European Journal of Neuroscience, Vol. 27, pp. 1710–1721, 2008
doi:10.1111/j.1460-9568.2008.06133.x
Generation recruitment and death of brain cells throughout
the life cycle of Sorex shrews (Lipotyphla)
Katarzyna Bartkowska,1 Rouzanna L. Djavadian,1 Jan R. E. Taylor 2 and Kris Turlejski1
1
Department of Molecular and Cellular Neurobiology, Nencki Institute of Experimental Biology, 3 Pasteur Street, 02-093 Warsaw,
Poland
2
University of Bialystok, Institute of Biology, Bialystok, Poland
Keywords: apoptosis, dentate gyrus, neurogenesis, olfactory bulb, subventricular zone
Abstract
Young shrews of the genus Sorex that are born in early summer reduce their body size before wintering, including a reduction of brain
weight of 10–30%. In the spring they mature sexually, double their body weight and regain about half of the loss in brain weight. To
investigate the mechanisms of brain weight oscillations we studied the rate of cell death and generation in the brain during the whole
life cycle of the common shrew (Sorex araneus) and pygmy shrew (S. minutus). After weaning, shrews generate new brain cells in
only two mammalian neurogenic zones and approximately 80% of these develop into neurones. The increase of the shrew brain
weight in the spring did not depend on recruitment of new cells. Moreover, adult Sorex shrews did not generate new cells in the
dentate gyri. Injections of 5-HT1A receptor agonists in the adult shrews induced neurogenesis in their dentate gyri, showing the
presence of dormant progenitor cells. Generation of new neurones in the subventricular zone of the lateral ventricles and their
recruitment to olfactory bulbs continued throughout life. TUNEL labelling showed that the rate of cell death in all brain structures,
including the proliferation zones and olfactory bulb, was very low throughout life. We conclude that neither cell death nor recruitment
significantly contributes to seasonal oscillations and the net loss of brain weight in the Sorex shrews. With the exception of dentate
gyrus and olfactory bulb, cellular populations of brain structures are stable throughout the life cycle of these shrews.
Introduction
Sorex shrews live in the colder zones of Eurasia and North
America. They are born in the spring or summer and, when
3–4 weeks old, disperse and live a solitary life as subadults. They
mature sexually only after wintering, breed in the second spring and
summer, and die out before the next winter (Churchfield, 1990;
Rychlik, 1998). These shrews have unique wintering adaptations; in
the autumn they reduce the size of their bodies, including reduction
of the brain weight and the capacity of the brain case by 10–30%,
depending on geographical area, and then in the spring they
markedly increase their body weight and regain about half of the
brain weight and volume loss (Dehnel, 1949; Siivonen, 1954;
Pucek & Markov, 1964; Pucek, 1965a; Hyvarinen, 1969; Churchfield, 1990). Seasonal changes of weight are highest in the
hippocampus and neocortex, and lowest in the olfactory bulbs
(OBs), brainstem and cerebellum (Yaskin, 1994). Seasonal oscillations in the volume of some brain structures were also found in
birds (Nottebohm, 1981; Sherry et al., 1993; Barnea & Nottebohm,
1994) and were explained by seasonal changes in the rate of
neurogenesis and cell death (Goldman & Nottebohm, 1983; Kirn &
Nottebohm, 1993).
In all previously investigated mammalian species, post-developmental neurogenesis was found in two brain structures, the dentate
gyrus (DG) and subventricular zone (SVZ) of the lateral ventricles
Correspondence: Dr K. Turlejski, as above.
E-mail: [email protected]
Received 9 July 2007, revised 16 January 2008, accepted 7 February 2008
(Altman, 1962; Eriksson et al., 1998; Kornack. & Rakic, 1999;
Gage, 2000; Gould, 2007). In rodents 70–85% of the newly
generated cells differentiate into neurones that are incorporated
into the DG and OB (Kempermann et al., 1997; Cameron &
McKay, 2001). Adult neurogenesis was also observed in the
neocortex of the golden hamster (Huang et al., 1998) and
macaque (Gould et al., 1999). In addition, seasonal modulation of
the rate of proliferation in the DG and SVZ was found in the
golden hamster (Huang et al., 1998) and meadow vole (Galea &
McEwen, 1999). Therefore, the hypothesis of cell proliferation
being a mechanism of seasonal oscillations of the brain weight in
shrews was tested.
Serotonin, agonists of the 5-HT1A receptor and physical exercise
increase the rate of proliferation in the DG (Tanapat et al., 1999; Van
Praag et al., 1999; Santarelli et al., 2003), whereas stress lowers it
(Gould et al., 1998). The influence of oestrogens and androgens on the
neurogenesis is complex and may depend on the species, strain, sex,
age and other variables (Ormerod & Galea, 2001; Perfilieva et al.,
2001; Hajszan et al., 2007).
We aimed to answer the question of whether the seasonal
oscillations of brain weight in Sorex shrews could depend on changes
in the number of brain cells. Furthermore, are there an autumnal wave
of cell death and a spring wave of recruitment in any brain structure,
similar to that observed in birds (Kirn & Nottebohm, 1993)?
Additionally we investigated whether some of the factors altering
the rate of neurogenesis in other mammals similarly affect shrews and
therefore whether their precursor cells are regulated in a similar
manner (Grote & Hannan, 2007).
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
Cell generation and death in the brain of shrews 1711
Materials and methods
Animals
Forty-nine pygmy shrews (Sorex minutus) and 64 common shrews
(S. araneus) were used in this research. They were trapped in the
north-eastern part of Poland under licences from the Polish Ministry of
the Environment to J.R.E.T. and K.T. All experimental procedures
were approved by the State Committee for the Ethics of Animal
Experimentation and were compatible with the standards of the Polish
Law on Experimenting on Animals, which implements the European
Communities
Council
Directive
of
24 November 1986
(86 ⁄ 609 ⁄ EEC), as well as with the NIH Guide for the Care and
Use of Laboratory Animals. Animals were captured in north-eastern
Poland by permission of the Polish Ministry of Environment to
J.R.E.T. Shrews were collected over a 2.5-year period that covered the
whole life span of two consecutive age cohorts. They were classified
into six age groups: Summer I (June–August), Autumn I (September–
October), Winter (December–February), Spring (April–May),
Summer II and Autumn II. The groups from Summer I until Spring
consisted of five to seven animals, whereas older groups consisted of
only three to five shrews because of the scarcity of shrews of that age.
In the first half of their life the Sorex shrews are sexually immature and
their sex may be unequivocally determined only through section.
Therefore, some of these groups were sexually unbalanced. In
older groups the evidently gravid females were released at the place
of capture. Two females captured at a very early phase of pregnancy
and perfused 2 days later were excluded after the post-mortem
checkup.
randomly chosen series were used for BrdU and TUNEL labelling and
three for double-labelling with BrdU and phenotypic markers. The
remaining series were used for controls or sometimes to confirm the
results of the labelling.
TUNEL labelling
This technique allows for labelling of the free 3¢ ends of DNA and the
number of such ends is very high when DNA is fragmented in both
apoptosis and necrosis (Otsuki et al., 2003). Therefore, nuclei of
dying cells were labelled and visualized. Sections were washed in two
changes of phosphate-buffered saline, soaked for 30 min in 0.3%
H2O2 in methanol and then permeabilized for 20 min in ethanol ⁄ acetic acid (2 : 1) at 4 C. Sections were then washed twice in
phosphate-buffered saline and incubated for 1 h in the TUNEL
reaction mixture (Roche Diagnostics, IN, USA) containing 5 mL of
the terminal deoxynucleotidyl transferase and 45 mL of deoxyuridine
triphosphate marked with fluorescein. Afterwards, the tissue was
washed with phosphate-buffered saline, mounted on slides and
coverslipped with Fluoromount (Vector Laboratories, CA, USA).
Positive TUNEL controls were performed on separate sections treated
with DNAse I [500 mg ⁄ mL (Roche Diagnostics) in TrisCl with
MgCl2 at 37 C] to induce DNA breaks and then labelled with
TUNEL. In control sections, labelled nuclei were visible in all brain
structures and the cell density was similar to that seen in the Nisslstained material. For negative controls the DNAse I-treated sections
were incubated for 1 h with deoxyuridine triphosphate but without
terminal deoxynucleotidyl transferase, which resulted in a complete
lack of labelling.
Animal care and treatment
Except for the Winter groups, all shrews were given an i.p. injection of
50 mg ⁄ kg of bromodeoxyuridine (BrdU) (Sigma) in saline shortly
after their capture and the injection was repeated 2 h later. Winter
groups were captured in the autumn (September–October) and kept
under controlled conditions in the laboratory until January–February
and then injected with BrdU. Shrews were fed ad libitum and kept at a
constant temperature (+2 C) and on a light ⁄ dark cycle matching that
in Poland at that time. In each age group, shrews survived for either 2
or 14 days after injections. Additional groups of common shrews from
the Winter and Summer II groups were injected i.p. with 5-HT1A
receptor agonists, either 8-hydroxy-2-(di-n-propylamino)-tetralin
(8-OH-DPAT) (0.4 mg ⁄ kg, Tocris Cookson, Ellisville, MO, USA) or
buspirone (1.0 mg ⁄ kg, Sigma), 1 h before the first injection of BrdU.
One group of wintering shrews was trained to run on a treadmill for
1 h ⁄ day over four consecutive days, during which time BrdU
injections were performed on the third day at the cessation of the
day’s exercise. All drug-injected and trained shrews survived for
14 days after BrdU injections.
Perfusion and histological procedures
Animals were injected with pentobarbital (200 mg ⁄ kg, i.p.) and
perfused transcardially, first with saline and then briefly with 4%
paraformaldehyde in 0.1 m phosphate buffer. Brains were cut off the
spinal cord at the level of foramen magnum, removed from the brain
cases and weighed with 0.1-mg accuracy. Tissue was then post-fixed
for 2 weeks in 4% paraformaldehyde, cryoprotected with 30% sucrose
and cut coronally on a cryostat into 40-lm sections. All sections of a
brain were collected and arranged into 10 parallel series, of which one
was Nissl stained to allow for identification of brain structures. Two
Immunohistochemistry for BrdU
Bromodeoxyuridine is a thymidine analogue that is permanently
incorporated into DNA in the S-phase of mitotic division and may be
detected immunohistochemically in the nuclei of daughter cells (del
Rio & Soriano, 1989; Nowakowski et al., 1989). Therefore, the nuclei
of those cells that were generated at the time that BrdU was present in
the body after injection (40–90 min) are permanently labelled and may
be visualized. Free-floating sections were washed in twice-concentrated saline-sodium citrate, permeabilized for 2 h with 50% formamide in 2 · saline-sodium citrate at 60 C, rinsed for 5 min in
2 · saline-sodium citrate, denatured in 2 N HCl at 37 C for 30 min
and rinsed for 10 min in 0.1 m boric acid (pH 8.5). The sections were
then treated for 30 min with 1% H2O2 in Tris-buffered saline to block
endogenous peroxidases and washed in Tris-buffered saline with 0.1%
Triton X-100 and Tris-buffered saline with 0.1% Triton X-100 with
0.05% bovine serum albumin (TBS-B). After soaking for 1 h in 10%
goat serum in TBS-B, sections were incubated overnight at room
temperature (22C) with the primary anti-BrdU antibody (Boehringer
Mannheim, Germany, 1 : 1000 in TBS-B) and then rinsed in Trisbuffered saline with 0.1% Triton X-100 and TBS-B followed by
incubation with biotinylated goat anti-mouse secondary antibody
(Sigma, 1 : 100 in TBS-B) for 45 min. After further rinsing, the
extravidin-biotin-peroxidase complex (Sigma) was applied for 1 h and
peroxidase visualized with the 3,3¢-Diaminobenaidime kit (Vector
Laboratories). Finally, after rinsing in phosphate-buffered saline,
sections were mounted on gelatinized slides, air dried, dehydrated and
coverslipped with DePeX Mounting Medium (Serva, Heidelberg,
Germany). Control sections taken from another series were processed
in the same way but either without the primary or secondary antibody,
or both. Labelled cell nuclei were absent in all control tissue.
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
1712 K. Bartkowska et al.
Immunohistochemical double labelling
To determine the phenotypes of newly generated cells, the tissue was
double labelled for BrdU in combination with the neuronal marker
(NeuN) (neurones), glial fibrillary acidic protein (astrocytes) or
CNPase (oligodendrocytes) using immunofluorescent markers. BrdU
staining was performed as described above but using rat anti-BrdU
(1 : 500; Accurate Chemical, NY, USA) and a fluorescent secondary
antibody Alexa fluor 488 or Alexa fluor 568 (1 : 200; Molecular
Probes, OR, USA) after which the sections were immunolabelled with
primary antibodies for either NeuN (mouse, 1 : 200; Chemicon
International, Temecula, CA, USA), glial fibrillary acidic protein
(rabbit, 1 : 500; DAKO, Glostrup, Denmark) or CNPase (mouse,
1 : 200; Sigma), followed by a fluorescent secondary antibody.
Double-labelled cells and nuclei were photographed and counted
using a confocal microscope.
Quantification of the data
The TUNEL-labelled nuclei were counted and photographed under
510-nm fluorescent illumination using light and confocal microscopes. BrdU labelling was quantified with the light microscope under
20· and 40· objectives, and only labelled nuclei larger than about
1.0 lm were included in order to avoid counting artefacts. Each brain
structure on both sides of the brain was examined separately and all
TUNEL- or BrdU-labelled nuclei present were counted throughout a
whole series of sections (i.e. every tenth brain section). To establish
total numbers of BrdU-positive cells, BrdU-labelled nuclei in the
SVZ, rostral migratory stream and OB of both hemispheres were
pooled together, as were the numbers of labelled nuclei in the
subgranular and granular layers of the DG. These totals were
multiplied by 10 and the product taken as the number of labelled
nuclei contained in the whole structure. No difference in the rate of
BrdU or TUNEL labelling was found between males and females in
any group and therefore the data for both sexes were merged. For each
group of shrews, the means and SDs were calculated and any
significant differences between groups were assessed using the twotailed Student’s t-test (P < 0.05).
Results
General morphology of the shrew brain differs from
that in rodents (Fig. 1A and B)
The whole brain is compressed antero-posteriorly, so the shape of the
OB, cerebral hemispheres and cerebellum taken together is close to a
section of a sphere. The OBs are short and broad, with the anterior
recesses of the lateral ventricles open in their centres (Fig. 3A and B).
The anterior and posterior poles of the hemispheres are compressed
and therefore the antero-posterior dimension of the hemispheres is
shortened. As a result of the compression and bending of the posterior
pole of the hemisphere in shrews, the anterior part of their
hippocampus is placed relatively posterior, dorsally to the posterior
part. Therefore, when the brain is cut coronally, the anterior (dorsal)
and ventral parts of the hippocampus appear in the same sections
(Fig. 3C).
Seasonal changes in brain weight
We found no significant difference in the brain weight between
male and female shrews in any age group. For example, in the first
summer the mean brain weight of the common shrews was 212 mg
Fig. 1. Brain shape and weight in the investigated species of shrews.
(A) Common shrew (S. araneus); (B) pygmy shrew (S, minutus). Scale
(between the longer lines), 1 mm. (C) Seasonal changes in the brain weight
(the Dehnel’s effect) in the investigated common (s) and pygmy (d) shrews.
In both species differences between the Summer I (S-I) and Winter (W) groups
(reduction of brain weight) were highly statistically significant
(***P < 0.0002). Differences between the W and Summer II (S-II) groups
(recovery of brain weight) were also significant (*P < 0.02). A-I, Autumn I; S,
Spring; A-II, Autumn II.
in both males and females (n ¼ 8 in each group), whereas in the
winter groups of the same size the means were 194 mg for males
and 189 mg for females (P £ 0.46). In other groups of both species
the significance of differences in brain weight varied from P £ 0.19
to P £ 0.46, depending on the group. Therefore, data for males and
females were merged. Shrews that were investigated in our
experiments showed seasonal differences in brain weight
(Fig. 1C). In the youngest common shrews the average brain
weight was 212 ± 17 mg. In the wintering shrews of this species
the total brain weight was 10% lower (P < 0.0003) and 79% of the
loss was regained by the second summer (P < 0.02). Brains of the
subadult pygmy shrews weighed 122 ± 11 mg. In the winter their
weight was 17% lower (P < 0.0001) and 45% of the loss was
recovered by the second summer (P < 0.02). There was no
statistical difference between the brain weight of the two age
groups of common shrews living in the summer (Summer I and
Summer II), whereas the brain weight of adult pygmy shrews
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
Cell generation and death in the brain of shrews 1713
Fig. 2. TUNEL-labelled cell nuclei in two brain structures of the common and pygmy shrews. (A) Example of labelled nuclei in the OBs of a young common
shrew. (B) TUNEL-labelled nucleus in the DG of a young common shrew. (C) TUNEL-labelled nucleus in the OB of a common shrew photographed with the
confocal microscope. Scale bars: A and B, 50 lm; C, 5 lm. (D) Numbers (mean ± SD) of TUNEL-labelled nuclei in SVZ ⁄ OB of the two species in various
seasons. (E) Numbers (mean ± SD) of TUNEL-labelled nuclei in the DG of the two species. Season symbols as in Fig. 1.
(Summer II) was significantly lower (10%, P < 0.04) than that of
subadults (Summer I). Therefore, our data confirm the presence of
the Dehnel’s effect in the investigated shrews.
Cell death numbers
The TUNEL labelling showed that the rate of cell death in all brain
structures including DG, SVZ and OB was very low throughout life.
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
1714 K. Bartkowska et al.
Fig. 3. BrdU-labelled cell nuclei in brain sections of young pygmy shrews (Summer I) that were injected with BrdU 2 days (A) and 14 days (B–E) before
perfusion. Micrographs were taken from 40-lm sections that were not counterstained. (A) Horizontal section across the telencephalon. The majority of the new cells
were generated in the SVZ of the lateral walls of lateral ventricles and migrated along the rostral migratory stream (RMS) into the OBs. (B and D) Photomicrographs
of the OB showing the BrdU-labelled cell nuclei. Arrow in B points to the rostral extension of the lateral ventricle penetrating OB. In the OB a large majority of the
labelled nuclei were found in the granule cell layer and a minority in the periglomerular layer. (C) Coronal section across the hemisphere showing the anterior
(above) and posterior (below) parts of the DG. A fragment of the anterior part is shown in E at a higher magnification. Almost all BrdU-labelled nuclei are seen in the
granule cell layer of the DG. Scale bars: A–C, 500 lm; E, 10 lm, refers to D and E.
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
Cell generation and death in the brain of shrews 1715
In both species we found TUNEL-labelled nuclei in only four brain
structures: SVZ ⁄ OB (Fig. 2A and D), DG (Fig. 2B and E), cerebellum
and neocortex. In the common shrews the highest numbers of
TUNEL-positive nuclei were always observed in the SVZ ⁄ OB,
peaking in both autumn groups (on average about 3000 and 2000
TUNEL-labelled nuclei ⁄ brain in the Autumn I and Autumn II groups,
respectively). In the pygmy shrews (Fig. 2D) about twice the number
of TUNEL-labelled nuclei were found in the SVZ ⁄ OB during the first
autumn (average about 5700) compared with the Autumn II group
(average about 2200). In the Winter and Spring groups of both species
the number of nuclei was much lower in the SVZ ⁄ OB (Winter, on
average 60 labelled nuclei in the common shrews and 250 in the
pygmy shrews, Fig. 2D).
In the DG, the highest numbers of TUNEL-labelled nuclei were
observed in the ageing Autumn II group of common shrews (average
65, Fig. 2E) and in the Summer I group of pygmy shrews (average 80,
Fig. 2E). In the neocortex and cerebellum of both species we found
very low numbers of TUNEL-labelled nuclei. For example, in the
whole neocortex of the common shrews there were on average only
10 ± 14 TUNEL-labelled nuclei in the Summer I group and later the
TUNEL-labelled cells were found only sporadically, with no labelled
cells found in the Summer II group. In other brain structures of both
species, particularly in areas CA1–CA3 of the hippocampus, single
TUNEL-labelled nuclei were found only sporadically.
Rate of neurogenesis and phenotypes of newly generated cells
Except for very young shrews, in all other age groups BrdU-labelled
nuclei were visible exclusively in the SVZ ⁄ OB and DG (Fig. 3). In
very young shrews BrdU-labelled nuclei were also present in the white
matter and, although scarce, were observed in the cortex and other
brain nuclei. Double labelling showed that these BrdU-labelled cells
expressed CNPase and were therefore oligodendrocytes (Fig. 4E
and F). In this group BrdU-labelled nuclei outside the proliferation
zones were not counted. In older shrews (starting from Autumn I)
there were no cell divisions outside the two proliferation zones
(SVZ ⁄ OB and DG) and it is of particular note that there was no
proliferation in those cortical structures of the brain that are markedly
increasing their weight during the spring (Yaskin, 1994).
The rate of proliferation in the DG of the youngest shrews was very
high. In the subadult common shrews captured in June–August (group
Summer I), at 2 days after BrdU injections there were on average over
8000 labelled nuclei in both DGs (Fig. 5A). At 14 days after BrdU
injection (Fig. 5A) the number of labelled nuclei was 38% higher,
showing that some of the newly generated cells had divided again
during that period, while the rate of their apoptosis was not yet
significant. At that time, 78% of the BrdU-positive nuclei (74 out of
95) that were recruited into the DG were immunoreactive for NeuN
and were therefore classified as neurones (Fig. 4A). Only 3% of the
BrdU-positive cells expressed the astroglial marker (glial fibrillary
acidic protein) and 4% were oligodendrocytes expressing CNPase
(Fig. 4C). By the autumn, just 4 months later, the rate of proliferation
in the DG decreased by more than 73% and fell further to reach a level
of approximately 200 labelled nuclei ⁄ brain after 2 days’ survival
resulting in 400 cells recruited after 14 days in the Winter group
(Fig. 5A). Starting from spring, proliferation in the DG of both species
ceased altogether. Out of the 12 investigated common shrews captured
in the spring (two different subgroups) we found BrdU-labelled nuclei
in the DG of only one male (110 labelled nuclei ⁄ brain) and one female
(70 labelled nuclei). In older age groups of the common shrews
(Summer II and Autumn II, n ¼ 12 each) we found no labelled nuclei
in the DG (Fig. 5A).
In the young pygmy shrews the numbers of labelled nuclei in the
DG were even higher than in the common shrews (in the range of
10 000 after 2 days) and the age-dependent changes of rate of DG
neurogenesis were the same (Figs 3C and E, and 5B).
New cells generated in the SVZ of the lateral ventricles migrated to
the OB for more than 1 week. At 2 days after injections, BrdUlabelled nuclei were found in the SVZ covering a large part of the
lateral ventricles, extending from the ventricle recess in the centre of
the OB to their posterior parts (Fig. 3A). At 14 days after the
injections the majority of BrdU-labelled nuclei were found in the
granular and periglomerular layers of the OB (Fig. 3B and D), whereas
only a small proportion of these cells were still present in the SVZ. In
the OB, 83% of the BrdU-labelled nuclei (79 out of 95) expressed
NeuN (Fig. 4B), whereas only 2% expressed glial fibrillary acidic
protein (Fig. 4D) and one cell expressed CNPase.
The number of cells generated in the SVZ also changed with
seasons and age but the pattern of neurogenesis and recruitment was
different from that found in the DG. In the young common shrews
(Summer I group) numbers of the BrdU-labelled nuclei in SVZ ⁄ OB
were similar after both survival times (on average about 20 000 ⁄ brain)
(Fig. 6A). In the autumn, the number of BrdU-labelled nuclei
decreased by 65% and stayed at that low level throughout winter and
spring. The lowest number of labelled nuclei (average about
8000 ⁄ brain) was observed in the Winter group of the common shrews
at 2 days after the injections of BrdU. By the second summer the rate
of neurogenesis in the SVZ ⁄ OB of these shrews increased to match
that in the Summer I group. In the young pygmy shrews the rate of
proliferation in the SVZ was similar (15 000–25 000 ⁄ brain) and
similarly decreased in the winter (Fig. 6B). However, there was no
rebound of neurogenesis after wintering and therefore the rate of
proliferation in the SVZ of this species decreased with age although it
continued at a substantial level (3000–4000 ⁄ brain) even into senility.
Experimental procedures changing the rate of neurogenesis
Some of the external factors (agonists of the serotonergic receptors
5-HT1A, exercise) known to increase proliferation in the brain were
tested on separate groups of wintering common shrews, with a waning
proliferation in the DG. Two groups of these shrews were injected
with agonists of the 5-HT1A receptors, either 8-OH-DPAT or
buspirone, before the injection of BrdU (see Materials and methods)
and were investigated 14 days later (Fig. 7). These injections did not
significantly change the numbers of BrdU-labelled nuclei recruited
into the SVZ ⁄ OB, as compared with the untreated control group.
However, both drugs significantly increased the numbers of cells
recruited into DG. The most significant increase of their numbers
(over fivefold) was observed in the 8-OH-DPAT-treated animals
(P < 0.01), whereas the buspirone-treated animals had over twice the
number of BrdU-labelled cell nuclei in the DG compared with the
controls (P < 0.05). Another group of wintering common shrews
received physical training on a treadmill (Fig. 7). In this group
recruitment of new cells into DG was significantly lower than in
the controls (11 ± 11 vs. 402 ± 244, nuclei, P < 0.02). Injections of
8-OH-DPAT were also performed in the group of four common shrews
from the Summer II group and these were compared with four control
adult common shrews captured at the same time. After 2 days of
survival we found on average over 70 BrdU-labelled nuclei in both
DGs of the 8-OH-DPAT-injected shrews, whereas in the control
shrews there were none (P < 0.005). This shows that precursors in the
DG of the adult shrews are still present but dormant. Again, the rate of
neurogenesis in the SVZ ⁄ OB was not changed by the 8-OH-DPAT
injection.
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
1716 K. Bartkowska et al.
Fig. 4. Confocal images of double-labelled cells in the DG, OB and white matter of young common shrews. Double-labelled BrdU- (green) and NeuN- (red)
positive cell nuclei in DG (A) and OB (B). (C and D) Double labelling for BrdU (red) and glial fibrillary acidic protein (GFAP) (green) in the DG and OB,
respectively. Note that the majority of BrdU-positive cell nuclei did not colocalize with the GFAP-expressing cells. (E) In the white matter (corpus callosum) BrdUpositive cell nuclei (green) belonged to cells expressing CNPase (red), whereas in the OB (F) these two markers did not colocalize.
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
Cell generation and death in the brain of shrews 1717
Fig. 7. Influence of injections of the 5-HT1A receptor agonists [8-OH-DPAT
(DPAT), buspirone] or of physical exercise on the numbers of BrdU-labelled
nuclei in the DGs of the Winter groups of common shrews killed 14 days after
the injections. Numbers were calculated as in Fig. 5. Serotonergic drugs
significantly increased, whereas enforced running significantly decreased the
numbers of BrdU-labelled nuclei in the DG. None of these factors significantly
changed the numbers of labelled nuclei in the SVZ ⁄ OB (see text for details).
*P < 0.05; ***P < 0.01.
Fig. 5. Average numbers (mean ± SD) of BrdU-labelled nuclei in the DGs of
the common [S. araneus (SA)] and pygmy [S. minutus (SM)] shrews in the
consecutive seasons of their life cycle. Numbers were calculated by bilaterally
counting all labelled nuclei on every tenth 40-lm section through the DG and
multiplying the sum by 10. (A) Numbers of BrdU-labelled nuclei in the DG of
the common shrews. (B) Numbers of BrdU-positive nuclei in the DG of the
pygmy shrews. Groups consisted of animals that survived either 2 or 14 days
after BrdU injections. Season symbols as in Fig. 1.
Fig. 6. Average numbers (mean ± SD) of BrdU-labelled nuclei in the
SVZ ⁄ OB of common and pygmy shrews. (A) Common shrews [S. araneus
(SA)]; (B) pygmy shrews [S. minutus (SM)]. Data were collected from the
same animals shown in Fig. 5. Note that the numbers of cells generated in the
SVZ are two times higher than in the DG and that proliferation there continues
at a relatively high rate throughout life. Season symbols as in Fig. 1.
Discussion
Seasonal changes of brain weight in Sorex shrews
Our results showed that changes in the number of brain cells of any
particular phenotype do not significantly contribute to the seasonal
changes of brain weight that are a part of the Dehnel’s effect and that
were confirmed in our population of the Sorex shrews. The amount of
cell death in the brains of the investigated shrews was too low to
contribute significantly to the autumnal reduction of the brain weight
and no seasonal wave of cell death was observed. In particular, the
amount of cell death did not increase in the neocortex and
hippocampus although these structures are significantly reducing their
weight in the autumn (Yaskin, 1994). The majority of the TUNELlabelled nuclei were found in those structures of the brain that generate
and recruit new cells (SVZ, OB and DG), and in these structures the
dynamics of cell death roughly paralleled the dynamics of cell
generation. We estimated that the percentage of cell loss in the autumn
could have been in the range of 1% in the cerebral cortex and less than
1% in other structures. Therefore, the observed amount of cell death
could not reduce the number of brain cells to the degree that would
measurably change its weight.
Similarly, as the BrdU labelling showed, the spring increase in the
weight of the shrew brain did not depend on the generation of new
cells. At that time BrdU-positive cells were absent in all brain
structures outside the SVZ ⁄ OB. In particular, there were no newly
generated cells in the neocortex and hippocampus, areas that increase
their weight the most in the spring (Yaskin, 1994). As a consequence,
in spite of the unusual seasonal oscillations of their brain weight, the
pool of brain cells in the Sorex shrews (of all phenotypes) remained
stable throughout life, from weaning until senility. Cellular populations in the brains of other mammals also seem to be resistant to an
age-related decline (for review cf. Turlejski & Djavadian, 2002).
Little is known about the external factors and mechanisms, such as
hormonal stimuli, that trigger seasonal changes of the body and brain
weight in shrews. However, various facts, like the active process of
osteoclasis of the skull and vertebrae (Pucek, 1957) and differential
rate of reduction of the internal organs (Pucek, 1965a) and brain
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
1718 K. Bartkowska et al.
structures (Yaskin, 1994) in the autumn, speak for an active and
precisely regulated process. The selective reduction and increase of the
mass of brain parts may be achieved via several complementary
mechanisms, e.g. changes in the volume of neuropil, cell bodies
and ⁄ or extracellular space. One of them, i.e. changes of water content,
may be excluded, at least as the main mechanism of the oscillations.
The water content of the shrew brain decreases by only 3–4% from
their first summer to winter, thus excluding seasonal changes in tissue
hydration as a main factor in the Dehnel’s effect, especially in those
populations of shrews and brain structures for which the weight
reduction during the winter exceeds 25–30% (Pucek, 1965b; Myrcha,
1969; Yaskin, 1994). The absolute weight of the brain lipids increases
between summer and winter in the subadult shrews in spite of the
reduction of brain volume (Pucek, 1965b; Myrcha, 1969), which
probably depends on the progressing myelinization. Therefore, the
slight reduction of the percentage of water content with the parallel
increase of the content of lipids in the shrew brain may be a passive
reflection of the processes of brain maturation.
The volume of extracellular space is in the range of 20–25% in the
mouse and may fall to 5–9% in pathological conditions (Mazel et al.,
2002; Sykova, 2004). Therefore, it is possible that in shrews the
extracellular space of the brain may be somewhat reduced in the
winter by as yet unknown factors, contributing to the reduction of
brain weight. However, the most probable mechanism is the
oscillation of the volume of neuropil and ⁄ or cell bodies. The dendritic
spines are well known to be dynamically changing structures (Alvarez &
Sabatini, 2007) and the dendritic and axonal trees may also be
remodelled (Cesa & Strata, 2005).
Absence of neurogenesis in the DG of adult Sorex shrews
and its consequences
An unexpected finding of our experiments is that adult Sorex shrews
do not generate new neurones in their DG throughout the second part
of their life (from spring to autumn). Already by the late autumn and
winter the rate of cell division in their DG is very low and such low
rates of progenitor divisions tend to generate new progenitors rather
than differentiating cells (Seaberg & van der Kooy, 2002). The
presence of proliferation in the SVZ ⁄ OB of all shrews is an internal
control indicating that our negative findings in the DG are not due to
inappropriate methodology. Therefore, we conclude that Sorex shrews
terminate neurogenesis and then all cell divisions in the DG while
reaching adulthood and not old age. Few neurones, if any, are added to
their DG after the end of the first autumn of their life until their death
almost 1 year later.
The lack of DG neurogenesis in the adult individuals makes Sorex
shrews an exception among the investigated mammalian species but
not the only one. Recently it was shown that neurogenesis was absent
in the DG of nine out of 12 African nectar- and fruit-eating bat species
(Amrein et al., 2007). European bats of closely related species have
variable rates of neurogenesis, in some species being close to nil
(Djavadian et al., 2006). Bats are extremely long-lived species (up to
30 years; Brunet-Rossinni & Austad, 2004) and it is probable that in
other long-living mammals the decrease of the DG proliferation in
adulthood and old age is profound. In contrast, shrews are very shortlived mammals. In addition, even in the closely related shrews of the
genus Neomys and also shrews of the genus Crocidura, proliferation in
the DG is present in the adults (our unpublished results).
The biological advantages of that elimination of proliferation in the
DG of the Sorex shrew, whereas robust proliferation in their SVZ
continues throughout life, are not clear. One possibility is that this
difference may be a side-effect of the mechanisms of the unique brain
volume oscillations in these species. However, in that case it would
have to be at least neutral as a selection factor in evolution.
An overwhelming majority of newly generated cells in the DG
differentiate into neurones and replace older neurones of that structure
(Cameron & McKay, 2001; Van Praag et al., 2002). This is also true
for young Sorex shrews. It was first postulated for birds, and then
mammals, that neuronal replacement is the mechanism for creating or
remodelling a function, like seasonal bird singing or spatial memory
(Goldman & Nottebohm, 1983; Kirn & Nottebohm, 1993; Barnea &
Nottebohm, 1994; Shors et al., 2001; Kempermann, 2002).
Lack of neurogenesis in the DG of adult Sorex shrews and probably
also in some species of bats poses problems for the general validity of the
hypothesis of ‘spatial memory change through turnover of DG granule
cells’ in relation to mammals. In the case of bats, the general topography
of their environment may be rather stable in the macroscale but seasonal
changes in the presence of food resources may be dramatic, requiring
frequent relearning of the placement of resources that is vital for
survival. This is equally true for shrews. They experience rapid changes
between winter and summer environments, and their food demands,
especially of pregnant and lactating females, are exceptionally high
(Genoud & Vogel, 1990). Because of an extremely high metabolic rate
(Ochocinska & Taylor, 2005; for review see Taylor, 1998) these shrews
may die of starvation after only 4–5 h of fasting and so must respond to
and overcome changing food resources very effectively.
The case of the Sorex shrews excludes one explanation that may
assist bats, i.e. their communal life with its opportunities for mutual
observation and learning. Sorex shrews are solitary animals that are
territorial for most of their lives and exploit extensive territories
(S. minutus, 1100–1800 m2; S. araneus, 500–600 m2, for review see
Rychlik, 1998), chasing out all conspecifics and therefore they must
depend only on their own memory.
The substantial reduction of the rate of DG neurogenesis and then
its cessation in the middle of the Sorex shrew’s life or earlier, rather
than in old age, raises one more problem for explanation. At that age
shrews are at the peak of their performance and not behaviourally
impaired due to senility. The timing of that decrease is similar to what
was found in rats and marmosets (Rao et al., 2006; Leuner et al.,
2007). Reduction of the DG neurogenesis was also postulated for
middle-aged and older humans (Eriksson et al., 1998; Fahrner et al.,
2007). In macaques proliferation in the DG decreases but is preserved
until old age (Gould et al., 1999).
A recent finding throws new light on these phenomena. It was found
that the DG neurones generated during development still dominate the
DG of the adult mice (Muramatsu et al., 2007). The authors’
conclusion is that the newly generated neurones are short-lived and
undergo a quick turnover, whereas the pool of neurones generated
during development is rather stable. The transient character of the new
DG neurones in macaques (Gould et al., 2001) leads to a similar
conclusion. If that transient character of newly generated neurones is
confirmed in other species, then this would mean that the pool of new
neurones generated in the DG of adult mammals (unlike the pool
generated during development) undergoes a quick turnover and that
the reduction of neurogenesis starting in mid-life influences only (or
mainly) that transient pool. Therefore, Sorex shrews and some bat
species would just be extreme cases of the reduction of turnover of the
transient pool of DG neurones, whereas a less radical reduction
starting in the middle of the life cycle would be a common feature.
The general validity of the hypothesis of the neurogenesisdependent hippocampal memory was also questioned by some
authors who concluded that the hippocampal neurogenesis may be
involved in some, but not all, hippocampus-dependent memory tasks
(Shors et al., 2002; Saxe et al., 2007). Meshi et al. (2006) reported
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
Cell generation and death in the brain of shrews 1719
that the hippocampal neurogenesis did not mediate the effects of
environmental enrichment on spatial learning. Thus, it seems that
wild mammals such as shrews or bats, as well as laboratory rodents
(Merrill et al., 2003), in reaction to environmental changes are able
to modify at least some important aspects of their spatial memory
without adding or removing new cells from their DG. It is probable
that they do it by modifying the synaptic input to and output from
the DG without exchanging granule cells (Ramı́rez-Amaya et al.,
1999; Frank et al., 2006; Rekart et al., 2007) or by some
hippocampus-independent spatial memory systems, like path
integration (Alyan & McNaughton, 1999).
The possibility of the prevalence of the extrahippocampal mechanisms of spatial memory learning in adult shrews raises another
question. The hippocampus of shrews (and of other small mammals) is
a proportionally very large structure and if, in these animals, spatial
maps are created and modified by extrahippocampal mechanisms, then
other types of important hippocampus-dependent aspects of behaviour
should be postulated for this structure. This refers to many small
mammals that have large hippocampi, especially if they have poor
vision or live a predominantly nocturnal or fossorial life (e.g. mole
rats, cf. Nemec et al., 2008), as they have to use mainly non-visual
cues in learning of their space maps. Participation of the hippocampus
in the limbic circuitry and its extensive involvement in the emotional
behaviour is a possible explanation.
Factors influencing adult neurogenesis in shrews
Seasonal oscillations in the rate of neurogenesis in mammals were
rarely investigated and have only been confirmed in the DG of two
species of rodents. In the golden hamster (Mesocricetus auratus) the
highest rate of proliferation was seen in the autumn, at the beginning
of the season of vegetation growth in the Near East (Huang et al.,
1998). In the meadow vole (Microtus pennsylvanicus) the rate of
proliferation in the DG decreased in females when they started to
breed, whereas it did not change seasonally in males. However, in the
hippocampal hilus of breeding males the rate of proliferation increased
(Galea & McEwen, 1999).
These seasonal oscillations of the rate of proliferation in the DG,
although induced by changes in day length, may depend on the
hormonal status and may differ in males and females. Such sex
differences in the rate of proliferation in either DG or SVZ were not
observed in the Sorex shrews in any season. For half of their life (until
spring) these shrews are sexually immature, so their hormonal status
cannot influence neurogenesis in any structure. Then, from the time
that they become sexually mature, proliferation in the DG ceases
completely in both sexes. In the second part of the shrew’s life,
proliferation in the SVZ is high but again we found no sex differences.
When animals reach sexual maturity, the levels of oestrogens and
androgens in their bodies rise dramatically. Oestrogens were found to
stimulate proliferation in the DG of some rodents (Tanapat et al.,
1999; Galea, 2007) but in other rodent species the rate of DG
neurogenesis decreased in breeding females (Galea & McEwen, 1999;
Ormerod & Galea, 2001; Perfilieva et al., 2001). In addition, in the
rats and meadow voles the rate of proliferation and ⁄ or the rate of new
cell survival in the DG were higher in males (Galea & McEwen, 1999;
Perfilieva et al., 2001; Galea, 2007). An important pool of oestrogens
is synthesized locally in the DG of both males and females, where the
hormones act locally, influencing the rate of neurogenesis (Hajszan
et al., 2007). These local oestrogens are necessary for the hippocampal
granule cell proliferation, apoptosis and synaptic plasticity (Banasr
et al., 2001; Kretz et al., 2004; Fester et al., 2006). Therefore, the
regulatory influence of sex steroids on the DG neurogenesis is a
complex process that is, in large part, independent of hormones
produced by the gonads and the effects of changes of sex hormone
levels in the blood may vary from species to species.
Similar to other mammalian species (Santarelli et al., 2003; Banasr
et al., 2004), in shrews the rate of division of the DG precursor cells
increased after injections of 5-HT1A agonists. Moreover, the 5-HT1A
agonists were able to induce proliferation in the DG of adult shrews
when physiologically it does not exist. This confirmed that a pool of
neural precursor cells was still present in the DG of the adult shrews,
although reduced and dormant.
In contrast to findings in adult mice (Van Praag et al., 1999), we
found that forced physical exercise reduced the rate of cell division in
the shrew DG. One possible explanation of our results is that the
forced exercise could have been more stressful for wild shrews
compared with laboratory mice and therefore the inhibitory influences
of stress on the DG proliferation (Gould et al., 1997; Dranovsky &
Hen, 2006) prevailed over the stimulating effects of physical exercise.
Consistent with this possibility, the extensive, prolonged training of
rats in a water maze increased levels of circulating corticosterone and
resulted in a reduction of the number of newborn hippocampal
neurones (Meshi et al., 2006; Mohapel et al., 2006).
As in other mammals, neuronal precursor cells proliferating in the
SVZ of the Sorex shrews migrate to the OB. It was shown in other
species that these cells are becoming GABAergic and dopaminergic
inhibitory interneurones of the OB (Saino-Saito et al., 2004; Hack
et al., 2005). These interneurones help to cross-relate information
transferred from the more than 1000 types of relatively unspecific
olfactory receptors that segregate their inputs to glomeruli of the OB.
When neurogenesis in the SVZ is reduced, the detection threshold for
single odours and short-term olfactory memory is normal. However,
discrimination between odours in a mixture is impaired (Gheusi et al.,
2000). The SVZ precursor cells seem to have a very different
physiology; they proliferate throughout life and, after the winter
depression, they are just as active in very old shrews as in younger
animals. Life-long persistence of the SVZ neurogenesis also indicates
that the lack of neurogenesis in the DG was not caused by general
senility of these animals (Luo et al., 2006).
The seasonal modulation of the rate of neurogenesis in the SVZ of
Sorex shrews is probably the first such finding in mammals. Reduction
of the SVZ neurogenesis in the winter may reflect a lower level of
olfactory stimulation and ⁄ or its less complex character in that period.
Similar reduction of the SVZ neurogenesis was observed in mice after
blocking one of their nostrils (Corotto et al., 1994; Mandairon et al.,
2006). This is an interesting example of regulation of the brain
neurogenesis by external stimuli. Its mechanisms are difficult to grasp,
as the precursor cells that must be influenced do not have any direct
contact with the olfactory system.
Proliferation in the SVZ was not significantly influenced by our
experimentally introduced factors, which did increase or reduce the
rate of proliferation seen in the DG. These differences are compatible
with findings concerning the different biology of the two types of
precursor cells in the brains of rodents (Seaberg & van der Kooy,
2002).
Conclusions
The changes in the number of brain cells do not significantly
contribute to the unusual seasonal changes of brain weight in Sorex
shrews. Sorex shrews are an exception (but not the only one) among
the investigated mammalian species showing lack of neurogenesis in
the DG of adults. However, substantial reduction of the DG
neurogenesis with age may be a more common phenomenon. Further
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
1720 K. Bartkowska et al.
studies are needed for better understanding of the survival advantage
of the seasonal brain changes in Sorex shrews, factors that block the
cell cycle of progenitor cells in the shrew DG and the interconnection
between these two phenomena, as well as the importance of the
presence or absence of neurogenesis for the behaviour of the adult
mammals.
Acknowledgements
This research was supported by the Polish Ministry of Science and Higher
Education grant no. 0496 ⁄ P04 ⁄ 2005 ⁄ 29 and by statutory grant of the Polish
Ministry of Science and Higher Education to the University of Bialystok. The
authors thank Dr Tom FitzGibbon and Dr Sara Churchfield for correcting the
English language and useful comments.
Abbreviations
BrdU, bromodeoxyuridine; CNPase, 2¢,3¢-cyclic nucleotide 3¢-phosphodiesterase; DG, dentate gyrus; 5-HT1A, serotonin receptor 1A; NeuN, neuronal
marker; OB, olfactory bulb; 8-OH-DPAT, 8-hydroxy-2-(di- n-propylamino)tetralin; SVZ, subventricular zone; TBS-B, Tris-buffered saline with 0.1%
Triton X-100 with 0.05% bovine serum albumin; TUNEL, terminal
deoxynucleotidyl transferase-mediated dUTP-biotin nick and labelling.
References
Altman, J. (1962) Are new neurons formed in the brains of adult mammals?
Science, 135, 1127–1128.
Alvarez, V.A. & Sabatini, B.L. (2007) Anatomical and physiological plasticity
of dendritic spines. Annu. Rev. Neurosci., 30, 79–97.
Alyan, S. & McNaughton, B.L. (1999) Hippocampectomized rats are capable
of homing by path integration. Behav. Neurosci., 113, 19–31.
Amrein, I., Dechmann, D.K.N., Winter, Y. & Lipp, H.P. (2007) Absent or low
rate of adult neurogenesis in the hippocampus of bats (Chiroptera). Plos One,
5, e455.
Banasr, M., Hery, M., Brezun, J.M. & Daszuta, A. (2001) Serotonin mediates
oestrogen stimulation of cell proliferation in the adult dentate gyrus. Eur. J.
Neurosci., 14, 1417–1424.
Banasr, M., Hery, M., Printemps, R. & Daszuta, A. (2004) Serotonininduced increases in adult cell proliferation and neurogenesis are mediated
through different and common 5-HT receptor subtypes in the dentate
gyrus and the subventricular zone. Neuropsychopharmacology, 29, 450–
460.
Barnea, A. & Nottebohm, F. (1994) Seasonal recruitment of hippocampal
neurons in adult free-ranging black-capped chickadees. Proc. Natl Acad. Sci.
U.S.A., 91, 11 217–11 221.
Brunet-Rossinni, A.K. & Austad, S.N. (2004) Ageing studies on bats: a review.
Biogerontology, 5, 211–222.
Cameron, H.A. & McKay, R.D.G. (2001) Adult neurogenesis produces a large
pool of new granule cells in the dentate gyrus. J. Comp. Neurol., 435, 406–417.
Cesa, R. & Strata, P. (2005) Axonal and synaptic remodeling in the mature
cerebellar cortex. Prog. Brain Res., 148, 45–56.
Churchfield, S. (1990) The Natural History of Shrews. Cornell University
Press, Ithaca, New York.
Corotto, F.S., Henegar, J.R. & Maruniak, J.A. (1994) Odor deprivation leads to
reduced neurogenesis and reduced neuronal survival in the olfactory bulb of
the adult mouse. Neuroscience, 61, 739–744.
Dehnel, A. (1949) Studies on the genus Sorex L. Ann. Univ. M. Curie-Sklod., 4,
18–102.
Djavadian, R.L., Ghazaryan, A., Yavrouyan, E.G. & Turlejski, K. (2006)
Generation of brain cells in adult microbats. FENS 5th Forum of European
Neuroscience, Vienna, 2006. Abstract 3A, 086.8.
Dranovsky, A. & Hen, R. (2006) Hippocampal neurogenesis: regulation by
stress and antidepressants. Biol. Psychiat., 59, 1136–1143.
Eriksson, P.S., Perfilieva, E., Bjork-Eriksson, T., Alborn, A.M., Nordborg, C.,
Peterson, D.A. & Gage, F.H. (1998) Neurogenesis in the adult human
hippocampus. Nat. Med., 4, 1313–1317.
Fahrner, A., Kann, G., Flubacher, A., Heinrich, C. & Freiman, T.M. (2007)
Granule cell dispersion is not accompanied by enhanced neurogenesis in
temporal lobe epilepsy patients. Exp. Neurol., 203, 320–332.
Fester, L., Ribeiro-Gouveia, V., Prange-Kiel, J., von Schassen, C., Bottner, M.,
Jarry, H. & Rune, G.M. (2006) Proliferation and apoptosis of hippocampal
granule cells require local oestrogen synthesis. J. Neurochem., 97, 1136–
1144.
Frank, L.M., Brown, E.N. & Stanley, G.B. (2006) Hippocampal and cortical
place cell plasticity: implications for episodic memory. Hippocampus, 16,
775–784.
Gage, F.H. (2000) Mammalian neural stem cells. Science, 287, 1433–1438.
Galea, L.A. (2007) Gonadal hormone modulation of neurogenesis in the
dentate gyrus of adult male and female rodents. Brain Res. Rev., epub ahead
of print.
Galea, L.A. & McEwen, B.S. (1999) Sex and seasonal differences in the rate of
cell proliferation in the dentate gyrus of adult wild meadow voles.
Neuroscience, 89, 955–964.
Genoud, M. & Vogel, P. (1990) Energy requirements during reproduction and
reproductive effort in shrews (Soricidae). J. Zool. Lond., 220, 41–60.
Gheusi, G., Cremer, H., McLean, H., Chazal, G., Vincent, J.D. & Lledo, P.M.
(2000) Importance of newly generated neurons in the adult olfactory bulb for
odor discrimination. Proc. Natl Acad. Sci. U.S.A., 97, 1823–1828.
Goldman, S.A. & Nottebohm, F. (1983) Neuronal production, migration, and
differentiation in a vocal control nucleus of the adult female canary brain.
Proc. Natl Acad. Sci. U.S.A., 80, 2390–2394.
Gould, E. (2007) How widespread is adult neurogenesis in mammals?
Neurosci. Nat. Rev., 8, 481–488.
Gould, E., McEwen, B.S., Tanapat, P., Galea, L.A. & Fuchs, E. (1997)
Neurogenesis in the dentate gyrus of the adult tree shrew is regulated by
psychosocial stress and NMDA receptor activation. J. Neurosci., 17, 2492–
2498.
Gould, E., Tanapat, P., McEwen, B.S., Flugge, G. & Fuchs, E. (1998)
Proliferation of granule cell precursors in the dentate gyrus of adult
monkeys is diminished by stress. Proc. Natl Acad. Sci. U.S.A., 95, 3168–
3171.
Gould, E., Reeves, A.J., Graziano, M.S. & Gross, C.G. (1999) Neurogenesis in
the neocortex of adult primates. Science, 286, 548–552.
Gould, E., Vail, N., Wagers, M. & Gross, C.G. (2001) Adult-generated
hippocampal and neocortical neurons in macaques have a transient existence.
Proc. Natl Acad. Sci. U.S.A., 98, 10 910–10 917.
Grote, H.E. & Hannan, A.J. (2007) Regulators of adult neurogenesis in the
healthy and diseased brain. Clin. Exp. Pharmacol. Physiol., 34, 533–545.
Hack, M.A., Saghatelyan, A., de Chevigny, A., Pfeifer, A., Ashery-Padan, R.,
Lledo, P.M. & Götz, M. (2005) Neuronal fate determinants of adult olfactory
bulb neurogenesis. Nat. Neurosci., 8, 865–872.
Hajszan, T., Milner, T.A. & Leranth, C. (2007) Sex steroids and the dentate
gyrus. Prog. Brain Res., 163, 399–415.
Huang, L., DeVries, G.J. & Bittman, E.L. (1998) Photoperiod regulates
neuronal bromodeoxyuridine labeling in the brain of seasonally breeding
mammal. J. Neurobiol., 36, 410–420.
Hyvarinen, H. (1969) On the seasonal changes in the skeleton of the common
shrew (Sorex araneus L.) and their physiological background. Aquilo Series
Zool., 7, 1–32.
Kempermann, G. (2002) Why new neurons? Possible functions for adult
hippocampal neurogenesis. J. Neurosci., 22, 635–638.
Kempermann, G., Kuhn, H.G. & Gage, F.H. (1997) Genetic influence on
neurogenesis in the dentate gyrus of adult mice. Proc. Natl Acad. Sci. U.S.A.,
94, 10 409–10 414.
Kirn, J.R. & Nottebohm, F. (1993) Direct evidence for loss and replacement of
projection neurons in adult canary brain. J. Neurosci., 13, 1654–1663.
Kornack, D.R. & Rakic, P. (1999) Continuation of neurogenesis in the
hippocampus of the adult macaque monkey. Proc. Natl Acad. Sci. U.S.A., 96,
5768–5773.
Kretz, O., Fester, L., Wehrenberg, U., Zhou, L., Brauckmann, S., Zhao, S.,
Prange-Kiel, J., Naumann, T., Jarry, H., Frotscher, M. & Rune, G.M. (2004)
Hippocampal synapses depend on hippocampal estrogen synthesis.
J. Neurosci., 24, 5913–5921.
Leuner, B., Kozorovitskiy, Y., Gross, C.G. & Gould, E. (2007) Diminished
adult neurogenesis in the marmoset brain precedes old age. Proc. Natl Acad.
Sci. U.S.A., 104, 17 169–17 173.
Luo, J., Daniels, S.B., Lennington, J.B., Notti, R.Q. & Conover, J.C. (2006)
The aging neurogenic subventricular zone. Aging Cell, 5, 139–152.
Mandairon, N., Sacquet, J., Jourdan, F. & Didier, A. (2006) Long-term fate and
distribution of newborn cells in the adult mouse olfactory bulb: Influences of
olfactory deprivation. Neuroscience, 141, 443–451.
Mazel, T., Richter, F., Vargova, L. & Sykova, E. (2002) Changes in external
space volume and geometry induced by cortical spreading depression in
immature and adult rats. Physiol. Res., 51, S85–S93.
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721
Cell generation and death in the brain of shrews 1721
Merrill, D.A., Karim, R., Darraq, M., Chiba, A.A. & Tuszynski, H.T. (2003)
Hippocampal cell genesis does not correlate with spatial learning ability in
aged rats. J. Comp. Neurol., 459, 201–207.
Meshi, D., Drew, M.R., Saxe, M., Ansorge, M.S., David, D., Santarelli, L.,
Malapani, C., Moore, H. & Hen, R. (2006) Hippocampal neurogenesis is not
required for behavioral effects of environmental enrichment. Nat. Neurosci.,
9, 729–731.
Mohapel, P., Mundt-Petersen, K., Brundin, P. & Frielingsdorf, H. (2006)
Working memory training decreases hippocampal neurogenesis. Neuroscience, 142, 609–613.
Muramatsu, R., Ikegaya, Y., Matsuki, N. & Koyama, R. (2007) Neonatally born
granule cells numerically dominate adult mice dentate gyrus. Neuroscience,
148, 593–598.
Myrcha, A. (1969) Seasonal changes in caloric value, body water and fat in
some shrews. Acta Theriol., 14, 211–227.
Nemec, P., Cvekova, P., Benada, O., Wielkopolska, E., Olkowicz, S.,
Turlejski, K., Burda, H., Bennett, N.C. & Peichl, L. (2008) The visual
system in subterranean African mole-rats (Rodentia, Bathyergidae): Retina,
subcortical visual nuclei and primary visual cortex. Brain Res. Bull., 75,
356–364.
Nottebohm, F. (1981) A brain for all seasons: cyclical anatomical changes in
song control nuclei of the canary brain. Science, 214, 1368–1370.
Nowakowski, R.S., Lewin, S.B. & Miller, M.W. (1989) Bromodeoxyuridine
immunochistochemical determination of the length of the cell cycle and the
DNA-synthesis phase for an anatomically defined population. J. Neurocytol.,
18, 311–318.
Ochocinska, D. & Taylor, J.R.E. (2005) Living at the physiological limits: field
and maximum metabolic rates of the common shrew (Sorex araneus).
Physiol. Biochem. Zool., 78, 808–818.
Ormerod, B.K. & Galea, L.A. (2001) Reproductive status influences
cell proliferation and cell survival in the dentate gyrus of adult female
meadow voles: a possible regulatory role of estradiol. Neuroscience, 102,
369–379.
Otsuki, Y., Li, Z. & Shibata, M.A. (2003) Apoptotic detection methods — from
morphology to gene. Prog. Histochem. Cytochem., 38, 275–339.
Perfilieva, E., Risedal, A., Nyberg, J., Johansson, B.B. & Eriksson, P.S. (2001)
Gender and strain influence on neurogenesis in dentate gyrus of young rats.
J. Cereb. Blood Flow Metab., 21, 211–217.
Pucek, Z. (1957) Histomorphologische Untersuchungen uber die Winterdepression des Schadels bei Sorex L. und Neomys Kaup. Ann. Univ. M.
Curie-Sklod. Sect. C, 10, 399–428.
Pucek, Z. (1965a) Seasonal and age changes in the weight of internal organs of
shrews. Acta Theriol., 10, 369–438.
Pucek, M. (1965b) Water content and seasonal changes of the brain-weight in
shrews. Acta Theriol., 10, 353–367.
Pucek, Z. & Markov, G. (1964) Seasonal changes in the skull of the common
shrew from Bulgaria. Acta Theriol., 9, 363–366.
Ramı́rez-Amaya, V., Escobar, M.L., Chao, V. & Bermúdez-Rattoni, F. (1999)
Synaptogenesis of mossy fibers induced by spatial water maze overtraining.
Hippocampus, 9, 631–636.
Rao, M.S., Hattiangady, B. & Shetty, A.K. (2006) The window and mechanisms
of major age-related decline in the production of new neurons within the
dentate gyrus of the hippocampus. Aging Cell, 5, 545–558.
Rekart, J.L., Sandoval, C.J., Bermudez-Rattoni, F. & Routtenberg, A. (2007)
Remodeling of hippocampal mossy fibers is selectively induced seven days
after the acquisition of a spatial but not a cued reference memory task. Learn.
Mem., 14, 416–421.
del Rio, J.A. & Soriano, E. (1989) Immunocytochemical detection of
5¢-bromodeoxyuridine incorporation in the central nervous system of the
mouse. Brain Res. Dev. Brain Res., 49, 311–317.
Rychlik, L. (1998) Evolution of social systems in shrews. In Wojcik, J.M. &
Wolsan, M. (Eds), Evolution of Shrews. Mammal Research Institute PAS,
Bialowieza, pp. 347–406.
Saino-Saito, S., Sasaki, H., Volpe, B.T., Kobayashi, K., Berlin, R. & Baker, H.
(2004) Differentiation of the dopaminergic phenotype in the olfactory system
of neonatal and adult mice. J. Comp. Neurol., 479, 389–398.
Santarelli, L., Saxe, M., Gross, C., Surget, A., Battaglia, F., Dulawa, S.,
Weisstaub, N., Lee, J., Duman, R., Arancio, O., Belzung, C. & Hen, R.
(2003) Requirement of hippocampal neurogenesis for the behavioral effects
of antidepressants. Science, 301, 805–809.
Saxe, M.D., Malleret, G., Vronskaya, S., Mendez, I., Garcia, A.D., Sofroniew,
M.V., Kandel, E.R. & Hen, R. (2007) Paradoxical influence of hippocampal
neurogenesis on working memory. Proc. Natl Acad. Sci. U.S.A., 104, 4642–
4646.
Seaberg, R.M. & van der Kooy, D. (2002) Adult rodent neurogenic regions: the
ventricular subependyma contains neural stem cells, but the dentate gyrus
contains restricted progenitors. J. Neurosci., 22, 1784–1793.
Sherry, D.F., Forbes, M.R.L., Khurgel, M. & Ivy, G. (1993) Females have a
larger hippocampus than males in the brood-parasitic brown-headed
cowbird. Proc. Natl Acad. Sci. U.S.A., 90, 7839–7843.
Shors, T.J., Miesegaes, G., Beylin, A., Zhao, M., Rydel, T. & Gould, E. (2001)
Neurogenesis in the adult is involved in the formation of trace memories.
Nature, 410, 372–375.
Shors, T.J., Townsend, D.A., Zhao, M., Kozorovitskiy, Y. & Gould, E. (2002)
Neurogenesis may relate to some, but not all types of hippocampaldependent learning. Hippocampus, 12, 578–584.
Siivonen, L. (1954) Uber die Grossenvariationen der Saugetiere und die Sorex
macropygmaeus Mill-Frage in Fennoskandien. Ann. Acad. Sci. Fenn., 4,
Series A, 1–24.
Sykova, E. (2004) Diffusion properties of the brain in health and disease.
Neurochem. Int., 45, 453–466.
Tanapat, P., Hastings, N.B., Reeves, A.J. & Gould, E. (1999) Estrogen
stimulates a transient increase in the number of new neurons in the dentate
gyrus of the adult female rat. J. Neurosci., 19, 5792–5801.
Taylor, J.R.E. (1998) Evolution of energetic strategies in shrews. In Wojcik,
J.M. & Wolsan, M. (Eds), Evolution of Shrews. Mammal Research Institute
PAS, Bialowieza, pp. 309–346.
Turlejski, K. & Djavadian, R.L. (2002) Life-long stability of neurons: a century
of research on neurogenesis, neuronal death and neuron quantification in
adult CNS. Prog. Brain Res., 36, 39–65.
Van Praag, H., Kempermann, G. & Gage, F.H. (1999) Running increases cell
proliferation and neurogenesis in the adult mouse dentate gyrus. Nat.
Neurosci., 2, 266–270.
Van Praag, H., Schinder, A.F., Christie, B.R., Toni, N., Palmer, T.D. & Gage,
F.H. (2002) Functional neurogenesis in the adult hippocampus. Nature, 415,
1030–1034.
Yaskin, V. (1994) Variation in brain morphology of the common shrew. In
Merritt, J.F., Kirkland, G.L. Jr & Rose, R.K. (Eds), Advances in the Biology
of Shrews. Special Publication. Carnegie Museum of Natural History 18,
Pittsburgh, pp. 155–161.
ª The Authors (2008). Journal Compilation ª Federation of European Neuroscience Societies and Blackwell Publishing Ltd
European Journal of Neuroscience, 27, 1710–1721