Download Mouse Embryology Module, Woods Hole 2010

Mouse Embryology Module, Woods Hole 2010
Paul Trainor
([email protected])
Stowers Institute, 1000 E. 50th Street Kansas City, MO, 64110 USA
Amanda Barlow
([email protected])
Stowers Institute, 1000 E. 50th Street Kansas City, MO, 64110 USA
Angelo Iulianella
([email protected])
Dalhousie University, 5850 College Street, Halifax, B3H 1X5, Canada
Reference books for techniques in mouse embryology
Manipulating the Mouse Embryo: A Laboratory Manual 3rd edition (eds A. Nagy, M. Gersenstein, K.
Vintersten, and R. Behringer), Cold Spring Harbour Laboratory Press, New York.
Guide to Techniques in Mouse Development: In Methods in Enzymology, Vol 225 (eds P. W.
Wassarman and M. L. DePamphilis) 1993. Academic Press, San Diego.
Molecular Embryology, Methods and Protocols: In Methods in Molecular Biology, Vol 97 (eds P
Sharpe and I Mason), 1999. Humana Press, Totowa.
Postimplantation Mammalian Embryos: A Practical Approach (eds A. J. Copp and D. L. Cockroft)
1990. Oxford University Press, Oxford.
Mouse Develpoment: Patterning, Morphogenesis, Organogenesis (Janet Rossant and Patrick Tam,
eds) 2002. Academic Press, New York.
1
Introduction
Many techniques and protocols can be used interchangeably on chick and mouse
embryos (DiI labelling, in situ hybridisation, antibody staining, electroporation, etc).
To avoid redundancy we have omitted listing some techniques here that are
described in the chick manual. In some cases, we have listed specific protocols
below but generally you should feel comfortable using the chick protocols on mice.
If in doubt, or if there is something unusual you want to try, please don’t hesitate to
ask and we’ll be happy to advise.
1. Skeletal Preparations- Cartilage and Bone staining
3
1.1 Cartilage staining in mouse embryos from E13.5
3
1.2 Bone and cartilage stain for embryos 14.5 days or older
3
2. Immunohistochemistry on whole mouse embryos
4
2.1 Wholemount immunofluorescence for neuronal differentation (TuJ1) and organ
development (glucagon-pancreas)
4
2.2 Neurofilament Staining (2H3 antibody)
5
2.3 Endothelial cell staining (PECAM1 antibody)
6
3. RNA in situ hybridization of mouse embryos
7
4. Isolation and in vitro culture of 6.5dpc-9.5dpc embryos
10
4.1 Isolation and in vitro culture of embryos 6.5dpc-9.5dpc
10
4.2 Whole embryo culture
13
4.3 Lineage tracing with DiI
13
4.4 In vitro culture with morphogens or inhibitors or teratogenic agents such drugs or
ethanol
13
5. Detection of apoptotic cell death
14
5.1 “LysoTracker Red” staining for apoptotic cells in mouse embryos
14
5.2 Acridine orange staining for apoptotic cells in mouse embryos
15
5.3 Nile blue staining for apoptotic cells in mouse embryos
15
6. Visualizing vasculature by injection of India Ink
16
7. Detection of primordial germ cells
16
8. X-gal staining of lacZ transgenic embryos
17
9. Organ cultures of explanted organ and tissue
18
10. Tissue explants cultured under the kidney capsule for long term differentiation
analyses
20
11. Mouse embryo DNA electroporation
21
11.1 In vitro mouse embryo DNA electroporation
21
11.2 In vivo/ex utero mouse embryo DNA electroporation
22
12. Interspecies transplantations (somites, neural tube, neural crest etc)
22
13. Transgenic and Mutant Mouse lines
22
14. Snakes and Lizards
23
15. Inhibitors, Antagonists and Antibodies
27
16. Mouse Development references
28
16.1 Mouse Development timeline
28
16.2 Whole mount embryos at representative stages of development
29
16.3 Mouse Embryo Staging Criteria – from Edinburgh Mouse Atlas website
30
16. Sample Photographs of Results of the Above Protocols
33
2
1. Skeletal Preparations- Cartilage and Bone staining
1.1 Cartilage staining in mouse embryos from E12.5 - E13.5
Dissect out embryos, rinse in 1 x PBS, and place in ice-cold 95% Ethanol for 1 hour.
Change to fresh 95% ETOH and rock overnight at room temperature (RT).
Change 95% Ethanol and begin the staining alternatively you can store the embryos at 4°C until you
are ready to stain.
Note: no deskinning/ removal of organs needed for E15.5 and younger.
Stain overnight at RT in freshly prepared stain solution (see recipe below).
Briefly rinse the embryos in water.
Dissolve tissues by adding KOH for the following times:
10 mins in 1%KOH for E12.5 to E13
20 mins in 1%KOH for E13.5
Rinse in 0.25% KOH for 30 mins.
Clear in 20% glycerol / 0.25% KOH for 1 hr.
Clear in 33% glycerol / 0.25% KOH for 1 hr.
Clear in 50% glycerol / 0.25% KOH overnight.
Change solution to fresh 50% glycerol / 0.25% KOH further clearing and storage.
1.2 Bone and cartilage stain for embryos 14.5 days or older
Prenatal and neonatal foetuses and postnatal pups all need to be anesthetized by hypothermia in ice
cold PBS until there is no response then euthanized by submerging in ice cold ethanol (not cervical
dislocation). For embryos at E17.5 or older, after euthanasia, carefully remove as much of the skin
and viscera as possible.
Dissect out embryos, rinse in 1x PBS, and place in ice-cold 95% Ethanol for 1 hour.
Change to fresh 95% Ethanol and rock overnight at RT. (The embryos can be stored at this stage in
95% at 4°C until you are ready to do the stain). Note: no deskining/ removal of organs is needed for
E15 and younger. Skin and remove organs for E17.5 and older.
Stain overnight with rocking at room temperature (RT) in freshly prepared stain solution (see recipe
below).
Briefly rinse the embryos in water.
Dissolve tissues by adding KOH for the following times:
30 mins in 1%KOH for E14.5
3 hrs in 2%KOH for E15.5
6 hrs in 2%KOH for E17.5
Rinse in 0.25% KOH for 30 mins.
Clear in 20% glycerol / 0.25% KOH for 1 hr.
Clear in 33% glycerol / 0.25% KOH for 1 hr.
Clear in 50% glycerol / 0.25% KOH overnight.
3
Change solution to fresh 50% glycerol / 0.25% KOH further clearing and storage.
Recipe for stain solution:
5 ml
0.4% Alcian Blue 8 GX in 70% ETOH
5 ml
glacial acetic acid
70 ml 95% ETOH
20 ml water
final volume is 100ml
When ready to do stain add 100ul of 0.5% Alizarin red to 10 ml of stain solution.
0.4% Alcian Blue /70% ETOH:
Alcian Blue does not readily dissolve in 70% ETOH, but does dissolve in 50% ETOH. To make 100
ml of 0.4% Alcian Blue stock, add 0.4g Alcian Blue to 10 ml 50% ETOH in a 250 ml bottle. Swirl and
place in 37°C water bath. Swirl occasionally. When dissolved, add 25 ml water and 65 ml 95%
ETOH.
0.5% Alizarin Red S/ water:
Add 0.5g alizarin red S to 100ml water. Swirl to dissolve.
2. Immunohistochemistry on whole mouse embryos
2.1 Wholemount immunofluorescence for neuronal differentiation (TuJ1) and organ
development (glucagon-pancreas)
adapted from: Jonas Ahnfelt-Rønne et al, J. Histochem. Cytochem.2007
Day 1
Fix embryos or tissue in 4% PFA in PBS over night at 4°C.
Dehydrate embryos through to 100% Methanol by rinsing for 10 minutes at RT in PBS, then 25%
methanol.PBS, 50% methanol/PBS, 75% methanol/PBS and then into 100% MeOH with gentle
agitation.
Embryos and tissues can be stored in Methanol for extended periods at -20°C until you are ready to
proceed alternatively incubate the embryos in Dent’s bleach (MeOH:DMSO:30%H2O2, 4:1:1) for 2
hours at room temperature. (This is a critical step which promotes antibody penetration and quenches
auto-fluorescence (Alanentalo et al. 2007)).
Remove Dent’s bleach and replace with absolute Methanol and rinse for 10 minutes at RT rocking.
Equilibrate the embryos to PBS through a series of descending Methanol/PBS concentrations,
(100%Methanol, 75% methanol/PBS, 50% methanol/PBS, 25% methanol/PBS, PBS), 10 minutes
each. The tissue may become sticky during this procedure so be aware not to suck it into pipettes etc.
Place the embryos into blocking solution (3% BSA in PBS + 0.1% Triton) and rock for two hours at
RT.
Dilute the primary antibodies (TuJ1-1:1000; Glucagon-1:500) in blocking solution and incubate
overnight at 4°C with gentle rocking.
Day2
Wash 3 x 10 minutes with PBS and then 3 x 1 hour with PBS.
4
Apply the secondary antibodies, usually diluted 1:500, in blocking solution and rock over night at 4°C.
Day3
Wash 3 times 10 minutes with PBS and then 2 x 1 hour in PBS at RT with rocking.
Then place into 25% glycerol/PBS and rock at RT for 1 hour.
Replace with 50% glycerol/PBS and rock for 1 hour at RT, then replace this with Vectashield
containing DAPI to stain all of the nuclei.
Visualize the embryos under a fluorescent microscope.
2.2 Neurofilament Staining (2H3 antibody)
For embryos up to E10, fix in 4% PFA at RT for 1 hour.
Rinse the embryos in PBS 3 x 5 minutes at RT. Place one embryo per tube in order to attain good
immunostaining
Remove the PBS and then add the blocking solution (3% BSA in PBS + 0.1% Triton), rock for 1 hour
at RT.
Dilute the neurofilament 2H3 antibody 1:1000 in the blocking solution and add to the embryos, then
either rock at RT for 4 hours or place at 4°C and rock overnight.
Rinse the embryos in PBS + 0.1% Triton 3 x 5 minutes at RT, then rock in PBS + 01% triton for I hour
at RT.
Dilute the fluorescent secondary antibody (anti-mouse Alexa Fluor 1:500) in blocking solution and
rock the embryos in this IN THE DARK either overnight at 4°C or at RT for 4 hours.
Rinse the embryos in PBS + 0.1% Triton 3 x 5 minutes at RT, then rock in PBS + 01% triton for I hour
at RT.
Rinse the embryos in PBS 3 x 5 minutes and then place into 25% glycerol/PBS and rock at RT for 1
hour.
Replace with 50% glycerol/PBS and rock for 1 hour at RT and then replace with Vectashield
containing DAPI to stain all of the nuclei.
Visualize the embryos under a fluorescent microscope.
For embryos older than E10
Day 1
Dissect out the embryos and then fix in 4 % PFA at 4°C overnight.
Day 2.
5
Rinse the embryos in PBS 3 x 5 minutes at RT. Then dehydrate the embryos through a series of 25%
methanol/PBS, 50% methanol/PBS, 75% methanol/PBS into 100% methanol each step for 10
minutes at RT.
( Note, that the embryos can be stored in the freezer at this stage at -20°C until you are ready to
continue).
Place one embryo per tube in order to attain good immunostaining.
Remove the methanol and incubate the embryos in Dent’s bleach (4:1:1 Methanol:DMSO:Hydrogen
peroxide) for 2 hours at RT. ( Make this solution just prior to using).
Rinse the embryos in 100% methanol for 15 minutes at RT.
Equilibrate the embryos through a descending series of Methanol concentrations in PBS (75%, 50%,
25%), for 10 minutes each rocking at RT. The embryos may become sticky during this procedure and
so be careful when adding or removing solutions not to suck the embryos into the pipettes.
Place the embryos into 1 x PBS for 10 minutes at RT and then into blocking solution. Rock the
embryos at RT for 2 hours.
Add the primary antibody (1:1000 dilution in blocking solution) to the embryos and incubate overnight
at 4°C with gentle rocking.
Day 3.
Wash the embryos in PBS 3 x 5 minutes and then 3 x 1 hour at RT rocking.
Apply the secondary antibody diluted 1: 500 (anti-mouse Alexa Flour) in blocking solution and rock
the embryos overnight at 4°C IN THE DARK.
Day 4.
Wash the embryos 3 x 5 minutes with PBS and then for 1 hour at RT rocking.
Then place into 25% glycerol/PBS and rock at RT for 1 hour.
Replace this with 50% glycerol/PBS and then add Vectashield containing DAPI to stain all of the
nuclei.
Visualize the embryos under a fluorescent microscope. The embryos can be stored in the dark at
4°C.
2.3 Endothelial cell staining (PECAM1 antibody)
Materials:
Use Tween20 (not Triton) in all solutions.
FBS-HI = Heat inactivated fetal bovine serum: heat FBS 1hour at 56°C (no more than 1h, otherwise it
turns to jelly)
Antibodies:
Rat anti-mouse PECAM1 . Mec13.3 clone (BD Pharmingen, 553370) – use @ 1:500
Goat anti-rat Cy3 (Jackson, 112-165-167) – use @ 1:400
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Method:
DAY1
Dissect embryos in PBS.
Fix in 4%PFA at RT for not more than 1hour with gentle agitation.
Wash 3 x 10min in PBS + 0.1% Tween20 (PBT) at RT with gentle agitation.
Place one embryo per tube in order to attain good immunostaining. Incubate for 2 hours at RT in
blocking solution (PBS + 0.1% Tween20 + 10% FBS-HI).
Incubate with primary antibody in block solution, 1hour at RT then overnight at 4°C with gentle
agitation.
DAY2
Wash with PBT 3 x 5 minutes and then 2 x 1 hour at RT with gentle agitation.
Incubate with secondary antibody in block solution IN THE DARK, overnight at 4°C with gentle
agitation.
DAY3
Wash with PBT 3 x 5 minutes and then 2 x 1 hour at RT with gentle agitation.
Place the embryos into 25% glycerol/PBS and rock for 1 hour at RT.
Replace this with 50% glycerol/PBS and rock for 1 hour at RT.
Place embryos into Vectashield with DAPI and visualize on a fluorescent microscope. The embryos
can be stored in the dark at 4°C.
3. RNA in situ hybridization of mouse embryos
This protocol can be used for both mouse and chick embryos
Synthesis of RNA probe (riboprobe):
Mix the following at room temperature
Sterile distilled water (DEPC-H2O)
5X transcription buffer
100mM DTT
DIG 10X nucleotide mix
Linearised plasmid (1ug/ul)
Rnasin ribonuclease inhibitor
SP6, T3 or T7 polymerase
11.5ul
5.0ul
2.5ul
2.5ul
1.5ul
0.5ul
1.5ul
Total 25ul
Incubate at 37°C for 2 hours.
7
Remove 1ul aliquot and run on a 1% TAE gel to check synthesis. Expect to see RNA band 10 fold
more intense than plasmid band suggesting 10-15ug probe synthesized.
Add 2ul of DNAse1 (ribonuclease free) and incubate at 37°C for 15 minutes.
Add 50ul dH20, 25ul 10M ammonium acetate and 200ul of 100% ethanol.
Mix and leave on dry ice for 30 minutes or at –70°C overnight.
Spin in centrifuge at 4°C for 20 minutes at 13-15000rpm.
Wash pellet in 50ul 70% ethanol and spin at 4 C for 5 minutes at 13-15000rpm.
Air dry pellet for 10-15 minutes until obvious traces of ethanol have evaporated.
Re-dissolve pellet in 50ul DEPC-H2O and store at –20°C.
Use 2ul of probe for each 1ml of hybridization mix.
Note: all solutions should be DEPC treated and autoclaved or prepared with DEPC-H2O
Embryo preparation
Dissect embryos in PBS and fix in 4% Paraformaldehyde (in PBS) overnight.
Rinse embryos in PBS + 0.1% Tween 20 (PBT) at room temp for 10 minutes
Dehydrate the embryos through a graded series of methanol diluted in PBT
25% methanol
10 minutes
50% methanol
10 minutes
75% methanol
10 minutes
100% methanol
10 minutes
The embryos can be stored at –20°C for a very long time.
Rehydrate the embryos through a graded series of Methanol diluted in PBT
75% methanol
10 minutes
50% methanol
10 minutes
25% methanol
10 minutes
Wash in PBT
5 minutes
Treat embryos with 10ug/ml Proteinase K in PBT for 5-15 minutes or longer at room temp, depending
on the size of the embryo. All probes should be tested for optimum digestion time prior to routine use.
(Proteinase K stock is 10mg/ml, therefore use 2ul in 2ml of PBT)
Proteinase K at 10ug/ml in PBT incubate at RT without agitation.
E8.5
5 minutes
E9.5
10 minutes
E10.5 15 minutes
Wash in PBT
5 minutes
Re-fix embryos in 4% Paraformaldehyde / 0.25% Glutaraldehyde for 20 minutes at RT
(Glutaraldehyde stock is 25%, therefore use 20ul in 2ml)
8
Wash in PBT
10 minutes
Transfer embryos to 2ml eppendorf tube (round bottom). 1-3 embryos can be processed in a single
eppendorf tube.
Hybridisation of embryos
Prehybridise embryos (incubate) for at least 1.5 hours in 1-2ml of pre-warmed Hybridisation buffer at
65°C.
Hybridise embryos (incubate) by adding 2ul of RNA probe per 1ml of hybridisation buffer at 65°C
overnight in a waterbath (1ml of buffer is sufficient for each tube).
Post-hybridisation of embryos
Start the washes first thing in the morning so that the antibody washes can be performed overnight.
Pre-warm 2x SSC/0.1%CHAPS and 0.2 x SSC/0.1% CHAPS at 65°C for around 30 minutes.
Wash embryos twice in 2ml of 2xSSC/0.1% Chaps for 45 minutes at 65°C.
Wash embryos once in 2ml of 0.2xSSC/0.1% Chaps for 30 minutes at 65°C.
Wash in KTBT for 5 minutes at RT.
Antibody and colour development
Incubate the embryos in 2ml of blocking solution (20% goat serum or lamb serum in KTBT) for at
least 1 hour at 4°C with continual rocking.
Add 1:2000 dilution of DIG-Alkaline Phosphatase antibody in blocking solution and incubate at 4°C
overnight with continual rocking.
Rinse the embryos 3 x 1 hour in KTBT and then wash overnight at 4°C in 2ml of KTBT.
In the morning rinse the embryos 2 x 10 minutes with KTBT.
Wash the embryos in 1ml of Alkaline Phosphatase buffer for 3 x 10 minutes at RT.
Add 3.375ul of NBT (100mg/ml in DMF) and 3.5ul of BCIP (50mg/ml in DMF) in 1ml Alkaline
Phosphatase buffer and allow the reaction to occur with shaking at room temperature in the dark (ie
wrap the eppendorfs in foil or place inside a dark container).
Stop the reaction once the desired colour intensity is achieved by fixing the embryos in 4%
paraformaldehyde. This can take from 15minutes to overnight, however 1-2 hours is more usual.
Continual observation/checking of the colour development is not advisable as the background signal
will increase.
9
Alternative post-hybridisation washes in MABT
For some genes/riboprobes which consistently give high background staining, it may be preferable to
wash the embryos in MAB instead of SSC/SDS.
After hybridising the embryos overnight at 65°C, wash the embryos in a 1:1.5 mix of hybridization
buffer and MABT at 65°C for 1 hour.
Then wash the embryos for 30 minutes at 65°C in MABT.
Incubate the embryos in blocking solution (20% goat serum or lamb serum in MABT) for at least 1-1.5
hours at 4°C with continual rocking.
Add 1:2000 dilution of DIG-Alkaline Phosphatase antibody in blocking solution and incubate at 4°C
overnight with continual rocking.
Rinse the embryos at least twice in MABT and then wash overnight at 4°C in 2ml of MABT.
In the morning wash the embryos at 2 x 10 minutes with MABT.
Wash the embryos in 1ml of Alkaline Phosphatase buffer for 3 x 10 minutes at RT
Add 3.375ul of NBT (100mg/ml in DMF) and 3.5ul of BCIP (50mg/ml in DMF) into 1ml Alkaline
Phosphatase buffer and allow the reaction to occur with shaking at RT in the dark (ie wrap the
eppendorfs in foil or place inside a dark container)
Stop the reaction once the desired colour intensity is achieved by fixing the embryos in 4%
paraformaldehyde. This can take from 15minutes to overnight, however 1-2 hours is more usual.
Continual observation/checking of the colour development is not advisable as the background signal
will increase.
4. Isolation and in vitro culture of 6.5dpc-9.5dpc embryos
4.1 Isolation and in vitro culture of embryos 6.5dpc-9.5dpc
For a detailed description of this procedure please refer to Chapter 5 protocol 2 “Isolating
Postimplantation Embryos” from Manipulating the Mouse Embryo: a Laboratory Manual 3rd edition
(Eds: Nagy, A., Getsenstein, M., Vintersten, K., Behringer, R.) Cold Spring Harbor Laboratory Press,
from which the following diagrams were copied
Materials
DMEM/F12-Glutamax (Dulbecco’s modified eagles medium) supplemented with penicillin and
streptomycin
Fetal calf serum
Rat serum
CO2 incubator
Incubator and rotating drum
Special gas mixtures 5%O2, 5%CO2, 90%N2; mixtures 20%O2, 5%CO2, 75%N2
10
Dissecting decidua + diagrams
Sacrifice pregnant mice by cervical dislocation and place the mouse in a supine position. Wet the fur
over the abdomen with 70% ethanol and make a V-shaped incision through the skin and underlying
muscle. Lift the uterine horns clear of the peritoneal cavity and trim the mesometrium (blood supply)
and any adhering fat, along the length of the horns. Explant the uteri by cutting their connection with
the oviduct into DMEM. The uterine horns are opened by splitting the muscular wall along the
antimesometrial side. Push the uterine walls apart to expose the decidua which can then be freed by
carefully sliding a pair of forceps underneath the decidua and teasing or pinching them off the uterine
wall. Transfer the decidua to clean DMEM medium plus 10% fetal calf serum (which reduces
stickiness) with a pasture pipette. Care should be taken during the process to avoid excessive
compression which could deform the embryo and compromise its subsequent development.
Dissection of gastrulating stage embryos (6.5-7.5dpc) + diagrams
Primitive streak stage embryos (6.5-7.5dpc) are cylindrical structures contained within the
antimesometrial half (thinner end) of the pear shaped decidua. Dissection therefore starts with the
broader end to avoid damaging the embryo. Hold the deciduum lightly with a pair of forceps and
insert the points of another pair of forceps through the decidua in the midplane at about 1/3 distance
from the broad end. The deciduum is then split into halves by allowing the arms of the forceps to
open. Pull the deciduum halves apart by grasping and pulling apart the flaps of tissue. The egg
cylinder, with the reddish ectoplacental cone and a fluffy trophoblastic covering is usually left
imbedded in one half of the deciduum. The half deciduum is then pinned down with a holding pair of
11
forceps, and the egg cylinder is scooped out by teasing the tissues surrounding the egg cylinder with
the closed points of a pair of forceps. The final and most difficult step is to remove Reichert’s
membrane and the associated trophoblasts and parietal endoderm. Grasp the membrane at the
junction between the embryonic and extra-embryonic portions of the egg cylinder and pin the egg
cylinder down on the petri dish with another pair of forceps positioned immediately proximal to the
first pair. Reichert’s membrane is then torn by gently bringing the dissecting forceps toward the distal
tip of the egg cylinder. Once reflected over the tip of the cylinder, Reichert’s membrane will retract on
its own accord onto the extra-embryonic portion of the egg cylinder. The loosened membrane is then
trimmed away from the edge of the ectoplacental cone to prevent the embryo from sticking to the
cultural vessel or to another embryo during culture. Embryos can be kept in DMEM+10%fetal calf
serum for 45 minutes at room temperature without any significant effect on developmental potential,
however it is preferable to transfer dissected embryos into pre-equilibrated culture medium.
Dissection of early organogenesis stage (8.5-9.5dpc) + diagrams
These embryos are at least 3 times the size of the egg cylinder and now have an expanded visceral
yolk sac and amniotic cavity. Care should be taken not to compress the fluid filled yolk sac and
amnion or to damage the epithelial lining as any leakage of fluid will cause the yolk sac to collapse.
Therefore the technique involved is slightly different to 6.5dpc and7.5dpc embryos as it requires more
of a pinching and slicing mechanism as well as more patience.
The first step is to isolate the 8.5-9.5dpc decidua from the uterus and this can be done in a similar
manner to 6.5-7.5dpc embryos. Begin the dissection of the decidual mass (in DMEM + 10% fetal calf)
at the junction of the ectoplacental cone and the yolk sac junction. This region is distinguished by the
transistion from a red tissue mass to a clearer more translucent tissue which overlies the fluid filled
conceptus. Hold the decidua steady with one pair of forceps and pierce the decidual tissue with the
points of dissecting forceps. Carefully remove the dull red decidual tissue by cutting along the
equator. This is achieved by pinching off a piece of tissue with one pair of forceps and sliding the
arms of another pair of forceps against the first pair which creates a slicing or cutting mechanism. Do
not try to tear the tissue away with both pairs of forceps as this will lead to distortion and ultimately
collapse of the yolk sac.
Having removed the dull red tissue by cutting all the way around the equator (in a manner similar to
opening a can), follow the same principle along the longitudinal axis until the conceptus is finally free
of all adhering pieces of decidual tissue. The final step is to remove Reichert’s membrane and this
invloves pinching the membrane with both pairs of forcepts and carefully pulling the forceps apart to
expose the visceral yolk sac. Reichert’s membrane is then trimmed back to the edge of the
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ectoplacental cone and the embryo is ready for lineage tracing, grafting, exposure to exogenous
agents etc and in vitro culture.
4.2 Whole embryo culture
Mouse embryos are cultured in chemically defined media supplemented with rat serum (preferably) or
a mixture of rat and human foetal cord serum. The media is maintained at physiological pH (7.3-7.4)
and temperature 37oC. The level of O2 changes in accordance with the metabolic requirements of the
embryos at different developmental stages (see table1 below). The general consenus is that mouse
embryos develop best when they are explanted at the late primitive streak (7.5dpc) to early somite
stages (8.5dpc)in continuous gas exchange conditions. Generally in over 90% of embryos cultured
from these stages, tissue growth and morphological development is relatively normal for 36-48 hours.
8.5dpc embryos can be pushed to approximately 11.5dpc. However at this point developmental
retardation and morphological abnormalities occur. Postimplantation embryo culture can be
commenced as early as 6.5dpc and as late as 12.5 or 13.5dpc, however the older embryos (from
9.5dpc) can only be properly ustained for up to 24 hours.
Table 1. Conditions for mouse embryo culture
Age
Medium
O2 Level
6.5-7.5dpc
DRH
5%O2
IYS
static culture
7.5-8.5dpc
DR50
5%O2
IYS
roller culture
8.5-9.5dpc
DR50
5%O2
IYS
roller culture
9.5-10.5dpc
DR75
20-40%O2
IYS or OYS
roller culture
10.5-11.5dpc
R100*
40-65%O2
OYS
roller culture
11.5-12.5dpc
R100*
65-95%O2
OYS
roller culture
12.5-13.5dpc
R100*
95%O2
OYS
roller culture
D, DMEM; H, human cord serum; R, rat serum; *, supplemented with 2mg/ml glucose; IYS, intact yolk
sac; OYS, open yolk sac
DRH= 25%DMEM, 50%R, 25%H
DR50= 50%DMEM, 50%R
DR75= 25%DMEM, 75%R
R100= 100% R
4.3 Lineage tracing with DiI
Focal populations of cells can be labelled in cultured embryos using vital dyes DiI (red) and DiO
(green). These fluorescent dyes are lipid soluble but water insoluble such that they partition to cell
membranes and do not diffuse in culture medium. DiI (1,1′-dioctadecyl-3,3,3′,3′tetramethylindocarbocyanine perchlorate; Molecular Probes, Eugene, OR) is suspended in
Ethanol/sucrose and stored at 4oC, protected from light. Prior to labeling, a microdroplet is placed in
a petri dish, covered with Tyrodes or culture medium to prevent evaporation. Make a small focal
injection of DiI into desired tissue. Culture embryos as described above.
4.4 in vitro culture with morphogens or inhibitors or teratogenic agents such drugs or ethanol
Any number of exogenous agents (eg retinoic acid, FGFs, BMPs, dominant-negative receptors,
ethanol) at various concentrations can be added directly to the culture medium or injected into the
amniotic fluid in tests for teratogenicity or other effects.
For example, retinoic acid is a direct regulator of Hox gene expression. Culturing 8.5dpc Hox-lacZ
transgenic embryos in the presence of retinoic acid should result in an anterior expansion of the Hox
domain of expression and thereby posteriorize the hindbrain, neural crest cells, spinal cord and
somites. Remember to keep a control embryo to compare the final expression patterns between
retinoic acid treated and untreated embryos. All you have to do is culture the embryos overnight in the
presence of retinoic acid and assay the effects by X-gal staining for lacZ
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5. Detection of apoptotic cell death
Cell death has been proposed as a mechanism for the apoptotic elimination of cranial neural crest
cells from rhombomeres 3 and 5 in avians. In contrast, in the neural tubes of mice and zebrafish cell
death is extremely dynamic and does not appear to be specifically restricted to odd or even
rhombomeres. The differences maybe significant in terms of the evolution of muscle attachment sites.
Three cell death stain methods are described below. Each is based on detection of lysosomal activity
that occurs in response to the nuclear fragmentation which occurs early in the apoptotic programme.
We recommend either the acridine orange or LysoTracker methods but the protocol for Nile Blue
staining is included as an alternative.
5.1 “LysoTracker Red” staining for apoptotic cells in mouse embryos
Materials
LysoTracker Red (Molecular Probes, Eugene OR) stock 1mM in DMSO
HBSS medium
clearing agent BABB or
Dilute 2.5 ul LysoTracker stock solution into 0.5 ml HBSS to yield a final concentration of 5 uM.
Incubate 1-3 embryos for 30 minutes at 37oC in incubator with rocking. Alternatively, incubate 1 hour
in 2uM stain solution.
After stain incubation, wash embryo extensively in HBSS. Transfer to 2% Paraformaldehyde and fix
overnight at 4oC.
Wash several time with PBS to remove PFA, embryos can be stored in PBS at this step.
Visualize apoptosis staining using a fluorescent microscope
LysoTracker Red DND99 Abs 577nm, Em 590nm
For clearing of the embryos:
Dehydrate embryos via a series of methanol washes (15 minutes each). Start with 25%, then 50%,
70%, 95% and finally three washes in 100% methanol (v/v in distilled H2O). As the embryos
dehydrate through the methanol washes they become very brittle and fragile. Wash by removing
medium from vial with transfer pipette rather than manipulating embryos.
Optional: embryos can be cleared with 1:2 (v/v) benzyl alcohol:benzyl benzoate solution (BABB).
(WARNING- BABB is a powerful solvent that will damage microscopes if it is not carefully contained
in a glass dish or depression slide sealed with fingernail polish.) Incubate embryos in a solution of
1:1 MeOH:BABB. Replace with 100% BABB and incubate for at least 30 minutes. Remove most of
BABB solution and carefully transfer embryos with transfer pipette to slide.
14
5.2 Acridine orange staining for apoptotic cells in mouse embryos
Materials
Acridine orange stock solution of 5mg/ml in water
Culture medium such as DMEM (+ 10% Foetal Bovine Serum)
Procedure
Dissect mouse embryos (7.5-10.5dpc) in DMEM removing all extra-embryonic membranes.
Incubate embryos in a 5ug/ml acridine orange solution in DMEM at 37oC for approximately 30-40
minutes. Monitor the embryos every 15-20 minutes and avoid stopping the reaction too early.
Although the embryo will have a faint background, the cell death staining should be intensely blue.
Stop the reaction by washing the embryos in DMEM for 5 minutes.
Fix the embryos in 4% paraformaldehyde in PBS to prevent tissue degradation and analyse the
specimens immediately under rhodamine epifluorescence. This is a vital stain which will fade
overnight
5.3 Nile blue staining for apoptotic cells in mouse embryos
(this technique can be difficult to get to work properly, but we include it here for the sake of
completeness.)
Materials
Nile Blue stock solution 10% in water, culture medium such as DMEM (+10% serum)
Procedure
Dissect mouse embryos (7.5-10.5dpc) in DMEM removing all extra-embryonic membranes where
possible.
Add 2-5ul of Nile Blue stock to 2ml of DMEM (sufficient for 10 embryos, final solution should be deep
blue in colour)
o
Incubate embryos in NileBlue/DMEM at 37 C for approximately 30-45 minutes with shaking if
possible. Monitor the embryos every 15 minutes and avoid stopping the reaction too early. Although
the embryo will have a faint background, the cell death staining should be intensely blue.
Stop the reaction by washing the embryos in DMEM for 5 minutes.
It is best to photograph the embryos as soon as possible as nile blue stain will fade. However if you
want to photograph later rather than sooner, fix and store the embryos in 4% paraformaldehyde
overnight. These fixed embryos can then be processed for in situ hybridisation although the nile blue
staining will be subsequently lost.
15
6. Visualizing vasculature by injection of India Ink
Reagents and equipment:
PBS or Tyrodes
India ink
spin down to pellet larger carbon particles
dilute supernate 1/10 in PBS or Tyrodes
Warming plate or incubator at 37oC
Mouth pipette
Embryos should be harvested quickly and carefully so that ink can be injected while the circulation is
still active. Therefore, prepare equipment, ink and pre-warm PBS or Tyrodes prior to euthanizing
pregnant female.
Euthanize pregnant female and quickly dissect out embryos with yolk sacs intact.
Place embryos immediately into Petri dish with warm PBS or Tyrodes. Keep at 37oC on warming
plate or incubator.
Inject ink very slowly and gently into a branch of the vitelline vein of the yolk sac. It may be helpful to
cut into the vein with a ~26gauge hypodermic needle prior to inserting mouth pipette.
The carbon particles in the ink will be distributed through the arterial system by the beating heart.
When the desired labeling is accomplished, dissect embryo from yolk sac and fix in 4%
paraformaldehyde.
Embryos may be cleared in BABB or glycerol.
7. Detection of primordial germ cells
Primordial germ cells (PGC’s) arise in the vicinity of the posterior end of the primitive streak region
around 7.5dpc. PGC’s will accumulate towards the base of the allantois, then migrate along the
hindgut into the mesentry at the level of the genital ridges before colonising the genital ridges. The
number of PGC’s increase from approximately 100 at 7.5dpc to nearly 25000 by 12.5dpc. The most
convenient marker for assaying PGC’s and their proliferation is the tissue non-specific alkaline
phosphatase stain.
Materials
4% paraformaldehyde
alkaline phosphatase staining solution (prepare fresh as detailed below)
NBT/BCIP (see in situ hybridisation protocol)
NTMT (see in situ hybridisation protocol)
Procedure
Dissect embryos and wash in PBS.
Fix in 4% PFA, 4 degrees, 15 minutes for 7.5dpc, 20 minutes for 8.5dpc,
25 minutes for 9.5dpc
Wash 3x in NTMT (see protocols for in situ hyb), 10 minutes each, room temp.
Stain in NBT/BCIP mix in alkaline phosphatase buffer
Germ cell stain should appear in 15 minutes.
16
8. X-gal staining of lacZ transgenic embryos
STAIN FOR BETA-GAL EXPRESSION In Tissue
using prepared reagents from Specialty Media (Chemicon)
Reagents Required:
1. Tissue Fixative - Catalog #BG-5-C (100ml)
2. Tissue Rinse Solution A - Catalog #BG-6-B (500ml)
3. Tissue Rinse Solution B - Catalog # BG-7-B (500ml)
4. Tissue Stain Base Solution - Catalog #BG-8-C (100ml)
5. X-Gal Stock Solution - Catalog #BG-3-G (1ml)
Protocol
Complete B-Gal Tissue Stain Solution must be prepared FRESH for each staining.
Prepare as follows:
A. Warm the volume of Tissue Stain Base Solution needed for the experiment
in a 37°C water bath, keeping it in the DARK.
B. Thaw the X-Gal Stock Solution. Make a 1:40 dilution of the X-Gal Stock Solution
in warm Tissue Stain Base Solution, mix with a pipet and hold in the DARK
at 37°C until used. Dilution of X-Gal Stock Solution into cold Tissue Stain
Base will cause the X-Gal to precipitate.
Fix tissue in Tissue Fixative for 45 minutes to 1.5 hours on wet ice.
Rinse tissue in Tissue Rinse Solution - A.
Wash tissue in Tissue Rinse Solution - A for 30 minutes at room temperature.
Rinse tissue in Tissue Rinse Solution - B.
Wash tissue in Tissue Rinse Solution - B for 5 minutes at room temperature.
Drain tissue and add COMPLETE B-Gal Tissue Stain Solution. Incubate at 37°C in the dark. Stain
should be visible after 1 hour, but may need a longer incubation period (overnight is OK).
17
9. Organ cultures of explanted organ and tissue
Many tissues and organs can be cultured in vitro for several days. In some cases the tissues or
organ morphology is retained and develops in culture, in other cases morphology is lost but
development and differentiation can be assessed by in situ hybridization or immunohistochemistry.
Examples of organs and tissues that can be cultured include lung bug, allantois, gential ridge, kidney,
heart, intestine, forebrain, and branchial arches. Explants can be treated with morphogens, inhibitors,
teratogens or drugs. Some tissues requires several days of culture to allow for development of
relevant structures.
General organ explant culture protocol:
Here we present a generalized protocol for culture or organs or embryo fragments. Optimal
conditions for culturing each specific tissue or organ type usually vary from the following protocol.
Isolate organ or tissue explant fragment and remove unwanted excess tissue by dissection in
Tyrodes or culture medium. Use care to avoid contamination and rinse tissues in sterile Tyrodes or
culture medium prior to transfer to culture dish. Prepare culture chamber by equilibrating medium
o
(DMEM:F12 + Glutamax/ 15% FCS plus antibiotics) and nucleopore filter insert at 37 C with 5%CO2.
Use 2ml medium per well of 6-well plate. Filter membrane should be completely wet but organ
explant tissues should not be submerged. Multiple isolated organs can be cultured on a single filter
insert.
Examples of organs and embryo fragments suitable for explant organ culture
suggested
morphogens,
antagonists
things you could look for
suggested
culture
duration
lung bud
E10.5 –
E11.5
retinoic acid
FGFR inhibitor
branching morphogenesis
24-72 hours
allantois
E7.5
Notch inhib
VEGFR inhib
vasculogenesis (PECAM1 Ab)
24-72 hours
gut
E8.5E9.5
E11.5
retinoic acid
pancreas morphogenesis
neurogenesis (2H3 Ab)
2+ days
genital
ridge/gonad
E11.5E16.5
retinoic acid
Male gonad T-spermatogonia
in mitotic arrest
1+ days
metanephros
E11.5E12.5
E8.5E9.5
FGFR inhibitor
formation of uteric branches
2-5 days
Wnt inhibitor
Hedgehog inhibitor
cartilage (Alcian blue)
tooth condensations
2-10 days
E11.5
E14.5
Hedgehog inhibitor
formation of cochlear loop
morphogenesis, neurogenesis
1-3 days
branchial arches
otocyst
18
Isolation of lung buds for explant culture can be isolated from e10.5 or e11.5 embryos. Cut
embryo transversely at branchial arches and interlimb region to remove head and lower trunk.
Expose lung buds by carefully moving heart aside from thoracic cavity. Lung bud explants usually
develop good branching morphology after 24-72 hours of culture.
Branchial arch explant culture
Isolate mandible and maxillary primordia (as shown in figure) from embryos at e9.5–e11.5.
Gonad organ cultures
Collect mouse embryos between 11.5 and 16.5 dpc from (1) a wild type and (2) a transgenic strain
expressing a constitutive marker in all its cells such as b-gal (Gt(ROSA)26Sortm1Sor/J) or EGFP
(FVB.Cg-Tg(GFPU)5Nagy/J). You will need to know the sex of embryos from the wild type strain. For
sexing embryos at 11.5 dpc, before sexual dimorphism is apparent, see amnion prep below. This is a
quick method to identify XX samples by the presence of cells with a condensed chromatin body. Save
a bit of each embryo’s tail for later confirmation by PCR.
Dissect the entire urogenital complex in association with the dorsal aorta from all embryos and place
in individual wells in warmed Dulbecco’s Minimal Eagle’s Medium (DMEM)/10% fetal calf serum
–1
(FCS) (Gibco)/50 mg.ml ampicillin and keep at 5% CO2 at 37°C. Once you have sexed the wild type
samples, begin fine dissection and assembly. Using a 27 gauge needle, separate the mesonephroi
from the urogenital complexes of the marked embryos and place the mesoneprhoi in prepared
grooves cast in 1.5% agar blocks (see protocol below).
Next separate mesonephroi and gonads from the wild type sexed urogenital complexes. It is easier to
handle one pair at a time, immediately assembling gonads with a mesonephros already positioned on
the block. Work with a pulled glass pipette of a size just larger than the gonad diameter. Removing
medium from the grooves will promote adhesion between the mesonephroi and gonads. After
assembly is complete, incubate agar blocks in 35 mm tissue culture dishes with 400 ml DMEM/10%
FCS/ 50 mg.ml–1 ampicillin medium added to the bottom of the plate at 37°C in 5% CO2.
Replacement of medium each day may help with contamination problems. Cultures maintain good
morphology for at least 48 hours.
19
10. Tissue explants cultured under the kidney capsule for long term
differentiation analyses
For this protocol see the attached pages below copied from Manipulating the Mouse Emryo: a
Laboratory Manual 3rd edition (Eds: Nagy, A., Getsenstein, M., Vintersten, K., Behringer, R.) Cold
Spring Harbor Laboratory Press, from which the following protocol was copied.
1. Weigh and anesthetize the recipient mouse
2. Wipe the back of the recipient mouse with 70% ethanol. Shave fur if required by local legislation.
Then make an incision ~1-cm long as for embryo transfer (see p. 254). Slide the incision to one side
and cut the body wall just above the level of the ovary. Use blunt fine forceps to pull out the kidney
by is fat pad. Immobilize the kidney in Desmarres chalazion forceps (Fig. 6.5). Allow the surface of
the kidney to air-dry for a few minutes. (This enables the capsule to be picked up with the
watchmaker’s forceps.)
3. Use the watchmaker’s forceps to make a small transverse tear in the exposed capsule membrane
and then moisten the capsule with sterile
saline of PBS. Use moistened
watchmaker’s forceps to make a pocket
underneath the capsule and then use
forceps or a pipette to insert the tissue to
be grown. The function of the braking
pipette is to prevent the backflow of
blood into the pipette, which could result
in the pipette opening becoming
clogged.
4. Push the tissue underneath the
capsule as far away from the tear
as possible. Release the kidney
from the Desmarres chalazion
forces and replace it in the body
cavity with blunt fine forceps.
Sew up the body wall with one
or two stitches and close the
skin with wound clips.
5. At the end of the
procedure, place the mouse
in a clean cage and keep it
warm under a 50W light
bulb (taking care to cover
the eyes) or by placing
the cage on a warming
plate or heating pad
until the mouse
recovers from an
injected anaesthetic.
Follow local animal
care committee
regulations for
more details.
20
11. Mouse embryo DNA electroporation
11.1 In vitro mouse embryo DNA electroporation
Dissect embryos as above in Tyrodes or DMEM (+HEPES), leaving the necessary membranes intact.
Place dissected embryos into one of the culture bottles containing DR50. Insert the bottle and its
rubber holder into one of the slots on the roller in the BTC whole embryo culture chamber. Leave the
embryos for a few hours to recover from the dissection.
It is not necessary to mix your DNA with fast green but it sure helps visualising just how much DNA
you have injected and where. There have been some concerns raised about whether fast green
inhibits successful electroporation of morpholinos and if in doubt phenol red is another alternative to
labelling your solution. DNA concentrations for mouse electroporation should be at least 2ug/ul, but
even higher is preferable. Too high ie 5ug/ul can be toxic to the embryos and too viscous, so you may
need to test this in your own lab situations.
For electropotation of the neural plate, depending on the age of the embryos you need to inject either
the amniotic cavity in 8.5dpc embryos (open neural plate) or the closed neural tube in 9.5dpc
embryos with as much DNA as possible but try not to distort the entire shape of the embryo (it will
burst like a balloon).
Electroporation in mouse embryos seems to work a little less consistently than in chick embryos and
other non-mammalian species. This may be due to a whole host of reasons including time
dependencies, morphological and structural differences and less receptiveness to DNA uptake in
mammalian embryos.
The optimum time for neural plate electroporation seems to be within the 1-3 somite stage. Therefore
we recommend isolating your embryos in the afternoon and electroporating them in the evening.
Prepare your injection needles with your DNA/fast green mix (aliquots kept on ice) before you remove
your embryos from the incubator. You can use any of the same plasmids that you have been using
for chick experiments.
The injection needles are pulled on the horizontal pipette puller and you can break the needle tip with
a pair of forceps. Remove embryos from the culture chamber and inject the DNA into the desired
location (ie cavity or node, or endoderm, mesoderm, ectoderm, somites etc). Quickly orient the
embryo between the electrodes and zap.
The following conditions are used for mouse embryo electroporation
20V, 50ms pulse width, 1 second gap between pulses and 5 pulses. You can try higher voltages but
beware of frying your embryos. You may see blebbing in the endodermal or yolk sac. This is normal if
it is minor and should not be detrimental to your embryo culture.
21
11.2 In vivo/ex utero mouse embryo DNA electroporation
This protocol describes a basic method for in vivo electroporation in the nervous system of embryonic
mice. Delivery of electric pulses following microinjection of DNA into the brain ventricle or the spinal
cord central canal enables efficient transfection of genes into the nervous system developing mouse
embryos.
For a detailed description of this protocol, please refer to the following reference:
http://www.nature.com/nprot/journal/v1/n3/abs/nprot.2006.276.html
In vivo electroporation in the embryonic mouse central nervous system
Tetsuichiro Saito (2006) Nature Protocols 1, - 1552 - 1558 (2006)
12. Interspecies transplantations (somites, neural tube, neural crest,
rhombomeres etc)
Transplanting specific tissues between different species can shed a lot of light on the evolution of cell
types. For instance, avians lack teeth, however, teeth can be generated in an avian host by
homotopically transplanting first arch neural crest cells from mice.
What happens if you transplant anterior somites that generate the forelimbs in mice into avian hosts.
Do you get a wing, a leg, a weg or a ling?
13. Transgenic and Mutant Mouse lines
Hoxb2-lacZ (labels rhombomere 4, second arch neural crest cells, trunk neural tube and somites)
Hoxa2-lacZ (labels rhombomeres 3 and 5, second arch neural crest cells and somites
Pax3-GFP a targeted knock in of GFP into the Pax3 gene. In this line GFP is expressed in the neural
crest, dorsal neural tube, somites and migratory muscle progenitor cells.
Tcof mutant mice. The Tcof1/Treacle protein is required for formation and proliferation of neural crest
cells. Mice heterozygous for mutations in the murine Tcof1 gene exhibit distinctive phenotypes
reminiscent of human Treacher-Collins syndrome. Neonates have severe craniofacial malformations
owing to apoptosis of neural crest cells. (note: these phenotypes are observed on a mixed
background of DBA and C57BL/6, other backgrounds show more severe phenotypes.)
22
14. Snakes and Lizards
Inland Bearded Dragon
Pogona vitticeps
Family: Agamidae
Size: adults are typically 18-23 inches in length, weighing at least 250 g; hatchlings are just under 4
inches in length
Longevity: 8 - 10 years in captivity
Habitat: hot, arid regions from dry forests and scrublands to sandy deserts
Food: omnivores; adults consume predominantly vegetable matter, preferring soft leaves and flowers
when available; a voracious insect predator
Behavior: diurnal, largely terrestrial lizards. Bearded dragons exhibit a wide range of social
interactions including arm-waving, head-bobbing and tail curling. They can inflate a beard-like
extension of the throat in mating displays and when threatened.
Reproduction: Sexual maturity is reached between 1 and 2 years of age. Mating and breeding occur
in spring and summer months. Females can store sperm. They lay many clutches in one season of 8
to 30 eggs each time. Eggs hatch from 55 to 75 days depending on temperature.
Range: widely distributed in Australia throughout the non-coastal areas of the eastern states, through
the eastern half of south Australia and north to southeastern Northern Territory
23
Desert Grassland Whiptail
Aspidoscelis uniparens
(formerly Cnemidophorus uniparens)
Family: Teiidae
Size: 16 - 24cm; head to body max. 7 cm
Identification: black to reddish brown having 6 or 7 light longitudinal stripes; no spots; slender lizard
with narrow head; tail bluish-green to olive-green; blue in juveniles
Habitat: desert and mesquite grassland, ascending river valleys into lower mountainous areas
Food: mostly insects, especially termites
Active: April to October; alert, active forager, basking morning and evening. On cloudy days may be
active all day.
Hibernation: this species hibernates in underground burrows during the colder months
Reproduction: all-female parthenogenetic species. Oviparous. Clutches of 1 to 4 eggs laid June to
August, hatching after 50-55 days. In captivity, eggs take approximately 58 days when incubated at
28ºC. Pseudocopulatory behavior has been observed in some females.
Range: central Arizona to extreme West Texas, central to SW New Mexico and south into Mexico
Status: locally common within range
24
Pantherophis guttatus
(formerly Elaphe guttata guttata)
Corn Snake
Family: Colubridae
Size: usually 30-48 inches (76 to 122 cm) with a record of 72 inches (183 cm)
Identification: checkered ventral pattern with black on white; ground color varies from grey to brown to
orange; orange-red dorsal blotches outlined in black; weakly keeled scales
Habitat: fields, pine barrens, woody lots, rocky hillsides; frequently found close to farming and
habitation, or in abandoned buildings
Food: non-venomous constrictors; hatchlings feed largely on small lizards and tree frogs; adults on
rodents and birds
Behavior: climbs well, but mostly terrestrial; crepuscular, preferring to be active around dusk; often
found resting or prowling in rodent burrows
Reproduction: mature at 2 to 3 years of age; mating takes place from March to May and eggs are
generally laid 31 to 45 days after mating, 12-18 being most common. In the wild, corn snakes deposit
their eggs out of sight in holes, piles of leaves, or in dead trees.
Incubation: a range of temperatures from 21ºC to 32ºC is well tolerated and eggs incubated at 28ºC
hatch in approximately 60 days.
Range: southern New Jersey through Florida and the Keys, and west to Louisiana
25
Chihuahuan Spotted Whiptail
Aspidoscelis exsanguis
(formerly Cnemidophorus exsanguis)
Family: Teiidae
Size: 24 - 31cm; head to body max. 10 cm
Identification: brown or red-brown having 6 light longitudinal stripes overlaid with pale yellow to white
spots; slender lizard with narrow head; tail blue-grey to green
Habitat: desert grassland to mountain woodlands; often found in canyon bottoms or dry washes
Food: Carnivorous: lepidopteran larvae, ants, beetles, spiders, scorpions, grasshoppers; often
observed foraging around shrubs and vegetative litter for insects
Active: April to October; alert, active forager, basking morning and evening.
Hibernation: this species hibernates in underground burrows during the colder months
Reproduction: all-female parthenogenetic species. Oviparous. Clutches of 2 to 6 eggs depending on
the size of the mother laid June to August. Offspring are genetically identical to the mother, appearing
about a month and a half after laying. In captivity, eggs take approximately 62 days when incubated
at 28ºC.
Range: extreme SE Arizona and central New Mexico south into West Texas and Mexico
Status: locally common within range
26
15. Inhibitors, Antagonists and Antibodies
Inhibitors:
1. VEGF receptor 2 kinase inhibitor III
Calbiochem
www.emdbiosciences.com
Cat#676487
1 mg
Comments: M.W. 238.3.
Make stock at 15 mM in DMSO and use at 10-20 uM in culture overnight.
2. CKI-7 casein kinase (Wnt inhibitor)
US Biological
Cat#P9104
www.usbio.net
Comments: M.W. 285.74. Soluble in MeOH or EtOH.
We made in 100% EtOH at concentration of 50 mM and use at concentration of 200 uM.
3. DAPT (Notch inhibitor)
Sigma-Aldrich
Cat#D5942-25MG
25 mg
Comments: Resuspend in DMSO at concentration of 50 mM and use at 100 uM.
4. SU5402 (FGFR inhibitor)
Calbiochem
www.emdbiosciences.com
Comments: M.W. 269.3. Make stock at 25 mM in DMSO, use at 25-120 uM.
Cat#572630
500 ug
6. Noggin (BMP anatagonist)
R&D Biosystems
Cat#719-NG-050
Comments: Suspend in PBS with at least 0.1% BSA, stock concentration 1mg/ml,
use in culture in range of 150ng/ml to 1ug/ml
7. Cyclopamine (Shh inhibitor)
Toronto Research Chem
Cat#C988400
1 mg
Comments: suspend in 100% Ethanol, use at 5uM final
Antibodies
Neurofilament Ab (2H3)
rat brain membrane
2
nd
Ab: donkey anti- mouse HRP
Vascular-endothelial Antibody PECAM1
Rat anti-mouse
2nd Ab: goat anti-rat Cy3
University of Iowa
Cat#2H3
Jackson Immuno
Cat#715-035-150
BD Biosciences Cat#553370
0.5 mg/ml
Jackson Immuno
27
Cat#112-165-167
0.5 mg
16. Mouse Development references
16.1. Mouse Development timeline
For a detailed description of mouse development see Chapter 2 from Manipulating the Mouse
Embryo: a Laboratory Manual 3rd edition (Eds: Nagy, A., Getsenstein, M., Vintersten, K., Behringer,
R.) Cold Spring Harbor Laboratory Press, from which the images below were copied
28
16.2 Whole mount embryos at representative stages of development
29
16.3 Mouse Embryo Staging Criteria – from Edinburgh Mouse Atlas website
The Edinburgh Mouse Atlas, from which the following pages are copied, is an excellent resource for
information about mouse embryology.
http://genex.hgu.mrc.ac.uk/Atlas/intro.html
Mouse embryos can be staged according to a variety of criteria, the most general of which are those
described by Theiler in "The House Mouse: Atlas of Mouse Development" (Springer-Verlag, New
York, 1989). Theiler's criteria are too broad to distinguish many of the important phases of early
development and must therefore be supplemented by others, for example, cell number, somite
number or those charcteristics used by Downs and Davies (1993) Development, 118, 1255. We have
therefore combined these different criteria in the table1 below which defines a new set of stages
based on the numbered Theiler series, but with intermediate divisions indicated by non-integer stage
numbers. Embryos of the same gestational age may differ in their stage of development. We have
therefore included in the table an indication of the expected range of gestational ages (days of
gestation, dpc) over which each developmental stage may be found. Different mouse strains develop
at different rates and, in some cases show differences in the relative rates of development of different
organs. Strictly, the stages recognised by Downs and Davies apply to outbred mice of the PO strain.
The data in the remainder of the table below refer to embryos of crosses between F1 hybrid (C57BL
X CBA) mice.
Each Theiler stage is linked (click on the number) to its corresponding diagram, with more details of
the defining features for that stage. A brief text or pictorial index to the diagrams is also provided.
Somite Cell
Theiler
(C57BLxCBA)F1 mice4
dpc (range)2
number
No.3
Stage
PO mice5
1
0-0.9
(0 -2.5)
1
One-cell egg
2
1
(1 -2.5)
2-4
Dividing egg
3
2
(1-3.5)
4-16
Morula
4
3
(2-4)
16-40
Blastocyst, Inner cell mass apparent
5
4
(3-5.5)
Blastocyst (zona-free)
6
4.5
(4-5.5)
Attachment of blastocyst, primary endoderm covers
blastocoelic surface of inner cell mass
7
5
(4.5-6)
Implantation and formation of egg cylinder Ectoplacental
cone appears, enlarged epiblast, primary endoderm lines
mural trophectoderm
8
6
(5-6.5)
Differentiation of egg cylinder. Implantation sites 2x3mm.
Ectoplacental cone region invaded by maternal blood,
Reichertís membrane and proamniotic cavity form
9
6.5
(6.25-7.25 )
Pre-streak (PS), advanced endometrial reaction,
ectoplacental cone invaded by blood, extraembryonic
ectoderm, embryonic axis visible,
PS
Early streak (ES), gastrulation starts, first evidence of
mesoderm
ES
9a
30
10
7
(6.5-7.75)
Mid streak (MS), amniotic fold starts to form
MS
10a
Late streak, no bud (LSOB), exocoelom
LS
10b
Late streak, early bud (LSEB), allantoic bud first appears,
node, amnion closing
Neural plate (NP), head process developing, amnion
complete
OB
11a
Late neural plate (LNP), elongated allantoic bud
EB/ LB
11b
Early head fold (EHF)
EHF
11c
Late head fold (LHF), foregut invagination
LHF
11
12
7.5
(7.25-8)
8
(7.5-8.75 )
12a
1-4
1-4 somites, allantois extends, 1st branchial arch, heart
starts to form, foregut pocket visible, preotic sulcus (at 2-3
somite stage)
5-7
5-7 somites, allantois contacts chorion at the end of TS12
Absent 2nd arch, >7 somites
13
8.5
(8-9.25)
8-12
Turning of the embryo, 1st branchial arch has maxillary
and mandibular components, 2nd arch present
Absent 3rd branchial arch
14
9
(8.5-9.75)
13-20
Formation & closure of ant. neuropore, otic pit indented
but not closed, 3rd branchial arch visible
Absent forelimb bud
15
9.5
(9-10.25)
21-29
Formation of post. neuropore, forelimb bud, forebrain
vesicle subdivides
Absent hindlimb bud, Rathke's pouch
16
10
(9.5-10.75)
30-34
Posterior neuropore closes, Formation of hindlimb & tail
buds, lens plate, Rathke's pouch; the indented nasal
processes start to form
Absent thin & long tail
17
10.5
(10-11.25)
35-39
Deep lens indentation, adv. devel. of brain tube, tail
elongates and thins, umbilical hernia starts to form
Absent nasal pits
18
11
40-44
(10.5-11.25)
Closure of lens vesicle, nasal pits, cervical somites no
longer visible
Absent auditory hillocks, anterior footplate
19
11.5
(11-12.25)
45-47
Lens vesicle completely separated from the surface
epithelium. Anterior, but no posterior, footplate. Auditory
hillocks first visible
Absent retinal pigmentation and sign of fingers
20
12
(11.5-13)
48-51
Earliest sign of fingers (splayed-out), posterior footplate
apparent, retina pigmentation apparent, tongue well-defined,
brain vesicles clear
Absent 5 rows of whiskers, indented anterior footplate
21
13
(12.5-14)
52-55
Anterior footplate indented, elbow and wrist identifiable, 5
rows of whiskers, umbilical hernia now clearly apparent
Absent hair follicles, fingers separate distally
22
14
(13.5-15)
56-~
60
Fingers separate distally, only indentations between digits
of the posterior footplate, long bones of limbs present, hair
follicles in pectoral, pelvic and trunk regions
Absent open eyelids, hair follicles in cephalic region
31
23
15
Fingers & Toes separate, hair follicles also in cephalic
region but not at periphery of vibrissae, eyelids open
Absent nail primordia, fingers 2-5 parallel
24
16
Reposition of umbilical hernia, eyelids closing, fingers 2-5
are parallel, nail primordia visible on toes
Absent wrinkled skin, fingers & toes joined together
25
17
Skin is wrinkled, eyelids are closed,umbilical hernia is
gone
Absent ear extending over auditory meatus, long whiskers
26
18
Long whiskers, eyes barely visible through closed eyelids,
ear covers auditory meatus
27
19
Newborn Mouse
28
Postnatal development
Bard, J.B.L., Kaufman, M.H., Dubreuil, C., Brune. R.M., Burger, A., Baldock, R.A., Davidson, D.R.
(1998). An internet-accessible database of mouse developmental anatomy based on a systematic
nomenclature. Mechanisms of Development, in press.
Days post conception, with the morning after the vaginal plug is found being designated 0.5 dpc (or
E0.5). For detailed discussion see Kaufman (1994). The Atlas of Mouse Development (2nd printing),
pp. 515-525. London: Academic Press.
The figure given refers to the number of the most caudal somite. No account is taken of somites
partitioning into dermomyotomes and sclerotomes, nor of their subsequent differentiation.
Adapted from Theiler (1989) [The House Mouse: Atlas of Embryonic Development. New York:
Springer-Verlag] and Kaufman (1994); detailed staging for Theiler stages 9-12 courtesy of K. Lawson
[personal communication].
From Downes, K.M. and Davies, T. (1993). Staging of gastrulating mouse embryos by morphological
landmarks in the dissecting microscope. Development, 118, 1255 - 1266.
Comments
General comment on timing (dpc): In judging the lower and upper ranges of dpc equivalent to a
particular Theiler stage, we have generally followed Theiler's book and, in most cases, have given a
wider range than Theiler, because the numbers of embryos he cites are small. We have given a
larger range at the maximum than the minimum because, in general, embryos are more likely to be
retarded by their environment or genetic constitution than made to proceed more quickly through
development. In most cases, however, the resulting dpc range is an estimate that is consistent with
the results of Theiler, but not based on additional evidence.
Comment on somite numbers: The range of somite numbers for each stage is given only as a guide
to what might be expected of typical embryos. As can be seen from Theiler (1989)4 the true range can
be much wider. Therefore, for all stages after TS12, the somite number should not be taken as a
reliable global indicator of the overall embryo stage.
Richard Baldock, Jonathan Bard, Duncan Davidson and Kirstie Lawson, 7th May 1998
Please mail comments to Jonathan Bard
Dr. D. R. Davidson & Dr. R. A. Baldock (MRC
project leaders)
Dr. J. B. L. Bard & Prof. M. H. Kaufman (Dept.
Anatomy)
Dr. Richard Baldock (Web/DB problems)
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Document last modified:Friday, 25-Oct-2002 14:11:27 BST
32
Sample photographs of results obtained in the Trainor laboratory using the protocols
contained in this book.
33