Stem Cell Rev and Rep (2012) 8:1011–1020 DOI 10.1007/s12015-012-9377-4

Optimized Protocol for Derivation of Human Embryonic Stem Cell Lines María Vicenta Camarasa & Víctor Miguel Galvez & Daniel Roy Brison & Daniel Bachiller

Published online: 22 May 2012 # Springer Science+Business Media, LLC 2012

Abstract For the past 12 years, the biology and applications of human embryonic stem cells (hESCs) have received great attention from the scientific community. Derivatives of the first hESC line obtained by J. Thomson’s group (Science 282 (5391):1145–1147, 1998) have been used in clinical trials in patients with spinal cord injury, and other hESC lines have now been used to generate cells for use in treating blindness (Lancet 379(9817):713–720, 2012). In addition to the classical protocol based on mouse or human feeder layers using open culture methods (In Vitro Cellular & Developmental Biology Animal 46(3–4):386–394, 2010; Stem Cells 23(9):1221–1227, 2005; Nature Biotechnology 24(2):185–187, 2006; Human Reproduction 21(2):503–511, 2006; Human Reproduction 20 (8):2201–2206, 2005; Fertility and Sterility 83(5):1517–1529, 2005), novel hESC lines have been derived xeno-free (without using animal derived reagents) (PLoS One 5 (4):1024–1026, 2010), feeder-free (without supporting cell monolayers) (Lancet 365(9471):1601–1603, 2005), in microdrops under M. V. Camarasa (*) : V. M. Galvez : D. Bachiller Caubet-Cimera Fundation, Centre for Advanced Respiratory Medicine, Recinte Hospital Joan March, Ctra Sóller km 12, 07110 Bunyola, Illes Balears, Mallorca, Spain e-mail: [email protected] M. V. Camarasa e-mail: [email protected] D. R. Brison North West Embryonic Stem Cell Centre, Faculty of Life Sciences, Core Technology Facility, University of Manchester, 46 Grafton Street, Manchester M13 9NT, UK D. Bachiller Consejo Superior de Investigaciones Científicas (CSIC), Mallorca, Spain

oil (In Vitro Cellular & Developmental Biology - Animal 46 (3–4):236–41, 2010) and in suspension with ROCK inhibitor (Nature Biotechnology 28(4):361–4, 2010). Regardless of the culture system, successful hESC derivation usually requires optimization of embryo culture, the careful and timely isolation of its inner cell mass (ICM), and precise culture conditions up to the establishment of pluripotent cell growth during hESC line derivation. Herein we address the crucial steps of the hESC line derivation protocol, and provide tips to apply quality control to each step of the procedure. Keywords Human embryonic stem cells . Derivation . Blastocyst . Culture optimization

Introduction To date, hundreds of human embryonic stem cell (hESC) lines have been derived from surplus embryos after assisted reproduction techniques. Reported derivation efficiencies range from 7 to 100 %. This wide span probably reflects the diversity of the methodologies involved. Many variables of the procedure such as embryo grade, timing of embryo culture, zona pellucida (ZP) removal and inner cell mass (ICM) isolation and type of feeder layer, account for the differences among laboratories. Nevertheless, despite the variability, all hESC lines derived share growth properties and expression of common stem cell markers, indicating that diverse culture conditions select for unique characteristics. The International Stem Cell Initiative II has analysed the genetic stability of more than one hundred hESC lines using consensus protocols for karyotyping and genotyping. This echoes the importance of defining robust and common criteria for the assessment of both existing and newly derived cell lines, and for the validation of advances in culture conditions. [13].

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This protocol focuses on the main first steps of the derivation of hESC lines, i.e. embryo culture and grading, and precise manipulation of their ICMs and first outgrowths of pluripotent stem cells. The Notes section includes alternative procedures as well as commentaries and clarifications on the techniques described in the Methods section. Assuming experience in mammalian basic tissue culture technique, this protocol should aid anyone wishing to derive and culture hESCs to set up a rapid and reliable method. This protocol reflects continuous work optimizing the derivation, culture and characterization of hESC lines over more than 10 years, comprising activities carried out in four different laboratories. Advantages of the present protocol over others previously published include: optimization of feeder cell yield, increase of ICM cell production and adjustment of the derivation protocol according to embryo grade. Feeder cell yield is optimised by culturing tissue aggregates until they grow as monolayers. Embryo culture is optimized so that growth of the pluripotent cell population is favoured over that of the non-pluripotent trophectoderm (TE) cells. Finally, the fire pulling of glass Pasteur pipettes to form thin open and closed ends that match the size of the ICM or initial outgrowth to isolate, increase effectiveness of mechanical splitting techniques.

Materials Materials and Reagents 1. Bacteriological Petri dishes, 100 mm Ø, Fisher Scientific #09-720-500. 2. Centre-well Organ Culture Dish, 60 mm tissue culture treated, BD #353037. 3. Long glass Pasteur pipettes, Fisher # 1367820C. 4. Mr Frosty® Nalgene, Sigma #C1562-1EA. 5. Serological pipette 1 ml, Corning #4485. 6. Serological pipette 5 ml, Corning #4487. 7. Serological pipette, 10 ml, Corning #4488. 8. Serological pipette, 25 ml, Corning #4489. 9. Serological pipette, 50 ml, Corning #4490. 10. Tissue culture dishes, 150 mm Ø, Corning #430599. 11. Tissue culture flasks F100, 100 cm2 Corning #3816. 12. Tissue culture plates, 4-well, BD #353654. 13. Tissue culture plates, 6-well, Corning #3516. 14. Basic Fibroblast Growth Factor, #45103P-100. 15. Beta-mercaptoetanol, Sigma #M-7522. 16. Collagenase IV, Invitrogen #17104-019. 17. DMEM medium, Lonza #12-614F. 18. DMSO, Sigma # D2650. 19. Fetal bovine serum (FBS), Australian origin Lonza #DE14-701F. Test batches for fibroblast cell growth. Best origins are US and US approved, followed by Australian

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20. 21. 22. 23. 24. 25. 26. 27.

28. 29. 30. 31. 32. 33. 34.

and Australian/USDA approved. Poor quality serum is a confirmed cause of feeder cell failure in hESC support. G1, Vitrolife #10128. G2, Vitrolife #10132. Gelatin, porcine, Sigma #G-1890. Knockout (KO) DMEM, Invitrogen #10829-018. Knockout serum replacement (KO-SR), Invitrogen #10828-028. L-Glutamine, Lonza #17-605E. MEM Non-essential aminoacids (NEAA), Lonza # 13-114E. Mitomycin C, Sigma #M0503. Hazard: this substance is very toxic and procedures for safe handling and disposal need to be in place. Do not breathe dust. Avoid contact with eyes, on skin, on clothing. Avoid prolonged or repeated exposure. OVOIL, Vitrolife #10029. PBS with Ca+2/Mg+2, Lonza #17-513F. PBS, without Ca+2/Mg+2, Lonza #BE17-516F. Pregnant MF-1, CF-1 or MF-1 × CD-1 females, 12.5–13.5 dpc, Charles River Laboratories. Thawkit I, Vitrolife #10067. Trypsin-EDTA, Cambrex # CC-5012. Tyrode’s solution, acidic. Sigma #T1788.

Solutions 1. Complete DMEM medium: DMEM plus 10 % v/v FBS, 1 % v/v L-Glutamine 200 mM, 1 % NEAA. 2. Freezing medium: 90 % v/v FBS plus 10 % v/v DMSO. 3. Gelatin solution: 0,1 % w/v gelatin in tissue culture grade water. To avoid contaminants and residues (detergent, traces of other chemical, etc.), use a new 500 ml Pyrex bottle. Autoclave the gelatin solution and store it at RT. 4. HES medium: KO-DMEM supplemented with 20 % KO-SR, 2 mM L-Glutamine, 1× NEAA, 50 mM β-Mercaptoethanol and 8 ng/ml human basic fibroblast growth factor.

Methods ESCs are derived from preimplantation stage embryos. The most common procedure involves culturing embryos to the blastocyst stage (day five to seven after fertilization), isolating their ICMs, and culturing the latter on mitotically inactivated fibroblast cell layers. The resulting pluripotent cell populations are selected by morphological and growth criteria. Novel hESC lines are then established after continuous subculture of the initial outgrowth. A hESC population can be considered to be an established line when it reaches passage 8 with approximately 3×107 cells, as defined by the UK National Clinical human Embryonic Stem Cell Forum [14].

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After eight to ten consecutive passages, the novel line is sufficiently expanded to check its viability by freezing and thawing. Table 1 defines the term ‘one volume’ for each of the culture vessels described throughout the manuscript. All incubations are performed in conditions of 37°C, 95 % humidity and 5 % CO2 in air (See Note 1). Figure 1 depicts the expected time schedule to complete a hESC derivation protocol, provided that the feeder source has been previously validated. Feeder Layer Preparation Primary cultures of feeder cells have tended to be used for derivation of novel hESC lines, but routine culture after derivation can be performed on feeders produced from established cell lines, or primary feeder cells which have been immortalised. Alternatives to the use of primary mouse embryonic fibroblasts (pMEFs) as feeder layers for hESC culture are discussed in Note 2. Regardless of the type of feeders used, every feeder layer must be conditioned before stem cells are seeded on top of it. To do so, feeder culture medium is discarded; feeders are then washed three times with one volume of PBS each time and once with half a volume of stem cell medium (HES). Plates are ready for use, from 30′ to several hours afterwards. MEF Isolation 1. Sacrifice a pregnant female (12.5–13.5 dpc) by cervical dislocation. See Note 3 for tips on strain choice. 2. Dissect out the uterine horns and place them into a 100 mm Ø Petri dish containing one volume of sterile PBS 1×. Transfer the plate to a laminar flow hood to continue with the procedure. 3. Cut open the whole length of the uterus, so viteline sacs with embryos and placentas are partially released. 4. Separate each embryo from its placenta and out of the viteline sac and surrounding membranes, and transfer them all to a new Petri dish with one volume of fresh PBS. Table 1 Definition of the vessels and volumes used throughout culture procedures Dish/plate 4-well plate 24 well plate One-well 60 mm plate 6-well plate 60 mm Ø dish 100 mm Ø dish F100 flask 150 mm Ø dish

Area cm2/well 1.4 2.0 2. 9 9.5 21.3 58.1 100.0 176.7

One Volume ml/well 0,5 1 1 3 5 20 25 50

5. Wash the embryos and transfer them again to a new Petri dish with one volume of fresh PBS. 6. Decapitate the embryos and separate the visceral tissues from the bodies. Collect carcasses aside in the working plate. Aspirate and discard the PBS and the visceral tissues carefully with a 10 ml serological pipette. Leave the rest of the tissues in the plate. 7. Mince body walls into small pieces until a homogeneous suspension is formed. 8. Add 5 ml of trypsin-EDTA 1x to the plate and transfer all contents to a conical 50 ml tube. 9. Incubate digestion at 37°C and 5 % CO2 for 30 min. Homogenize the cell suspension by adding 45 ml of fresh culture medium and pipetting up and down for ten times, and centrifuge to recover the cells (700 g for 2 min). 10. Discard supernatant, add 50 ml of fresh culture media and resuspend cells. 11. Centrifuge the cells at 700 g for 2 min and discard supernatant. 12. Resuspend the cells in 5 ml of complete DMEM medium. 13. Seed cell suspension at a rate of three embryos per F100 flask. (Generally four F100 will be set up at the beginning of the procedure per dissected pregnant female). 14. Incubate at 37°C and 5 % CO2 until 90 % confluence is reached (approximately two to 4 days). 15. Split the cells twice by trypsinisation, seeding them onto new flasks/dishes between 1:3 and 1:6 split according to growth rate. After two passages, the cells should be growing as monolayers. Freeze 90 % of passage 2 cells in vials containing four to ten millions units each. The frozen cells will constitute a stock of active pMEFs for future use. To continue with the production of inactivated feeder cells, expand the remaining 10 % of the original culture up to passage four. Seed the cells onto new flasks/ plates between 1:3 and 1:6 split according to growth rate each passage. Then inactivate and freeze passage four cells as explained below. See Note 4 for tips on these steps. Generation of Batches of Frozen Feeder Cells by Mitomycin C Inactivation 1. Treat 80 % confluent, exponentially growing, passage four pMEF populations (between 48 and 96 P150 plates), with 20 ml of 10 μg/ml Mitomycin C in complete DMEM medium, for 2–3 h at 37°C and 5 % CO2.

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Fig. 1 Time schedule for the completion of a full derivation experiment

2. After inactivation, discard the supernatant and wash cell monolayers 3 times with one volume of PBS without Ca+2/Mg+2. 3. Trypsinise the cells (2 ml per F100 flask or 3 ml per P150 plate) for 2–7 min at room temperature (RT). 4. Tap the sides of the flasks/dishes to dislodge the cells, and harvest them into 50 ml conical centrifuge tubes. Pool the contents of two to three P150s in one tube. 5. Add 44 ml of complete medium per tube. 6. Centrifuge tubes at 700 g for 2 min. 7. Break pellet manually by finger vortexing (see Note 5). Resuspend each pellet in 30 ml of complete DMEM and count in a Neubauer chamber. 8. Count viable cells and calculate the number of cryovials needed, at a rate of 1–4 million cells per vial. 9. Centrifuge cell suspensions at 700 g during 2 min. 10. Label cryovials with batch number, amount of cells and method of inactivation, date and user ID. 11. Discard supernatants from step 9 above and break pellets carefully by finger vortexing. 12. Add cold freezing medium to the pellets at 0,5 ml per vial. Dispense cell suspension into labelled cryovials and transfer them immediately to a precooled Mr Frosty® container, and then into an −80°C freezer (see Note 6). Feeder Layer Plating for ICM Culture 1. Dispense one volume of gelatin 0.1 % onto the required number of dishes/wells, and incubate them at 37°C for 30 min. 2. Equilibrate 10 ml of fresh complete DMEM medium, in a 15 ml conical tube, in the CO2 incubator per each 1–2 vial(s) to be thawed (see Note 7). 3. Once the coated dishes and the media are equilibrated, thaw the required number of cryovials to plate 0.2–0.3 millions of inactivated pMEFs per one-well 60 mm dish. Spray the frozen vials with 70 % ethanol. Roll them in your hands, until only the last small piece of ice remains. Transfer the content of the vials into the equilibrated complete DMEM. In order to avoid cell damage it is not advised to thaw more than four vials at a time.

4. Centrifuge the tubes at 700 g for 2 min. Discard the supernatant and gently break the pellet by finger vortexing. 5. In order to eliminate residual DMSO, repeat step 4. 6. During the second centrifugation, aspirate the gelatin solution from the plate(s) and dispense half a volume of prewarmed complete DMEM medium on each one. 7. Break pellets from step 5 above by finger vortexing, and resuspend them in half a volume of the well(s) to plate. 8. Dispense cells dropwise onto the medium-containing well(s). 9. Place plate(s) at 5 % CO2 at 37°C, and carefully shake them horizontally in all directions to distribute the cells evenly. 10. Incubate feeders from 6 h to overnight to let them attach. Optimal feeder density corresponds to a confluent monolayer; higher and lower densities will result either in reduced hESC growth and increased differentiation, respectively. See Note 8 for comments on feeder layer preparation. Embryo Culture Surplus embryos from clinical in vitro fertilization (IVF) procedures can be supplied, fresh or frozen, at any stage from day one to day six post-fertilization. It is firmly established now that low-grade blastocysts produce hESC lines successfully [15, 16]. Therefore, the derivation of stem cell lines is almost guaranteed for any Research Programme established in collaboration with an IVF clinical department. In this section of the protocol we will detail the steps involved in embryo culture for hESC derivation. It is advisable to use the same clinical grade embryo culture reagents that are used in the clinical procedures. Alternatively, it is simple to prepare them from their components [17]. See Note 9 for tips on embryo culture. We will detail Vitrolife® protocols, and advise to follow manufacturer instructions in case of different brand reagents. The culture system used was a two-stage one, i.e. changing culture media formulation on day 3 or six-eight cell stage of development (from G1 to G2 series), until blastocyst formation.

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Embryos are cultured in microdrops under oil, to prevent evaporation and to minimise pH changes. The following protocol corresponds to the culture of fresh or thawed cleavage-stage embryos. In the case of receiving frozen embryos, it is crucial that the thawing is performed in the reagents corresponding to their freezing. Follow manual instructions from the appropriate thawing kit. After thawing, follow the protocol below for embryo culture. 1. Equilibrate OVOIL® washed with 1:10 volume of G1 or G2 for 6 h to overnight at 37°C and 5 % CO2. 2. Equilibrate G1 or G2 media as required (see Table 2) for a maximum of 3 h. 3. To set up embryo culture plates, pipette 25 μl of the equilibrated G1 or G2 media per embryo on embryo culture dishes/plates. Cover the drop(s) with equilibrated OVOIL®, and add 25 μl more of the medium to the formed drop(s). 4. Assess embryo development daily (see Fig. 2 for a diagram of growth pattern and corresponding images of embryo development), and transfer them to fresh media drops every second day (see Table 2 for media change guideline). Prepare new drops as described in steps 1 to 3 above. If embryo culture started in G1, remember that once embryos reach day 3 or the six-eight-cell stage, they must be cultured in G2 medium. 5. When embryos reach day six of development, or the ICM of the blastocyst is visible, it is time to set up the feeder plates to seed the former. ICM Isolation and Seeding During denudation and ICM isolation it is advised to process each embryo individually. Pulled glass Pasteur pipettes or capillaries are recommended, or commercially available embryo handling pipettes used for clinical IVF procedures. Condition the feeders before the denudation as follows: between 1 and 3 h before the seeding eliminate feeder medium. Wash three times with 1 ml of PBS with Ca+2/ Mg+2 and once with 0.25 ml of HES medium. Add 0.5 ml of HES medium and equilibrate for at least 20 min at 37°C and 5%CO2. At this point the new feeders are ready to receive the ICM. Zona Pellucida Removal 1. Equilibrate separately acid tyrode’s (AT) 1×, G2 medium and HES medium at 37°C and 5 % CO2, for a minimum of 20 min and a maximum of 6 h.

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2. Before any manipulation is performed, images of the blastocysts at different planes (×40) should be taken for further assessment of its grade. Afterwards return it to the incubator. 3. Pull 2 long sterile glass Pasteur pipettes with the aid of a Mecker or Bunsen burner. Leave one of them opened and rounded, and close the other one leaving a small rounded ball at the tip (Fig. 4). Check the thickness of the tips under the stereomicroscope and repeat the procedure until a couple of pipettes whose section match the diameter of the blastocyst are produced. 4. Set three drops of AT forming a row in a 60 mm tissue culture dish, and mark their position at the rim of the dish. Add another three drops of G2 medium in a second row at the middle of the dish, and three drops of HES media at the bottom, again aligned in a row (See Fig. 3 for drop distribution). 5. Set another three drops of approximately 100 μl of HES medium in a new 60 mm dish. 6. Pick the embryo up in G2 medium with a 275 μm capillary micropipette, and transfer it to the left AT drop of the plate prepared in step 5 above. Stop dispensing G2 medium into the AT drop as soon as the embryo has been released. Empty the pipette in the border of the dish to discard residual G2 medium. 7. Wash the pipette twice in the second AT drop. 8. Return to the first drop, pick up the embryo and transfer it to the third drop. Monitor under the microscope the dissolution of the ZP (0–2 min). 9. When the trophectoderm (TE) of the embryo is exposed by the dissolving ZP, transfer the embryo sequentially to the three G2 drops. Use the first drop as a pre-wash, dispensing the minimum amount of AT into the G2, the second to load the pipette with fresh G2 medium, and the third to wash the blastocyst. 10. Transfer the zona-free (denuded) blastocyst to the drops of the HES media using the washing procedure as described above for G2. 11. Immediately transfer the blastocyst to the drops of the HES media prepared in a separate plate in step 2. Follow the same procedure of transferring to the left drop, washing the pipette and loading it in the second intermediate drop and transferring the embryo to the third drop. Place the plate into the CO2 incubator. 12. Using the images taken in step 2 above, assess the derivation grade of the denuded blastocyst (see embryo grading scheme in reference [18]) and apply Table 3 for determining the best method of ICM isolation. Depending of the outcome, go to protocol A or B below. See Note 10 for comments on these procedures.

1016 Table 2 Embryo culture protocol

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Day of development

Stage

Medium

Image recording

Media change

1 2 3 4 5 6 7

Zygote (2pn) 2–4 cells 6–8 cells Compacting morula Expanding Blastocyst Hatching

G1 G1 G2 G2 G2 G2 G2

− Yes Yes Yes − Yes Yes

G1 − G1 to G2 − Fresh G2 − −

ICM Seeding by Pipetting (A) 1. Choose an open pulled long Pasteur pipette of the same diameter as the denuded blastocyst (see shape in Fig. 4a). 2. Load half of the tip with fresh warm HES medium from the drops set up to equilibrate before denudation (step 5 on zona pellucida removal protocol). Fig. 2 Dagram of early human embryo development. a-a’ two-cell stage embryo, 1 day after fertilization; b-b’ four-cell stage embryo, 2 days after fertilization; c-c’ eight-cell stage embryo, 3 days after f ertilization; d early expanding blastocyst; d’ morula containing more than 16 cells, 4 days after fertilization; e early blastocyst, 5 days after fertilization; f hatching blastocyst, around 7 days after fertilization. ZP: Zona Pellucida. ICM: Inner Cell Mass. TE: Trophectoderm

3. Aspirate the denuded embryo and pipette it in and out while checking for loosening of the TE cells. 4. Aspirate the isolated cell clumps and dispense them separately on the feeder plate. 5. Take an image of the seeded fragments of the blastocyst. Pay attention to locate the ICM and record its position.

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1017 Table 3 Choice of method for ICM isolation according to blastocyst grade ICM TE 0 1 2 3

0

A

1

2

3

A A B

A A B B

A A/B B B

TE score - 0: no cell layer formed. 1: few big cells forming a loose layer. 2: cells forming a medium coherent layer. 3: many cells in a tight cellular layer. ICM score - 0: no visible inner cell mass, inner to the TE in any plane of the blastocyst. 1: ICM with less than 10 compacted cells, or loosened mass with up to 20 cells in the central focused plane of the blastocyst. 2: ICM with more than 10 compacted cells, or more than 20 loosened cells in the central focused plane of the blastocyst under the stereomicroscope. 3: ICM with more than 20 compacted cells, or more than 25 loosened cells in the central focused plane of the blastocyst. Method A refers to pipetting for ICM isolation. Method B consists of mechanical cutting of the ICM

Fig. 3 Shapes of long glass pipette tips for ICM and stem cell colony processing. a transferring pipette with open rounded end; b cutting pipette with open bevelled end; c holding pipette with a fine rounded closed end; d curved hook with closed end for outgrowth scraping; e-g flat hooks for colony cutting and scraping. h detail of the tip of the holding pipette depicted in C

6. Transfer the plate to the incubator, avoiding abrupt movements that may separate the seeded cells from the feeders. 7. Inspect the plates daily. Change the medium every other day. 8. Prepare new feeder cell plates as soon as pluripotent growth is detected (see Note 11 for derivation timelines and Fig. 5 for images on derivation progress). 9. Split part of the emerging pluripotent colony when residual TE cells begin detaching from the bottom of the plate. Follow Section 4 below to do so.

drop will diminish up to the point that the embryo flattens and sticks to the plastic. At this point you should be able to locate the ICM: it is much brighter and less flattened than the dim TE. Save the pipette without emptying it for following cutting steps. 3. Pick a closed pulled (holding) pipette (Fig. 4c) with the left hand (right for left-handed) and hold TE by pressing its border against the plate. 4. Pick the open (cutting) pipette still containing medium with the right hand (left for left-handed). With its edge, cut out as much transparent TE as possible. Immediately release the HES medium to avoid damaging the ICM. 5. Scrape loose the ICM containing piece, which will be adhered to the plastic. Monitor the ICM while it

ICM Seeding by Mechanical Cutting (B) 1. Aspirate the denuded blastocyst with a pulled open pipette (Fig. 4b) and dispense drops of approximately 20 μl on the dry area of the 60 mm dish containing fresh HES drops, until the blastocyst is dispensed in one of them. 2. Drag the denuded blastocyst across the dish, away from the HES drops. The volume of the embryo-containing

Fig. 4 Derivation plate. Embryos are dispensed in AT in drop 1, and transferred serially according to the flow of the arrows. AT: Acid Tyrodes. G2: embryo culture medium. HES: Human Embryonic Stem cell medium

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Fig. 5 Micrographs showing evolution of the pluripotent and TE cells during a derivation experiment. a ICM 1 day after seeding. b ICM cell expansion after 4 days in culture. Arrow points to the pluripotent outgrowth, which looks like a clump at this stage; c colony outgrowth 2 days after splitting the outgrowth in panel B: arrow points to the expanding pluripotent cells out of the initial hESC clump; d residual TE cells 3 days after ICM seeding; e TE differentiated cells 4 days after seeding; f TE cells detached from the plate 9 days after seeding; g detail of the emerging pluripotent colony of picture C; h early hESC colony at passage 4; i regular colony of an established line at passage six

6.

7. 8. 9.

detaches and floats. Quickly aspirate medium from the HES drops in the plate and add them to the ICMcontaining drop to compensate for the evaporation occurred during cutting. Aspirate the floating piece(s) and spread them onto the feeder dish. Try to avoid pure TE fragments, recognized by their transparency. Take an image of the fragments seeded, recording the position of the ICM piece on the dish. Inspect plates daily, feeding them each second day until pluripotent cell growth is obvious. Prepare new feeder cell plates as soon as pluripotent growth is detected (see Note 11 for derivation timelines and Fig. 5 for images on derivation progress). Go to Section 4 below for the first split of the pluripotent cells.

Split of First Pluripotent Outgrowth When the pluripotent outgrowth(s) reach the stage depicted in Fig. 5c, they are ready to be split. Mechanical cutting can be performed with a glass pipette of any of the shapes A, D, E or F depicted in Fig. 4. In any case, prepare feeder plates the day before splitting. If the pluripotent outgrowth is isolated from the residual TE, a closed pipette and a scraping method could be more effective. If stem cells are growing in clumps surrounded by TE, an open pipette to cut some of them will be preferred. It is wise to leave some outgrowth(s) behind, until split cells are successfully growing in the new feeder dish.

Notes Isolation and culture of mouse embryonic fibroblasts and feeder preparation Note 1. We describe here the derivation of hESC lines in atmospheric oxygen; however, the use of physiological concentrations of oxygen is currently gaining support. Newly published data report a higher efficiency in stem cell derivation and lack of chromosome X inactivation in hESC lines established under such conditions [19]. Note 2. Primary MEFs are the first cells to be used as feeder layers for hESC culture. Alternatively, either transformed [20] or naturally immortalised [21] MEFs, as well as human fibroblasts from several origins, including foreskin [22] and placenta primary cells [8] and an immortalised placental fibroblast cell line [23] have been used successfully as feeders for hESC derivation and culture. Primary lines from human foreskin fibroblasts (HFF-1) have also been used reproducibly to culture hESC lines. In the case of HFF-1 cells, irradiation is the preferred method of inactivation. All methods described in this paper are equally valid regardless of the type of feeders used. Note 3. Mice strain has proven to be an important factor for the quality of the feeder cells derived from

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Note 4.

Note 5.

Note 6.

Note 7.

Note 8.

them. In our hands, embryos from the CF-1 and MF-1 lines, and from the cross between MF-1 and CD-1, are much better than inbred CD-1 embryos. Isolated pMEFs can be frozen at passage zero, and expanded at a later time to prepare feeders. Nevertheless, the expansion up to passage two before freezing has the advantage of increasing the number of cells obtained at passage four. It has to be noted that in the isolation procedure described in the Methods section, and differently from other published protocols [24], tissue chunks are not discarded before seeding after first trypsinization. This procedure yields significantly higher number of cells. It is advised to culture chunks and clumps separately from single cell suspensions, and trypsinise them until they produce monolayers. A couple of passages from one isolation/mouse will normally be sufficient to produce enough number of vials for culturing up to six new stem cell lines during 1 year. Finger vortexing is a simple way to mix a solution or disaggregate a pellet in a test tube. Hold the top of the tube securely in one hand and draw the ring, middle and index finger of the other hand sequentially towards you, tapping the tube. This creates a whirlpool effect inside the tube, which can be adjusted in intensity by speed. It is milder than mechanical vortexing and yields healthier cells after centrifugation. Freezing fibroblasts with a slow freezing method is not an issue, and they can be stored at −80°C for months without a decrease in subsequent plating efficiency. The protocol described herein uses the slow rate freezing container Mr Frosty, which has been validated and used for nearly 20 years in mammalian cell freezing [25]. It is a cheap and time saving option when compared to alternative methods such as controlled-rate freezing and vitrification, which may be suitable for more delicate materials like embryos and hESCs. It is convenient to dilute freezing medium in complete DMEM medium at a rate of 1:10 to avoid toxicity and differentiation during culture. Mitomycin C is the preferred method to inactivate MEFs, and detailed steps to perform the procedure are included in the text. Alternatively, γ- or X-ray irradiation can be used. In any case, the hESC supporting capacity of each new batch of feeders has to be tested on known hESC lines before considering them validated for use. Once the new batch has been validated for hESC culture, it can be used in subsequent derivation experiments.

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Embryo and ICM Culture Note 9.

The procedures detailed here are based on the use of high quality clinical grade embryos, but frequently, the quality of the embryos donated for research does not correspond to the standards presented in Fig. 2. If by day 6 of development, compacted cells as in Fig. 2e–e are not formed, extending embryo culture in complete HES medium for 1–2 additional days will selectively favour the growth of the ICM versus the TE. This approach is supported in the most recent literature [15]. Note 10. Regarding ICM isolation, some authors use an extended acid tyrode’s treatment to weaken the TE [26], thus facilitating the spreading of blastocyst onto the feeders. However, in our hands, the efficiency of this method depends greatly on the TE grading. Applying the same AT treatment to all embryos usually results in severe damage to some of them. Therefore, fitting the method for TE reduction to the grade of each embryo (Table 2) helps maximizing the recovery of viable cells from the ICM. When TE is relatively small compared to the ICM, or when the ICM is not compact, a pipetting method (Method A) is preferable. Alternatively, if the TE is robust and the ICM is compact, mechanical cutting (Method B) should be the method of choice. Speed during embryo denudation and ICM isolation is critical, even if the manipulations are performed on a heated stage. Note 11. If healthy, the first seeding of the cells from the ICMs will attach in 6 h to overnight. In our experience, those clumps needing more time to attach are not robust enough to yield derivations. The first outgrowth after the attachment of the clump can be seen from days 3 to 16. At this point, the undifferentiated cells need the support of fresh feeder layers to establish pluripotent cell growth. TE cells evolve invariably to form syncytial cells that invade the feeder layer and die leaving gaps in it. It is critical to have fresh feeders ready by the time the TE cells and derivatives detach from the plate. Acknowledgments This protocol is the result of work funded by the North West Development Agency (NWDA) in the UK and the MICINN-PLE2009-0091, IPT-20011-1402-900000 and FPI-CAIB Grant FPI10 grants in Spain. Conflict of interest interest.

The authors declare no potential conflicts of

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