Working in the tissue culture lab
General lab etiquette
- Wear gloves and lab coat.
- Label your plates: initials + date + 20.309 + cell line (+ # cells + other comments).
- Bring nothing from regular lab into cell culture lab, and vice versa.
- Leave all hoods empty.
- Never user pipetters (= pipet pens) in stock bottles; rather only use glass serological pipets.
- The end-point is the large red burn box (medical waste), BUT no liquids, no sharps belong therein.
- Anything that has been in the hood must be discarded in burn box!!! (gloves, paper wraps, towels too)
- In Sharps container, dispose only of really sharp needles, Pasteur pipets, razor blades, …,
- while pipet tips can go directly in burn box (spray down temporary sharp waste container in hood).
- Trypan blue, if diluted with cells (1:10 typically), if in small amounts (< 1 mL), and if capped well (in eppendorf tube typically) can now be tossed in big red burn box.
- To recycle media bottles or tip boxes: Bleach a lot, then soap a lot, then recycling is OK.
- Water at the bottom:
- Autoclaved water only + 2 drops of algicide
- Some, but level below diamond
- Replace every semester all water
- Don't talk into incubator!
- The water bath in the 20.109 tissue culture room warms up fast (10 min), don't leave it on.
- Don't leave items inside.
- Good amount of algicide (20 drops).
- When you turn it on, press Enter arrow until 37 displayed; now it's heating.
- Only warm up aliquots of media, PBS, etc (not the entire 500 mL stock bottle:
- Medium gets basic when in contact with air, plus L-glutamine in cell culture media spontaneously degrades.
- So first open the hood, give it 3 minutes to settle on steady air flow and be sterile, next aliquot adequate amounts in the hood, and then warm this smaller amount in water bath.
- The rcf units are wrong in 20.109 tissue culture room. Therefore dial more g than required by protocol for the real spin to take place.
- Sleeves for different sizes of Falcon tubes, and balances are available.
- Blower always on, vented at the bottom (20.109 students, beware: this type of hood is different from regular ones at MIT!)
- Open slash to just above red arrow mark.
- Give it 3 minutes to settle on steady air flow and be sterile before working in the hood.
- If blower was off in the first place, then open the slash, wait 10 minutes before working in hood.
- Clean hood: wipe with 70% ethanol (some bacteria can leave in 100% ethanol environment), everywhere, even under stuff, wipe with paper towel (Kimwipes expensive…).
- Air curtain: the air comes down, then gets split to front and back of the hood.
- Try not to disturb air flow, yet still work!
- Everything (except cells) gets sprayed with ethanol and wiped (no dripping) before it gets into the hood.
- No UV (only once in a while overnight, only efficient for 10 hours!)
- Work 6" inward, away from glass.
- Limit hands in-out travels.
- Place nothing on grids, nor pushed all the way to the back.
- Spray 70% ethanol onto everything that goes in.
- Spray down your gloves regularly.
- Vacuum tube: Wipe from top to bottom, and add Pasteur pipet to aspirate liquids.
- Fluid waste (~maybe once a week): 10% bleach (in conical side-arm tube) for 20 minutes, then down the drain.
- If pipet aid stops working, check filter first. Don't sip up too fast! (especially with 2 mL pipets).
- Keep containers closed unless adding to / taking from them.
- Finish with bleach at the very end: aspirate through vacuum tubing: 50-50 bleach-water.
For 1 L of DMEM++
- In the hood, combine
- 1 L DMEM
- 100 mL FBS
- 10 mL Penicillin/Streptomycin/Glutamine
- 10 mL non-essential amino acids
- 10 mL sodium pyruvate
- Filter into sterile container.
- Label as DMEM++ and store at 4°C.
- One 16% ampoule
- 3.3 mL 1x PBS
Store in dark at room temperature.
0.1% Triton X-100
- 0.05 g Triton X-100
- 50 mL PBS
Store at room temperature.
Store at 4°C.
10 μM cytochalasin D
- 50 μL 2mM cytoD
- 10 mL PBS
Store at -20°C.
Alexa Fluor dye solution
- 7 μL dye in methanol
- 200 μL PBS
- For 10 mL of bead solution at a concentration of 5e5 beads/mL: Pipet 1 μL of bead solution and 99 μL of molecular-grade water into centrifuge tube. Vortex.
- Centrifuge the microspheres at 1300 g for 10 minutes to clear the supernatant.
- Remove and discard supernatant. Resuspend the microspheres in 100 μL of water and vortex.
- Centrifuge at 1300 g for 10 minutes.
- Pipet out supernatant and bring into tissue culture hood. Resuspend microspheres in 1 mL of DMEM++ to get concentration of 5e6 beads/mL. (Check in hemocytometer.) Allocate 9 mL of DMEM++ into Falcon tube. Vortex the beads and transfer into the same Falcon tube. Vortex the 10 mL solution. Beads should be at a concentration of 5e5 beads/mL.
- Before pipetting onto cells: vortex the solution, sonicate for 10 minutes to break up aggregates, and warm in hot water bath.
Note from Spring 2016: Beads were not cleaned. Sonicate and vortex prior to adding to cells.
Bead endocytosis with 3T3 cells
For Fall 2019: Grow cells in dish for 1 day before adding beads.
Add 100e6 beads (total) in 2mL of media to 60,000 3T3 cells ~15-24 hours prior to the students' use.
- Spherotech 0.84um Nile Red beads, 1% w/v (keep sterile)
- Polystyrene density = 1.05g/mL
- Calculated bead concentration: 3.07e10 beads/mL
- Make stock solution of 100e6 beads/mL in medium so you can add 1 mL of bead stock to 1 mL of cell solution into Mattek dish
- 32.6uL of sonicated beads from spherotech bottle
- 10mL medium (DMEM++)
- Vortex before adding to cells
Preparing slides of immobilized microspheres
(JS edited 3/14/17)
- 3.26 um red = Spherotech FP-3056-2, Nile Red 1% w/v
- 0.84 um red = Spherotech FP-0856-2, Nile Red 1% w/v
- PSF beads = PS-Speck orange 0.175 μm beads 540/560 nm ex/em
- Note that PSF beads bleach easily and should be remade each semester
- 18 x 18 mm coverslips
- microscope slides
- Sonicate bead stock for 2 minutes
- For 3.26 and 0.84 um beads, prepare 1:50 dilution of beads in ethanol (20 μL beads in 1 mL ethanol), use PSF beads undiluted
- Spread 20 μL of bead dilution on coverslip, air blow to dry
- Add ~5 ul glycerol on slide
- Sandwich and let glycerol spread to edge of cover slip (> 15 minutes)
- Seal with nail polish.
- Turn on 37°C water bath, open tissue culture hood, turn on light, and spray counter with 70% ethanol. Also wipe (downward) vacuum tube with ethanol. Periodically spray gloves with ethanol throughout the procedure.
- In the hood, to split a single T75 into a single T75, pipette 125 mL of DMEM++ into a separate sterile container. Warm the DMEM++, PBS, and trypsin ontainers in the bath.
- After warming up for 20 minutes, take DMEM++, PBS, and trypsin out of bath. Dry them, wipe with ethanol, and bring into hood.
- Add 25 mL of DMEM++ into the new T75 flask.
- Fetch flask of cells from incubator and make sure they look OK under the inverted microscope. Spray with ethanol and bring into hood.
- Connect a glass Pasteur pipette to the vacuum tube. Use the glass pipette to drain the medium from a corner of the old flask. With the 5 mL pipette, rinse old flask with 4 mL of PBS. Drain with vacuum.
- Pipette 2 mL of trypsin into old flask and close the cap. Set timer for 2 minutes and put in incubator.
- Use the microscope to verify cell detachment.
- Spray flask with ethanol and bring into hood. Pipette 25 mL of DMEM++ into flask of trypsinized cells. Make sure to spread the medium and wash the cells from the walls by pipetting up and down. Using the same pipette, transfer less than 1 mL of cells into an eppendorf tube. Take eppendorf out of hood.
- Transfer ~90 μL of cells and 10 μL of Trypan Blue dye into new eppendorf. Load each chamber of the hemacytometer with ~10 μL of the cell-dye mixture.
- There should be 9 large squares in the field of view. Count the number of cells in the 4 large corner squares and the large center square. Repeat with second chamber and sum the two counts. Multiply by 1000 to estimate the number of cells/mL.
- Calculate the volume required for the desired number of reseeded cells, and pipet that volume into new T75 flask (Can seed 300,000-500,000 cells in new T75 flask in ~10mL medium, for the flask to be passaged again in 2-3 days). To plate the MatTek dishes, first add more DMEM++ into the flask of cells such that there are approximately 5x10$ ^4 $ in 0.5 mL. Add 1.5 mL of medium into each MatTek dish. Pipet 0.5 mL of cells into each MatTek. Close the caps, label, and place in incubator.
- Vacuum remainder of cells. Drain 50 mL Falcon tube of 50% bleach to wash the system. Disconnect vacuum tubes.
- Put extra DMEM++ and PBS in the fridge. Trysin goes in the -20°C freezer.
- Wipe the counter and close the hood. Clean hemacytometer. Toss eppendorf, old flasks, and gloves in the biowaste container.
To seed multiple MatTek dishes to be ready on different days, you can try the following (assuming a doubling time ~20 hrs):
|For use in (# days after seeding)
|| Seed at:
Fall 2016 note: DMEM++ without glutamine, doubling time is rougly 24 hours
- ~80,000 cells for use in 1 day
- ~40,000 cells for use in 2 days
Imaging the actin cytoskeleton
The mushroom toxin phalloidin binds to polymerized filamentous actin (F-actin) much more tightly than to actin monomers (G-actin), and stabilizes actin filaments by preventing their depolymerization. You will take advantage of this strong affinity between phalloidin and F-actin to image the cell's actin cytoskeleton. Phalloidin conjugated with the fluorescent dye Alexa Fluor 568 will be introduced in Chinese hamster ovarian (CHO), mouse fibroblast (NIH 3T3), or mouse embryonic fibroblast (mEF) cells.
|Example of a Chinese hamster ovarian cell labeled with Alexa Fluor phalloidin
||Excitation and emission spectra for Alexa Fluor 568
||Wear gloves when you are handling biological samples.
Procedures for fixing and labeling cells
You are provided with mEF cells, which were prepared as follows:
Cells were cultured at 37°C in 5% CO$ _2 $ in standard T75 flasks in a medium referred to as DMEM+++. This consists of Dulbecco's Modified Eagle Medium (DMEM - Invitrogen) supplemented with 10% fetal bovine serum (FBS - Invitrogen), 1% of the antibiotic penicillin-streptomycin (Invitrogen), and 1% of non-essential amino acids (NEAA). The day prior to the fixing/labeling experiments, fibroblasts were plated on 35 mm glass-bottom cell culture dishes (MatTek, equipped with coverslip suited for optical microscopy studies).
Below is the protocol to stain cells with Alexa Fluor 568 phalloidin:
- Start with cells about 60% confluent. This is about the optimum percentage of cell population. If cells are too crowded, they will not stretch properly and show their beautiful actin filaments. Note also that these cells remain alive until the addition of formaldehyde, therefore requiring that any buffer/media added be pre-warmed.
- Pre-warm 3.7% formaldehyde solution and phosphate buffered saline at pH 7.4 (PBS) in a 37°C water bath. Keep the formaldehyde wrapped in foil to protect from light.
- A key technique to keep in mind when working with live cells - to avoid shocking them with "cold" at 20°C - is to be sure that any solutions you add are pre-warmed to 37°C. We keep a warm-water bath running for this purpose.
- Remove the medium with a pipette and wash the dish 2X with 2 mL of pre-warmed PBS. Pipet up and down into the dish gently to avoid washing away cells.
- Carefully pipet 400 μL of 3.7% formaldehyde solution onto the cells in the central glass region of the dish and incubate for 10 minutes at room temperature. This "fixes" the cells, i.e. cross-links the intracellular proteins and freezes the cell structure.
- Wash the cells 3X with 2 mL PBS (note that this PBS solution no longer needs to be pre-warmed as the cells are dead).
- Extract the dish with 2 mL 0.1% Triton X-100 (a detergent) for 3-5 minutes. (Extraction refers to the partially dissolution of the plasma membrane of the cell.)
- Wash the cells 3X with 2 mL PBS.
- Incubate the fixed cells with 2 mL 1% BSA in PBS for 20 minutes. (BSA blocks the nonspecific binding sites.)
- Wash the cells 2X with PBS.
- Add 200 μL of Alexa Fluor 568 phalloidin solution (pre-mixed in methanol and PBS). Carefully pipet this just onto the center of the dish, cover with aluminum foil, and incubate for 45 minutes at room temperature.
- Wash 3X with PBS.
- You can now store the sample at +4°C (regular refrigerator) in PBS for a few days, wrapped in parafilm and foil. It can also be stored in mounting medium for up to 1 year.
Culturing KF95 E.Coli "Spinners" for Optical Trap
KF95 are stored in the -80 C freezer (upper left corner) in the 109 lab.
They are Ampicillin resistant.
- LB--Borrow from 109 lab
- Ampicillin (Amp)--Borrow from 109 lab
- TB: 5g Tryptone + 2.5g NaCl + 500mL clean water --> use vacuum filtration with 0.2um filter
To thaw spinners:
- Prepare two autoclaved glass tubes with 5mL LB + 5uL Amp
- Use an autoclaved wooden stick to scrape the frozen stock and drop it into the tube. Use one stick per glass tube.
- Place the tubes in the spinning rack on speed 7 or 8 in the big incubator in the 109 lab
To subculture spinners:
- Prepare 5mL LB + 5uL Amp in an autoclaved glass vial
- Add 10uL of spinners from previous day
To prep spinners for optical trap:
- The evening before the bugs are needed, prepare a glass tube with TB and 5% LB:
- 5mL TB + 250uL LB + 5uL Amp
- Add 200uL of the bacteria to the tube
- Create double sided tape glass chambers
- Pipette in the cells
- Let sit so cells stick
- Rinse cells with PBS or HBSS for optical trap
Bacteria for aerotaxis studies
- For them all, we should filter the 2216 media with 0.2-μm filters.
- Grow at 30°C.
- We should shake it at 300 rpm all week long.
- From the stock mixture:
- Mix 3 mL of 2216 media with 60 μL of stock culture.
- Shake for two hours.
- Then we can monitor the bugs under the microscope.
- From the frozen stock:
- Incubation condition: 30°C, 200-300 rpm.
- Culture medium: 1/2x 2216 (BD Difco) media (for 1/2x media you can simple dilute 1x with MilliQ water).
- Incubation time: overnight (~20-24 hours) and the cells will be good for the following day.
- Culture volume: 2-5 mL in a 15 mL culture tube.
- Spin the cells at <3,000 rcf to conserve their motility for staining.
- For subsequent culturing from the tube:
- Inoculate 20 μL into 2 mL of the culture medium (1/100 dilution).
- Incubate overnight (same as above).
- Do this daily to retrieve best motility.
- Grow at 37°C.
- Instead of 2216 media, use CAM buffer.
- Their aerotactic response is routinely and convincingly observed in the Stocker lab.
Staining live bacteria
- Obtain two vials of FM 4-64 dye (Invitrogen # T13320) from the center drawer in the 16-352 prep room.
- Obtain one eppendorf of desired bacteria, diluted as-desired.
- Obtain ample fresh warm media for the bacteria, generally at least the same volume as the bacterial sample.
- Obtain 15 mL of HBSS in a new Falcon tube. Sterilize the HBSS as necessary.
- Spin down the two FM 4-64 vials at highest RPM for at least 2 minutes.
- Remove the FM 4-64 vials and spin down the eppendorf of bacteria, and another of water for balance, at highest RPM for 1 minute.
- While the bacteria are spinning, pipette at least 200 uL of HBSS from the Falcon tube into one of the FM4-64 vials (the other was just for balance in the centrifuge) and use the HBSS/FM 4-64 solution to wash and transfer the dye to the Falcon tube.
- Repeat as necessary to extract all dye from one vial.
- Note: This is about 30% greater concentration than specified in the FM product information, but is convenient and works well.
- Remove supernatant from the bacteria and discard.
- Add FM4-64 solution to re-suspend the bacteria pellet. Gently pipette up and down to break up the pellet.
- About 10 times should suffice.
- Incubate at room temperature for 10 minutes.
- Spin down the bacteria at highest RPM for 1 minute.
- Remove supernatant from the bacteria and discard.
- Add fresh warm media to re-suspend the bacteria pellet. Gently pipette up and down to break up the pellet.
- Dispose of waste.
- Return extra FM4-64 concentrate vial to drawer.
- Save FM4-64 dye solution in refrigerator in 20.309 lab.
Optical microscopy lab
Code examples and simulations
Actin Staining Protocol