INTRODUCTION
Human embryonic stem cells (hESCs) are derived from the inner cell mass (ICM)
of surplus one week old in vitro fertilized embryos. The first human embryonic stem cell
lines were derived in 1998 by plating isolated ICMs from blastocysts on a layer of
mitotically inactivated mouse primary embryonic fibroblasts (Thomson et al., 1998).
Since then, groups around the world have derived lines using numerous conditions
(Mateizel et al., 2006; Cowan et al., 2004; Chen et al., 2005; Kim et al., 2005; Simon et
al., 2005; Klimanskaya et al., 2006; Zhang et al., 2006). A variety of growth substrates
have also been reported to support the derivation of pluripotent hESC lines, including
several types of human feeder cells and feeder-free systems (Lee et al., 2005; Oh et al.,
2005; Fletcher et al., 2006; Klimanskaya et al., 2005; Hovatta and Skottman, 2005).
Finally, different media formulations have been reported to support the growth of hESCs
and progress has recently been made toward deriving hESCs using chemically defined,
xeno-free medias (Ludwig et al., 2006; Inzunza et al., 2005; Genbacev et al., 2005;
Hovatta, 2006; Hong-mei and Gui-an, 2006; Ellerstrom et al., 2006; Fletcher et al., 2006;
Chen et al., 2007).
Research with hESC lines may be subject to local governmental restrictions and
require approval from an Institutional Review Board. In the United States of America,
the National Academies of Sciences has released guidelines for conducting human
embryonic stem cell research, including recommendations concerning obtaining proper
informed consent from donors of embryos for the derivation of human embryonic stem
cells (http://dels.nas.edu/bls/stemcells/).
Described here are basic methods for growing hESCs.
FEEDER PREPARATION
Preparing Primary Mouse Embryonic Fibroblasts (MEFs)
On the bench: Observe clean and sterile tissue technique. Sterilize all instruments needed
for the surgery.
1. Sacrifice pregnant females 12 days after the morning the vaginal plugs were
observed (E12.5).
2. Swab the abdomen with 70% ethanol.
3. Grasp the skin of the abdomen with forceps and make a large incision through the
skin with scissors.
4. Grasp the body wall (peritoneum) with fine forceps and make an incision with
scissors.
5. Remove the uterus with blunt forceps.
6. Embryos can be released from the uterus by inserting the tip of fine tipped
scissors into the anterior end of one of the uterine horns, then cutting along the
length of the uterus.
7. Release each embryo and place them into a 10cm dish with 1xPBS.
8. Remove the yolk sac and placenta and place the embryo into a new 10cm dish
with PBS.
9. Remove the head and the intestines with forceps (all the red parts of the embryos)
and place into a new 10cm dish with PBS. This dish is still unsterile.
Cut off
anterior half
of embryo.
Gently remove
intestines (red).
(Cebra-Thomas., 2002)
10. Using forceps, transfer the embryos from the unsterile dish to a new, sterile dish
under a laminar flow hood. When transferring, make sure the forceps do not
touch the contents of the sterile dish.
Under the hood: Observe sterile technique.
11. Wash 2 times with PBS by transferring each embryo from the same litter from
one 1xPBS dish to a new dish with sterile 1xPBS. Make sure the forceps do not
touch the contents of the dish to which you are transferring, in order to go
from “clean” to “cleaner.”
12. Transfer the embryos to a new 10cm dish with 5ml of 0.5% Trypsin-EDTA.
13. Mince the embryos with sterilized scissors.
14. Leave the 5ml trypsin/ minced embryos dish tilted into the 37°C incubator for a 5
minute incubation.
.
15. Bring the dish back under the hood and pipette vigorously to disperse the cells.
16. Add fibroblast media (MEF media, see Appendix 2) to neutralize the trypsin. Plate
approximately 2 embryos per T175 flask. If desired, the largest clumps of tissue
can be settled out in a 50ml conical tube prior to plating.
17. Change the media the day after plating. Incubate flasks until confluent, usually 12 days.
18. When the T175 are confluent, split MEFs (see Passaging Feeders) or freeze down
2 vials per T175 (see Freezing Feeders), which would be passage 1.
Passaging Feeders
Once the MEFs are confluent, they either need to be split for expanding or frozen down.
For either, start by removing the cells from the flasks.
1. Remove media from flasks and wash with PBS to remove all traces of FBS.
2. Trypsinize the cells:
Add enough trypsin to cover the bottom of the flask (5-7ml in a T175 flask).
Incubate for ~5 minutes.
Check the cells under a microscope to make sure they are all detached.
Inactivate the trypsin by adding media containing FBS (at least the same volume
as the trypsin originally added).
3. Transfer the contents of the flask to a tube to spin down.
4. Spin at 1000 rpm for 5 minutes.
5. Aspirate supernatant, resuspend in MEF media and move to new flask. MEFs are
commonly split in 1:3 to 1:5 ratios.
Freezing Feeders
Feeders can be cryopreserved in liquid nitrogen after slow freezing in isopropanolcontaining jars. We routinely use 90% FBS/10% DMSO as the freezing media.
1. Trypsinize the cells (see Passaging Feeders).
2. Spin down.
3. Remove supernatant and resuspend the cell pellet in freezing media (see Appendix
2).
4. Freeze 2 vials of cells per flask, 1ml per vial.
Mitotically Inactivating Feeders
If feeders are used, they must be mitotically inactivated either by treating with mitomycin
C or by irradiation. We have successfully derived hESC lines using both irradiated human
foreskin fibroblasts (ATCC) and mitomycin C treated E12.5 primary embryonic
fibroblast from CF1 mice (Charles River).
Prior to mitotically inactivating the cells, we typically expand the cells to the largest
number of flasks we can manage, typically around 36-48 T175 flasks per person
performing the prep. In addition, we try to inactivate primary mouse embryonic
fibroblasts by passage 4 or 5. Therefore, for a routine prep, we would thaw 2 vials of
primary mouse embryonic fibroblasts into 2 T175 flask (P1), pass 1:3 to 6 flasks (P2),
pass 1:3 to 18 flasks (P3), then pass 1:3 into 54 flasks (P4) and then pass 1:3 into 162
flasks (P5). We grow the cells in media containing antibiotics for the first two passages
and switch to antibiotic-free media upon the third passage. Although gelatin coating is
always advisable when inactive MEFs are plated (in order to promote adhesion of the
cells to the culture dishes), growing MEFs DO NOT require it.
» Inactivation with Mitomycin C
1.
2.
3.
4.
5.
6.
7.
8.
9.
Remove media from a confluent flask or dish of cells.
Add 10ug/ml Mitomycin C in media to the cells (see Appendix 2).
Incubate 2.5 hours.
Wash with 2xPBS.
Trypsinize the cells (see Passaging Feeders).
Spin down.
Repeat step #7 two times.
Resuspend in media and count the cells. Spin down.
Resuspend the cells in freezing media at a convenient density (we usually freeze
the cells into aliquots containing 0.5, 1.5, 4.5 and 9 x 106 cells).
10. Aliquot the cells into cryovials and freeze (see Freezing Feeders).
» Inactivation by Irradiation
1.
2.
3.
4.
5.
6.
Wash with 1xPBS.
Trypsinize the cells (see Passaging Feeders).
Irradiate the cells to 6000 rads.
Count the cells. Spin down.
Resuspend the cells in freezing media at a convenient density (see #10 above).
Aliquot the cells into cryovials and freeze (see Freezing Feeders).
Thawing Feeders
1. Fill a 15ml conical tube with ~10ml of MEF media (see Appendix 2).
2. Place the vial of frozen feeders in a 37ºC water bath. When only a thin piece of
ice remains in the vial, remove the vial from the water bath.
3. Resuspend the MEFs in ~1ml of MEF media. (Release only a few drops of the
MEF media at a time. Use the pipette tip to gently stir the contents of the vial.)
4. Transfer resuspended MEFs from vial to the 15ml conical tube.
5. Pellet the cells in a conical tube at ~1000 rpm for 3-5 minutes.
6. Remove supernatant. Resuspend the cells in the desired volume and transfer to a
cell culture dish (previously gelatinized). The MEFs should be inactive.
Plating Feeders to Support hESC Growth
Approximate Feeder Plating Densities
Format
Area
(per well)
Minimum
(2x104
cells/cm2)
Optimal
(2.5x104
cells/cm2)
Maximum
(3x104
cells/cm2)
For conditioned
media
(5.6x104
cells/cm2)
35 mm dish
8
1.6x105
2.0x105
2.4x105
4.5x105
60 mm dish
21
4.2x105
5.3x105
6.3x105
1.2x106
10 cm dish
56
1.1x106
1.4x106
1.7x106
3.1x106
T-25 flask
25
5.0x105
6.3 x105
7.5x105
1.4x106
T-75 flask
75
1.5x106
1.9x106
2.3x106
4.2x106
24-well
plate
2
4.0x104
5.0x104
6.0x104
12-well
plate
4
8.0x104
1.0x105
1.2x105
6-well plate
9.5
1.9x105
2.4x105
2.9x105
1. Gelatinize tissue culture grade dishes with 0.25% gelatin for 30 minutes.
2. Thaw a vial of mitotically inactivated feeders and resuspend in at least 5 volumes
of feeder media.
3. Pellet the cells in a conical tube at ~1000 rpm for 3-5 minutes.
4. Remove supernatant. Resuspend the cells and plate at a density between between
2x104 to 3x104 cells per cm2 (see table above).
5. Incubate the feeder cells overnight before plating hESCs.
6. Prior to plating hESCs (at least 30 minutes before, but up to 6 hours prior),
remove feeder media. Wash two times with 1xPBS. Then add hESC media.
hESC CULTURE
Growing hESCs in Feeder-Free Conditions
» Growth in Chemically Defined, Feeder-Free Media
All steps described in the following are specific for hESC culture under feeder-free
conditions in chemically defined media.
Day1:
Thawing hESCs
1. Fill a 15ml conical tube with ~10ml of mTeSR1 media (see Appendix 2).
2. Place the vial of frozen hESCs in a 37ºC water bath. When only a thin piece of ice
remains in the vial, remove the vial from the water bath.
3. Slowly resuspend the hESCs in ~1ml of mTeSR1 media. (Release only a few
drops of the mTeSR1 media at a time. Use the pipette tip to gently stir the
contents of the vial.)
4. Transfer resuspended hESCs from vial to the 15ml conical tube.
5. Pellet the cells in a conical tube at ~1000 rpm for 3-5 minutes.
6. Remove supernatant. Slowly resuspend the cells in the desired volume.
Plating hESCs
1.
2.
3.
4.
Be sure to prepare culture plates coated with matrigel
Bake matrigel coated plates in incubator for at least 30 minutes
Aspirate matrigel and wash once with 1xPBS
Plate hESCs (resuspended in mTeSR1 media) in appropriate media
What to Expect
As the hESC lines begin to expand, spontaneous differentiation will be observed.
Undifferentiated hESC colonies have clearly defined borders, with uniform, tightly
packed cells that exhibit a shiny, refractory morphology under the phase contrast
microscope. Under higher power, hESCs have a high nucleus to cytoplasm ration with
prominent nucleoli. If the number of differentiated colonies becomes greater than
approximately 40% of the total colonies, they should be removed by aspiration. If the
number of undifferentiated colonies becomes extremely low compared to differentiated
colonies, the undifferentiated colonies can be individually dissected out and plated into a
smaller dish. In general, hESCs are more difficult to grow in synthetic media when
compared to standard hESC media, exhibiting a higher rate of spontaneous
differentiation.
Recovery rates following the first cryopreservation can be extremely low. If only a few
small colonies are visible one week after thawing, additional feeders can be plated onto
the existing culture to allow further growth of the hESCs (not in feeder-free conditions).
Passaging hESCs
Approximately once per week, between 4-8 days, hESC colonies are passed either
mechanically or enzymatically to fresh plates of feeders or matrigel. Colonies can be
mechanically dissected using borosilicate Pasteur pipettes that have been pulled into fine
needles and blunted over a flame. Mechanical passage is best performed within a laminar
flow hood outfitted with a dissection microscope (stereomicroscope).
» Mechanical Passage
1. Heat the middle of a sterile borosilicate Pasteur pipette over a flame. Stretch the
pipette so it forms a narrow bore where heated.
2. Bend the pipette, snapping the glass where it narrows, so that the end creates a
needle with a fine point. Polish the end of the needle over a flame.
3. Using either a single needle or a pair of needles, cut the colonies into
approximately 2 to 4 pieces.
4. Transfer dissected colonies to a new plate with fresh feeders.
» Enzymatic Passage
hESCs have been successfully passaged with collagenase, dispase, trypsin and
TrypLE. When we pass the cells enzymatically, we routinely use collagenase as
described below for hESCs on feeders and dispase or accutase for hESCs on matrigel. Described
here is the passaging on feeder-free conditions with accutase.
1.
2.
3.
4.
5.
6.
7.
Remove the old media.
Wash cells once in mTeSR1 media.
Add accutase: 1 ml per well (6well format).
Incubate at 37 C for 2-4 min (depending on cell line).
Cells should be lifting up and colonies loosening out, check under microscope.
Dilute/Neutralize accutase by adding 1 ml warm mTeSR1 media.
Pipette with P1000 carefully to release cells from dish and transfer to 15 ml
conical (use scraper also).
8. If cells are left in the dish, wash with mTeSR1 media to collect those cells and
transfer to 15 ml conical.
9. Centrifuge at 1000 rpm, 4 minutes to pellet cells.
10. Resuspend in fresh mTeSR1 media and distribute to new plate (see Plating
hESCs).
11. Shake well to distribute, check under scope to see how cells look.
12. IMPORTANT: if passaging cells with accutase to a new plate, make sure to
add Thaizovivin at 1:5000 dilution in mTeSR1 (this allows for better
attachment and survival of single cells).
Freezing hESCs
hESCs can be cryopreserved in liquid nitrogen using slow freezing. We routinely use
90% FBS + 10% DMSO (see Appendix 2) as the freezing media or more recently, we
have successfully utilized BAMBANKER, a serum-free freezing medium (sold by
Wako). We have also successfully substituted human serum albumin for FBS. When
hESCs are dissociated in preparation for cryopreservation, we try to maintain the average
size of the colonies larger than when we passage the colonies. The recovery rate of
hESCs following cryopreservation is low. Therefore, we freeze down cells at a high
concentration and thaw at a high density. For master stocks, we freeze one 10cm dish into
three vials and thaw each vial into a single 60mm dish.
1.
2.
3.
4.
5.
6.
7.
Remove the old media.
Wash cells once in mTeSR1 media.
Add accutase: 1 ml per well (6well format).
Incubate at 37 C for 2-4 min (depending on cell line).
Cells should be lifting up and colonies loosening out, check under microscope.
Dilute/Neutralize accutase by adding 1 ml warm mTeSR1 media.
Pipette with P1000 carefully to release cells from dish and transfer to 15 ml
conical (use scraper also).
8. If cells are left in the dish, wash with mTeSR1 media to collect those cells and
transfer to 15 ml conical.
9. Centrifuge at 1000 rpm, 4 minutes to pellet cells.
10. Resuspend in Bambanker freezing medium and transfer to cryovial (750ul).
Day 2:


Observe and evaluate passaged hESCs
Observe and evaluate thawed hESCs
Students will focus on the hESC culture section during the lab sessions. The rest of the
protocol is additional information and useful to understand the basics of hESC culture in
both, feeder-free and feeder-based culturing.
Appendix 1: MATERIALS LIST
Reagents
*Invitrogen has different catalog numbers for single bottles and cases.
REAGENT
ALSO KNOWN
MANUFACTURER
AS
Basic Fibroblast Growth
FGF-basic, FGF-b, Invitrogen
Factor
FGF-2, bFGF
BAMBANKER Freezing
Lymphotec
Media
(sold by Wako)
55mM
βInvitrogen
Betamercaptoethanol
mercaptoethanol,
(1000x)
β-ME, BME, 2mercaptoethanol,
2-ME
Collagenase, Type IV
Invitrogen
Dimethyl sulfoxide
DMSO
Sigma
DMEM with GlutaMAX
KODMEM:F12,
KODMEM/F12
FBS
Knockout Serum Replacer
KOSR, KO-SR,
KSR
Matrigel,
Growth Factor Reduced
Mitomycin C
mTeSR1
302-14681 (120ml)
21985-023 (50ml)
Invitrogen
Invitrogen
26140-079 (500ml)
ATCC
SCRC-1041
Sigma
Invitrogen
Invitrogen
G1890
25030-081 (100ml)
25030-164
(20x100ml)
10828-028 (500ml)
BD Biosciences
354230 (10ml)
M4287-2MG (2mg)
05850 (1kit)
10mM MEM Non-Essential
Amino Acids (100x)
NEAA
Sigma
STEMCELL
Technologies
Invitrogen
Phosphate Buffered Saline
(1x)
PBS
Invitrogen
Penicillin/Streptomycin
(100x)
Pen/Strep, P/S
Invitrogen
Trypsin-EDTA, 0.05%
MMC
PHG0263 (1mg)
17104-019 (1g)
D2438 (50ml OR
5x10ml)
10569-010 (500ml)
10569-044
(10x500ml)
12660-012 (500ml)
Invitrogen
Knockout DMEM:F12
medium
Fetal Bovine Serum,
Qualified
Human Foreskin
Fibroblasts
Gelatin Powder
200mM L-Glutamine
(100x)
CATALOG #
Invitrogen
11140-050 (100ml)
11140-076
(20x100ml)
10010-023 (500ml)
10010-049
(10x500ml)
15140-122 (100ml)
15140-163
(20x100ml)
25300-062 (500ml)
Supplies
EQUIPMENT
Cell Scraper
Cryo 1°C Freezing Container
0.22um Sterile Filter
(250ml bottle)
0.22um Sterile Filter
(500ml bottle)
T175 Flasks
60mm Ultra-Low Adhesion
Culture Dish
MANUFACTURER
BD Falcon
Nalgene
Millipore
CATALOG #
353085 (case of 100)
5100-0001
SCGPU02RE (case of 12)
Millipore
SCGPU05RE (case of 12)
BD Biosciences
Corning
353112 (case of 40)
3261 (case of 20)
Appendix 2: MEDIA PREPARATION
MEF Media
Recipe: 90% DMEM + 10% FBS + P/S
Preparation: Filter reagents through a 0.2um sterile filter unit. (For the purposes of this
course, we will add FBS and P/S to the original 500ml bottle of DMEM.)
250ml MEF Media
REAGENT
DMEM with
GlutaMAX
FBS
100x Pen/Strep
VOLUME
225ml
25ml
2.5ml
STOCK
WORKING CONC.
90%
10%
hESC Media
Several hESC media variations based on Thomson’s original recipe have been reported to
successfully support the growth of hESCs (Thomson et al., 1998; Genbacev et al., 2005;
Inzunza et al., 2005; Hong-mei and Gui-an, 2006). Most use Knockout Serum
Replacement from Invitrogen and FGF-2 (basic FGF). Our standard hESC Media recipe,
“W8,” is a modification of the media recommended by WiCell, “W4,”
(http://www.wicell.org) with twice as much FGF-2 (8ng/ml vs. 4ng/ml). Since Knockout
Serum Replacement contains bovine serum albumin, efforts have been made to develop
completely xeno-free media for growing hESCs. Recently, xeno-free, chemically defined
media has been used to derive hESCs (Ludwig et al., 2006).
» W8 Media
Recipe: (see below)
Preparation: Filter sterilize media through a 0.2um filter unit. Thaw KOSR immediately
before use and be careful not to re-treat repeatedly as KOSR is heat-sensitive.
250ml W8 Media
REAGENT
Knockout DMEM:F12
KOSR
100x Glutamine
100x NEAA
100x Pen/Strep
β-ME
FGF-2
VOLUME
192.5ml
50ml
2.5ml
2.5ml
2.5ml
455ul
200ul
STOCK
WORKING CONC.
200mM
10mM
20%
1%
1%
1%
55mM
10ug/ml
8ng/ml
Varying Volumes W8 Media
REAGENT
KODMEM:F12
KOSR
GLUTAMINE
NEAA
PEN/STREP
β-ME (*uL)
FGF-2 (*uL)
VOLUME (*mL unless otherwise noted)
50
100
150
200
250
300
mL
mL
mL
mL
mL
mL
350
mL
400
mL
450
mL
500
mL
650
mL
38.5
10
0.5
0.5
0.5
91
40
269.5
70
3.5
3.5
3.5
637
280
308
80
4
4
4
728
320
346.5
90
4.5
4.5
4.5
819
360
385
100
5
5
5
910
400
500.5
130
6.5
6.5
6.5
1183
520
77
20
1
1
1
182
80
115.5
30
1.5
1.5
1.5
273
120
154
40
2
2
2
364
160
192.5
50
2.5
2.5
2.5
455
200
231
60
3
3
3
546
240
» W4 Media
Recipe: (see below)
Preparation: Filter sterilize media through a 0.2um filter unit. Thaw KOSR immediately
before use and be careful not to re-treat repeatedly as KOSR is heat-sensitive.
250ml W4 Media
REAGENT
Knockout DMEM:F12
KOSR
100x Glutamine
100x NEAA
100x Pen/Strep
β-ME
FGF-2
VOLUME
192.5ml
50ml
2.5ml
2.5ml
2.5ml
455ul
100ul
Freezing Media
Recipe: 90% FBS + 10% DMSO
Preparation: Filter sterilize media.
STOCK
WORKING CONC.
200mM
10mM
20%
1%
1%
1%
55mM
10ug/ml
4ng/ml
50ml Freezing Media
REAGENT
FBS
DMSO
VOLUME
45ml
5ml
STOCK
WORKING CONC.
90%
10%
Mitomycin C Solution
Recipe: 10ug/ml Mitomycin C in media
Preparation: Standard bottle will make 200ml Mitomycin C solution. Filter sterilize.
Matrigel Solution
Recipe: Matrigel diluted in KODMEM:F12
Preparation: Recipe will vary depending on batch of Matrigel. (See bottle for relevant
instructions.) Filter sterilize.
Collagenase Solution
Recipe: 200 units/ml collagenase in hESC media
Preparation: Recipe will vary depending on the stock concentration of the batch of
collagenase (supplied as a lyophilized powder) at hand. It will be easier to adjust the
volume of hESC media used than adjusting the mass of collagenase powder used. The
target working concentration of collagenase is 200 units per ml of media.
200 units = (stock concentration) units/mg * (mass collagenase) mg
1 ml
(vol. hESC media) ml
FOR EXAMPLE: If the stock concentration is 247 units/mg, using ~40.5 mg of
collagenase will yield ~50ml of 200 units/ml collagenase solution.
1. After figuring out the volume of hESC media corresponding to the mass of
collagenase powder to be used, transfer the collagenase (unsterile) from the
weighing paper to a conical tube. Transfer tube to a laminar flow hood.
2. Add the appropriate volume of hESC media to the tube. Make sure collagenase
dissolves completely.
3. Filter sterilize the collagenase solution through a syringe filter.
Appendix 3: PHOTOS
Optimal Feeder Density for H9 Cells
Feeder Density Too High for H9 Cells
Optimal H9 Morphology
Sub-Optimal H9 Morphology
Sub-Optimal H9 Morphology
Optimal H9 Morphology (Feeder-Free Cultures)
Appendix 4: COUNTING CELLS
A Neubauer hemocytometer is used to count cells under a microscope. Two 3x3mm
counting grids, or “chambers,” are engraved onto the hemocytometer. Each chamber is
composed of 9 smaller squares (outlined in red in the diagram below), each measuring
1x1mm. These squares themselves are again subdivided into smaller and smaller squares.
(Woermann, 2000), modified
A hemocytometer chamber.
1. Place a cover slip over both chambers of the hemocytometer.
2. To ensure your cell suspension is homogenous, pipette the suspension up and
down several times.
3. Either using a 20ul pipette or a glass Pasteur pipette, draw a small amount of
suspension into the pipette. (Capillary action will cause liquid to enter the glass
Pasteur pipette.) Form a droplet of liquid at the tip of the pipette (for the glass
pipette, gently tap the open end).
4. Lightly touch the droplet to the edge of the cover slip, where the slip meets the
chamber. Do the same at the opposing edge for the second chamber. (Capillary
action will cause the liquid to spread out under the cover slip, across the chamber.
Add only enough liquid to fill the chambers.)
5. Count the cells in 4-5 of the 9 square subdivisions (outlined in red in the
diagrams) per chamber, for a total of 8-10 counts.
*During the count, all cells that lie on or just touch the left and the lower
borders are counted. The cells on or touching the right or upper borders are
not. (Or vice versa – just remember to use the same two borders for all
counts.)
**Counts per square should be in the 50-150 range to ensure proper cell
count. If less, the suspension is too dilute – respin and resuspend in a lower
volume. If greater, the suspension requires further dilution.
(Woermann, 2000), modified
Blue cells are counted.
6. Average the 8-10 counts (per square) to get an average count per square.
Multiply the average count by 104 to get the number of cells per ml. (Each redoutlined square is 1mm x 1mm with a depth of 0.1mm, giving each square a
volume of 0.1mm3. There are thus 104 squares in 103 mm3 = 1 ml.) Multiply this
by the volume (ml) of the original suspension and by the dilution factor (if
relevant) to get the total cell count.
FOR EXAMPLE: In the diagram above, there are 74 cells in the square. Suppose
other squares yielded counts of 85, 71, 68, 80, 81, 77 and 76, and suppose the
original cell suspension had a volume of 8ml.
Average count per square: 76.5 cells/square
Count per ml: 76.5 cells/square * 104 squares/ml = 765,000 cells/ml
Total count: 765,000 cells/ml * 8 ml = 6,120,000 cells
Appendix 5: REFERENCES
Behr, B, et al. (1999). 'Preliminary clinical experience with human blastocyst
development in vitro without co-culture.', Hum Reprod, 14 (2), 454-57.
Behr, B (1999). 'Blastocyst culture and transfer.', Hum Reprod, 14 (1), 5-6.
Cebra-Thomas, J (2002, December 15). Mouse embryo staging. Retrieved from
Swarthmore College website:
http://www.swarthmore.edu/NatSci/sgilber1/DB_lab/Mouse/mouse_stage.html.
Chen, H, et al. (2005). 'The derivation of two additional human embryonic stem cell
lines from day 3 embryos with low morphological scores.', Hum Reprod, 20 (8),
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