Sperm
Cryopreservation
Protocol
Cryopreserving Sperm
Applications
• Sperm cyopreservation is a quick and inexpensive method to cryopreserve strains
• Sperm cryopreservation only requires 2 sperm donor mice
• It is best used for cryopreserving strains on common inbred backgrounds
• Heterozygous and/or wild type mice can be produced from cryopreserved sperm
Key Points
• Perform dissections and sperm collection as quickly as possible.
• Use male mice at 10-14 wks of age.
• Handle loaded straws carefully. If a straw is jostled and the sperm aliquots combine,
discard the straw and fill a new one.
• Follow laboratory safety practices when handling liquid nitrogen (LN2).
• Instead of using a Styrofoam box for freezing, you may use any apparatus that will
cool sperm from −10 to −60°C at a rate of 37±1°C/min.
• Do not allow cassettes to fall into LN2 prior to completing the 10 minute exposure
to LN2 vapor. If this occurs, these samples should be discarded.
• Use a new tissue culture dish and a new 16-gauge needle for each strain, and clean
dissection tools with 70% EtOH between strains to prevent cross contamination of
sperm samples.
Materials
• Dissecting instruments:
o Sharp-sharp 3” scissors
o Sharp-sharp 5” scissors
o 2 pair of micro-dissecting forceps
o 2 pair of Dumont forceps #5
• Styrofoam box with 3.6 cm thick walls (27.5 cm x 22.6 cm x 27.7 cm)
• Styrofoam box lid
• Styrofoam raft (polystyrene insulation board,18.3 cm x 13.4 cm x 3.6 cm)
• 37°C slide warmer
• 37°C water bath
• LN2 personal protective equipment
• 5 mm tissue culture dishes
• 0.25 ml French straws
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Cassettes for straws
Large forceps (25 cm)
Data sheet
Liquid Nitrogen (LN2)
Timer
16 gauge needles and 1 ml syringe
1 ml pipettor and tips
Monoject syringes (1 cc TB regular Luer tip)
10X dissecting microscope with transmitted light base
Heat sealer or Critoseal™ straw sealing powder
2 male mice (10-16 weeks of age) per strain
Cryoprotective medium (CPM)
70% Ethanol (EtOH) in spray bottle
2
Method
1. Prepare straws (Figure 1):
a. Draw one line 0.6 cm from the open end of the straw.
b. Draw a second line 1.5 cm from the open end of the straw.
c. Label straws adjacent to the cotton plug.
2. Label the cassettes with the strain ID and any other identifying information.
0.6 cm
1.5 cm
Figure 1. Marks are made on the straw at 0.6 cm and 1.5 cm from the open end of the straw.
3. Fill the Styrofoam freeze box to the fill line (9 cm from bottom of box) LN2
(Figure 2).
4. Make a sperm dish by pipetting 2 ml of warmed (37°C) cryoprotective medium
(CPM) into a 35 mm dish (if only one male is available for cryopreservation,
decrease the amount of CPM in the sperm dish to 1 ml).
5.
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Place the Styrofoam raft in the freezing box so that it floats on top of the LN2 and
replace the lid.
3
A
B
Figure 2. Sperm Freezing Apparatus. A. The sperm freezing apparatus includes a Styrofoam box, lid and raft. B. The Styrofoam box is filled with 9
cm of LN2 .
6. Euthanize two male mice.
7.
Collect sperm:
a. Remove both vasa deferentia and cauda epididymides from each male
(Figure 3).
b. Place the vas deferentia and cauda epididymides from both males into the
sperm dish (Figure 4a).
c. Make 7-8 cuts in each epididymis using a sterile 16-gauge needle or a pair of
fine dissection scissors (Figure 4b-c).
d. Squeeze sperm out of each vas deferens using the needle or Dumont forceps.
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Quick Guide for Dissecting Cauda Epididymis
70% EtOH
A
B
C
cauda epididymis
bladder
seminal vesicle
testicular
fat pad
D
caput
epididymis
testis
CUT HERE
E
vas deferens
F
blood vessel
CUT HERE
CUT HERE
G
H
I
Figure 3. A. Abdomen with 70% EtOH. B. Skin being pulled in opposite directions to expose peritoneum. C-D. Open body cavity exposing the
testicular fat pads. E. Pull on a testicular fat pad to expose the attached testical. F. Locate the cauda epididymis and vas deferens. Cut the ligament
adjacent to the cauda epididymis. G. Cut the vas deferens near the bladder. H. Gently pull the vas deferens away from the body cavity leaving behind
connective tissue and blood vessel. I. Cut just below the cauda epididymis to liberate the tissue.
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A
B
C
Figure 4. Sperm Collection. A. Vasa deferentia and cauda epididymides in CPM. B-C. Epididymis secured with forceps and cut with
a 16 gauge needle.
8. Incubate at 37°C for 10 minutes on the slide warmer.
9. Complete steps 10-13 in ten minutes or less.
10. Remove and discard tissues from the sperm dish and gently swirl the dish to
distribute sperm and create a sperm solution.
11. Immediately begin loading the straws:
a. Attach a Monoject syringe to the cotton/PVA plug end of a straw (Figure 5a).
b. Aspirate sperm solution to the second line on the straw (Figure 5a-I). This
creates ballast. CPM without sperm may also be used for ballast.
c. Aspirate air to the second line (Figure 5a-II).
d. Aspirate sperm solution to the first line, creating the first sperm aliquot (Figure
5a-III).
e. Repeat steps c and d, alternating aspiration of sperm and air, until you have
four individual sperm aliquots separated by air (Figure 5b-V through IX).
f. Aspirate air until the CPM ballast wets the cotton/PVA plug (Figure 5a-X).
g. Remove the straw from the syringe.
12. Seal the open end of each straw with the heat sealer or Critoseal™.
13. Place five straws into each of four cassettes, sealed end first (Figure 5c). Slide the
cassette plunger containing the straws into the clear sleeve until the cassette is
completely closed.
14. Cryopreserve the samples:
a. Position the cassettes on the raft in LN2 vapor (Figure 6) and
replace the box lid.
b. All the cassettes are to remain in the vapor at least 10 minutes, but not longer
than 30 minutes. Do not remove the lid or jostle the box during this time.
c. Use long forceps to tip the raft so the cassettes are plunged
directly into LN2.
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A
Syringe
I.
II.
III.
IV.
V.
VI.
VII.
VIII.
IX.
X.
B
C
Figure 5. Filling straws. A. Diagram of straw loading steps I-X showing aspirated CPM ballast, air and sperm samples relative to the marks on
the straw. B. Straw loaded with four sperm aliquots (separated by air pockets) and sealed with straw sealing powder. C. Straws being loaded into a
cassette, straw seal end first.
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d. Repeat steps a-c until all strains are cryopreserved, monitoring the
LN2 level in the freezing apparatus at the start of each newstrain.
e. Transfer cassettes into a LN2 storage tank. Be sure the straws do
not warm during the transfer.
Raft
Cassette
Figure 6. Freezing samples. Cassettes placed flat on top of the raft to cool in LN2 vapor.
Reference
Ostermeier GC, Wiles MV, Farley JS, Taft RA. 2008. Conserving, Distributing and
Managing Genetically Modified Mouse Lines by Sperm Cryopreservation. PLoS ONE
3(7): e2792 doi:10.1371/journal.pone.0002792
1
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Thawing Sperm
Applications
• Use cryopreserved sperm for in vitro fertilization to recover live mice
• Test a sample of a cryopreserved set to assess cryopreservation success
Key Points
• Follow laboratory safety practices when handling liquid nitrogen (LN2).
• Keep sperm straws submerged in LN2 until just prior to thaw.
• When transferring straws from long-term storage to a benchtop Dewar, do not
allow straws to be exposed to air for longer than 2 seconds.
Materials
• Liquid Nitrogen (LN2)
• 3L LN2 Dewar
• 37°C incubator with mixed gas (5% O2, 5% CO2, balanced with N2)
• 37°C water bath
• LN2 personal protective equipment
• Cryopreserved sperm samples in straws and cassettes
• 10X dissecting microscope with transmitted light base
• Forceps
• Small scissors
• Disposable wipes
• Metal push rod
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Method
1.
Fill the LN2 Dewar 90% full with LN2.
2. Quickly transfer the cassette(s) containing the sperm straws from the long-term
storage tank to the LN2 Dewar.
3. Using forceps to keep the cassette and remaining straws submerged in LN2, remove
a straw from the cassette.
4. Submerge the straw in the 37°C water bath until ice crystals are no longer visible
(approximately 10 seconds).
5. Remove the straw from the water bath and carefully dry the straw with a disposable
wipe.
6. Cut the ends of the straw with scissors (Figure 7).
7. Insert the metal push rod into the cotton plug end of the straw, and push gently to
dispense an aliquot of sperm.
8. If using the sperm for IVF, dispense one aliquot into each IVF dish fertilization
drop.
Insert Rod
in this end
Cut here
Cut here
Figure 7. Cutting straw. Insert metal push rod in the end with the cotton plug.
Reference
Ostermeier GC, Wiles MV, Farley JS, Taft RA. 2008. Conserving, Distributing and
Managing Genetically Modified Mouse Lines by Sperm Cryopreservation. PLoS ONE
3(7): e2792 doi:10.1371/journal.pone.0002792
1
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Preparing Hormones and Superovulating Mice
Applications
• Produce large quantities of oocytes from a female mouse.
• Produce oocytes for IVF.
• Mate superovulated females to male mice and collect embryos via flushing.
• Mate superovulated females to collect fresh sperm from naturally mated mice.
Key Points
• PMSG mimics FSH (follicle stimulating hormone) and causes follicles to grow.
• PMSG dosage is strain dependent (see “PMSG doses” below). 5 IU PMSG is likely
to work for most inbred strains, but 2.5 IU works best for F1 hybrids. Depending on
your source of hormone, higher doses of PMSG may be needed.
• PMSG injection time depends on the time oocytes are to be collected (see “Sample
Timeline” below).
• hCG mimics LH (lutenizing hormone) and stimulates ovulation.
• 5 IU hCG is commonly administered, although 2.5 IU may be sufficient to reach
the threshold necessary to cause ovulation.
Materials
• PMSG (pregnant mare’s serum gonadotropin) (Sigma G-4877)
• hCG (human Chorionic Gonadotropin) (Sigma CG10)
• Culture grade water
• Vials with caps
• 25G 5/8” needle
• 30G 1/2" needle
• 1ml syringe
• Mice (12-15g or 8 weeks and older)
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Method
1.
Prepare and store hormones:
a. Reconstitute lyophilized hormone to a concentration of 2.5IU per 0.1ml
b. Store in aliquots at -20°C for up to 3 months (store at -80°C for up to 6
months)
c. Once thawed, store at 4°C for up to 3 days
2. If using frozen hormone, thaw at room temperature, 4°C or in a 37°C water bath.
Do not thaw using hot water.
3. Invert vial slowly several times directly prior to use. Do not shake.
4. Draw hormone into syringe.
5. Inject mice intraperitoneally using a 30G needle with 2.5 IU to 5 IU PMSG.
6. 46-50 hrs after administration of PMSG, inject females with 2.5 IU or 5 IU hCG.
7. Collect oocytes 13-16 hours post-hCG.
PMSG Doses
2.5 IU
5.0 IU
NOD
C57BL/6J
BALB/cJ
129/Sv
F1 hybrids
C3H/HeSnJ
CBA/J
DBA/2J
FVB
SJL/J
Sample Timeline
Monday
Administer PMSG at
6:00pm
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Tuesday
Wednesday
Thursday
Administer hCG at 7:00pm
(46-50 hours after administration of PMSG)
collect clutches of eggs
between 8:00-11:00am
(13-16 hours after administration of hCG)
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Expected Response to Superovulation
Oocytes per female (mean ± SEM)
Strain
No. of females
treated with eCG
and hCG
No. COCs1
Total Oocytes2
Normal
Frag3
Dead
129S1/SvImJ
45
72
1650
39.5 ± 1.7 a
0.2 ± 0.7 b
1.6 ± 0.3 a
A/J
44
80
226
5.4 ± 2.0 f
0 ± 0.8 b
0 ± 0.3 b,c
BALB/cByJ
55
99
1999
33.7 ± 1.5 a
0.8 ± 0.6 b
1.9 ± 0.2 a
BALB/cJ
45
88
679
14.0 ± 1.6 e
0.7 ± 0.6 b
0.3 ± 0.3 b,c
C3H/HeJ
45
86
700
14.9 ± 1.6 d,e
0.5 ± 0.6 b
0.2 ± 0.3 b,c
C57BL/6J
40
80
1027
25.0 ± 1.7 b,c
0.3 ± 0.7 b
0.2 ± 0.3 b,c
DBA/2J
30
57
946
31.1 ± 2.0 a,b
0.4 ± 0.8 b
0 ± 0.3 b,c
FVB/NJ
43
84
838
18.9 ± 1.6 c,d,e
0.3 ± 0.6 b
0.1 ± 0.3 c
NOD/LtJ
45
90
1400
22.0 ± 1.6 c,d
7.7 ± 0.6 a
1.4 ± 0.3 a,b
SJL/J
45
80
560
12.4 ± 1.6 e,f
0.1 ± 0.6 b
0 ± 0.3 c
COCs: Cumulus oocyte complexes collected and used in IVF procedure.
Total oocytes (normal, frag, and dead) evaluated.
3
Frag: Fragmented oocyte.
1
2
The number of normal, fragmented, and dead oocytes ovulated per female are
expressed as mean ± SEM.
Within a column, superscript letters represent statistical differences. Those data not
connected by the same letter are different (p<0.05).
Reference
Byers SL, Payson SJ, 2005. Taft RA. Performance of ten inbred mouse strains following
assisted reproductive technologies (ARTs). Therio 65:1716-1726
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In Vitro Fertilization
Applications
•
IVF is used to produce 2-cell embryos for cryopreservation, embryo transfer, or
culture.
•
IVF is used to recover frozen sperm
•
IVF is used as a quality control check of cryopreserved sperm
Key Points
•
Temperature and pH are critical to IVF success. Keep everything at 37°C and under
mixed gas as much as possible. Place small gas incubators immediately beside your
microscopes so that IVF dishes have minimal exposure to room temperature and
atmosphere.
•
Volatile organic compounds in the air of the laboratory can greatly affect success of
IVF and the viability of embryos. It is important to maintain good air quality.
•
IVF dishes may be made the day prior to the IVF, but GSH medium must be kept at
4 degrees Celsius until just prior to use.
•
Work quickly when collecting egg clutches (5 minutes per IVF dish)
•
Be sure to avoid spraying egg collection and IVF dishes when dispensing 70% EtOH
during mouse dissection (EtOH has been shown to cause parthenogenesis).
•
In the IVF dish fertilization drop, a final concentration of 1 – 2.5 x 106 total sperm
per ml is ideal.
•
When washing presumptive zygotes, be careful not to wash or culture zygotes in
the oocyte preincubation drop.
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Materials
•
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Dissecting instruments
o sharp-sharp 3” scissors
o sharp-sharp 5” scissors
o 2 pair of micro-dissecting forceps
o 2 pair of dumont forceps #5
Tissue culture dishes
o 1- 60 x 100mm Falcon dish for every 3 females
o 1- 35 x 10mm Falcon dish for each male
70% EtOH in spray bottle
Cull bag
Paper towels
IVF datasheet
Permanent marker
Stereomicroscope with a transmitted light base
Bench top 37o incubator with mixed gas source (Type 37900 Culture incubator)
Billups-Rothenberg modular incubator chambers (AKA “satellite”)
37°C incubator
Mineral oil, 10mL pipette, and pipettor
Fertilization medium (Cook RVF)
Mixed gas (5% O2, 5% CO2, balanced with N2)
Sperm preincubation medium - TYH Medium + 0.75mM Methyl-β-cyclodextrin
(TYH+MBCD)
GSH
Mouth micropipette
Male mouse 3-6 months old or cryopreserved sperm straws in liquid nitrogen (LN2)
15
Method
1. Prepare sperm dish (if using live males):
a. Pipette 1000µl of TYH+MBCD medium into a 35 x 10mm dish and cover with
mineral oil.
b. Incubate at37°C with 5% CO2, 5% O2, 90% N2 mixed gas for at least 30
minutes.
2. Prepare IVF dish (Figure 1):
a. Pipette one 1000 µl drop (“fertilization drop”) and four 150 µl-drops of Cook
RVF medium into each 60 x 15 mm dish.
b. Cover with oil and incubate
at 37° C with 5% CO2, 5% O2,
90% N2 mixed gas
A
for at least 30 minutes.
c. Use a permanent marker to
C
B
circle one of the 150 µl wash
drops, identifying it as the
C
D
oocyte preincubation drop.
d. Pipette 50 µl of GSH medium
in the circled oocyte
preincubation drop.
Figure 1. IVF dish. A. Fertilization drop B. Preincubation drop. C.
e. Prepare one dish for every
Wash drop. D. Culture drop.
3 females.
3. Prepare cumulus oocyte complex (egg clutches) collection dish:
a. Pipette ~2ml of PBS into a 35 x 10mm dish (do not cover with oil).
b. Incubate at 37°C, but do not gas.
c. Prepare one collection dish for each IVF dish.
4. If using frozen sperm:
a. Thaw sperm following the “Thawing Cryopreserved Sperm” protocol 5-10
minutes before the females reach 13-15 hours post-hCG.
b. Dispense one aliquot of thawed sperm into the 500 µl fertilization drop for
each IVF dish.
5. If using live males:
a. Euthanize sperm donor mouse 5-10 minutes before the females reach 13-15
hours post-hCG injection.
b. Spray the abdomen with 70% EtOH.
c. Dissect the cauda epididymides and vas deferentia (Figure 2). Avoid collecting
fat or other tissue.
d. Place tissue on a dry paper towel to remove excess blood before
placing in the sperm collection dish.
e. Make several cuts in the tissue and squeeze any sperm out of the vas deferens.
f. Immediately return the dishes to the incubator.
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Quick Guide for Dissecting Cauda Epididymis
70% EtOH
A
B
C
cauda epididymis
bladder
seminal vesicle
testicular
fat pad
D
caput
epididymis
testis
CUT HERE
E
vas deferens
F
blood vessel
CUT HERE
CUT HERE
G
H
I
Figure 3. A. Abdomen with 70% EtOH. B. Skin being pulled in opposite directions to expose peritoneum. C-D. Open body cavity exposing the
testicular fat pads. E. Pull on a testicular fat pad to expose the attached testical. F. Locate the cauda epididymis and vas deferens. Cut the ligament
adjacent to the cauda epididymis. G. Cut the vas deferens near the bladder. H. Gently pull the vas deferens away from the body cavity leaving behind
connective tissue and blood vessel. I. Cut just below the cauda epididymis to liberate the tissue.
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6. Allow sperm to preincubate for 40-60 minutes.
7. Immediately after preparing sperm, collect cumulus-oocyte complexes
(egg clutches): See Figure 4.
a. Euthanize 2 to 5 females at 13-15 hours post-hCG injection.
b. Spray the abdomen with 70% EtOH and use dissecting
instruments to open the peritoneal cavity.
c. Using a single cut, dissect out the ovary, oviduct, and a small part
of the uterus (Figure 3), and place into the collection dish
containing ~ 2ml of PBS. See Figure 4a.
oviduct
ovary
CUT HERE
Uterine horns
Figure 3.
Figure
A 4.
B
C
d. Tear each ampullae to release the clutches (Figure 4b-c).
e. Transfer the clutches in the smallest volume of medium possible
to the 200 µl preincubation drop of the IVF dish.
8. Preincubate the clutches in the fertilization drop for 30-60 minutes.
9. Pipette 10µ of sperm into the 1000 µl drop of each IVF dish
10. Confirm the aliquot has entered the IVF drop and did not float into
the oil.
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11. Transfer the clutches in the smallest volume of medium possible from the 200 µl
preincubation drop to the corresponding fertilization drop.
12. Incubate the IVF dishes at 37°C under mixed gas (5% CO2 5% O2 90% N2) for a
total of 2-6 hours.
NOTE: You may use a Billups-Rothenberg modular incubator chambers (AKA
“satellites”), which can be sealed, gassed, and transferred to a large 37° incubator.
13. Wash presumptive zygotes:
a. Using a mouth pipette, transfer the presumptive zygotes from the
IVF drop to the first 150 µl wash drop.
b. Transfer the zygotes to the second wash drop.
c. Leave any fragmented or dead cells in the second drop and
transfer the zygotes to the third wash drop.
d. Record the number of live cells in the third drop and record the
number of fragmented and dead cells left behind in the second drop.
A
C
B
C
A
B
C
D
D
Figure 6. IVF dish. A. Fertilization drop B. Preincubation drop. C. Wash drop. D. Culture drop.
Figure 7. Grading embryos. A. One cell embryos. B. Two cell embryos. C.
Dead cells. D.Fragmented cells.
14. Incubate the dishes under mixed gas overnight.
15. The following day, evaluate and count embryos:
a. Record the number of one cell, two cell, dead, and fragmented
embryos.
16. Viable two cell embryos may now be cultured further in KSOM plus amino acids,
cryopreserved, or transferred into pseudopregnant recipients.
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