet. arhiv 72 (4), 213-220, 2002

Z. E. Özkan: Macro-anatomical investigations on the skeletons of hedgehog (Erinaceus

europaeus L.). II. ossa membri pelvini

phalanges were arched and pointed to accommodate the curved nails. The
comparative lengths of nails were: II>III> IV>V>I.

Discussion
Any sexually dimorphic character can be used to distinguish males

from females, including differences in genitalia, body size, pelage,
ornamentation, behaviour. In practice, males are 20% larger than females
on average (

KUNZ et al., 1996

). In the present study, certain measurements

were taken in male hedgehogs.

In the Erinaceidae family, symphysis was formed by a cartilage or

interpubic ligament (

LESSERTISSEUR and SABAN, 1967

) and in the order

Insectivora the symphysis is sometimes non-existent, and always weak, as
in Erinaceus for example, where it is confined to the pubis (

SAUNDERS and

MANTON, 1969

). In the present study, an interpubic ligamentous form was

observed in hedgehogs.

The presence of a large obturator foramen bounded by the pubis and

ischium is characteristics of mammals (

WEICHERT, 1970

). In hedgehogs,

this foramen was also large and had an almost hemicycle form. Average
sagittal length and width of the foramen obturatum were 11.4 mm and 6.6
mm, respectively.

ROMER (1970)

reported that the fourth trochanter in the femur is absent

in mammals, and

SAUNDERS and MANTON (1969)

mentioned that the femur

of the Insectivora has a third trochanter which is particularly well developed
in Erinaceus and Centetes. In this study, three trochanter in the femur in
hedgehogs were observed: the greater (trochanter major), the lesser
(trochanter minor) and the third (trochanter tertius).

The fovea capitis on the caput ossis femoris and fossa supracondylaris

are absent in porcupines (

YILMAZ et al., 1999

). The fovea capitis on the

caput ossis femoris is absent in the African rhinocerus ( Diceros bicornis
L.), too (

GOMERČIĆ and HUBER, 1982

). Similar findings were observed in

this study.

The fibula is a slender bone and is usually seperated from the tibia but

is, however, fused at the distal end in Erinaceus (

SAUNDERS and MANTON,

1969; DEMÝRSOY, 1998

). In porcupines the fibula is fused with the tibia at the

219

Vet. arhiv 72 (4), 213-220, 2002

Z. E. Özkan: Macro-anatomical investigations on the skeletons of hedgehog (Erinaceus

europaeus L.). II. ossa membri pelvini

proximal portion

(YILMAZ et al., 1999)

. In our study the slender fibula was

fused only at the distal half of the tibia in hedgehogs.

In some species of the Erinaceidae family the pedis is comprises four

digits (

KURU, 1999

). In our study, the pedis was complete with five digits.

References

ANONYMOUS (1994): Nomina anatomica veterinaria. 4th ed. By the World Association

of Veterinary Anatomists.

BARTELS, T. H., W. MEYER (1991): Eine schnelle und effektive Methode zur Mazeration

von Wirbeltieren. Dtsch. Tierärztl. Wschr. 98, 407-409.

DEMÝRSOY, A. (1996): Genel ve Türkiye Zoocođrafyasý, Hayvan Cođrafyasý, Meteksan

A. Ţ., Ankara.

DEMÝRSOY, A. (1997): Türkiye Omurgalýlarý. Memeliler, Meteksan A. Ţ., Ankara.
DEMÝRSOY, A. (1998): Yaţamýn Temel Kurallarý. Meteksan A. Ţ., Ankara.
DÝNÇ, G., A. AYDIN, Ö. ATALAR (1999): Macro-anatomical investigations on the

skeletons of otter (Lutra lutra) II. ossa membri pelvini. Fýrat Univ. J. Health Sci. 13,
229-232.

DUBRUL, E. L. (1950): Posture, locomotion and the skull in Lagomorpha. Am. J. Anat. 87,

277-314.

DURSUN, N., S. TIPIRDAMAZ (1989): Etudes macro-anatomiquement sur les os du

squelete du vison (Mustela vison). J. Fac. Vet. Med. Univ. Selçuk. 5, 13-27.

GÝRGÝN, A., H. KARADAĐ, S. BÝLGÝÇ, A. TEMÝZER (1988): A study on the macro-

anatomical differences of the skeletons of wolf and fox as compared with the skeleton
of dog. J. Fac. Vet. Med. Univ. Selçuk. 4, 169-182.

GOMERČIĆ, H., Đ. HUBER (1982): Articulus coxae u Africkog nosoroga. XIX th. Congress

of Yugoslav Association of Anatomists. Abstracts. Prishtine, 13-15. 09. 1982. p. 34.

HIDAKA, S., M. MATSUMOTO, H. HIJI, S. OHSAKO, H. NISHINAKAGAWA (1998):

Morphology and morphometry of skulls of raccoon dogs, Nyctereutes procyonoides
and badgers, Meles meles. J. Vet. Med. Sci. 60, 161-167.

KUNZ, T. H., C. WEMMER, V. HAYSSEN (1996): Standard Methods for Mammals.

Measuring and Monitoring Biological Diversity. (D. E. Wilson, F. R. Cole, J. D.
Nichols, R. Rudran, M. S. Foster, Eds.). Smithsonian Institution Press, Washington,
London.

KURU, M. (1999): Omurgalý Hayvanlar. Palme Yayýncýlýk, Feryal Matbaacýlýk San. Ltd.

Ţti., Ankara.

LESSERTISSEUR, J., R. SABAN (1967): Generalites sur le Squelette. Traite’de Zoologie,

Anatomie, Systematique, Biologie. Publie’ Sous la Direction de Grasse’, P. P. Masson
et Cie, Paris.

220

Vet. arhiv 72 (4), 213-220, 2002

Z. E. Özkan: Macro-anatomical investigations on the skeletons of hedgehog (Erinaceus

europaeus L.). II. ossa membri pelvini

Received: 5 July 2002
Accepted: 29 August 2002

ROMER, A. S. (1970): The Vertebrate Body. W. B. Saunders Company, Philadelphia,

London, Toronto.

SAUNDERS, J. T., S. M. MANTON (1969): A manual of practical vertebrate morphology,

ed., Clarendon Press. Oxford.

TAŢBAŢ, M., S. TECÝRLÝO

GLU (1966): Maserasyon tekniđi üzerinde araţtýrmalar. J.

Fac. Vet. Med. Univ. Ankara 12, 324-330.

TECÝRLÝO

GLU, S. (1983): Makro-anatomische Untersuchungen über die Skelettknochen

von Hunden der Hyäne. I: Truncus. J. Fac. Vet. Med. Univ. Ankara. 30, 149-166.

VAUGHAN, T. A. (1972): Mammalogy. W. B. Saunders Company, Philadelphia, London,

Toronto.

YILMAZ, S., Z. E. ÖZKAN, D. ÖZDEMÝR (1998): Macro-anatomical investigations on

the skeletons of porcupine (Hystrix cristata) I. ossa membri thoracici. Tr. J. Vet. Anim.
Sci. 22, 389-392.

YILMAZ, S., G. DÝNÇ, A. AYDIN (1999): Macro-anatomical investigations on the

skeletons of porcupine (Hystrix cristata) II. ossa membri pelvini. Tr. J. Vet. Anim. Sci.
23, 297-300.

WEICHERT, C. K. (1970): Anatomy of the Chordates. Mc Graw-Hill Book Company.

New York

ÖZKAN, Z. E.: Makroanatomska istra

živanja kostura ježa (Erinaceus europaeus

L.). II. ossa membri pelvini. Vet. arhiv 72, 213-220, 2002.

SAŽETAK

Istražena je anatomska građa kostiju stražnje noge u tri odrasla ježa (Erinaceus europaeus

L.). Nedostajala je spina iliaca ventralis caudalis. Symphysis pelvis je bila oblikovana od symphysis
pubis, a ustanovljen je interpubični ligament. Prosječni Vialleton kut iznosio je 8

. Prosječna

udaljenost između središta acetabuluma i bočne kvrge iznosila je 39,2 mm, a prosječna udaljenost
između središta acetabuluma i sjedne kvrge bila je 13,8 mm. Na bedrenoj kosti su ustanovljena
tri trochantera. Goljenica i lisnjača su bile spojene u distalnoj polovici. Utvrđeno je 7 tarzalnih
kostiju, a stopalo je bilo potpuno s pet prstiju.

Ključne riječi: jež, Erinaceus europaeus L., kosti stražnje noge

PATTERNS & PHENOTYPES

Sonic Hedgehog Is Required for the Assembly
and Remodeling of Branchial Arch Blood
Vessels

Hana Kolesova

´ ,

Henk Roelink,

* and Milosˇ Grim

Sonic hedgehog (Shh) is a morphogen involved in many developmental processes. Injection of cells (5E1)
that produce a Shh-blocking antibody causes an attenuation of the Shh response, and this causes vascular
malformations and impaired remodeling characterized by hemorrhages and protrusions of the anterior
cardinal vein and outﬂow tract, delayed fusion of the dorsal aortae, impaired branching of the internal
carotid artery, and delayed remodeling of the aortic arches. Distribution of smooth muscle cells in the vessel
wall is unchanged. In 5E1-injected embryos, we also observed impaired assembly of endothelial cells into
vascular tubes, particularly in the sixth branchial arch, around the anterior cardinal vein and around the
dorsal aorta. In 5E1-treated embryos, increased numbers of macrophage-like cells, apoptotic cells, and a
decreased level of proliferation were observed in head mesenchyme. Together, these observations show that
Shh signaling is required at multiple stages for proper vessel formation and remodeling. Developmental
Dynamics 237:1923–1934, 2008.

2008 Wiley-Liss, Inc.

Key words: Shh; blood vessels; branchial arches; 5E1 hybridoma cells; quail embryo; endothelium

Accepted 8 May 2008

INTRODUCTION

The inductive events controlling the
formation and remodeling of the ﬁrst
intra-embryonic vessels are not well
understood. Classical embryological
experiments have demonstrated that
signals derived from endoderm can in-
duce vessel formation in adjacent me-
soderm (Pardanaud et al., 1989), and
it appears that the Hedgehog (Hh) sig-
naling mediates at least some of this
endoderm-derived activity since Smo
null embryos, which cannot respond to
Hedgehog (Hh), exhibit severe vascu-
lar defects (Byrd et al., 2002). Em-
bryos treated at later stages with the

Smo inhibitor cyclopamine show de-
fects in vascular remodeling (Nagase
et al., 2006), indicating an ongoing re-
quirement for Hh signaling.

Among the ﬁrst intraembryonic ves-

sels induced by endodermally derived
signals are the vessels of the branchial
(pharyngeal) region. Mouse embryos
lacking Shh have hypoplastic ﬁrst
branchial arches that prematurely
fuse in the midline (Yamagishi et al.,
2006). The second and third branchial
arches are hypoplastic, while the
fourth and sixth arches do not appear
to develop at all (Washington Smoak
et al., 2005). Smo null embryos die too

early to assess the role for Hedgehog
signaling for pharyngeal vessel devel-
opment (Zhang et al., 2001; Wijgerde
et al., 2002).

Vessels of the branchial region un-

dergo extensive remodeling in stages
15 to 23. At stage 15, caudal parts of
the paired aortae fuse to form a single
descending aorta, while rostrally they
become the distal parts of the left and
right internal carotid arteries. The
ventral aorta after the branching into
the aortic arches continues as the left
and right external carotid arteries.

The ﬁrst to sixth aortic arches de-

velop in a cranio-caudal gradient (Hi-

Institute of Anatomy, First Faculty of Medicine, Charles University, Prague, Prague, Czech Republic

Department of Molecular and Cell Biology, University of California at Berkeley, Berkeley, California

Grant sponsor: Charles University; Grant number: GAUK 54/203209; Grant sponsor: NIH; Grant number: 1R01HD042307; Grant sponsor:
The Ministry of Education of The Czech Republic; Grant number: MSM 0021620806.
*Correspondence to: Henk Roelink, Department of Molecular and Cell Biology, University of California at Berkeley, Berkeley,
CA 94720-3204. E-mail: roelink@berkeley.edu

DOI 10.1002/dvdy.21608
Published online 21 June 2008 in Wiley InterScience (www.interscience.wiley.com).

DEVELOPMENTAL DYNAMICS 237:1923–1934, 2008

2008 Wiley-Liss, Inc.

ruma and Hirakow, 1995) from stages
12 to 23. They form in branchial arch
mesenchyme from cords of angioblasts
around the foregut, which subse-
quently become luminized and serve
as a communication between the ven-
tral and dorsal aorta. In ﬁsh and am-
phibian larvae, most branchial arches
develop into the gill arches (Kolesova
et al., 2007); in amniotes, aortic arches
undergo signiﬁcant remodeling.

The ﬁrst, second, and ﬁfth aortic

arches undergo regression that starts
at stage 21. The third, fourth, and
sixth aortic arches are gradually rear-
ranged. The third aortic arch is incor-
porated into the common and internal
carotid arteries. In birds, the right
fourth arch contributes to the arch of
aorta. The remainder of the left one
incorporates in the subclavian artery.
The sixth aortic arch becomes part of
the pulmonary artery. The main veins
of the cranial region are the paired
anterior cardinal veins, which drain
blood from the head and neck to the
common cardinal vein. The anterior
cardinal veins develop at stage 12.

The endothelial cells lining the ves-

sels in the branchial region originate
from paraxial mesoderm (Noden and
Trainor, 2005; Evans and Noden,
2006). Initially, presumptive vessels
consist of an endothelial lining, which

is subsequently covered with a layer of
smooth muscle cells. The dorsal aorta
is covered by sclerotome-derived cells
(Wiegreffe et al., 2007), the aortic
arches by cells derived from the neu-
ral crest (Le Lievre and Le Douarin,
1975). Shh can induce angiogenic fac-
tors such as VEGFs and Angiopoietins
in mesenchyme (Pola et al., 2001) and
thus affect the smooth muscle distri-
bution and vessel stabilization (van
Tuyl et al., 2007). The anterior cardi-
nal veins have no smooth muscle
layer.

The aortic arches are located within

the corresponding branchial arches,
which are formed from neural crest
cells and cells of paraxial mesoderm
and are lined with an ectoderm on the
outside, and endoderm on the surfaces
surrounding the developing pharynx.
Endothelial cells and striated muscle
cells are derived from mesoderm,
while other tissues in the branchial
arches are of neural crest origin
(Evans and Noden, 2006). According
to some observations, mesenchymal
cells of branchial arches are stimu-
lated to proliferate and prevented
from apoptosis by Shh (Ahlgren and
Bronner-Fraser, 1999; Jeong et al.,
2004).

We tested the requirement for Shh

for the correct development of the ves-

sels associated with the branchial
arches, and found that decreased lev-
els of Shh signaling result in angio-
genic malformations. Ongoing devel-
opment of existing vessels is disrupted
by attenuated Shh signaling. These
vessels lose their ability to remodel,
fuse, and form branches. The vessel
walls, in particular of the anterior car-
dinal veins, appear to be malformed;
hemorrhages are common in this area,
and these vessels cannot contain ink.
The area around the anterior cardinal
veins has increased levels of apoptotic
cells and macrophage-like cells. More-
over, new vessel formation is im-
paired, and endoderm cells can often
been seen lining incomplete vessels, or
as aggregates. Altogether, our results
demonstrate a varied and continual
requirement for Shh signaling in the
development of the vessels in and
around the branchial arches.

RESULTS

Vessel Formation in
Branchial Arches

The development of vessels in the
branchial region was extensively de-
scribed in chicken embryos (Hiruma
and Hirakow, 1995), and is very sim-
ilar in the quail embryo, although the

Fig. 1. A–D: Normal development of head and neck vessels in quail embryos. Vessels stained with QH1 Ab. E–F: Expression of Shh, visualized with
5E1 Ab. A: A stage-15 embryo with a formed ﬁrst aortic arch (I), which connects the ventral (VA) with the dorsal aorta (DA). The anterior cardinal vein
is also indicated (CV). B: Stage 18, with the second (II), third (III), and the fourth (not visible on this section) aortic arches indicated. The anterior part
of the dorsal aorta (DA) continues as the internal carotid artery (IC). C: At stage 21, the fourth (IV) aortic arch is indicated. D: Stage 23, with the ﬁfth
(V) and sixth (VI) aortic arches. E,F: Shh expression visualized with anti-Shh antibody in normal embryos. Stage 15 (E), stage 18 (F). Shh expression
is visible in the notochord, ﬂoor plate (arrows), and foregut (E, arrowheads) and parts of the branchial arch endoderm (F, arrowheads). Transversal
sections. D–F are counterstained with hematoxylin. Scale bar

⫽ 400 ␮m in all panels.

Fig. 2. Inhibition of the Shh response by 5E1 (anti-Shh) Ab produced by hybridoma cells. A,B: Sections of stage-18 embryos injected at stage 11 with
hybridoma cells. Sections incubated with secondary antibody. A: An embryo injected with control 12CA5 hybridoma cells. No staining is visible in the
embryo. Secondary antibody reacts with hybridoma cells (arrowheads). B: An embryo injected with 5E1 hybridoma cells producing anti-Shh antibody.
Shh-expressing structures, ﬂoor plate, notochord, and endoderm of the branchial arches are indicated (arrows). Hybridoma cells are also stained
(arrowheads). Counterstained with hematoxylin. Scale bar

⫽ 400 ␮m in both panels. C,D: Expression of Ptch1, which is induced in response to Shh

signaling. C: An embryo injected with 12CA5 hybridoma cells shows normal distribution of Ptch1 in the ventral part of the neuroepithelium, in the
endoderm of branchial arches, and in adjacent mesenchyme (arrows). D: An embryo injected with 5E1 hybridoma cells shows a decreased level of
Ptch1 expression. Residual expression is observed only in the ventral part of the nervous system and in a few areas of branchial arch endoderm
(arrows). No expression is detected in the mesenchyme. Scale bar

⫽ 200 ␮m in both panels.

Fig. 3. Ink injection into blood vessels at stage 18. A: In a 12CA5-injected control embryo, the ﬁrst and the second aortic arches have regressed and
formed capillary plexuses (white arrows I, II). The main artery conducting blood from the heart to the dorsal aorta is the third aortic arch in this stage
(white arrow III). The fourth aortic arch starts to develop (white arrow IV). Normal capillary plexuses drained into anterior cardinal vein are visible in the
head. B: Embryo injected with 5E1 hybridoma cells has a smaller head compared to the control. Development of the aortic arches is delayed. The ﬁrst
and the second aortic arches are still present and form functional communication between the heart and the dorsal aorta (white arrows I, II). The third
and the fourth aortic arches are normally developed (white arrows III, IV). A signiﬁcant amount of ink has leaked from the tributaries to the anterior
cardinal vein (white arrowheads). Scale bar in A and B

⫽ 500 ␮m. C, D: Parafﬁn sections of the anterior cardinal vein region, stained for acid

phosphatase. N, neuroepithelium. C: Red blood cells (arrowheads) are visible only in vessels; macrophage-like cells are indicated (arrow). D: Red blood
cells localized extravascularly (arrowheads), macrophage-like cells are more numerous than in control embryos (arrow). Scale bar in C and D

⫽ 200

␮m.

1924 KOLESOVA

´ ET AL.

timing of their development is some-
what different. Generally, in quail em-
bryos the aortic arches remodel faster;
they form approximately one stage
later, and they regress a few stages
earlier than in chick embryos. At

stage 15, the ﬁrst aortic arch is
present as a vessel connecting the
ventral and dorsal aorta, and the an-
terior cardinal vein is already devel-
oped (Fig. 1A). Immediately after the
ﬁrst aortic arch has formed, the sec-

ond arch starts to develop. The third
aortic arch develops around stage 18
(Fig. 1B) and the fourth one develops
at stage 20. At stage 21, the ﬁfth and
sixth aortic arches start to develop
and are fully formed at stage 23 (Fig.
1C,D). The ﬁrst and second aortic
arches start to regress at stage 18.
The ﬁfth aortic arch, a minor bypass of
the sixth aortic arch, starts to regress
at stage 24. While the ﬁrst, second,
and ﬁfth aortic arches are transient
structures, the third, fourth, and sixth
arches persist.

At stage 15, only the aortic arches

Fig. 1.

Fig. 2.

Fig. 3.

SHH AND BRANCHIAL VESSEL DEVELOPMENT 1925

supply the branchial arches (Fig. 1A),
while at stage 18, the dorsal aorta, the
aortic arches, as well as the anterior
cardinal

vein

give

off

smaller

branches and capillaries (Fig. 1B). At
stages 21 and 23, the development of
small vessels and capillaries contin-
ues in the whole region, resulting in a
dense capillary network in branchial
arch mesenchyme and in the regions
surrounding the brain and eyes (Fig.
1C,D).

Besides luminized vessels and cap-

illaries, we also observed an increas-
ing number of isolated angiogenic
cells at stages 18 –23 (Fig. 1B–D).
These cells are mainly present around
the anterior cardinal vein and in
branchial arches, while in the area in
the vicinity of the dorsal aorta, only a
few of these cells are detected. These
isolated angiogenic cells are evenly
distributed in the mesenchyme.

Inhibition of Shh Signaling
With Anti-Shh Antibodies
Produced by 5E1 Hybridoma
Cells

In general, we detected Shh in a pat-
tern and timing consistent with that
observed in the chick (Roelink et al.,
1995). The notochord is a prominent
site of early expression, and starting
at stage 15, Shh is expressed in
endoderm lining the branchial arches
and the foregut (Fig. 1E,F). To condi-
tionally attenuate Shh signaling, we
injected 5E1 (anti-Shh) hybridoma
cells under the vitelline membrane of
stage-10 –12 embryos, which were an-
alyzed at stages 18 –23. 5E1 hybrid-
oma cell-derived anti-Shh antibodies
distribute widely in injected embryos.
Visualizing the 5E1 antibodies by sim-
ply using an anti-IgG secondary anti-
body on sectioned embryos, staining is
detected at the sites of Shh expres-
sion, such as the notochord, ﬂoor
plate, and endoderm of the branchial
arches and foregut. The secondary an-
tibodies also react with 5E1 hybrid-
oma cells, which always remain at the
embryonic surface (Fig. 2A,B), just as
the control 12CA5 hybridoma cells
(Fig. 2C,D).

To verify if the Shh response is efﬁ-

ciently blocked after 5E1 injection, we
analyzed the expression of the gene
coding for its receptor Ptch1 by mRNA
in situ hybridization. Ptch1 expres-

sion is invariably upregulated in re-
sponse to Shh signaling (Marigo and
Tabin, 1996). In control embryos,
Ptch1 is expressed in areas adjacent to
Shh sources, such as branchial arch
mesenchyme, around the foregut, in
the ventral part of the neural tube,
and around domains of Shh produc-
tion in the brain (Fig. 2C). In embryos
injected with 5E1 cells, Ptch1 expres-
sion in the neural tube and endoderm
is decreased and no Ptch1 expression
is found in the abutting mesenchyme
(Fig. 2D), demonstrating a signiﬁcant
attenuation of the Shh response. Re-
sidual expression of Ptch1 could be the
result of incomplete inhibition, but
also be caused by other Hh ligands,
which are not recognized by 5E1
(Goodrich et al., 1997; Carpenter et
al., 1998).

Embryos injected with 5E1 and

with control 12CA5 hybridoma cells
develop slightly slower than unin-
jected embryos, and staging was per-
formed based on anatomical land-
marks. The 5E1 antibody-injected
embryos largely exhibit a normal
gross morphology, although at least
half of them have a smaller head com-
pared to control (Fig. 3B). Similar ce-
phalic phenotypes have been reported
(Ahlgren and Bronner-Fraser, 1999).
Embryos injected with control 12CA5
hybridoma cells have a macroscopic
and microscopic anatomy identical to
untreated embryos.

Vessel Malformations in
Anti-Shh Antibody-Treated
Embryos

Even at the anatomical level, the ef-
fect of inhibiting Shh on vessel devel-
opment is remarkable. Generally, in
embryos injected with 5E1 hybridoma
cells development of the aortic arches
is delayed. While in stage-18 control
embryos, the ﬁrst and the second aor-
tic arches start to regress into capil-
lary plexi, these aortic arches are still
present in 5E1-injected embryos, indi-
cating a delay in remodeling (Fig.
3A,B).

A consequence of Shh inhibition is

the failure of the anterior cardinal
vein and its branches to form func-
tional vessel walls. Hemorrhages were
observed frequently (Fig. 3A–D), and
these vessels were permeable to ink,
unlike the control hybridoma-injected

animals, which were able to contain
the ink within the vessels (Fig. 3A,B).
In addition to the ink-permeability,
the lumen of either anterior cardinal
vein in 5E1-injected embryos is sinu-
soidal with endothelium-lined protru-
sions (Fig. 4A–D). This demonstrates
that Shh plays an important role in
the establishment of a functional wall
in the anterior cardinal veins and its
tributaries.

Several arteries show abnormal de-

velopment as a consequence of 5E1
injection as well. The internal carotid
arteries are characterized by the pres-
ence of a transverse septum over a
length of up to 40

␮m (in 2 of 6 em-

bryos). This septum consists of two
layers of endothelium, with mesen-
chyme in between, dividing the inter-
nal carotid artery into two separate
vessels, which merge again further
rostrally (Fig. 4E–G). Similarly, an
abnormal septum is present in the
dorsal aorta (in 4 out of 6 embryos).
This aortic septum is usually about 60
␮m long, and is covered with endothe-
lium on both sides (Fig. 4H–J). Al-
though we assume that this septation
is a result of delayed fusion, it re-
mains possible that it has formed af-
ter the initial fusion of the left and
right dorsal aortae. The same domain
of the dorsal aorta in control embryos
is already fused and has a single lu-
men. Vessel abnormalities are also
found in outﬂow tracts of the heart.
Besides vessels with irregular lumina
and unusual invaginations (Fig. 4N–
P), we have also observed curving
strands of endothelial cells in succes-
sive sections, possibly malformed ves-
sels with an incomplete vessel wall
(Fig. 4K–M) (in 4 of 6 embryos). Em-
bryos injected with the control hybrid-
oma cells had no obvious vessel mal-
formations.

To further assess malformations in

vessel endothelium, we determined
the expression VEGFR2, which is ex-
pressed in endothelial cells (Jaffredo
et al., 1998) albeit not ubiquitously. At
stage 18, VEGFR2 is expressed in all
endothelial cells of small developing
vessels as capillaries, such as brain
capillaries, while only about half the
endothelial cells of bigger arteries ex-
press VEGFR2. Similarly, the dorsal
aorta is lined by VEGFR-positive
cells, but only in its ventrolateral as-
pect, where the aortic arches are con-

1926 KOLESOVA

´ ET AL.

nected.

Besides

endothelial

cells,

VEGFR2 is highly expressed in the
outﬂow tract myocardium, while out-
ﬂow tract endothelium contains only
few VEGFR2-positive cells. These
myocardial cells are probably derived
from endothelium (Wilting et al.,
1997). VEGFR2 was also expressed in
the notochord, as it was previously re-
ported (Nimmagadda et al., 2004). We
did not detect any difference in
VEGFR2 expression in 5E1-injected
embryos compared to control, indicat-
ing that VEGF is not a critical medi-
ator of the effects of Shh (not shown).

Following the formation of an en-

dothelial layer, the smooth muscle
cells start to surround the forming
vessels. At stage 18, smooth muscle
cells completely cover the endothe-
lial lining of the dorsal aorta and the
internal carotid arteries (Fig. 4A).
The aortic arches have smooth mus-
cle cells only on their lateral side,
while an incomplete layer of smooth
muscle cells is associated with the
outﬂow tract. Smooth muscle actin is
also present in the myotome (Fig.

Fig. 4.

Fig. 4. QH1 staining of vessel endothelium
showing lumen malformations in anti-Shh hy-
bridoma cell–injected embryos. A–D: Lumen
malformation in the anterior cardinal vein. A:
Control embryo. B: The anterior cardinal vein
exhibits irregularities and contains protrusions
of the endothelial layer extending into the ves-
sel lumen (arrow) at stage 18. C, D: Protrusions
in the anterior cardinal vein (arrow) are also
found in embryos injected with anti-Shh anti-
body and harvested at stage 23. E–G: Internal
carotid artery malformations at stage 18. E:
Control embryo. F: The internal carotid artery is
divided by a horizontal septum (arrow) into two
separate vessels. G: Detail of septum covered
in endothelium from both sides (arrow). H–J:
Dorsal aorta malformations, stage 18. H: Con-
trol embryo. I: Failure of complete fusion be-
tween the left and right dorsal aortae. The per-
sistent septum is indicated (arrow). J: Detail of
septum (arrow). K–M: Malformations of outﬂow
tract endothelium at stage 18. K: Control em-
bryo. L: The endothelium of the outﬂow tract
fails to form a luminized vessel (arrow), and
instead forms an isolated endothelial wall with
red blood cells on one side. M: Detail of patent
vessel endothelium (arrow) and adjacent mal-
formed vessel (arrow). N–P: Malformations of
the outﬂow tract at stage 18. N: Control em-
bryo. O: The outﬂow tract has developed a
protrusion of tissue surrounded by the endothe-
lial layer, which invaginates into the lumen (ar-
row). P: Detailed view of the endothelial protru-
sion (arrow). Transversal sections. B–D, K–M
are counterstained with hematoxylin. Scale
bar

⫽ 400 ␮m in all panels.

SHH AND BRANCHIAL VESSEL DEVELOPMENT 1927

5C). At stage 23, a continuous
smooth muscle layer surrounds aor-
tic arches and the outﬂow tracts
(Fig. 5E). Also, all branches of the
internal carotid arteries have a con-
tinuous layer of smooth muscle cells,
while the anterior cardinal veins and
their tributaries are devoid of
smooth muscle cells. The formation
of the smooth muscle cell lining of
the vessels is unaffected by injection
of 5E1 hybridoma cells (Fig. 5B,F),
despite the presence of obvious ves-
sel malformations. Smooth muscle
cells also line the abnormal aortic
septum (Fig. 5D). This is consistent
with our observation that the forma-
tion of the smooth muscle lining of
the dorsal aortae precedes the sub-
sequent fusion of these vessels. Alto-
gether, this indicates that the loss of
Shh signaling has little effect on the
process in which smooth muscle cells
form around new vasculature.

Angiogenic and Macrophage-
Like Cells in Anti-Shh
Antibody-Treated Embryos

The main effect of conditional Shh
inhibition on blood vessel develop-
ment in embryos injected at later
stages (injected in stage 13–15 and
harvested at stage 21–23) is the
presence of an increased number of
free, round endothelial cells, positive
for QH1. These cells are not inte-
grated into functional vessel lumina,
but

aggregate

into

multicellular

clusters. Such aggregates usually
are found around the anterior cardi-
nal veins, around the dorsal aorta,
and in the branchial arches around
the aortic arches. Such cell aggre-
gates are not present in control
12CA5 hybridoma-injected embryos,
where angiogenic cells are fewer and
isolated. The increased number of
aggregates is most signiﬁcant in the
sixth branchial arch (Fig. 6G–L),
around the anterior cardinal vein,
and just ventral to the dorsal aorta
(Fig.

7A–L),

while

the

branchial arches are the least af-
fected.

the

ﬁrst

and

second

branchial arch, we have not observed
any signiﬁcant difference compared
to control in the number of endothe-
lial cell aggregates (data not shown).
This might either indicate that at
the moment of injection the Shh re-

Fig. 5.

Distribution of vascular smooth muscle cells visualized with smooth muscle actin Ab. A,B:

Smooth muscle cells in a stage-18 embryo are present around the dorsal aorta, internal carotid
artery, and the lateral part of the aortic arches (arrows). 5E1-injected embryos show no difference
in the distribution of smooth muscle cells (B) compared to control injected embryos (A). Scale bar

⫽

400

␮m. C,D: Stage-18 embryos. Smooth muscle cells also cover the endothelial malformation

caused by 5E1 injection. C: Normal dorsal aorta. D: Persistent septum in the dorsal aorta (arrow).
Arrowheads show expression of the smooth muscle actin in the myotome. Scale bar

⫽ 400 ␮m.

E,F: Smooth muscle cells in a stage-23 embryo are present in the dorsal aorta, internal carotid
artery, aortic arches, and the outﬂow tracts (arrows). 5E1-injected embryos show no difference in
the distribution of the smooth muscle cells (F) compared to control-injected embryos (E). Scale
bar

⫽ 200 ␮m.

1928 KOLESOVA

´ ET AL.

Fig. 6.

QH1-positive cell abnormalities in branchial arches at stage 21 (A–D, F) and stage 23 (E, G–U); Characterization of QH1-positive cells:

macrophage-like cells stained for acid phosphatase (M–O); apoptotic cells positive for cleaved caspase-3 (P–R); proliferating cells positive for phospho
histone H3 (S–U). A–F: QH1-positive cells in the mesenchyme and ectoderm of third and fourth branchial arches at stage 21 (stage 23-E). A,B: Control
embryos. C,D: Larger aggregates of QH1-positive cells (arrows) in anti-Shh-injected embryos at stage 21 or (E) stage 23. F: Graph shows that
anti-Shh-treated embryos have a similar number of QH1-positive cells as the controls. However, QH1-positive cells are present in large aggregates
compared to single cells in control embryos. Y-axis: average number of QH1-positive cell aggregates in the third and fourth branchial arches. G–L:
Increased number of QH1-positive cells in the sixth branchial arch of stage-23 embryos injected with anti-Shh hybridoma cells. G,H: Control embryos.
I: QH1-positive cells (arrows) in the distal region of the sixth branchial arch and (J) detail. K: Increased number of QH1-positive cells (arrows) in the
proximal region of the branchial arch. L: Graph shows the increased number of QH1-positive cells in anti-Shh-injected embryos in the sixth branchial
arch. Y-axis: average number of QH1-positive cell aggregates in the sixth branchial arch. M–O: Very few macrophage-like cells are present in the
branchial arches. Anti-Shh injection has no signiﬁcant effect on the number of macrophage-like cells (O). P–R: No signiﬁcant difference in the number
of apoptotic cells (arrow) in response to anti-Shh injection (R). S–U: No signiﬁcant difference in the number of proliferating cells in the branchial arches
in anti-Shh injected embryos (arrows). Transversal sections. C–E, K, P, Q counterstained with hematoxylin. Scale bar

⫽ 200 ␮m in all panels. Graphs:

Standard deviation is indicated. *Signiﬁcant difference (P

⬍ 0.05); ns, no signiﬁcant difference.

SHH AND BRANCHIAL VESSEL DEVELOPMENT 1929

Fig. 7.

Abnormalities in the number of QH1-positive cells around the main vessel trunks at stage 23: the anterior cardinal vein and the dorsal aorta

(A–L); Characterization of QH1-positive cells: macrophage-like cells stained for acid phosphatase (M–O); apoptotic cells positive for cleaved caspase-3
(P–R); proliferating cells positive for phospho histone H3 (S–U). A–F: Anterior cardinal vein malformations. A, B: Control embryos. C: Increased number
of QH1-positive cells around the anterior cardinal vein (arrows) in anti-Shh hybridoma cell–injected embryos. D: Increased number of QH1-positive
cells around a branch of the anterior cardinal vein (arrows). E: Free QH1-positive cells are present around the anterior cardinal vein already at stage
18 (arrows). F: Graph showing a higher number of QH1-positive cells in the anterior cardinal vein region in embryos injected with anti-Shh antibody.
Y-axis: average number of QH1-positive cells per section. Standard deviation is indicated. G–L: Increased number of QH1-positive cells around the
dorsal aorta at stage 23. G,H: Control embryos. I: Aggregate of QH1-positive cells on the ventral side of the dorsal aorta (arrow) and (J) detail of the
aggregate (arrow). K: QH1-positive cells are present (arrow), but hematopoietic regions in dorsal aorta appear normal (arrowhead). L: Graph shows
the increased number of QH1-positive cells in anti-Shh-injected embryos in the dorsal aorta region. Y-axis: average number of QH1-positive cells per
ﬁeld. Standard deviation is indicated. M–O: Increased number of macrophage-like cells stained with acid phosphatase (arrows) in the region of the
anterior cardinal vein in anti-Shh-injected embryos. O: Graph indicating the increased number of macrophage-like cells. P–R: Increased number of
cleaved Caspase-3 (apoptotic) cells (arrows) around the anterior cardinal vein in anti-Shh-injected embryos. R: Graph showing the increased number
of apoptotic cells. S–U: Decreased number of dividing cells (arrows) in the anterior cardinal vein region in anti-Shh-injected embryos. U: Graph showing
the difference in proliferating cells. Transverse sections. C, E, P, Q, counterstained with hematoxylin; M, N, counterstained with Fast green. Scale bar

⫽

200

␮m in all panels. Graphs: Y-axis: average number of positive cells per ﬁeld. Standard deviation is indicated; *signiﬁcant difference (P ⬍ 0.05).

1930 KOLESOVA

´ ET AL.

quirement for initial vessel forma-
tion has passed, or that the absence
of de-novo vascularization associ-
ated with the regression of the ﬁrst
and second aortic arches results in
an absence of these aggregates of
QH1-positive cells.

In the third and fourth branchial

arches, Shh inhibition results in an
increased number of larger endothe-
lial aggregates compared to 12CA5-
injected control embryos. These aggre-
gates are mainly localized in the
mesenchyme, sometimes in close prox-
imity to branchial arch ectoderm and
endoderm. Such aggregates are not
present in control embryos, where we
observed only solitary QH1-positive
cells, in comparable numbers to the
solitary cells in anti-Shh-treated em-
bryos (Fig. 6A–F).

The sixth branchial arch is the most

affected as measured by the number of
QH1-positive cells not part of an obvi-
ous vessel wall. These cells are
present along the whole extent of the
arches and are concentrated in mes-
enchyme as aggregates and solitary
cells. In control embryos, only solitary
QH1-positive cells are observed. The
total number of nonintegrated QH1-
positive cells is signiﬁcantly higher in
5E1 hybridoma-injected embryos than
in controls (Fig. 6G–L).

The biggest increase of noninte-

grated endothelial cells as a conse-
quence of 5E1 injection is detected
ventral to the dorsal aorta (Fig. 7G–
L). It is possible that this abundance
of angiogenic cells is related to the
area of hemangiogenesis within the
wall of the ventral aorta (Jaffredo et
al., 1998). Increased numbers of endo-
thelial cell aggregates were also found
in the vicinity of the anterior cardinal
veins. The QH1-positive cells in this
area (Fig. 7A–F) are the only cells that
are predominantly identiﬁed as mac-
rophage-like cells, rather than angio-
genic cells (Fig. 6M–O). Macrophage-
like cells are the phagocytic cells of
the early embryo, are derived from he-
mangioblasts (Cuadros et al., 1992),
and are characterized by their expres-
sion of acid phosphatase (Fig. 7M–O).
Direct co-localization of acid phospha-
tase and QH1 staining is not possible
due to incompatible ﬁxation and pro-
cessing requirements.

Apoptosis and Proliferation
in Anti-Shh Ab-Treated
Embryos

In many instances during develop-
ment, loss of Shh leads to decreased
proliferation and increased apoptosis,
possibly explaining some of the vessel
malformations we observed as a con-
sequence of 5E1 injection.

A signiﬁcant increase in the number

of apoptotic, caspase-3-positive cells
(Fig. 7P–R) is observed around the an-
terior cardinal veins in 5E1-injected
embryos, suggesting a role for Shh in
cell survival in this region and, conse-
quently, an increased number of mac-
rophage-like cells is detected. In con-
trast, in the branchial arches and
around the dorsal aorta, the frequency
of cleaved caspase-3-positive apoptotic
cells do not differ from control em-
bryos (Fig. 6P–R). Since the domains
of apoptosis coincide with the areas
where vessel integrity is compro-
mised, it remains a possibility that the
hemorrhages increase apoptosis in the
surrounding tissues (Figs. 6,7S–U).

Coincident with the increased apo-

ptosis, we observed decreased prolifer-
ation near the anterior cardinal vein
(Figs. 5, 6S–U), but not in the other
regions studied.

DISCUSSION

The widespread expression of Shh in
endoderm, ﬂoor plate, and notochord,
as well as the proximity of the dorsal
aortae, anterior cardinal veins, and
aortic arches to these Shh-producing
structures, prompted us to assess the
role of Shh signaling in pharyngeal
vessel development.

Previous examinations of Shh null

embryos (Jeong et al., 2004; Yama-
gishi et al., 2006) have demonstrated
a

Shh

requirement

for

normal

branchial arch formation. However,
global defects in these animals pre-
vent assessment of the involvement of
Shh in later stages of branchial arch
development (Washington Smoak et
al., 2005). Using a method of inhibit-
ing Shh response at later developmen-
tal stages, we demonstrate a continual
requirement for Shh for the correct
formation of the pharyngeal vascula-
ture. Taking advantage of the ante-
rior-posterior developmental gradient
of the branchial arches, we could si-

multaneously study the effects of Shh
inhibition in the further developed an-
terior branchial arches (I–III) as well
as the newly formed posterior arches
(IV–VI). In the posterior arches, we
observed a failure of vessel luminiza-
tion, represented by formation of nu-
merous

aggregates

endothelial

cells. This shows Shh requirement in
the early stages of vessel develop-
ment. A similar effect that blocked
Shh signaling in the early stages of
vasculogenesis was observed before
mice and chick embryos (Vokes and
Krieg, 2002; Vokes et al., 2004).

Already formed vessels are affected

by Shh inhibition in their remodeling.
This effect is characterized by delayed
fusion, impaired branching, and un-
usual invaginations of the vessel
walls. Such malformations are ob-
served in the dorsal and ventral aorta,
the internal carotid arteries, and an-
terior cardinal veins, which are ac-
tively remodeled at that time. These
disturbances of vessel remodeling are
not known to be affected by Shh sig-
naling, although it has been observed
that cyclopamine treatment affected
fusion of the dorsal aortae (Nagase et
al., 2006). Alterations of vessel remod-
eling are found in arteries that have
already formed smooth muscle layer.
However, the wall of malformed ante-
rior cardinal veins does not contain
smooth muscle cells. This suggests
that the presence or absence of a
smooth muscle layer does not play a
critical role in the generation of vessel
malformations. These results show
that Shh is essential to maintain the
stability and coherence of the endo-
thelial layer of the veins, as we de-
tected in the anterior cardinal vein. A
similar phenotype is observed in ze-
braﬁsh with disrupted Shh signaling
(S.J. Childs personal communication).

In our study, we expected to ﬁnd

increased level of apoptosis and de-
creased proliferation as a consequence
of Shh signal attenuation, as it was
reported by previous observation at
earlier stages of avian development
(Ahlgren and Bronner-Fraser, 1999)
and in mice (Jeong et al., 2004; Wash-
ington Smoak et al., 2005). However,
our results do not generally show this
effect. We observed increased level of
apoptosis and decreased proliferation
only around the anterior cardinal
veins at stage 23. At stage 18, there

SHH AND BRANCHIAL VESSEL DEVELOPMENT 1931

was no signiﬁcant difference in num-
bers of proliferating and apoptotic cell
between control and Shh-inhibited
embryos (data not shown). It seems,
thus, that Shh signaling inﬂuences
cell proliferation and survival more
profoundly at earlier developmental
stages.

Interestingly, sources of Shh are not

necessarily in the direct vicinity of the
vessels affected by the application of
5E1 anti-Shh hybridoma cells. There
are several sites of Shh expression in
the vicinity of the anterior cardinal
veins, including the ﬂoorplate of the
hindbrain and midbrain, the domains
of Shh expression in the forebrain, as
well as the notochord and the pre-
chordal plate. Shh derived from the
notochord and ﬂoor plate inﬂuences
dorsal aortic development (Nagase et
al., 2006), and it remains unsolved
how Shh reaches these distant sites.
Intermediary factors such Angiopoi-
etin-1, VEGFs (Pola et al., 2001),
Fox1, and BMP4 (Jeong et al., 2004;
Astorga and Carlsson, 2007) could me-
diate the effects of Shh. However, di-
rect signaling of Shh to the endothe-
lial is likely to occur, consistent with
the relatively large area over which
Ptch1 expression is decreased after
Shh inhibition. Also, the Shh indepen-
dence of VEGFR2 expression indi-
cates that VEGF does not play a crit-
ical intermediary role between Shh
and its effect on the endothelial cells,
altogether supporting a model in
which Shh signals the vessel endothe-
lium directly. Furthermore, in experi-
ments with mutant mice, in which
neural crest– derived mesenchyme in
the branchial arches was rendered in-
sensitive to Hh signaling, initial vas-
cularization appeared normal (Sasai
et al., 2001; Jeong et al., 2004), show-
ing that a possible intermediary sig-
nal is not generated in crest-derived
mesenchyme. However, the possible
role of non-crest mesenchymal cells
remains to be determined.

In conclusion, our results demon-

strate a continual requirement for
Shh signaling for vascular develop-
ment and remodeling. We have ob-
served minor differences at best in
apoptosis and proliferation, and no
difference in smooth muscle actin and
VEGFR2 expression, supporting the
idea that Shh acts directly on vessel
endothelium and not via intermedi-

aries induced in the mesenchyme be-
tween the Shh sources and the ves-
sels.

Nevertheless,

the

molecular

events following the initial Shh re-
sponse, which instructs endothelial
cells to form blood vessels, remains to
be solved.

EXPERIMENTAL
PROCEDURES

Embryos

Fertilized quail (Coturnix coturnix ja-
ponica) eggs were obtained from the
Research Institute of Animal Produc-
tion, Prague, Czech Republic, and
from B&D Farm (Harrah, OK). For in
situ hybridization of Ptch1 mRNA, we
used chick embryos. Eggs were incu-
bated at 38°C and embryos ranging
from stage 6 to 23, as deﬁned by Ham-
burger and Hamilton (1951) (23–100
hr of incubation), were studied. We
injected over 150 embryos with 5E1
hybridoma cells. As a control, we used
approximately 100 embryos injected
with 12cA5 hybridoma cells and as an
additional control we used uninjected
embryos.

Immunohistochemistry and
Histochemistry

Shh was detected in cryostat sections
derived from embryos ﬁxed in 4%
Phosphate

Buffered

Paraformalde-

hyde using monoclonal antibody 5E1
(Hybridoma Bank) diluted at 1:50. As
a secondary antibody, Goat anti-
Mouse Biotin (Sigma B7264) (1:500)
was used and tertiary antibody was
Extravidin Px (Sigma 2886) (1:100)
and DAB (Sigma D5905) as a chromo-
gen.

The endothelium of vessels was vi-

sualized with monoclonal antibody
QH1 (Hybridoma Bank) in parafﬁn
sections (1:1,000). As a secondary an-
tibody, we used Goat anti-Mouse Px
IgG (Sigma A 4416) and the reaction
product was detected with DAB.
Alternatively, endothelium was visu-
alized with VEGFR2 monoclonal anti-
body (kindly provided by Dr. Eich-
mann; Eichmann et al., 1997) diluted
1:1 in the cryostat sections. For en-
hancing the signal Tyramide Signal
Ampliﬁcation system (TSA, Dako) was
used according to the manufacturer’s
recommendation except that Strepta-

vidin peroxidase was used rather than
Extravidin peroxidase as it decreased
background levels signiﬁcantly. As a
secondary

antibody,

Rabbit

anti

mouse IgG1-Biotin was used.

Apoptosis was detected with Anti-

Cleaved Caspase-3 monoclonal anti-
body (BD Pharmingen 559565) at
1:500, in parafﬁn sections and cryo-
sections. The secondary antibody was
Goat anti-Rabbit Biotin (1:500) and
tertiary antibody was Extravidin Px
(Sigma 2886) (1:100), and visualized
using DAB.

Proliferating cells were detected

with anti-Phospho Histone H3 poly-
clonal antibody (Upstate 06570) 1:200
in cryosections. As a secondary anti-
body we used Goat-anti Rabbit Rhoda-
mine (Cappel Pharmaceutical) (1:500).

Periendothelial smooth muscle cells

were detected using anti-smooth mus-
cle actin Ab (Sigma A2547) 1:500 and
Goat anti Mouse TRITC as a second-
ary Ab (1:150). Phagocytic function of
macrophage-like cells was detected
with histochemical staining for Acid
Phosphatase

resistant

Tartaric

Acid. Embryos were ﬁxed in acetone
overnight at 4°C, transferred into xy-
lene, and subsequently embedded in
parafﬁn. Sections were deparafﬁnized
with xylene and rehydrated through
acetone and acetone/distiled water (1:
1). Sections were incubated overnight
at room temperature in solution pre-
pared with 10 mg of Naphtol-AS-BI
phosphate (Sigma, 70491-Fluka) in
0.5 ml N,N-dimethyl formamide. This
solution was resuspended in 50 ml of
0.1M acetic acid buffer, pH 5.2 (Wal-
pore acetate), and 20 mg of Fast Red
Violet (Sigma, F3381) was added, as
well as a minimum of 140 mg of Tar-
taric Acid (Sigma Aldrich, 14314DE).

mRNA In Situ Hybridization

A plasmid containing chick Ptch1
(clone 200; a gift from M. Scott) (Xie et
al., 1997) was linearized with SalI and
transcribed using T3 polymerase. Hy-
bridization in situ on parafﬁn sections
was carried out as described (Nieto et
al., 1996; Nanka et al., 2006).

Vascular System Ink
Injection

The vascular system was visualized in
vivo by injecting black ink diluted 1:20

1932 KOLESOVA

´ ET AL.

in PBS via the vitelline vein using a
glass capillary. The beating heart dis-
tributed the ink completely through-
out the vascular system. After injec-
tion, embryos were collected, ﬁxed in
4% PFA, and analyzed.

Inhibition of Shh Function
With Anti-Shh (5E1)
Antibody

Just before the injection, around 10

hybridoma cells were collected by cen-
trifugation and resuspended in about
200

␮l of Liebovitz L-15 medium

(Sigma). This suspension was loaded
into a small capillary and injected un-
der the vitelline membrane near the
branchial region of quail and chick
embryos. Injecting the cells to the
paraxial mesoderm caused the same
level of Shh inhibition as with injec-
tion in the proximity of branchial
arches, which is less invasive. After
injection, embryos were incubated for
1 to 3 days and isolated for analysis.
Embryos

were

either

injected

stages 10 –12 and re-incubated either
until they reached stage 18 or 21–23,
or embryos were injected at stage
13–15 embryos and incubated until
stage 21–23. We used either 5E1 anti-
Shh mouse hybridoma or 12CA5
mouse hybridoma cells, which produce
an anti-HA antibody (Handley-Gear-
hart et al., 1994).

Quantiﬁcation

QH1-positive cell aggregates and cells
not integrated into the endothelial
layer, macrophage-like cells positive
for acid phosphatase, apoptotic cells
exhibiting cleaved caspase-3 Ab, and
proliferating cells positive for Phospho
Histone-3, were counted. Counting
was performed on serial 10-

␮m sec-

tions from the following morphological
regions: each branchial arch, dorsal
aorta, anterior cardinal vein, and out-
ﬂow tract regions. The dorsal aorta
region includes its surrounding struc-
tures ventrally to the sixth branchial
arch, the liver primordium, and dor-
sally up to the neural tube. The ante-
rior cardinal vein region includes all
anterior structures besides the eye
and brain, and the area ventral to the
ﬁrst branchial arch and the oral cav-
ity. The outﬂow tract region includes
area of vessels and mesenchyme,

starting caudally in the heart and con-
tinuing cranially to the branchial
arches.

Cells were counted in every fourth

section, at a minimum of 20 sections
per region. Six embryos were analyzed
for the number of QH1-positive cells,
four

embryos

for

macrophage-like

cells and for apoptotic cells and two
embryos for the number of proliferat-
ing cells. Counts were averaged and
the standard deviation was deter-
mined.

ACKNOWLEDGMENTS

We thank Ms. E. Kluza´kova´, M. Plesch-
nerova´, A. Kvasilova´, and Mr. M. Tsma
for excellent technical assistance. This
work was supported by GAUK 54/
203209 (Grant Agency of Charles
University) to Hana Kolesova´, by
MSM 0021620806 (project from The
Ministry of Education of The Czech
Republic) to Milosˇ Grim, and by NIH
grant 1R01HD042307 to Henk Ro-
elink.

REFERENCES

Ahlgren SC, Bronner-Fraser M. 1999. In-

hibition of sonic hedgehog signaling in
vivo results in craniofacial neural crest
cell death. Curr Biol 9:1304 –1314.

Astorga J, Carlsson P. 2007. Hedgehog in-

duction of murine vasculogenesis is me-
diated by Foxf1 and Bmp4. Development
134:3753–3761.

Byrd N, Becker S, Maye P, Narasimhaiah

R, St-Jacques B, Zhang X, McMahon J,
McMahon A, Grabel L. 2002. Hedgehog
is required for murine yolk sac angiogen-
esis. Development 129:361–372.

Carpenter D, Stone DM, Brush J, Ryan A,

Armanini M, Frantz G, Rosenthal A, de
Sauvage FJ. 1998. Characterization of
two patched receptors for the vertebrate
hedgehog protein family. Proc Natl Acad
Sci USA 95:13630 –13634.

Cuadros MA, Coltey P, Carmen Nieto M,

Martin C. 1992. Demonstration of a
phagocytic cell system belonging to the
hemopoietic

lineage

and

originating

from the yolk sac in the early avian em-
bryo. Development 115:157–168.

Eichmann A, Corbel C, Nataf V, Vaigot P,

Breant C, Le Douarin NM. 1997. Ligand-
dependent development of the endothe-
lial and hemopoietic lineages from em-
bryonic mesodermal cells expressing
vascular endothelial growth factor recep-
tor 2. Proc Natl Acad Sci USA 94:5141–
5146.

Evans DJ, Noden DM. 2006. Spatial rela-

tions between avian craniofacial neural
crest and paraxial mesoderm cells. Dev
Dyn 235:1310 –1325.

Goodrich LV, Milenkovic L, Higgins KM,

Scott MP. 1997. Altered neural cell fates
and medulloblastoma in mouse patched
mutants. Science 277:1109 –1113.

Hamburger V, Hamilton HL. 1951. A se-

ries of normal stages in the development
of the chick embryo. J. Morphol. 88:49 –
90.

Handley-Gearhart

PM,

Stephen

AG,

Trausch-Azar

JS,

Ciechanover

Schwartz AL. 1994. Human ubiquitin-
activating enzyme, E1. Indication of po-
tential nuclear and cytoplasmic subpopu-
lations

using

epitope-tagged

cDNA

constructs. J Biol Chem 269:33171–33178.

Hiruma T, Hirakow R. 1995. Formation of

the pharyngeal arch arteries in the chick
embryo. Observations of corrosion casts
by scanning electron microscopy. Anat
Embryol (Berl) 191:415– 423.

Jaffredo T, Gautier R, Eichmann A, Diet-

erlen-Lievre F. 1998. Intraaortic hemo-
poietic cells are derived from endothelial
cells during ontogeny. Development 125:
4575– 4583.

Jeong J, Mao J, Tenzen T, Kottmann AH,

McMahon AP. 2004. Hedgehog signaling
in the neural crest cells regulates the
patterning and growth of facial primor-
dia. Genes Dev 18:937–951.

Kolesova H, Lametschwandtner A, Rocek

Z. 2007. The evolution of amphibian
metamorphosis: insights based on the
transformation of the aortic arches of Pe-
lobates fuscus (Anura). J Anat 210:379 –
393.

Le Lievre CS, Le Douarin NM. 1975. Mes-

enchymal derivatives of the neural crest:
analysis of chimaeric quail and chick em-
bryos. J Embryol Exp Morphol 34:125–
154.

Marigo V, Tabin CJ. 1996. Regulation of

patched by sonic hedgehog in the devel-
oping neural tube. Proc Natl Acad Sci
USA 93:9346 –9351.

Nagase T, Nagase M, Yoshimura K,

Machida M, Yamagishi M. 2006. Defects
in aortic fusion and craniofacial vascula-
ture in the holoprosencephalic mouse
embryo under inhibition of sonic hedge-
hog signaling. J Craniofac Surg 17:736 –
744.

Nanka O, Valasek P, Dvorakova M, Grim

M. 2006. Experimental hypoxia and em-
bryonic angiogenesis. Dev Dyn 235:723–
733.

Nieto MA, Patel K, Wilkinson DG. 1996. In

situ hybridization analysis of chick em-
bryos in whole mount and tissue sec-
tions. Methods Cell Biol 51:219 –235.

Nimmagadda S, Loganathan PG, Wilting

J, Christ B, Huang R. 2004. Expression
pattern of VEGFR-2 (Quek1) during
quail development. Anat Embryol (Berl)
208:219 –-224.

Noden DM, Trainor PA. 2005. Relations

and interactions between cranial meso-
derm and neural crest populations. J
Anat 207:575– 601.

Pardanaud L, Yassine F, Dieterlen-Lievre

F. 1989. Relationship between vasculo-
genesis, angiogenesis and haemopoiesis
during avian ontogeny. Development
105:473– 485.

SHH AND BRANCHIAL VESSEL DEVELOPMENT 1933

Pola R, Ling LE, Silver M, Corbley MJ,

Kearney M, Pepinsky RB, Shapiro R,
Taylor FR, Baker DP, Asahara T, Isner
JM. 2001. The morphogen Sonic hedge-
hog is an indirect angiogenic agent up-
regulating two families of angiogenic
growth factors. Nature Medicine 7:706 –
711.

Roelink H, Porter JA, Chiang C, Tanabe Y,

Chang DT, Beachy PA, Jessell TM. 1995.
Floor plate and motor neuron induction
by different concentrations of the amino-
terminal

cleavage

product

sonic

hedgehog autoproteolysis. Cell 81:445–
455.

Sasai N, Mizuseki K, Sasai Y. 2001. Re-

quirement of FoxD3-class signaling for
neural crest determination in Xenopus.
Development 128:2525–2536.

van Tuyl M, Groenman F, Wang J, Kulis-

zewski M, Liu J, Tibboel D, Post M. 2007.
Angiogenic factors stimulate tubular
branching morphogenesis of sonic hedge-
hog-deﬁcient lungs. Dev Biol 303:514 –
526.

Vokes SA, Krieg PA. 2002. Endoderm is

required for vascular endothelial tube
formation, but not for angioblast speciﬁ-
cation. Development 129:775–785.

Vokes SA, Yatskievych TA, Heimark RL,

McMahon J, McMahon AP, Antin PB,
Krieg PA. 2004. Hedgehog signaling is
essential for endothelial tube formation
during

vasculogenesis.

Development

131:4371– 4380.

Washington Smoak I, Byrd NA, Abu-Issa

R, Goddeeris MM, Anderson R, Morris J,
Yamamura K, Klingensmith J, Meyers
EN. 2005. Sonic hedgehog is required for
cardiac outﬂow tract and neural crest
cell development. Dev Biol 283:357–372.

Wiegreffe C, Christ B, Huang R, Scaal M.

2007. Sclerotomal origin of smooth mus-
cle cells in the wall of the avian dorsal
aorta. Dev Dyn 236:3191–3192.

Wijgerde M, McMahon JA, Rule M, McMa-

hon AP. 2002. A direct requirement for
Hedgehog signaling for normal speciﬁca-
tion of all ventral progenitor domains in
the

presumptive

mammalian

spinal

cord. Genes Dev 16:2849 –2864.

Wilting J, Eichmann A, Christ B. 1997.

Expression of the avian VEGF receptor
homologues Quek1 and Quek2 in blood-
vascular and lymphatic endothelial and
non-endothelial cells during quail em-
bryonic development. Cell Tissue Res
288:207–223.

Xie J, Johnson RL, Zhang X, Bare JW,

Waldman FM, Cogen PH, Menon AG,
Warren RS, Chen LC, Scott MP, Epstein
EH,

Jr.

1997.

Mutations

the

PATCHED gene in several types of spo-
radic extracutaneous tumors. Cancer
Res 57:2369 –2372.

Yamagishi C, Yamagishi H, Maeda J,

Tsuchihashi T, Ivey K, Hu T, Srivastava
D. 2006. Sonic hedgehog is essential for
ﬁrst pharyngeal arch development. Pedi-
atr Res 59:349 –354.

Zhang XM, Ramalho-Santos M, McMa-

hon AP. 2001. Smoothened mutants re-
veal redundant roles for Shh and Ihh
signaling including regulation of L/R
asymmetry by the mouse node. Cell
105:781–792.

1934 KOLESOVA

´ ET AL.

DEVELOPMENTAL BIOLOGY 180, 35 –40 (1996)
ARTICLE NO. 0282

Surgical Removal of Limb Bud Sonic hedgehog
Results in Posterior Skeletal Defects

Sylvia M. Pagan,*

Maria A. Ros,†

Cliff Tabin,* and John F. Fallon‡

*Department of Genetics, Harvard Medical School, Boston, Massachusetts 02115;
†Departamento de Anatomia y Biologia Celular, Universidad de Cantabria, 39011 Santander,
Spain; and

‡Department of Anatomy, University of Wisconsin, Madison, Wisconsin 53706

Using Sonic Hedgehog (Shh) as a marker for polarizing region cells we have repeated the experiments of MacCabe et al.
(1973) and Fallon and Crosby (1975) in an attempt to reexamine the question of a continuous role for the polarizing region
during limb development. We report that the earlier experiments probably left Shh-expressing cells after surgery. Our
results show that Shh-expressing cells do not regenerate and complete removal of the polarizing region results in truncations
along the anteroposterior (A –P) axis; further, A – P patterning cannot be restored when a bead soaked in FGF is implanted
in the limb bud mesenchyme to maintain outgrowth after extirpation of the polarizing region. However, in order to
reproducibly remove all Shh-positive cells, it is possible that cells with posterior limb skeletal fate also must be removed.
Therefore, microsurgical approaches do not permit an unequivocal answer to the question raised in this and the earlier
papers and it remains a reasonable possibility that at least up to stage 20 –21 the polarizing region plays a continuous role
in patterning of the limb bud during its development.

q 1996 Academic Press, Inc.

INTRODUCTION

of the limb. Arguing that these data could also reflect the
regeneration of polarizing activity after the surgeries, Fallon
and Crosby (1975) took posterior tissue from wing buds 24

The polarizing region (zone of polarizing activity or ZPA)

and 48 hr after polarizing region removal and assayed it for

was operationally defined more than 25 years ago as a re-

polarizing activity in a host limb bud. Negative results for

stricted area of posterior limb mesoderm that caused mirror

the presence of polarizing activity in the operated buds al-

image duplications of digit patterns along the anteroposter-

lowed them to conclude that the polarizing region was not

ior (A –P) axis when grafted into the anterior margin of a

regenerated following removal. They also suggested that,

host wing bud (Saunders and Gasseling, 1968; Tickle et al.,

given the fact that normal wing development still occurred

1975). It was assumed that the polarizing region played an

in about 30% of the cases, this indicated that ‘‘if the polariz-

important role in the anteroposterior patterning of the de-

ing zone had any role during limb development, it must be

veloping limb. One way to test this hypothesis was to surgi-

at an early stage, as during limb induction, and any informa-

cally remove it, expecting either truncations along the A –

tion from the zone is further acted upon throughout the

P axis or uniform skeletal elements along the A –P axis if

morphogenetic field.’’ Nevertheless, the presence of the po-

the polarizing region had an essential role in A – P specifica-

larizing region until nearly the end of limb development

tion during limb development. Results published by Mac-

has been taken as a strong suggestion of it playing an active

Cabe et al. (1973) argued against this conclusion: when the

role in patterning as the limb bud elongates (Tickle et al.,

polarizing region was removed from early limb buds, normal

1975). However, there is no conclusive evidence arguing

wings developed in about half of the cases, suggesting that

against the hypothesis that it exclusively acts early.

it was not involved in normal anteroposterior patterning

Three possibilities could explain why MacCabe et al.

(1973) and Fallon and Crosby (1975) obtained a high percent-
age of normal wings after polarizing region removals. First,

The first two authors contributed equally to this work.

all of the polarizing region was removed after the surgeries

To whom correspondence should be addressed at Department

and it is indeed dispensable for A– P patterning after limb

of Anatomy, University of Wisconsin, 1300 University Avenue,

induction stages, either because it has already fixed the posi-

Madison WI 53706. Fax: 608/262-2327. E-mail: jffallon@facstaff.
wisc.edu.

tional identity of the future wing elements or because it

0012-1606/96 $18.00
Copyright

q 1996 by Academic Press, Inc.

All rights of reproduction in any form reserved.

AID

DB 8384

6x14$$$361

10-28-96 20:43:12

dba

AP: Dev Bio

Pagan et al.

has already set the cascade of events that ultimately result

tissue in each successive experiment until we obtained no
Shh

signal in the limbs as tested by in situ hybridizations.

in A – P patterning. Second, all of the polarizing region was
removed but it eventually regenerates, resulting in restora-

We scored the amount of remaining Shh in the limbs and
compared these results with the wing patterns of Day 11

tion of anteroposterior pattern. This alternative would im-
ply that the negative results for polarizing activity obtained

embryos from the same batch.

When we performed more conservative surgeries, we ob-

by Fallon and Crosby (1975) were due to assays that were
not sensitive enough. Third, perhaps not all of the polarizing

tained results strikingly similar to those reported by Mac-
Cabe et al. (1973) and Fallon and Crosby (1975): at Day 11

region was removed with the surgeries and the remaining
amounts were sufficient to maintain normal A –P pat-

normal wings were obtained in about 30% of the cases, with
the rest exhibiting various postaxial defects, one of the most

terning of the developing bud, implying again that this re-
sidual activity must have been missed by the less sensitive

common being the presence of only a humerus, a radius,
and digits 2 and 3, as was also the case for Fallon and Crosby

polarizing activity assays. Discerning among these three
possibilities requires a molecular marker for the polarizing

(Table 1A, Fig. 1B). However, in situ hybridization of the
stage 20 –21 limbs revealed that only a small percentage

region cells, which did not exist at the time that the earlier
studies were carried out.

had no Shh left, and a large fraction exhibited a considerable
degree of remaining Shh signal in the operated limb (Table

Several lines of evidence suggest that Sonic Hedgehog

(Shh), a vertebrate homologue of the Drosophila hedgehog

1A, Fig. 1A). When we modified our surgeries such that
64% of the limbs had no Shh left and the remnant had very

gene, is a crucial component of the polarizing region signal-
ing pathway. Transcripts of Shh strongly localize to the

small amounts, the wing skeletal patterns of the Day 11
wings looked noticeably different: 100% exhibited extreme

polarizing region and its expression pattern strikingly corre-
lates with maps of the polarizing region throughout devel-

truncations along the A– P axis, most of them having only
a humerus, a radius, and a digit 2 (Table 1B, Fig. 1C).

opment (Riddle et al., 1993). Grafts of Shh-expressing cells
(Riddle et al., 1993; Chang et al., 1994) as well as SHH

To investigate whether Shh regenerated after the polariz-

ing region removals we performed in situ hybridizations of

protein in beads (Lo´pez-MartıBnez et al., 1995) have been
shown to induce polarized digit duplications along the ante-

limbs from embryos harvested 24 hr after the surgeries. Our
results show that the amount of Shh signal and the percent-

roposterior axis, which are indistinguishable from the clas-
sical polarizing region grafts. It thus seems likely that Shh

age of Shh-positive limbs after 24 hr is correlated with the
amount of Shh and percentage of positive limbs at 0 hr. In

is the molecule responsible for the patterning role of the
polarizing region in the developing limb and that it would

an experiment where we analyzed limbs at several times
after the surgery, it was clear that the Shh signal in the 24-

serve as an excellent marker for identifying the cells that
belong to the polarizing region. We have reinvestigated the

hr limbs came from Shh-expressing cells that had not been
removed during the surgery (data not shown). Table 1C

requirement for polarizing region tissue during limb devel-
opment using Shh expression as a marker.

summarizes an experiment where we removed a more distal
portion of the polarizing region, purposefully leaving proxi-
mal Shh-expressing cells. Almost 90% of the limbs har-
vested at 0 hr after the surgeries had high remaining

MATERIALS AND METHODS

amounts of proximal Shh, close to the junction of the limb
bud and body wall (Fig. 2). After 24 hr, 38% of the limb

White Leghorn chick embryos of stages 20– 21 (Hamburger and

Hamilton, 1951) were used for surgeries. The polarizing region was

buds still showed high amounts of Shh, 50% had low levels

excised with tungsten wire needles and embryos were either har-

of the signal, in each case confined to the proximal regions

vested immediately or allowed to develop for 24 hr or until the

of the limb buds (Fig. 2, n Å 8). In similar specimens, the

11th day of embryonic development. Whenever indicated, a bead

skeletal patterns of the Day 11 wings showed a relatively

(Affi-Gel Blue Gel, Bio-Rad) soaked in FGF-4 (0.85 mg/ml, a gift

high percentage (33%) of normal limbs; those remaining

from the Genetics Institute) was inserted in a cut made in the limb

formed a humerus, radius, ulna, and digit 2. When we com-

bud mesenchyme after the polarizing region removal. Embryos har-

pared these skeletal patterns with those obtained from our

vested 0 and 24 hr after the surgeries were fixed in 4% paraformal-

previous experiments, the most obvious difference was the

dehyde, dehydrated in a graded methanol series, and used for in situ

addition of an ulna in all limbs (compare Tables 1A and 1B

hybridization using the Shh probe as previously described (Riddle et

with Table 1C). This is consistent with the in situ analysis

al.,

1993). Embryos harvested at Day 11 were washed in PBS, fixed

in 4% paraformaldehyde, and stained for cartilage with Alcian blue.

of the 0- and 24-hr limbs, where the Shh signal was always
proximal, i.e., not near the digit area, and correlated with
the A – P pattern restoration at the zeugopod level. In con-

RESULTS AND DISCUSSION

trast, we never observed an ulna in truncated limbs at Day
11 from surgeries done that were similar to those where no
Shh

was left proximally.

Table 1 summarizes the results from surgeries performed

on stage 20– 21 chicken embryos. Our approach was to at-

We conclude from these experiments that Shh, and thus

the polarizing region, does not regenerate following polariz-

tempt to achieve a constant extent of cutting during each
individual experiment, progressively trying to remove more

ing region removal and that the degree of A – P patterning

q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

AID

DB 8384

6x14$$$361

10-28-96 20:43:12

dba

AP: Dev Bio

Shh and Limb Patterning

TABLE 1
Remaining Shh Signal after Polarizing Region Removal and Corresponding Wing Structures Present at Day 11

Goal of surgery:

A (Leave Shh)

B (Leave no Shh)

C (Proximal Shh)

D (No Shh / FGF)

Shh

expressions

0 hr

No Shh

17%

64%

100%

Low levels Shh

33%

29%

Some Shh

25%

11%

High levels Shh

25%

89%

n Å 12

n Å 14

n Å 9

n Å 6

Skeletal elements

11 days

10%

57%

R, 2

43%

90%

R, 2, 3

28.5%

R, U, 2

66%

43%

R, U, 2, 3

R, U, 2, 3, 4

28.5%

33%

n Å 7

n Å 5

n Å 3

n Å 6

Note.

During each surgery, great care was taken to be consistent in the amount of tissue removed in embryos of each batch (A, B, C,

and D). However, some variation is likely between surgeries done on different days; therefore, the phenotypic outcomes are only analyzed
relative to the amount of Shh left in representative limbs done in parallel, and numbers are not combined between experiments. We
performed a total of 203 polarizing region removals; a subset of representative experiments is shown here. The results obtained with those
experiments not shown are consistent with the data presented in the table. R, radius; U, ulna; 2, digit 2; 3, digit 3; 4, digit 4.

in the Day 11 wings is correlated with the amount of Shh

al.,

1993). Shh maintains FGF-4 expression necessary for

limb outgrowth and FGF-4, in turn, maintains Shh in the

left in the operated buds. We reason that MacCabe et al.
(1973) and Fallon and Crosby (1975) did not remove all of

posterior wing mesoderm. Hence, after limb induction
stages the positional fates along the A –P axis could have

the polarizing region in their experiments, such that enough
Shh

-expressing cells were left to maintain patterning of the

already been established, and the role of Shh— and the po-
larizing region — at that point would be to support limb

limbs. Negative assays for polarizing activity that allowed
Fallon and Crosby (1975) to conclude that the polarizing

outgrowth through its interaction with FGF-4 in the overly-
ing ectoderm. These hypotheses were tested by attempting

region was not regenerated seem to contradict our result
that Shh is found in some limbs 24 hr after polarizing region

to maintain limb outgrowth by implanting a bead loaded
with FGF in the limb mesoderm after polarizing region re-

removals. This inconsistency can be reconciled by assum-
ing that at the time, the assay for polarizing activity was

moval. Table 1D summarizes the results of one such experi-
ment, where a bead of FGF-4 was implanted in the meso-

not sensitive enough. Tickle (1981) has shown that placing
a tissue graft beneath an intact apical ridge constitutes a

derm of stage 20 – 21 limb buds after the polarizing region
had been removed. As is shown in the table, no Shh signal

significantly more sensitive test for polarizing activity, as
opposed to placing the graft in a notch, the procedure that

was detectable in 100% of the limbs harvested right after
the surgeries, and at Day 11 all of the wings showed extreme

Fallon and Crosby followed. We speculate that assaying the
24-hr postsurgery limb buds using the more sensitive assay

A –P truncations (Fig. 3). Interestingly, the ulna seemed to
be the only structure that was to some extent restored with

would have resulted in duplications along the A –P axis
indicative of the residual polarizing activity.

these experiments, since 43% of the limbs now had an ulna
in addition to the humerus, the radius, and digit 2 (compare

We envisioned two possible explanations of why remov-

ing all of Shh resulted in severe pattern defects along the

to 0% in experiment of Table 1B).

When we analyzed the limbs harvested 24 hr after polariz-

A– P axis of the limbs. One was that Shh could be necessary
to actively pattern structures along the A –P axis as the

ing region removal and FGF bead implants, we found that
100% showed Shh signal to various extents and always in

wing grows and develops, in which case removing it at early
stages results in postaxial defects. Alternatively, it was pos-

a proximal position, away from the digit region (Fig. 3, n Å
6). Given that we removed all of Shh at 0 hr and that the

sible that Shh has completed its A – P patterning role after
early limb induction stages and the reason truncations oc-

position of the Shh signal at 24 hr is always proximal, we
assume that these Shh-expressing cells belonged to the

cur is simply because of the disruption of a feedback loop
between Shh and FGF-4 (Laufer et al., 1994; Niswander et

flank at the time of polarizing region removal and that they

q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

AID

DB 8384

6x14$$$361

10-28-96 20:43:12

dba

AP: Dev Bio

Pagan et al.

FIG. 1. (A) Tissue from the polarizing region was surgically removed and remaining polarizing region cells were visualized by in situ
hybridizations with the Shh probe. Limbs in the figure show examples of what were categorized as low or high levels of remaining Shh
as classified in Table 1. Control: Shh expression in an unoperated normal limb at stage 20– 21. Arrows indicate remaining Shh signal in
the limb buds. (B) Representative Day 11 (Table 1A) limbs show the range of wing skeletal patterns obtained after incomplete polarizing
region removals. (C) When all or most of the polarizing region was removed the resulting Day 11 (Table 1B) wing skeletal patterns exhibited
extreme truncations along the A –P axis.

are brought into proximity with the FGF bead as the limb

We have shown that after complete polarizing region re-

moval, posterior truncations along the anteroposterior axis

bud grows and heals, inducing expression of Shh in a poste-
rior region of the limb bud where it normally is not ex-

result. The cause of truncations occurring in the absence of
Shh

remains to be addressed. First, it is possible that the

pressed at that stage of development. Similar results have
been reported by Yang and Niswander (1995), where a bead

surgery to remove all of Shh-expressing cells is radical
enough to result in the observed deletions. Second, cells

of FGF-4 placed close to the limb flank enables cells in this
region to express Shh when they normally would not. We

may no longer realize their fate without the signal from the
polarizing region. Third, cell death may be induced in these

propose that the proximal Shh expression permits regula-
tion of patterning at the level of the zeugopod and the pres-

cells after polarizing region removal, with the loss of poste-
rior structures being a direct result of this cell death. This

ence of an ulna in a high percentage of the wings. These
results parallel those from our previous experiments (com-

is the least likely since it would be expected that all the
digits would be affected (Todt and Fallon, 1987). In sum-

pare Tables 1C and 1D), where remaining proximal Shh
after polarizing region removal equally resulted in regula-

mary, we have resolved a discrepancy in the literature by
showing that the results previously obtained by MacCabe

tion of the ulna. In conclusion, our FGF experiments indi-
cate that Shh is probably needed to actively pattern the

et al.

(1973) and Fallon and Crosby (1975) probably were

achieved after incomplete polarizing region removals. In

A– P axis throughout development of the bud. When we
provided an FGF bead to maintain outgrowth, we did not

order to consistently remove all Shh-positive cells it is pos-
sible that cells with posterior skeletal fates are also re-

observe any restoration of A – P pattern other than the ap-
pearance of an ulna, which can be explained by an induction

moved. Our data indicate that microsurgery cannot unam-
biguously answer the question of a continuous role for po-

of Shh expression in cells close to the limb flank.

q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

AID

DB 8384

6x14$$$361

10-28-96 20:43:12

dba

AP: Dev Bio

Shh and Limb Patterning

FIG. 2. When we performed polarizing region removals such that Shh-expressing cells were left proximally when examined on the day
of surgery (0 D; D, day), the skeletal patterns were normal in a considerable number of wings, and the remaining wings showed extreme
A –P truncations at the autopod level, but restoration of the ulna at the zeugopod level (11 D). This phenotype correlated with continued
proximal Shh expression in a high degree of limbs analyzed 1 day after surgery (1 D). Control: Shh expression in unoperated limbs at 0
time and 1 day postsurgery. Arrows indicate remaining Shh signal in the limb buds.

FIG. 3. Implanting an FGF-soaked bead in the operated limbs to maintain outgrowth did not restore A –P pattern at the autopod level
(11 D). Regulation of the ulna in almost half of the cases correlates with induction of proximal Shh by FGF, as analyzed in limbs 1 day
after surgery (1 D). Arrows indicate induced Shh signal.

q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

AID

DB 8384

6x14$$8384

10-28-96 20:43:12

dba

AP: Dev Bio

Pagan et al.

M. A., Simandl, B. K., Beachy, P. A., and Fallon, J. F. (1995). Limb-

larizing region during limb development posed by MacCabe

patterning activity and restricted posterior localization of the

et al.

and Fallon and Crosby. Therefore, it remains a reason-

amino-terminal product of Sonic hedgehog cleavage. Curr. Biol.

able possibility that there is a continuous role for the polar-

5, 791 –796.

izing region and Shh in A– P patterning of the developing

MacCabe, A. B., Gasseling, M. T., and Saunders Jr., J. W. (1973).

limb at least up to stage 20– 21 of development.

Spatio-temporal distribution of mechanisms that control out-
growth and antero-posterior polarization of the limb bud in the
chick embryo. Mech. Ageing Dev. 2, 1– 12.

ACKNOWLEDGMENTS

Niswander, L., Tickle, C., Vogel, A., Booth, I., and Martin, G. R.

(1993). FGF-4 replaces the apical ectodermal ridge and directs
outgrowth and patterning of the limb. Cell 75, 579– 587.

This work was supported by Grant 95/0576 from FISS to M.A.R;

Riddle, R. D., Johnson, R. L., Laufer, E., and Tabin, C. (1993). Sonic

NIH Grant HD32443 to C.T; and NIH Grant HD32551 to J.F.F. We

hedgehog mediates the polarizing activity of the ZPA. Cell 75,

thank B. Kay Simandl and Won-Sun Kim for comments on the

1401 –1416.

manuscript.

Saunders, J. W., and Gasseling, M. T. (1968). Ectodermal-mesenchy-

mal interactions in the origin of limb symmetry. In ‘‘Epithelial
and Mesenchymal Interactions’’ (R. Fleischmajer and R. E. Bill-

REFERENCES

ingham, Eds.), pp. 78– 97. Williams & Wilkins, Baltimore.

Tickle, C. (1981). The number of polarizing region cells required

Chang, D. T., Lo´pez, A., von Kessler, D. P., Chiang, C., Simandl,

to specify additional digits in the developing chick wing. Nature

B. K., Zhao, R., Seldin, M. F., Fallon, J. F., and Beachy, P. A.

289, 295– 298.

(1994). Products, genetic linkage and limb patterning activity of

Tickle, C., Summerbell, D., and Wolpert, L. (1975). Positional sig-

a murine hedgehog gene. Development 120, 3339 –3353.

nalling and specification of digits in chick limb morphogenesis.

Fallon, J. F., and Crosby, G. M. (1975). Normal development of the

Nature

254, 199–202.

chick wing following removal of the polarizing zone. J. Exp. Zool.

Todt, W., and Fallon, J. F. (1987). Posterior apical ectodermal ridge

193, 449– 455.

removal in the chick wing bud triggers a series of events resulting

Hamburger, V., and Hamilton, H. L. (1951). A series of normal

in defective anterior pattern formation. Development 101, 501–

stages in the development of the chick embryo. J. Exp. Morphol.

515.

88, 49– 92.

Yang, Y., and Niswander, L. (1995). Interaction between the signal-

Laufer, E., Nelson, C. E., Johnson, R. L., Morgan, B. A., and Tabin,

ing molecules WNT7a and SHH during vertebrate limb develop-

C. (1994). Sonic hedgehog and Fgf-4 act through a signalling cas-

ment: Dorsal signals regulate anteroposterior patterning. Cell 80,

cade and a feedback loop to integrate growth and patterning of

939–947.

the developing limb bud. Cell 79, 993–1003.

LoBpez-MartıBnez, A., Chang, D. T., Chiang, C., Porter, J. A., Ros,

Received for publication June 6, 1996

q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

AID

DB 8384

6x14$$$361

10-28-96 20:43:12

dba

AP: Dev Bio

Wyszukiwarka

Podobne podstrony:
ANATOMIA I HISTOLOGIA JAMY USTNEJ
anatomia i histologia zebow mlecznych
Anatomiaa, Sem 2, Anatomia i histologia zwierząt, egzamin
Biomechanika ssż anatomia i histologia przyzębia
Anatomia i histologia zębów stałych niedojrzałych
anatomia 1 rok, fizjologia i anatomia
ANATOMIA I HISTOLOGIA JAMY USTNEJ
anatomia i histologia zebow mlecznych
PODSTAWY ANATOMII I FIZJOLOGII CZLOWIEKA
Anatomia i fizjologia układ mięśniowy (3)
Fizjologia i Anatomia wyklad I
Wyklad 3 pobudliwosc, Dietetyka, Anatomia i fizjologia człowieka, Fizjologia wykłady
Anatomia i fizjologia, Kulturystyka
Anatomia i fizjologia układ szkieletowy
Anatomia i fizjologia układu oddechowego

więcej podobnych podstron