Difference between revisions of "20.109(S08):Initiate cell culture (Day2)"
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==Introduction== | ==Introduction== | ||
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+ | Last time you proposed culture conditions for an investigation of chondrocyte phenotype maintenance, and today you will initiate said cultures. The cells you are using are freshly derived from bovine cartilage: cells from cows are often used in part because of their availability from abattoirs. In general, large animals are more useful for modeling human joint diseases such as osteoarthritis than are small animals, because the resting angle of their knee joints is more similar to that of humans. In this module, we will work with an ''in vitro'' culture model of cartilage-forming cells. | ||
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+ | Two of your three cell samples will be grown in alginate bead cultures. You have probably encountered alginates many times in your life, as thickeners in food and textiles, preservatives, and possibily at your dentist or in a pharmacy. Alginate is a polysacharride derived from seaweed, a co-polymer of mannuronic and guluronic acid. A single alginate molecule may contain long stretches of either acid (called M-blocks and G-blocks), as well as random and/or strictly alternating G/M sequences. The precise chemical composition of an alginate determines its mechanical properties, degradability, and other important characteristics. Qualities such as strength and viscosity are also influenced by the average length of the individual polymer chains (i.e., the molecular weight), and by alginate concentration. For example, high molecular weights correlate with increased viscosity. Alginates in general are shear-thinning, which is to say their viscosity decreases as shear rate increases (e.g., when quickly drawn into a syringe). | ||
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+ | [[Image: 20.109_alginate-crosslink.png|thumb|175px|'''Schematic of crosslinked-alginate.''' G-blocks are represented by dotted lines, M-blocks by curved solid lines, and calcium ions by green circles.]] | ||
+ | Cations such as calcium can cross-link alginate chains to form a network, or gel. The identity and concentration of the cross-linker influence the ultimate material properties. Only G-blocks can be linked to each other, while M- or MG-blocks cannot, but in turn provide flexibility (see figure). The resultant semi-solid structure has the capacity to hold a large amount of water, and the water-swollen structure is called a hydrogel. Hydrogels have several attractive properties for tissue engineering: they allow oxygen and nutrients to diffuse better than non-hydrated materials do; their mechanical and biochemical properties are readily varied by co-polymerization of multiple elements; they mimic the elasticity of natural tissues, and they often form rapidly and under mild conditions. Some gels can be injected into a patient in liquid form, then solidified within his or her body by heat or light. Such injectable gels have the advantage of easily filling an arbitrarily sized wound shape, which is difficult for implantable gels to do. Natural (e.g., alginate) and synthetic (e.g., poly(ethylene glycol)) hydrogels each have distinct advantages and disadvantages, as we will discuss in class. | ||
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+ | Today you will make alginate hydrogels in bead form, by slowly releasing alginate solution from a syringe into a bath containing calcium chloride. Next time you will see how well your cells survive in both alginate and monolayer culture. | ||
==Protocols== | ==Protocols== | ||
− | Half the class at a time will work in the tissue culture room today. The other half | + | Half the class at a time will work in the tissue culture room today. The other half of you will explore the NCBI bovine information site, and otherwise spend the time however you find useful (FNT assignment, notebook prep, or unrelated work). |
===Part 1: Chondrocyte cell culture=== | ===Part 1: Chondrocyte cell culture=== | ||
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#The digested cell solution is filtered into 2 tubes using a cell strainer with a 70 μm cutoff size. This is to remove undigested debris. | #The digested cell solution is filtered into 2 tubes using a cell strainer with a 70 μm cutoff size. This is to remove undigested debris. | ||
#A second filtration is done at a 40 μm cutoff size, to remove finer debris. (What else might be removed at this stage, for better or for worse?) | #A second filtration is done at a 40 μm cutoff size, to remove finer debris. (What else might be removed at this stage, for better or for worse?) | ||
− | #The cells are centrifuged for | + | #The cells are centrifuged for 6 min at 1900 g. |
#The enzymatic solution is aspirated, and the cells are initially resuspended in a 1 mL volume of sterile PBS, then combined in one tube at 25 mL total. (What would happen if you tried to resuspend the cells in a large volume right away?) | #The enzymatic solution is aspirated, and the cells are initially resuspended in a 1 mL volume of sterile PBS, then combined in one tube at 25 mL total. (What would happen if you tried to resuspend the cells in a large volume right away?) | ||
#The cells are spun down and resuspended in sterile PBS again. | #The cells are spun down and resuspended in sterile PBS again. | ||
#Finally, they are spun down and resuspended in pre-warmed culture medium, 20 mL total. | #Finally, they are spun down and resuspended in pre-warmed culture medium, 20 mL total. | ||
− | #We will now count the cells, diluting 50 μL in 200 μL PBS, then mixing 1:1 with Trypan blue. Each group will be given one-third of the cells. | + | #We will now count the cells, diluting 50 μL in 200 μL PBS, then mixing 1:1 with Trypan blue. Each group will be given one-third of the cells. |
====Cell culture==== | ====Cell culture==== | ||
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#The cells intended for your 2D culture probably do not need to be spun down again, but the cells for your 3D cultures must be directly resuspended in alginate rather than culture medium. While you spin down your alginate-bound cells, set up your 2D cultures according to the directions below. | #The cells intended for your 2D culture probably do not need to be spun down again, but the cells for your 3D cultures must be directly resuspended in alginate rather than culture medium. While you spin down your alginate-bound cells, set up your 2D cultures according to the directions below. | ||
#Label two T25 flasks with your group colour, initials, the date, and a description of the cells and culture conditions. Now make your cell dilution, using the example below as a guide - just change the volumes according to how many cells you want to add (always making 10 mL of culture total). | #Label two T25 flasks with your group colour, initials, the date, and a description of the cells and culture conditions. Now make your cell dilution, using the example below as a guide - just change the volumes according to how many cells you want to add (always making 10 mL of culture total). | ||
− | #* | + | #*Let s say you have 6M cells/mL in your conical tube, and want to use 600,000 cells to initiate your 2D cultures. Stand your T25 flasks upright and take the caps off. Start by adding 100 μL of chondrocytes against the back wall of each of your T25 flasks, making sure to touch only the sterile pipet tip to the flask, and to avoid touching the pipetman body inside this sterile area. (By back wall I mean what will become the bottom inside wall when you lay the flask down, which is the surface on which your cells grow.) |
#*Now add 9.9 mL of warm culture medium into each flask, and triterate to mix the cells. | #*Now add 9.9 mL of warm culture medium into each flask, and triterate to mix the cells. | ||
#*Finally, tighten the cap back onto the flask and put the cells in the incubator. | #*Finally, tighten the cap back onto the flask and put the cells in the incubator. | ||
#Now you are ready to initiate the alginate cultures. Label two six well plates with the same information as for your 2D cultures, using one plate per culture type (e.g., plate 1 = low cell density, plate 2 = high cell density). | #Now you are ready to initiate the alginate cultures. Label two six well plates with the same information as for your 2D cultures, using one plate per culture type (e.g., plate 1 = low cell density, plate 2 = high cell density). | ||
#Resuspend your cells in the appropriate amount of the type and concentration of alginate that you chose. | #Resuspend your cells in the appropriate amount of the type and concentration of alginate that you chose. | ||
− | #Using the syringe that has been prepared for you, very carefully pull up the cells, then release them drop-by-drop into the beaker full of calcium chloride solution. Recall that calcium effectively polymerizes the alginate, resulting in small gel beads filled with cells. '''Immediately discard the entire syringe into the sharps container - do not try to remove or recap the needle.''' | + | #Using the syringe that has been prepared for you, very carefully pull up the cells, then release them drop-by-drop into the beaker full of calcium chloride solution (20 mL). Recall that calcium effectively polymerizes the alginate, resulting in small gel beads filled with cells. '''Immediately discard the entire syringe into the sharps container - do not try to remove or recap the needle.''' |
#*Depending on the concentration of alginate that you chose, you may have between ~50-150 beads for 1 mL of alginate solution. | #*Depending on the concentration of alginate that you chose, you may have between ~50-150 beads for 1 mL of alginate solution. | ||
#Allow the polymerization to proceed for 10 min. at room temperature. Then pour your beads into a 50mL conical tube. | #Allow the polymerization to proceed for 10 min. at room temperature. Then pour your beads into a 50mL conical tube. | ||
#Remove the calcium chloride solution from your beads using a large serological pipet (to better avoid aspirating the beads), and put this solution in the temporary waste beaker in your hood. | #Remove the calcium chloride solution from your beads using a large serological pipet (to better avoid aspirating the beads), and put this solution in the temporary waste beaker in your hood. | ||
− | #Now fill the conical tube with sodium chloride, and gently shake it for 1-2 min. This is to remove excess calcium from the solution. | + | #Now fill the conical tube with sodium chloride (20 mL), and gently shake it for 1-2 min. This is to remove excess calcium from the solution. |
− | #Remove the NaCl using a fresh pipet, then wash the beads again with fresh NaCl. Finally, wash the beads two times with DMEM culture medium. | + | #Remove the NaCl using a fresh pipet, then wash the beads again with fresh NaCl. Finally, wash the beads two times with DMEM culture medium, first with the additive-poor and then with the additive-rich DMEM (20 mL each time). |
#For each of your two samples, transfer the beads to the two leftmost wells of a 6-well plate, using a sterile spatula. Try to put approximately equal numbers of beads in the two wells. | #For each of your two samples, transfer the beads to the two leftmost wells of a 6-well plate, using a sterile spatula. Try to put approximately equal numbers of beads in the two wells. | ||
#Finally, add 6 mL of warm culture medium to your four sample wells, then put the two well-plates in the incubator. | #Finally, add 6 mL of warm culture medium to your four sample wells, then put the two well-plates in the incubator. | ||
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Did you know that NCBI has a whole site devoted to [http://www.ncbi.nlm.nih.gov/projects/genome/guide/cow/ all things cow]? | Did you know that NCBI has a whole site devoted to [http://www.ncbi.nlm.nih.gov/projects/genome/guide/cow/ all things cow]? | ||
− | + | It s true! And today you will use this site to find the primers you need to perform RT-PCR on Day 4 of this module. Try searching for collagen types I and II (the alpha chain of each is fine) in the '''Map Viewer''' (upper right of page). What chromosomee is each collagen chain located on? See if you can make your way to the UniSTS entries for collagen, which list recommended primers for RT-PCR. How long are the expected PCR products if these primers are used? | |
− | Another option for finding primer suggestions is looking in the literature. Of course, this can be a risky proposition, but if you verify the primers against information in the NCBI database, it can be faster than making your own from scratch, and provide a feeling of security (someone, somewhere has succesfully amplified the sequence in question!). The paper by [ |Ikenooue et al.] lists primers recommended for collagen type II. What species are the primers for? If | + | Another option for finding primer suggestions is looking in the literature. Of course, this can be a risky proposition, but if you verify the primers against information in the NCBI database, it can be faster than making your own from scratch, and provide a feeling of security (someone, somewhere has succesfully amplified the sequence in question!). The paper by [ |Ikenooue et al.] lists primers recommended for collagen type II. What species are the primers for? If it's not bovine, you cannot use the primers directly. However, you can BLAST the primers against the bovine genome, similar to what you did in Module 2 to verify your mutagenized plasmids against the original. |
− | Go to the [http://www.ncbi.nlm.nih.gov/blast/Blast.cgi BLAST site] and select the bos taurus genome. Type in the primers from the journal article one at a time, then perform the BLAST as follows: select BLASTN, change the | + | Go to the [http://www.ncbi.nlm.nih.gov/blast/Blast.cgi BLAST site] and select the ''bos taurus'' genome. Type in the primers from the journal article one at a time, then perform the BLAST as follows: select BLASTN, change the |