Transgenic songbirds offer an opportunity to develop a
genetic model for vocal learning
The MIT Faculty has made this article openly available. Please share
how this access benefits you. Your story matters.
Citation
Agate, R. J. et al. “Transgenic songbirds offer an opportunity to
develop a genetic model for vocal learning.” Proceedings of the
National Academy of Sciences 106.42 (2009): 17963 -17967.
©2010 by the National Academy of Sciences.
As Published
http://dx.doi.org/10.1073/pnas.0909139106
Publisher
National Academy of Sciences
Version
Final published version
Accessed
Thu May 26 12:47:17 EDT 2016
Citable Link
http://hdl.handle.net/1721.1/58553
Terms of Use
Article is made available in accordance with the publisher's policy
and may be subject to US copyright law. Please refer to the
publisher's site for terms of use.
Detailed Terms
Transgenic songbirds offer an opportunity to develop
a genetic model for vocal learning
R. J. Agatea,1, B. B. Scottb, B. Haripala, C. Loisb,2, and F. Nottebohma,1,2
aLaboratory
of Animal Behavior, The Rockefeller University, 1230 York Avenue, New York, NY 10021; and bPicower Institute for Learning and Memory,
Department of Brain and Cognitive Science, Massachusetts Institute of Technology, 77 Massachusetts Avenue, Cambridge, MA 02139
Zebra finches are widely used for studying the basic biology of
vocal learning. The inability to introduce genetic modifications in
these animals has substantially limited studies on the molecular
biology of this behavior, however. We used an HIV-based lentivirus
to produce germline transgenic zebra finches. The lentivirus encoded the GFP regulated by the human ubiquitin-C promoter [Lois
C, Hong EJ, Pease S, Brown EJ, Baltimore D (2002) Science 295:868 –
872], which is active in a wide variety of cells. The virus was injected
into the very early embryo (blastodisc stage) to target the primordial germline cells that later give rise to sperm and eggs. A total of
265 fertile eggs were injected with virus, and 35 hatched (13%); 23
of these potential founders (F0) were bred, and three (13%)
produced germline transgenic hatchlings that expressed the GFP
protein (F1). Two of these three founders (F0) have produced
transgenic young at a rate of 12% and the third at a rate of 6%.
Furthermore, two of the F1 generation transgenics have since
reproduced, one having five offspring (all GFP positive) and the
other four offsping (one GFP positive).
lentivirus 兩 song system 兩 zebra finch
V
ocal learning is one of the most distinctive characteristics of
human behavior. It is not found in other living primates and
is rarely encountered in other mammals. It is common in
songbirds, however, in which it shares many characteristics with
the learning of speech in humans (1, 2). Songbirds are particularly advantageous for research purposes because the neural
circuit that mediates the acquisition and production of learned
song is anatomically distinct and highly accessible (3–6). Songbirds’ research value does not end there; they also provide an
excellent model for the study of sensitive periods for learning (7,
8), lateralization of brain function (9, 10), sexual dimorphism
(11–16), sensorimotor vocal learning (17–20), the role of sleep in
learning (21–24), and the production and replacement of neurons in the adult brain (25, 26).
Various molecular tools for exploring these issues in songbirds
are available, including a sequenced cDNA library and microarrays
of genes expressed in the zebra finch brain (27–30), the sequence
of the zebra finch genome (Songbird Genomics Organization;
http://songbirdgenome.org.), and a zebra finch BAC library (31).
Other molecular tools not developed specifically for songbirds have
been used fruitfully in them, including quantification of immediate
early genes to map pathway activation (32–34), laser capture
microdissection to compare gene expression between cell types
(35), and non–germline-mediated viral manipulation of gene expression to modify behavior (36). However, until now, the ability to
manipulate gene expression by using germline transgenic technology has not been available for songbirds. The value of this technology is clearly demonstrated by its use in other model organisms.
Here we report the experimental steps that we took to produce
transgenic zebra finches.
The zebra finch (Taeniopygia guttata), the most widely studied
songbird, is well suited for transgenic development because it
breeds year round in captivity and has a relatively short generation
time (3 months). Our effort started with attempts to duplicate in
zebra finches approaches that have been successful in other species.
www.pnas.org兾cgi兾doi兾10.1073兾pnas.0909139106
This involved using a lentiviral vector that was initially used to make
transgenic mice (1) and was later shown to be successful in making
transgenic chickens (37, 38) and quail (39, 40). Injection of the virus
followed a protocol that had produced good results in the quail
work. In this protocol, a single injection is made into the subgerminal cavity of embryos from freshly laid eggs (39, 40). At this stage,
the quail embryo is a flat disc ⬇2.0 mm in diameter comprising
thousands of cells. These cells give rise to extra-embryonic and
embryonic tissue. The disc also includes the primordial germline
cells (PGCs) that later give rise to sperm and oocytes. The PGCs,
or their precursors, are the intended targets of the viral infection.
The single injections given to quail embryos were efficient; out of
the 6 injected embryos that hatched, 5 became germline transgenic
founders (39). The success of the technique in quail as well as in
chickens (37–39) suggested that it could be readily used to make
transgenic zebra finches as well.
The protocol used in quail failed to produce transgenic zebra
finches, however. Single injections aimed slightly under the embryonic disc resulted in very sparse, if any, infection of embryonic
tissues. The reasons for this were not determined. Instead, we
manipulated a number of variables in a search for conditions that
might improve infection. Here, we report the conditions that
resulted in transgenic founders that produced transgenic offspring.
Results
To introduce the transgene, we used a self-inactivating lentiviral
vector encoding the GFP regulated by the human ubiquitin-C
promoter (1). We made viral injections into embryos (or blastodiscs) of 265 freshly laid zebra finch eggs, of which 35 hatched
and 26 were raised to sexual maturity (Table 1); the remaining
11 died soon after hatching. The embryos in the freshly laid eggs
were ⬇1.0 mm in diameter (Fig. 1) (approximately half the size
of the quail embryo) at the time of injection. We successfully
produced transgenic individuals under the following conditions:
1. Instead of making a single injection of virus into the subgerminal cavity below the embryo, as has been described for quail
(39), we made multiple injections (10–20) directly into the
embryo.
2. The volume of each injection was ⬇15 nL, for a total injected
volume of 150–300 nL.
3. The injections were very shallow and were concentrated in the
center of the embryo (Fig. 1B), which is where primordial
germ cells are located in chicken (41, 42).
4. Viral titers ranged from 0.8 to 3.0 ⫻ 106 TU/␮L, but only titers
at the upper end of this range produced transgenics.
5. Injected eggs were placed in an incubator for 1–3 days and
then returned to the nest.
Author contributions: R.J.A., C.L., and F.N. designed research; R.J.A., B.B.S., B.H., and C.L.
performed research; C.L. contributed new reagents/analytic tools; R.J.A. analyzed data; and
R.J.A. and F.N. wrote the paper.
The authors declare no conflict of interest.
1To
whom correspondence may be addressed. E-mail: agater@rockefeller.edu or
nottebo@mail.rockefeller.edu.
2C.L.
and F.N. contributed equally to this work.
PNAS 兩 October 20, 2009 兩 vol. 106 兩 no. 42 兩 17963–17967
NEUROSCIENCE
Contributed by F. Nottebohm, August 14, 2009 (sent for review June 17, 2009)
Table 1. Parameter differences of virally injected eggs
Viral
batch
Group
size
Hatched,
n (%)
Number of founders
(number of offspring)
A
B
C
D
Total
47
77
71
70
265
10 (21)
7 (9)
9 (13)
9 (13)
35 (13)
0
1 (5)
2 (4)
0
3
Viral batch A was proximately 1 ⫻ 106 TU/␮L, batches B and C were 2–3 ⫻
106 TU/␮L, and batch D was approximately 0.8 ⫻ 106 TU/␮L. Group size refers
to the number of eggs injected, and founders are those that produced
germline transgenic offspring. The number of offspring produced thus far is
given in parentheses.
These conditions resulted in numerous embryos that exhibited
infection (⬇60%) and survived to hatching. In the embryos
exhibiting infection, the extent of infection varied considerably,
from a few to thousands of cells.
Our initial inspection for GFP expression involved external
examination under fluorescence illumination of juveniles at
Table 2. Mosaic transgenic birds (age ⬇70 days) and tissues that
were GFP-positive when visualized under a fluorescence
stereomicroscope
Bird ID
Sex
Legs
Feather
base
Breast
Muscles
Eyes
Throat
Cloaca
Blk14
Blk3
Blk11
OR682
OR703
Blk13
Blk15
*Blk19
*Blk5
Blk2
Blk1
Blk12
*Blk16
OR702
OR701
Blk24
Blk25
Blk26
Blk65
Blk28
Blk32
Blk67
Blk69
Blk83
Blk82
Blk47
F
F
M
F
M
F
F
M
M
M
F
F
M
M
F
M
F
F
M
M
M
M
M
M
M
F
⫹
(LS)⫹
⫹
⫺
⫺
⫺
(LS)⫹
⫺
⫺
⫹
⫹
⫺
⫺
⫹
(LS)⫹
⫹
⫺
⫺
⫺
⫺
⫹
(LS)⫹
(LS)⫹
⫹
⫺
⫺
⫺
⫹
⫹
⫺
⫹
⫺
(LS)⫹
⫺
⫺
⫺
⫺
⫺
⫺
⫹
⫹
⫺
⫺
⫺
⫺
(LS)⫹
⫹
(LS)⫹
⫺
⫹
⫺
⫺
⫺
⫹
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫹
⫹
⫺
⫹
⫺
⫺
⫺
⫺
⫺
⫺
⫹
⫹
⫺
⫹
⫹
⫺
⫹
⫹
⫹
(RS)⫹
⫹
⫹
⫹
⫺
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫹
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫹
⫹
⫹
⫹
⫺
⫹
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫹
⫺
⫺
⫹
⫹
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫺
‘‘⫹’’ indicates the presence of any GFP-positive cells, and ‘‘⫺’’ indicates that
no positive cells were found in that tissue. LS, left side only; RS, right side only.
The ‘‘Feather base’’ column refers to the area in and around the base of the
primary feathers of the wings, including the papilla. Asterisks identify birds
that were not bred. Blk3, OR703, and OR703 were the germline founders.
Fig. 1. (A) View of the zebra finch egg (⬇1.5 cm long). A small hole, larger
than would normally be made, has been opened to show the embryo of a
freshly laid egg (arrow). (Scale bar: 1.0 mm.) (B) A higher-magnification view
of an embryo showing the central region in which viral injections were
concentrated (black circle). (Scale bar: 1.0 mm.)
17964 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0909139106
posthatch day 70; the results are presented in Table 2. Our initial
inspection did not attempt to quantify the extent of expression,
however. The data for these birds suggest that cells giving rise to
eye tissue had a higher probability of expressing GFP than cells
giving rise to any of the other tissues that we could observe
externally. Three of the birds (Blk16, Blk5, and Blk19) were
killed and examined in greater detail by opening the body cavity
and examining various organs (muscle, gonads, liver, kidney,
heart, syrinx, and brain). Blk19, which showed no GFP expression when viewed externally, also lacked GFP expression internally. In Blk16, external examination showed GFP-expressing
cells in eye tissue, but internal examination revealed GFP
expression in the brain but not in other tissues. Blk5 had a similar
external pattern of GFP expression as Blk16, along with expression in the brain, muscles, and testicular tissue (Fig. 2).
The remaining birds were mated with uninjected partners, and
their offspring were screened for expression of the GFP protein.
Of 26 potential mosaic founders, 23 were bred, and three (two
males and a female) produced respectively, three, five, and one
germline transgenic offspring (mean of 12% for two of them and
6% for the third; Fig. 3A). A PCR-based genetic test comparing
offspring from one of the founders revealed that all offspring
that inherited the transgene from their infected parent also
showed expression of the GFP protein under fluorescent illumination (Fig. 3B). Furthermore, Southern blot analysis of two
founders and their offspring indicated that a single copy of the
transgene was integrated into the genome (Fig. 3C). Although
more than one band was observed in the offspring from founder
1 (see lanes 2 and 5), the intensities of the bands differed
Agate et al.
A
A
B
Fig. 2. GFP expression in different tissues of Blk5, a mosaic transgenic mouse
(FO). Bright field images of tissues are shown in the left column, and the
corresponding fluorescent image is presented in the right column. (A) Whole
brain as seen from above, looking at the dorsal side of the brain with the
caudal end at the bottom of the image. (B) Testes. (C) Muscle tissue of the inner
side of the left leg.
significantly. Because all bands should be of equal intensity if
additional insertions of the transgene occurred, we did not count
the upper 2 less-intense bands as additional insertions of the
transgene. One possible explanation for the presence of these
bands is that they represent incompletely digested genomic
fragments. The slight difference in band size between founder 2
offspring (lanes 2 and 4) indicated the possibility of different
integration sites of the viral transgene in different infected
PGCs, although differences in how the DNA ran in these lanes
could be responsible as well. Of the remaining potential
founders, 3 have not yet produced eggs, 13 have produced
between 10 and 15 eggs each, and 5 have produced 20–30 eggs
each, all of which were negative for GFP expression. This shows
that 13% of the 23 injected embryos that were bred as adults
produced germline transgenic founders (F0). The time from the
viral injection to the birth of the first germline transgenic was 5–6
months for two of the transgenic founders and ten months for the
third (note: the third bird did not bond and mate until it was
months older than the other two founders). The song of the first
male germline transgenic (F1) that reached sexual maturity was
a good imitation of its tutor (Fig. 4). This male has since bred and
produced a single clutch of five babies (F2), all of which were
positive for GFP expression by external examination. In addition,
a female germline transgenic (F1) has also bred and produced a
clutch of four babies (F2), one of which was positive for GFP
Agate et al.
B
NEUROSCIENCE
C
C
Fig. 3. (A) Fluorescent illumination of 2 live offspring of a transgenic founder
1 day after hatching. The undersides of the hatchlings are shown; heads are at the
top. The soft down on the heads fluoresces orange-red. The hatchling on the left
is negative for GFP expression, but its sibling on the right is positive. (B) PCR
analysis of genomic DNA from blood. GFP primers (Lower band) test for transgene, and androgen receptor primers (Upper band) test for DNA quality. M,L ⫽
Quanti-ladder from Origene. (1) No DNA. (2 and 3) Non–viral-injected adult
female and male. (4, 5, 7, and 8) Four siblings from the same clutch (2 negative,
2 positive). All offspring are of the same mosaic founder, which is represented in
lane 6. (C) Southern blot analysis of members of 2 clutches of 5 and 3 individuals,
each of which was produced by a different F0 individual (represented in lane 1).
Note that each clutch has 2 GFP-positive individuals, and that their corresponding
lanes show a single dark band, evidence that the transgene is present as a single
copy (see Materials and Methods). The GFP-negative individuals lack this band.
The absence of a detectable band in the F0 individuals presumably results from
the fact that in mosaics, the total number of blood cells that carry the gene is
below the detectable limit of Southern blot analysis but not that of PCR analysis.
PNAS 兩 October 20, 2009 兩 vol. 106 兩 no. 42 兩 17965
20 GFP-positive colonies (dilution factor 1 ⫻ 105). This number was then used
to calculate the titer of virus used for injections.
Injection Apparatus. Glass pipettes (Drummond Science, catalog no. 5– 000-1001)
were pulled by using a Narishige model PE-2 pipette puller. Pipettes were polished to a 50- to 60-degree bevel (tip diameter ⬇10 –20 ␮m). Approximately 4 ␮L
of viral suspension plus 0.5 ␮L of phenol red (5% in PBS) were back-loaded into
the glass pipette, followed by 1–2 ␮L of mineral oil, after that a metal plunger
supplied with the glass pipettes was inserted. The position of the injecting needle
was determined by using a custom-made holder attached to a Narishige UM-3C
3-dimensional manipulator. Movement of the plunger that delivered the injection was controlled by a Narishige MO-10 hydraulic micromanipulator; further
details are available on request. This allowed us to position the glass pipette
properly and make precise injections of virus into the embryo.
Fig. 4. Insertion and expression of the transgene does not inhibit song
imitation. Shown are spectrograms of song from the tutor (A) and the germline transgenic male pupil (B) at age 4 months.
expression by external examination. Thus, the transgene was
passed to a second generation, and the F2 birds continued to
express it.
Discussion
The lentiviral vector that we used encodes for the GFP protein
regulated by the human ubiquitin-C promoter, which is active
in a wide range of cells. Mating of the mosaic transgenic
animals (F0) resulted in the transgene being inherited and
expressed by their offspring (F1) in some cases and, in two
cases at least, by the next generation (F2). The 13% efficiency
in producing the (F0) founders, although significantly less than
that seen in quail (83%) (39) or chickens (50%) (37), was
better than that reported in another study on producing
transgenic chickens (5%) (38). A mild improvement results if
efficiency is calculated based only on those embryos that
received injections of virus from batches B and C (13%), from
which the three transgenic founders were derived and that had
higher viral titers than the other batches. Because the proportion of injected embryos that completed development and
hatched was very similar in zebra finches, quail, and chickens,
the difference in the ability to produce transgenic zebra finches
is likely due to a lower efficiency in the infection of zebra finch
PGCs. This interpretation is supported by the need to make
multiple injections directly into the zebra finch embryo to get
F0 transgenics, whereas a single injection placed below the
embryo sufficed in quail and chickens. In addition, the viral
titers required to make transgenic quail and chickens (37, 39)
were lower than those required to produce transgenic zebra
finches. These observations suggest that zebra finch PGCs are
not as readily accessible to the virus or as readily infected by
the virus, and thus higher viral titers are needed to overcome
this. Determining the cause of this relative resistance to
infection will make it possible to increase the efficiency with
which transgenic zebra finches can be produced. We continue
to work on improving the efficiency of our protocol.
Materials and Methods
All animals were cared for in accordance with the standards set by the
American Association of Laboratory Animal Care and Rockefeller University’s
Animal Use and Care Committee.
Lentivirus Production. The following 3-plasmid system was used to generate
lentivirus: transfer vector, pFUGW; packaging vector, ⌬8.9; envelope glycoprotein, VSVg (1). Virus production and concentration were performed as
described in ref. 43. Titer was determined by making serial dilutions (10-fold
steps) of the virus in PBS and infecting HEK 293T cells grown in 96-well plates.
After a 72-h incubation, the number of GFP-positive colonies was counted in
the well with the most diluted sample of virus that still yielded a minimum of
17966 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0909139106
Injection Procedure and Incubation. An egg candler was used to visualize the
embryo through the eggshell. Each egg was placed lengthwise in a silicon
mold and oriented so that the embryo was just below the top surface of the
egg. A black marker was used to place a dot on the eggshell at this position.
With the aid of a stereo zoom microscope (Nikon SMZ800, with an Achromatic
0.5⫻ objective), a 32-gauge needle was used to make many small and shallow
holes in the eggshell surface. The holes were made in a circular pattern,
creating an approximate 1.0-mm-diameter perforated ring in the shell just
above the embryo. Sharp forceps were used to remove the circular piece of
shell and expose the embryo. To prevent the embryo from drying, a small
amount of albumin from another egg was applied directly to the window
opening. [Note: Although not quantified, using standard PBS (pH 7.4) for this
part of the protocol seemed to reduce the survival of embryos.] Between 10
and 20 shallow injections of virus (15 nL per injection; 150 –300 nL total) were
targeted in and around the center of the embryo (Fig. 1B). After the injection,
a piece of shell from another egg, slightly larger than the piece removed, was
used to patch the hole. Residual albumin around the edges of the patch, when
dried, served as a strong adhesive to hold the patch in place. (Note: The egg
was allowed to dry at room temperature.) The virally injected eggs were then
placed in an incubator for 1–3 days (37–38 °C, 40 –50% humidity), after
that they were returned to nests to be incubated by birds sitting on eggs of
similar age.
GFP Evaluation of Virally Injected Birds. Examination of GFP expression in
different tissues of potential founders (FO) was performed at 70 days. Two
different filter sets were used to evaluate GFP expression in tissues, one set to
visualize GFP and the other to visualize Texas Red (Chroma Technology 49002
and 49008). False-positives (i.e., non-GFP green spectrum emissions) were
readily distinguished from true GFP signals by examining whether the signal
also was visible when using the Texas Red filter set. (In most cases, falsepositives result from broad-spectrum autofluorescence and thus also appear
when viewed with a different spectral filter set, such as that for Texas Red.)
The tissues exhibiting the greatest autofluorescence were the intestines, the
crop (when full with seed), and the soft down of newly hatched chicks.
Genomic PCR Test for GFP. Genomic DNA was extracted from blood (⬇4 ␮L),
and the presence of the GFP transgene was established using the following
GFP-specific primers: forward, GCACGACTTCTTCAAGTCCGCCATGCC; reverse,
GCGGATCTTGAAGTTCACCTTGATGCC. The following androgen receptor
primers were used as controls: forward, CCTTGTGAGGTGGGAGAGCTTT; reverse, AAGGAGATGCTCAATCCAGGGC. PCR was performed by using an MJ
Research PTC200 instrument under the following conditions: 32 cycles of 95 °C
for 2 min, followed by 95 °C for 30 s, 67 °C for 1 min, and 68 °C for 1 min. All
samples were run on 3– 4% Nusieve 3:1 agarose gel.
Southern Blot Analysis of DNA from Blood. Approximately 75 ␮L of blood was
collected from the wing vein of each bird, and an equal volume of proteinase
K buffer [50 mM Tris䡠HCl (pH 7.6), 0.1 M EDTA (pH 8.0), 0.1 M NaCl, 1% SDS)
and 0.5 mg/mL proteinase K was added. The solution was incubated overnight
at 50 °C with shaking. A phenol/chloroform extraction was performed, and
the genomic DNA was precipitated with ethanol and then resuspended in TE
buffer. Then ⬇10 ␮g of DNA was digested with PstI, run on a gel, and
transferred to a nylon membrane. The membrane was probed with a 300-bp
fragment of the GFP coding sequence amplified using the primers listed above
in the presence of radiolabeled p32 dCTP. The membrane was then exposed
to radiographic film.
The transgene contains a single PstI restriction site upstream of the GFP coding
region. The next downstream PstI site in the zebra finch genome will occur where
the transgene integrates itself and will define the size of the band containing the
GFP sequence on Southern blot analysis. If more than one transgene is inserted,
Agate et al.
ACKNOWLEDGMENTS. We thank the Rockefeller Field Research Center staff
for their outstanding care of the birds in this study. This work was funded by
McKnight Technological Innovations in Neuroscience Awards, the Whitehall
Foundation, and The Rockefeller University.
1. Marler P (1970) Birdsong and speech development: Could there be parallels? Am Sci
58:669 – 673.
2. Doupe AJ, Kuhl PK (1999) Birdsong and human speech: Common themes and mechanisms. Annu Rev Neurosci 22:567– 631.
3. Nottebohm F, Stokes TM, Leonard CM (1976) Central control of song in canary, Serinus
canarius. J Comp Neurol 165:457– 486.
4. Bottjer SW, Miesner EA, Arnold AP (1984) Forebrain lesions disrupt development but
not maintenance of song in passerine birds. Science 224:901–903.
5. Bottjer SW, Halsema KA, Brown SA, Miesner EA (1989) Axonal connections of a
forebrain nucleus involved with vocal learning in zebra finches. J Comp Neurol
279:312–326.
6. Vates GE, Vicario DS, Nottebohm F (1997) Reafferent thalamo-‘‘cortical’’ loops in the
song system of oscine songbirds. J Comp Neurol 380:275–290.
7. Eales LA (1985) Song learning in zebra finches: Some effects of song model availability
on what is learnt and when. Anim Behav 33:1293–1300.
8. Marler P, Tamura M (1964) Culturally transmitted patterns of vocal behavior in sparrows. Science 146:1483–1486.
9. Nottebohm F (1971) Neural lateralization of vocal control in a passerine bird, I: Song.
J Exp Zool 177:229 –261.
10. Nottebohm F (1977) in Lateralization in the Nervous System, eds Harnad S, Doty R,
Goldstein L, Jaynes J, Krauthamer G (Academic, New York), pp 23– 44 .
11. Nottebohm F, Arnold AP (1976) Sexual dimorphism in vocal control areas of songbird
brain. Science 194:211–213.
12. Gurney ME, Konishi M (1980) Hormone-induced sexual differentiation of brain and
behavior in zebra finches. Science 208:1380 –1383.
13. Ball GF, et al. (2004) Seasonal plasticity in the song control system: Multiple brain sites
of steroid hormone action and the importance of variation in song behavior. Ann N Y
Acad Sci 1016:586 – 610.
14. Arnold AP, Bottjer SW, Brenowitz EA, Nordeen EJ, Nordeen KW (1986) Sexual dimorphisms in the neural vocal control system in song birds: Ontogeny and phylogeny. Brain
Behav Evol 28:22–31.
15. Bottjer SW, Johnson F (1997) Circuits, hormones, and learning: Vocal behavior in
songbirds. J Neurobiol 33:602– 618.
16. Wade J, Arnold AP (2004) Sexual differentiation of the zebra finch song system. Ann
N Y Acad Sci 1016:540 –559.
17. Solis MM, Doupe AJ (1997) Anterior forebrain neurons develop selectivity by an
intermediate stage of birdsong learning. J Neurosci 17:6447– 6462.
18. Konishi M (1965) The role of auditory feedback in the control of vocalization in the
white-crowned sparrow. Z Tierpsychol 22:770 –783.
19. Margoliash D, Konishi M (1985) Auditory representation of autogenous song in the
song system of white-crowned sparrows. Proc Natl Acad Sci USA 82:5997– 6000.
20. Williams H, Nottebohm F (1985) Auditory responses in avian vocal motor neurons: A
motor theory for song perception in birds. Science 229:279 –282.
21. Dave AS, Margoliash D (2000) Song replay during sleep and computational rules for
sensorimotor vocal learning. Science 290:812– 816.
22. Cardin JA, Schmidt MF (2003) Song system auditory responses are stable and highly
tuned during sedation, rapidly modulated and unselective during wakefulness, and
suppressed by arousal. J Neurophysiol 90:2884 –2899.
23. Deregnaucourt S, Mitra PP, Feher O, Pytte C, Tchernichovski O (2005) How sleep affects
the developmental learning of bird song. Nature 433:710 –716.
24. Hahnloser RH, Fee MS (2007) Sleep-related spike bursts in HVC are driven by the nucleus
interface of the nidopallium. J Neurophysiol 97:423– 435.
25. Goldman SA, Nottebohm F (1983) Neuronal production, migration, and differentiation
in a vocal control nucleus of the adult female canary brain. Proc Natl Acad Sci USA
80:2390 –2394.
26. Paton JA, Nottebohm FN (1984) Neurons generated in the adult brain are recruited into
functional circuits. Science 225:1046 –1048.
27. Wade J, et al. (2004) A cDNA microarray from the telencephalon of juvenile male and
female zebra finches. J Neurosci Methods 138:199 –206.
28. Wada K, et al. (2006) A molecular neuroethological approach for identifying and
characterizing a cascade of behaviorally regulated genes. Proc Natl Acad Sci USA
103:15212–15217.
29. Li X, et al. (2007) Genomic resources for songbird research and their use in characterizing gene expression during brain development. Proc Natl Acad Sci USA 104:6834 –
6839.
30. Replogle K, et al. (2008) The Songbird Neurogenomics (SoNG) Initiative: Communitybased tools and strategies for study of brain gene function and evolution. BMC
Genomics 9:131.
31. Luo M, et al. (2006) Utilization of a zebra finch BAC library to determine the structure
of an avian androgen receptor genomic region. Genomics 87:181–190.
32. Mello CV, Vicario DS, Clayton DF (1992) Song presentation induces gene expression in
the songbird forebrain. Proc Natl Acad Sci USA 89:6818 – 6822.
33. Jarvis ED, Nottebohm F (1997) Motor-driven gene expression. Proc Natl Acad Sci USA
94:4097– 4102.
34. Kimpo RR, Doupe AJ (1997) FOS is induced by singing in distinct neuronal populations
in a motor network. Neuron 18:315–325.
35. Lombardino AJ, Li XC, Hertel M, Nottebohm F (2005) Replaceable neurons and neurodegenerative disease share depressed UCHL1 levels. Proc Natl Acad Sci USA 31:8036 –
8041.
36. Haesler S, et al. (2007) Incomplete and inaccurate vocal imitation after knockdown of
FoxP2 in songbird basal ganglia nucleus area X. PLoS Biol 5:e321.
37. McGrew MJ, et al. (2004) Efficient production of germline transgenic chickens using
lentiviral vectors. EMBO Rep 5:728 –733.
38. Chapman SC, et al. (2005) Ubiquitous GFP expression in transgenic chickens using a
lentiviral vector. Development 132:935–940.
39. Scott BB, Lois C (2005) Generation of tissue-specific transgenic birds with lentiviral
vectors. Proc Natl Acad Sci USA 102:16443–16447.
40. Scott BB, Lois C (2006) Generation of transgenic birds with replication-deficient lentiviruses. Nat Protoc 1:1406 –1411.
41. Tsunekawa N, Naito M, Sakai Y, Nishida T, Noce T (2000) Isolation of chicken vasa
homolog gene and tracing the origin of primordial germ cells. Development 127:2741–
2750.
42. Nakamura Y, et al. (2007) Migration and proliferation of primordial germ cells in the
early chicken embryo. Poultry Sci 86:2182–2193.
43. Pease S, Lois C (2006) Mammalian and Avian Transgenesis: New Approaches (Springer,
New York).
Agate et al.
PNAS 兩 October 20, 2009 兩 vol. 106 兩 no. 42 兩 17967
NEUROSCIENCE
then the second PstI site likely will be located a different distance from the PstI in
the transgene, resulting in a different-sized band on the blot. Thus, the number
of bands detected by Southern blot analysis indicates how many copies of the
transgene have been integrated in the genome.