Human Stem Cell Manual: A Laboratory Guide

Stem cells are self-replicating and undifferentiated, meaning their function is not yet cell, tissue, or organ-specific. Due to the unique nature of these cells, research into their biology and function holds great promise for therapeutic applications through replacement or repair of diseased and damaged cells. This reader-friendly manual provides a practical "hands on" guide to the culture of human embryonic and somatic stem cells. By presenting methods for embryonic and adult lines side-by-side, the authors lay out an elegant and unique path to understanding the science of stem cell practice. The authors begin with a broad-based introduction to the field, and also review legal and regulatory issues and patents. Each experimental strategy is presented with an historical introduction, detailed method, discussion of alternative methods, and common pitfalls. This lab guide for researchers also serves as a textbook for undergraduate and graduate students in laboratory courses.

• Language: English
• Publisher: Academic Press
• Release date: Oct 10, 2011
• ISBN: 9780080549880

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Part I

Basic Methods in Stem Cell Culture

Outline

Chapter 1: Human Embryonic Stem Cell Culture

Chapter 2: Human Feeder Cells, Feeder-free, and Defined Culture Systems

Chapter 3: Mouse Embryonic Fibroblast Feeder Cells

Chapter 4: Cryopreservation of Human Embryonic Stem Cells

CHAPTER 1

Human Embryonic Stem Cell Culture

Rodolfo Gonzalez and Jeanne F. Loring,     Stem Cell Center Burnham Institute for Medical Research 10901 N. Torrey Pines Rd La Jolla, CA 92037, USA

Robin L. esselschmidt,     Center for Stem Cell and Regenerative Medicine, The University of Southern California 1501 San Pablo, ZNI 543 Los Angeles, CA 90033, USA

Philip H. Schartz,     Center for Neuroscience Research Children’s Hospital of Orange County Research Institute 455 South Main Street Orange, CA 92868-3874, USA

Publisher Summary

This chapter outlines protocols for the culture of human embryonic stem cells (hESCs). hESC lines ere originally derived using very similar culture medium, and conditions as those developed for the derivation, and culture of mouse embryonic stem cells (ESC) lines. Hoever, these methods ere suboptimal for hESCs, and have evolved considerably in the years since hESC lines ere derived. Compared ith mouse ESCs, hESCs are very difficult. Culturing hESCs requires a significant commitment of time, and resources. It takes eeks to establish a culture, and the cultures ill require daily attention. hESCs unlike mouse ESCs, do not survive ell hen dissociated to single cells. Therefore, the most reliable method for passaging undifferentiated hESC cultures is manual dissection of the colonies.for

INTRODUCTION

Culturing human embryonic stem cells (hESCs) requires a significant commitment of time and resources. It takes eeks to establish a culture, and the cultures ill require daily attention. Once hESC cultures are established, they can, ith skill and the methods described belo, be kept in continuous culture for years.

A ord of caution for those ith experience culturing mouse embryonic stem cells: they are not the same! Both mouse and human ESCs are diploid, they are pluripotent, and they are relatively stable in culture. Hoever, the stability of mouse ESC lines is regularly measured because the objective of almost all genetic manipulation is to make ne lines of mice. If an ESC line can generate a mouse, as e term it, go germline, e kno that it is clearly pluripotent. This has given us an operational definition for pluripotence and stability in culture for mouse ESCs.

hESC lines ere originally derived using very similar culture medium and conditions as those developed for the derivation and culture of mouse ESC lines. Hoever, these methods ere suboptimal for hESCs, and have evolved considerably in the years since hESC lines ere derived. Compared ith mouse ESCs, hESCs are very difficult to culture – they gro sloly, and most importantly, since e have no equivalent assays for germline competence, e cannot assume that the cells that e have in our culture dishes are either stable or pluripotent. This makes it far more critical to assay the cells frequently, using characterization methods such as the karyotyping, immunocytochemistry, gene expression analysis, and fluorescence activated cell sorting (FACS) methods provided in this manual.

OVERVIE

In this chapter e outline protocols for the culture of hESCs, starting as one ould usually do, by being handed a culture by an experienced colleague. Other chapters focus on cryopreservation and establishing hESC cultures from frozen stocks, and on the variety of culture conditions, including the preparation of various types of feeder layers, conditioned medium, and extracellular matrix substrata.

The methods e recommend are those that are the most straightforard and have orked ell in our hands; these are offered as the recommended methods and reagents. e also offer alternative methods and reagents that ork but are not routinely used in most laboratories. The key variables that e outline in this chapter are:

 Culture medium

– Rasai medium

– Serum or serum substitute

 Passaging cells

– Manual passage

– Non-enzymatic dissociation.

– Enzymatic dissociation.

While optimizing and standardizing conditions in your lab, it is important to keep in mind that changing one thing in a system may have unexpected impact on the entire system.

PROCEDURES

Tips for successfully culturing hESCs

 Feed cells every day, except for 1 or 2 days folloing passage.

 Examine the cultures every day under 4× and 10× phase contrast. This ill allo you to become familiar ith the morphologies of undifferentiated and differentiated cells and colonies.

 When they are cultured on feeder layers some hESC lines tend to undergo spontaneous differentiation in the centers of the colonies. hen passaging, take care to avoid passaging these differentiated centers to the ne culture.

 Most hESC lines double every 31–35 h.

 Store medium at 4°C and discard any unused medium after 10 days. Best results are achieved hen medium is prepared in small batches once a eek.

Recognizing hESC morphology

The single most important skill in successful culturing of hESCs may be the ability to recognize the morphology of undifferentiated cells under a variety of conditions (Figure 1.1).

FIGURE 1.1 Phase contrast micrographs from the same culture, 4 days after it as passaged onto a feeder layer (human foreskin fibroblasts, ATCC HS27). (A) Typical colonies ith smooth, phase-bright edges, ith the fibroblast feeder layer forming horls around the colonies (10× magnification). In contrast, in the same culture there are colonies ith obvious differentiation at the edges (B − 4× magnification) and in the center. (C − 10× magnification). In selecting colonies for passage and expansion, only the ones shon in (A) ould be acceptable. The others should not be passaged to the next culture dish.

For routine expansion of hESCs, e recommend that the cells be cultured at a relatively lo density so that individual colonies can be easily monitored and selected against differentiation. hESCs can be cultured to high density (Figure 1.2), but a higher proportion of differentiated cells must be expected.

FIGURE 1.2 hESCs can be cultured to high density.

Passaging hESCs

hESCs, unlike mouse ESCs, do not survive ell hen dissociated to single cells. Therefore, the most reliable method for passaging undifferentiated hESC cultures is manual dissection of the colonies. This method may seem tedious, but it is virtually foolproof and e recommend that novices use this method until they have familiarity ith the cells and can easily recognize differentiation in the culture. e also recommend manual passaging for producing cell banks of lo-passage hESCs. Enzymatic dissociation methods are provided in Alternative Procedures.

Note: Using the number of passages as a measure of the age of an hESC line is an unfortunate historical accident. Because of the inconsistencies in hESC culture procedures in different labs, cells are passaged at different time intervals, ranging from 4 to 7 days. Therefore the number of passages for one line might be tice that of another, even though the cells have been in culture for exactly the same amount of time. For example, in a year of continuous culture, a cell line could be passaged as fe as 52 and as many as 90 times. A better measure ould be the number of doublings, but to count the number of cells in a culture is difficult since the cells form tight clusters and are not passaged as single cells.

General guidelines

 The cells should be passaged at about 1:3 every 5–7 days.

 Prepare the feeder layer or extracellular matrix (ECM) substrata the day before passaging.

 Depending on the cell line, passaging on Friday may be a good routine. The cells can usually be left undisturbed for 2 days folloing passaging, hich allos them to settle don on the substrata, attach and begin dividing before the medium is changed.

 There ill be considerable variation in the size of colonies in a single dish. Compared ith their mouse counterparts, hESCs do not substantially pile up on each other, and their colonies can gro to a large diameter hile remaining undifferentiated. Culture conditions affect the flatness of the colonies, but as an approximation, they are ready to split hen the diameter fills the 10× field hen observed under the microscope. As shon in Figure 1.3, a colony about half the diameter of the 10× field contains about 4400 cells. A colony filling the field ould contain about 15 000 cells.

FIGURE 1.3 A colony about half the diameter of the 10× field contains about 4400 cells.

 For routine passaging by any method, do not make a single-cell suspension; dissociate the colonies into smaller colonies of a fe hundred cells.

 Examine the culture daily for colony morphology under the phase contrast or dissecting microscope.

 With experience, one can get a good overvie of colony morphology by holding the dish up to a light and looking at the bottom of the dish. The differentiated colonies ill have ragged edges and hollo centers.

 On the bottom of the dish, mark colonies that are badly differentiated or parts of the colony that you do not ish to transfer to a ne culture dish.

 To be certain that the colonies selected are undifferentiated, it is advisable to dissect the colonies hile vieing the dish under a dissecting microscope ith illumination from the base. But this is not absolutely necessary, and some prefer to passage the cells ithout magnification.

Mechanical dissociation

1. Evaluate the culture under 4× or 10× phase contrast optics.

2. The cells can be split among 3–6 dishes of the same size as the original culture, depending on the density of the original culture. If you ish to put the cells in different-sized dishes, calculate the amount of volume to add based on surface area of each type of dish.

3. Mark (or remove) overtly differentiated colonies so as not to disturb these during the dissociation process.

4. Remove the medium from the dish and replace ith fresh hESC medium.

5. Dissect the colonies by hand, either under a lo-poer dissecting microscope (in a horizontal flo hood) or ithout a microscope, in the tissue culture hood.

NOTE: Several implements can be used to slice up or break up the colonies. Because they are inexpensive and sterile, e recommend either a 20 μL pipettor hich has a sterile filter tip attached, or a sterile 23G needle.

6. Figure 1.4 shos the method used for slicing the colonies into about 100 pieces. The colony is cut into strips, and then into squares. Each piece of the colony has a fe hundred cells.

FIGURE 1.4 Frames from a movie, shoing the cutting up of an hESC colony for passaging.

7. Break up each colony by moving the tip around and across each colony in a crosshatch or a spiral motion.

NOTE: Since the colonies are large at the time of passage, it is relatively easy to see individual colonies on the plate and, ith practise, one can quickly dissociate an entire plate in less than 20 min.

8. After all of the colonies are dissected, use a 5 mL pipette to transfer the culture medium containing the dissected colonies to a 15 mL conical tube. Rinse the plate ith hESC medium and add this to the same 15 mL tube.

9. Bring up the final volume in the tube to 8–10 mL ith hESC medium.

10. Gently triturate the cell clumps using a sterile 10 mL pipette and divide the suspension into the prepared culture dishes on feeder layer or ECM-coated plates. Do not make a single-cell suspension; triturate gently, trying to achieve a relatively uniform suspension of cell clumps containing a fe hundred cells each.

ALTERNATIVE PROCEDURES

Enzymatic dissociation

Enzymatic dissociation methods vary idely, and the exact conditions need to be developed for each laboratory. Most importantly, cultures that have been maintained by manual passaging cannot be passaged by enzymatic dissociation unless exceptional care is taken to adapt and monitor the cells.

The type of enzyme used for dissociation is critical. For example, passaging ith trypsin appears to put more selective pressure on the cultures than other methods, resulting in a higher incidence of drift of hESC lines toard aneuploidy. But some hESC lines have been derived using trypsin from the outset; these lines can be rountinely passaged using hatever enzymatic technique is provided by the supplier.

Microbial collagenase is preferred by many laboratories, perhaps because of the ay in hich it is used. Collagenase is used to loosen the hESC colonies from the dishes, not to dissociate them to single cells, and the cell clumps have to be further dissociated by trituration.

Note: Keep in mind that enzymes are not highly purified recombinant products, and they may contain animal products. Trypsin is prepared from porcine (pig) tissue, and collagenase is a crude microbial product.

Collagenase dissociation

1. Remove the culture medium.

2. Rinse culture ith Dulbecco’s PBS (D-PBS).

3. Treat the culture ith 200 U/mL of collagenase IV for 5–10 min at 37°C until the edges of the colonies start to curl up – observe the culture under the microscope.

4. Remove the collagenase and replace ith 2 mL of hESC medium (if using a six-ell or 35 mm dish).

5. Using a 5 mL pipette, gently dislodge the good colonies from the plate and place them in a 15 mL conical tube. Alternatively, one could remove the differentiated colonies prior to treating the culture dish ith collagenase.

6. Gently triturate the cell clumps using a sterile 10 mL pipette and plate on feeder layer- or ECM-prcpared dishes. Try to achieve a relatively uniform suspension of cell clumps containing several hundred cells each.

7. The cells can be split among 3–6 dishes of the same size as the original culture, depending on density of the original culture. If you ish to put the cells in different sized dishes, calculate the dilution based on surface area of each type of dish.

Non-enzymatic cell dissociation

Ca²+- and Mg²+-free saline solutions containing EDTA or EGTA have not been as idely used for hESC dissociation as the methods described above, but they should offer advantages for assays that require intact cell surface proteins such as flo cytometry and immunocytochemistry. Commercial formulations are available, such as Cell Dissociation Buffer (Invitrogen catalog no. 13150016), hich contains glycerol as ell as a proprietary mixture of salts and chelators.

If you decide to try this method, remove all of the protein-containing medium and rinse the cells briefly ith the dissociation buffer. Add enough buffer to cover the cells and monitor them under the microscope until the edges of the colonies begin to lift, then triturate the cells gently to dissociate. If the cells are to be recultured, don’t dissociate them into single cells, and be certain to check the karyotype of the cells after 10 passages; until you prove otherise, you should assume that any untested passaging method is selecting for chromosomal abnormalities.

Other enzymes

Accutase and Accumax (Millipore/Chemicon catalog no. SCR005 and SCR006)

These products are proprietary mixtures of proteolytic and collagenolytic enzymes in EDTA that the manufacturer states is free of mammalian- or bacterial-derived products. Accumax also contains DNAse. If you test this method, start ith a 5-minute room temperature incubation and monitor the cells under the microscope. hile the manufacturer indicates that inactivation of the enzymes ith protein is not necessary, e recommend that protein-containing medium be used to dilute out the enzyme after the cells are dissociated, to prevent clumping and sticking of the cells to the pipettes.

Trypsin-like Enzyme (TrypLE Select, Invitrogen catalog no. 12563-029)

This is a single enzyme, a recombinant fungal serine protease ith trypsin-like activity. Anecdotal reports suggest that hESC line that have been mechanically passaged can be successfully transitioned to single-cell enzymatic passaging using TrypLE Select. If you decide to try this method, e recommend a saline rinse, then a 5-minute incubation in the 1× enzyme solution as provided by the manufacturer. Monitor the cells under the microscope and add protein-containing medium to the culture before triturating.

HyQTase (HyClone catalog no. SV30030.01)

This is a cell detachment solution in D-PBS ith EDTA. The composition is proprietary. According to the manufacturer, HyQTase is composed of a naturally derived complex of proteolytic and collagenolytic enzymes in D-PBS containing EDTA. According to the manufacturer it can be used for either serum-containing or serum-free cultures. The manufacturer states that it does not contain mammalian or bacterial derived products and is non-recombinant.

PITFALLS AND ADVICE

Monitoring drift in hESC cultures

Since hESC cultures are often kept in continuous culture for months, even years, it is very important to monitor for drift in the cultures. The best ay to avoid drift is to generate a large bank of frozen cells as soon as possible after the cultures are first expanded. The importance of this cannot be overemphasized; the value of discoveries based on hESC cells depends on the reproducibility of results. See Chapter 26 for methods for setting up an hESC lab.

Genetic drift

We know that hESCs acquire chromosomal abnormalities over long periods of culture, so karyotyping or other genetic analysis methods must be performed on a regular basis. For detailed information about ho to monitor genetic drifts, see Chapters 5–7 and 26.

Keep in mind that changes during the time the cells are cultured in your lab can only be detected if you analyze the cells very soon after you obtain them.

Developmental drift

hESCs can also drift toard a more differentiated state over periods of extended culture. Since there is no assay for pluripotence equivalent to germline transmission of mouse ESCs, surrogate markers, such as antibody markers, should be routinely checked, especially if the morphology of the cells seems to be different from the earlier cultures.

The gold standard for measuring the pluripotency of an hESC line is to transplant it to an immune-deficient mouse to form a teratoma tumor (Chapters 12 and 13). Keep in mind that it ill require histological expertise to identify cell types and tissues in the tumors.

In vitro, differentiation of hESCs using embryoid body culture ill allo at least a cursory analysis of hESC differentiation potential. Hoever, embryoid bodies never achieve the maturity of cells that develop in teratomas, and since the methods used to assess differentiation in vitro usually involve a small number of markers assayed by PCR (Chapter 10) or immunocytochemistry (Chapter 9), it is more difficult to judge the full range of pluripotence.

The best approach to monitoring developmental drift is to pick a particular method and differentiated cell type to check periodically (see Chapters 14 on embryoid body and neuroepithelial differentiation, as ell as the specific chapters on neuronal, cardiac, and hematopoietic cells, Chapters 15–18).

Contamination of cultures

hESCs are usually cultured ithout antibiotics; ith good culture technique, bacterial contamination should not be a problem. Hoever, e recommend that antibiotics be used hile ne investigators are being trained in the techniques. Antibiotics such as penicillin and streptomycin do not have any effect on mycoplasma. Mycoplasma is a serious problem in laboratories that culture multiple cell lines or have inadequately trained personnel. Cultures must be monitored for mycoplasma on a regular basis, and contaminated cultures destroyed. Methods for mycoplasma detection are provided in the quality control section of this chapter, and in Chapter