Note: A few final edits were completed by 3 pm on Wednesday (4/24) afternoon.
Facilitating cell life, and when appropriate, death, is a key part of tisssue engineering. When cells are put into contact with a biomaterial (or into any novel culture condition), their viability may be affected. Some materials are cytotoxic, i.e., deadly to cells. Cytotoxicity typically varies with the concentration of one or more chemicals (such as a cross-linker) comprising the biomaterial, and varies in severity for different cell types. Cell density within a culture is another factor affecting cell livelihood, notably when the number of cells exceeds the nutrient concentrations available in the culture medium. In a 3D culture such as an alginate bead, sufficient nutrients and even oxygen may not be able to diffuse to the center of the bead prior to depletion by cells on the outer rim, even when at a high concentration in the bulk fluid. Finally, note that most cells require certain soluble and/or contact-dependent signals to remain viable. For example, immune cells called naïve T cells require the cytokine IL-7 and contact with self-MHC proteins for survival.
LIVE/DEAD® assay example.
Cell viability was monitored using fluorescent dyes that differ in their cell permeance and nucleic acid affinity. Fluorescence emission in the green and red (left) and red alone (right) channels is shown for the same field of cells.
Many assays are available to monitor the numbers of live and dead cells in a culture. The kit you will use today is made by Molecular Probes, a company (now part of Life Technologies) that makes a plethora of fluorescent cell stains for tracking viability, calcium flux, and other cell characteristics. The principle exploited by the LIVE/DEAD® kit is the relative permeability of cell membranes when the cell is live (intact membrane) or dead (damaged membrane). Ethidium is a nucleic acid stain that you are familiar with from running agarose gels in modules 1 and 2; the ethidium homodimer-2 variant emits red fluorescence, and cannot diffuse past intact cell membranes. The dye SYTO 10, on the other hand, is membrane-permeant, and thus enters both live and dead cells; it emits fluorescence in the green channel. SYTO 10 has lower affinity for nucleic acids than does ethidium, and thus is excluded from dead cells over time, enabling one to distinguish between live (green) and dead (red) cells. Viability can be inferred by monitoring parameters other than cell permeability. For example, some membrane-permeable dyes are only activated to a fluorescent form inside cells that have active esterase enzymes, a good indicator of the cells' metabolic activity. Assays that measure cell potentials or redox activity are also available. In general, fluorescence assays are more sensitive than colorimetric assays. Along with sensitivity, factors such stability, toxicity, and ease of scale-up are important considerations when choosing an assay.
Cell proliferation assay example.
Cells were stained with CFDA-SE and monitored by flow cytometry after several days. (A. Stachowiak)
Cell vitality (or lack thereof) tells only one part of a cell culture’s story. For example, kits like the one we are using today cannot determine whether the cells assayed have divided or not. However, other dyes are available that specifically test for cell proliferation, or even distinguish cells based on what part of the cell cycle they are presently in. Proliferation assays are important for drug development, cancer research, and in tissue engineering. Total nucleic acid content is sometimes used as a measure of proliferation – Hoechst is a popular dye for this purpose. Active proliferation can be monitored by addition of 5-bromo-2'-deoxyuridine (BrdU) to cell cultures. BrdU will be incorporated only in recently synthesized DNA (S-phase cells), and can be assessed by antibody-detection after a time lag. For tracking multiple cell divisions, long-lived fluorescent dyes such as the fluorescein derivative CFDA-SE are used: about 6-10 divisions can be seen by flow cytometry (see figure at right).
Remember that cell death is just as important as cell life, and that the type of death also matters. Cells that die due to acute trauma or other tissue damage typically die by necrosis: they swell and finally burst, releasing their contents and often promoting inflammation. Under other circumstances, particularly in development and immunity, many cells undergo a programmed death called apoptosis. Unlike the more disruptive necrotic cells, apoptotic cells condense and then fragment, finally releasing membrane-contained cell bodies. Apoptosis gone awry is implicated in many diseases, and thus researchers are very interested in tracking apoptotic cells in various culture systems. Special dyes can be used to track nuclear fragmentation and other changes in early and late apoptotic cells.
Your objective today is to determine the viabilities of your two different cell cultures, and to gain experience with fluorescence assays. You are likely to encounter fluorescence and other microscopy techniques in many fields of biological engineering research.
Today you can stagger your arrivals to lab (see today’s Talk page). Only one group at a time will be able to work on the microscope, and assuming that cell culture setup takes ~ 1 hour, you will each have ~20-25 minutes to spend on the microscope. Please be respectful of your labmates’ time. Reading the protocol in advance will help you work more quickly, and is strongly recommended.
Part 1: Bead preparation for Live/Dead® fluorescence assay
- Retrieve your 2 six-well dishes from the incubator.
- The teaching faculty counted your beads during a recent media exchange (they are easiest to count in the absence of media). Based on the numbers written on your plate, decide how many beads (1-3 per sample) you can spare for today's assay. Ideally, for the three assays on Day 4 you want at least 45-60 beads total remaining (perhaps 30 or fewer for large beads). Be sure to take your bead(s) from only one of the two wells, just in case you contaminate it.
- Also take this time to describe bead uniformity in your notebook, as this feature may affect your eventual experimental outcomes. Some groups had more luck than others in keeping bead size consistent between and within their two samples.
- During a later incubation step, you might also take a look at your plate under the microscope, and focus in on cells within the beads. What is cell morphology and density like in each sample? Are there any cells growing under the beads, as a monolayer on the surface of the plate? Keep in mind that these will compete for media nutrients with the cells inside the beads.
- Using a sterile spatula, remove the beads (keeping the two samples separate) to two labeled Petri dishes. Do your best to keep the beads remaining in the culture wells sterile – the cells have to stay alive for 5 more days. Briefly dip the sterile spatula into the well, and immediately return your plate to the incubator, onto the shelf from which you took it.
- Within the Petri dish, cut your whole beads in half using a spatula or razor blade.
- Small beads may be difficult to cut in half – if so, look at the intact bead instead.
- Per dish, rinse the beads with 3 mL of warm HEPES buffered saline solution (HBSS).
- Aspirate the HBSS - this may be easiest/safest to do with a P1000 - then pipet 200 μL of dye solution right on the beads.
- Incubate for 15 min. with the TC hood light off.
- Remove the entire supernatant with a pipet, and expel it in the conical tube labeled Dye Collection. The dye waste will be disposed of by the teaching faculty. You should also throw the pipet tip into the container on the microscope bench; tips will later be disposed of as solid waste in the chemical fume hood. You do not need to throw any later tips away here, as the dye will then be very dilute.
- Rinse the cells with 3 mL HBSS buffer again. Pipet off as much liquid as possible, again into the Dye Collection tube.
- Soak in 3 mL of 4% glutaraldehyde solution for 15 minutes.
- Pipet off the solution, and then bring your Petri dish to the fluorescent microscope bench in the lab.
- For observation, place the half-bead on a glass slide and then cover with a coverslip -- don't press down too hard.
- You will probably want to look at the beads both flat side up (to see the core) and flat side down (to see the surface), time permitting.
- You can make a "map" of the beads in your notebook and/or on the white surface of the slide. For example, you might have one bead on the left that is core side up and another on the right that is surface side up.
Part 2: Microscopy
When observing your cells under fluorescence excitation, you should work with the room lights off for best results. A member of the teaching faculty will be with you to help you make the most of your 20-25 minutes.
- Prior to the first group using the microscope, the teaching faculty will turn on the microscope and allow it to warm up for 15-20 min. First, on the mercury lamp that is next to the microscope, the ‘POWER’ switch will be flipped. Next, the ‘Ignition’ button will be held down for about a second, then released.
- When you arrive, the lamp ready and power indicators should both be lit – talk to the teaching faculty if this is not the case.
- Place your first sample slide on the microscope, coverslip-side up, by pulling away the left side of the metal sample holder for a moment.
- Begin your observations with the 10X objective.
Fluorescent microscope, front view.
Fluorescent microscope, side view.
- Turn on the illumination using the button at the bottom left of the microscope body (on the right-hand side is a light intensity slider).
- Next, turn the excitation light slider at the top of the microscope to ‘DIA-ILL’ (position 4).
- Try to focus your sample. However, be aware that the contrast is not great for your cells, and you might not be able to focus unless you find a piece of debris. Whether or not you find focus, after a minute or two, switch over to fluorescence. Your cells will be easier to find this way.
- First, turn the white light illumination off.
- Next, move the excitation slider to ‘FITC’ (position 3). You should see a blue light coming from the bottom part of the microscope.
- This light can excite both the green and the red dye in the viability kit, and the associated filter allows you to view both colors at once.
- Finally, you must slide the light shield (labeled ‘SHUTTER’) to the right to unblock it. Now you can look in the microscope, and use the coarse focus to find your cells (which should primarily be bright green), then the fine focus to get a clearer view.
- You can also switch the excitation slider over to ‘EthD-1’ (position 2) to see only the red-stained cells. Some of your cells may appear to be dimly red, but the dead ones are usually obviously/brightly stained.
- Be aware that the dyes do fade upon prolonged exposure to the excitation light, so don’t stay in one place too long, and when you are not actively looking in the microscope, slide the light shield back into place.
- You can try looking at your cells with the 40X objective as well if you have time. As you move between objectives and samples, choose a few representative fields to take pictures of. As a minimal data set, try to get 3 fields at 10X of both of your samples.
- To take a picture, remove one eyepiece from the microscope, and replace it with the camera adaptor. Be sure to keep the light shield in place until you are ready to take the picture (to avoid photobleaching)!
- Note that 10X images will reveal a broader field, but 40X images may have better contrast.
- Check with the teaching faculty if you are having difficulty getting clear pictures.
- Later in the module, you will compare the average cell numbers in each sample using the statistical methods we discussed during Module 2.
- Post two well-captioned pictures to the wiki before leaving (one of each sample), so we can discuss the class data in our next lecture. Be sure to note whether the image is at the surface or core of the bead.
- If you are one of the last two groups to use the microscope, you may post your data within 24 hours instead.
For next time
- The first time this module was run, students created single-cell suspensions from their alginate beads by dissolving said beads in EDTA-citrate buffer, and only then stained the cells. What additional kind(s) of information about cell viability do you gain by staining whole constructs rather than cell isolates? Try to describe two.
- Read the editorial by Professor Alan Russell about standards in tissue engineering, and come to lecture next time prepared to discuss and/or write about your thoughts. You may find other articles at this link helpful.
- The primary assignment for this experimental module will be for you to develop a research proposal and present your idea to the class. For next time, please describe five recent findings that might define an interesting research question. You should hand in a 3-5 sentence description of each topic and cite an associated reference from the scientific literature. The topics you pick can be related to any aspect of the class, i.e. DNA, protein, or cell/biomaterial engineering. During lab next time, you and your partner will review the topics and narrow your choices, identifying one or perhaps two topics for further research.
- Note: for now, you do not have to have a novel research idea sketched out; you simply have to describe five recent examples of existing work. However, you can start to brainstorm how to build off of those topics into something new if you want to get ahead of the game.
- HEPES-buffered saline solution (HBSS)
- 135 mM NaCl
- 5 mM KCl
- 1 mM MgSO4
- 1.8 mM CaCl2
- 10 mM HEPES
- pH 7.4
- 4% glutaraldehyde (GAH) in HBSS
- original stock GAH at 50%, reagent grade
- Live/Dead Reduced Biohazard Viability/Cytotoxicity kit from Invitrogen
- SYTO 10 green nucleic acid stain
- ethidium homodimer-2 red nucleic acid stain
- original dye stocks in DMSO, diluted in HBSS 1:250