20.109(F18):Practice tissue culture and prepare microwell array (Day1)

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20.109(F18): Laboratory Fundamentals of Biological Engineering

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Fall 2018 schedule        FYI        Assignments        Homework        Class data        Communication
       1. Measuring genomic instability        2. Modulating metabolism        3. Engineering biomaterials              


Introduction

Most of the laboratory exercises you will complete for this module are based on the CometChip assay developed in the Engelward Laboratory. The CometChip assay is used to assess various types of DNA lesions, including base excisions, abasic sites, strand breaks, and crosslinks.

To measure DNA damage, the CometChip assay relies on gel electrophoresis. Electrophoresis is a technique used to separate molecules by size using an applied electrical field and a sieving matrix. DNA, RNA and proteins are often studied with this technique; agarose and acrylamide gels are the two most common sieves. The molecules to be separated enter the matrix through a well at one end and are pulled through the matrix when a current is applied. Because DNA and RNA are negatively charged molecules due to their phosphate backbone, they naturally travel toward the positive charge at the far end of the gel. Larger molecules are entwined in the matrix and are stalled; smaller molecules wind through the matrix more easily and travel further from the well. Over time fragments of similar length accumulate into 'bands' in the gel. The CometChip assay is based upon the principle that damaged DNA travels more readily compared to undamaged DNA.

CometChip assay results.
In the CometChip assay, cells are loaded into microwells that are 'stamped' onto an agarose gel sieving matrix. Specific treatments can then be applied to the cells within these microwells that induce DNA damage. The cells are then lysed to release the DNA into the microwell. Following cell lysis, the CometChip is incubated in an alkaline buffer that unwinds the DNA. This step allows for all types of DNA damage to be detected. Lastly, gel electrophoresis is used to separate the DNA fragments. DNA fragments migrate away from the microwell and generate a comet tail as shown in the image to the right (panel A). The distance that the DNA migrates (i.e. the length of the comet tail) is proportional to the amount of damage. For example, with cells not treated with a DNA damaging agent, there are no evident comet tails to the right of the microwell or 'head' (panel B, image on the left); however, in cells treated with a DNA damaging agent there are comet tails apparent (panel B, image on the right). Comet tail lengths can be compared across experimental treatments to determine the deleterious effects of chemicals and toxins on DNA stability. In addition, the CometChip assay can be used to study the rate and efficiency of repair in response to specific treatments.

For this module you will first complete a CometChip experiment in an effort to optimize the assay. Today you will complete the first steps of the CometChip assay by generating an agarose CometChip and preparing the cells that you will load into the microwells of your CometChip during the next laboratory meeting. We will use the data collected for this initial loading experiment to determine the conditions necessary to measure DNA damage in the +/- DNAPKcs cell lines.


Protocols

Part 1: Laboratory orientation quiz

Complete the orientation quiz with your partner. Though you are working with your partner, each student should record all answers on the provided quiz. If you disagree with your partner on an answer, you should write what you think is the correct answer on your quiz.

Good luck!

Part 2: Prepare CometChip

  1. Obtain a sheet of gelbond film from the laboratory bench at the front of the room. The paper is protecting the hydrophilic side of the gelbond film.
    • Be sure to keep the paper associated with the gelbond film so you know which side is which.
  2. Also obtain a special permanent marker from the front bench (Secureline Marker II).
    • If you use a marker from your drawer the ink will wash off during a later step in the CometChip assay protocol.
  3. Use the ruler in your team drawer and the Secureline permanent marker to draw a 6.5 x 4.5 cm rectangle near the center on the hydrophobic side of the gelbond film.
    GelBond marking for CometChip
    • Note: you are writing on what will be the bottom of the CometChip and may want to write backwards so the labels are clear when you look at the top of your CometChip.
  4. Prepare 20 mL of 1% normal melting point (NMP) agarose. Be careful as the agarose solution will be very hot!
    • Calculate the amount of NMP agarose powder needed for a 1% w/v solution. Check your math with the teaching faculty before you continue.
    • Obtain a small milk bottle from the front bench.
    • Weigh out the appropriate amount of NMP agarose and add it to the milk bottle.
    • Use a cylinder to measure 20 mL of 1x PBS and add it to the milk bottle with the NMP agarose powder.
    • Swirl to mix.
    • To melt the NMP agarose, microwave the solution for 20 seconds, swirl, then microwave for 3-second intervals until all crystals are in solutions. After each interval, remove the milk bottle and gently swirl while checking for unmelted agarose crystals. It is important that the solution does NOT boil as you will lose water to evaporation and the density of the agarose will be altered. If your solution starts to boil, immediately remove it from the microwave and gently swirl.
    • When no more crystals are visible in the solution take the milk bottle to your bench.
  5. Obtain a small rectangle dish (labeled "scraped lid") and the CometChip 'stamp' from the front bench.
  6. Add 2.5 mL of the agarose solution to the small dish, then quickly place the gelbond film in the dish with the marked hydrophobic side down. Remove the paper from the film.
  7. Add 13 mL of the agarose solution on top of the gelbond film.
  8. Slowly place the CometChip stamp on top of the agarose.
    • Lower the bottom left of the stamp first, then slowly allow the stamp to 'roll' into the agarose. Be sure to leave the top right corner of the small dish accessible.
    • Be careful not to introduce bubbles into the agarose and work quickly as the agarose will solidify as it cools.
  9. Allow the agarose to solidify, undisturbed, on your bench for 30 min.
  10. Add ~5 mL of 1x PBS to the small dish that contains your agarose CometChip.
    • Pipet in the 1x PBS using the accessible corner.
  11. Slowly pull from one corner of the stamp to lift it away from your CometChip in the dish.
    • If the CometChip sticks to the stamp, carefully peel it off using tweezers.
    • Discard the PBS in the sink.
  12. Remove excess agarose from the perimeter of your CometChip using a razor blade (obtain and return razor blade to front bench).
  13. Clean the agarose from the bottom of your CometChip (gelbond side) using a Kimwipe.
  14. Place your CometChip in the small dish containing 1x PBS for storage at 4 °C until next time.
    • Be sure the chip is completely submerged.
  15. Please return the stamp to the front bench. Be sure never to wipe the stamp, as that will ruin the microposts.

Part 3: Split mammalian cells

In the past century, we have learned a tremendous amount by studying the behavior of mammalian cells maintained in the laboratory. Tissue culture was originally developed about 100 years ago as a method for learning about mammalian biology. The term tissue culture was coined because people were doing exactly that, extracting tissue and letting it live in a dish for a short time. Today, most tissue culture experiments are done using isolated cells rather than whole tissues. Much of what we know about cancer, heritable diseases, and the effects of the environment on human health has been derived from studies of cultured cells.

Cells that are isolated from tissue are called primary cells, because they come directly from an animal. It is very difficult to culture primary cells, largely because primary cells that are placed in culture divide only a limited number of times. This limitation on the lifespan of cultured primary cells, called the Hayflick limit, is a problem because it requires a researcher to constantly remove tissues from animals in order to complete a study. Cell isolation processes can be quite labor-intensive, and also can complicate data analysis due to inherent animal-to-animal variation. To get around the first of these problems, researchers use cells that are immortal, which means they can divide indefinitely, though some inherent cell-to-cell variation still exists in such cells.

One familiar type of immortalized cell is the cancer cell. Tumor cells continuously divide, allowing cancer to invade tissues and proliferate. Cancer cells behave the same way in culture, and under the right conditions, cells can be taken from a tumor and divide indefinitely in culture. Another type of immortalized cell is the embryonic stem cell. Embryonic stem cells are derived from an early stage embryo, and these cells are completely undifferentiated and pluripotent, which means that under the right conditions, they can become any mammalian cell type.

The art of tissue culture lies in the ability to create conditions that are similar to what a cell would experience in an animal, namely 37 °C and neutral pH. Blood nourishes the cells in an animal, and blood components are used to feed cells in culture. Serum, the cell-free (and clotting-factor free) component of blood, contains many of the factors necessary to support the growth of cells outside the animal. Consequently, serum is frequently added to tissue culture medium, although serum-free media exist and support some types of cultured cells. Furthermore, cultured mammalian cells must grow in a germ-free environment and researchers using tissue culture must be skilled in sterile technique.

One major objective for this experimental module is for you to learn how to perform tissue culture using correct sterile techniques. Pay close attention to the demonstration provided by the teaching faculty!

Preparing tissue culture hood

  1. The tissue culture hood is partly set up for you. Finish preparing your hood according to the demonstration, first bringing in any remaining supplies you will need, then obtaining the pre-warmed reagents from the water bath, and finally retrieving your cells from the 37 °C incubator.
    • Don't forget to spray everything (except cells) with 70% ethanol!
    • One of the greatest sources for tissue culture contamination is moving materials in and out of the hood because this disturbs the air flow that maintains a sterile environment inside the hood. Think about what you will need during your experiment to avoid moving your arms in and out of the hood while you are handling your cells.

Collecting cells

  1. Obtain two ~48 h cultures of cells in T25 flask from the 37 °C incubator.
    • Each team should get one '+DNAPKcs' and one '-DNAPKcs' flask.
  2. Examine your cell cultures after you remove the flask from the incubator.
    • Look first at the color and clarity of the media. Fresh media is reddish-orange in color and if the media in your flask is yellow or cloudy, it could mean that the cells are overgrown, contaminated, or starved for CO2.
    • Next, look at the cells using the inverted microscope. Note their shape, arrangement, and how densely the cells cover the surface of the flask.
  3. After you look at your cells, take the flask to your tissue culture hood to begin the seeding procedure.
  4. Aspirate the media from the cells using a sterile Pasteur pipet.
  5. Wash the cells by adding 2 mL PBS using a 5 mL pipet. Slightly tip the flask back and forth to rinse the cells then aspirate the PBS with a fresh Pasteur pipet.
  6. To dislodge the cells from the flask, you will add trypsin, a proteolytic enzyme.
    • With a 2 mL pipet, add 0.5 mL of trypsin to the flask.
    • 2 mL pipets are tricky! They fill up quickly. Be careful not to pull up the liquid too quickly or it will go all the way up your pipet into the pipet-aid! If this happens, please alert the teaching faculty rather than returning the pipet-aid to the rack.
  7. Tip the flask in each direction to distribute the trypsin evenly then incubate the cells at 37°C for 2 minutes using a timer.
    • This is a great time to clear out your trash and read ahead!
  8. Retrieve your flask from the incubator and firmly tap the bottom to dislodge the cells.
    • Check your cells using the microscope to ensure they are dislodged. They should appear round and move freely.
    • If your cells are not detached from the flask, incubate at 37 °C for an additional minute.
  9. When your cells are dislodged, move your flask back into the tissue culture hood and add 3 mL of media to the cells then pipet the liquid up and down (“triturate”) to break up cells that are clumped together and suspend them in the liquid.
    • Note: do not take up or release all the liquid, in order to avoid bubbles.
  10. Transfer the suspended cells into a labeled 15 mL conical tube.
  11. Transfer 90 μL of your cell suspension from the 15 mL conical tube into a labeled eppendorf tube.
    • This aliquot will be used for Part 4.
    • Be sure to cap your conical tube and eppendorf tube after you transfer your cells.

Seeding cells

  1. Obtain two fresh T75 flasks and label them with '+DNAPKcs' or '-DNAPKcs'.
    • Include your team color and section information and the date.
  2. Add 9 mL of fresh media to the flask.
  3. Add 3 mL of your cell suspension to the flask.
    • You should mix your cell suspension by pipetting before distributing. The cells likely settled to the bottom of the conical tube while you were working.
  4. Finally, tilt your flask back and forth to distribute the cells evenly in the T75.
  5. Before moving your flask to the 37 °C incubator, use the microscope to visually confirm that cells are present in the T75.
    • Be sure to place your flask on the appropriate shelf.

Cleaning the tissue culture hood
The next group who uses your hood should find the surfaces wiped down and free of equipment.

  1. Aspirate any remaining cell suspensions.
  2. Dispose of all vessels that held cells in the biohazard waste box and be sure that all sharps are in the sharps jar.
  3. Remove any equipment or supplies that you transferred into the hood and return to the appropriate location.
    • Please leave the equipment that was already there.
  4. Spray the TC hood surface with 70% ethanol and wipe with paper towels.
    • Be sure the paper towels are disposed of in the biohazard waste box!
  5. Empty the benchtop biohazard bucket into the biohazard waste box.

Part 4: Introduction to hemocytometer

During your work in tissue culture, you will use a hemocytometer to count mammalian cells. More importantly, you will use the cell count information to determine the density of your cultures. A hemocytometer is a modified glass microscope slide that has a chamber engraved with a grid. Stained mammalian cells are loaded into the chamber, which is manufactured such that the area within the gridlines is known and the volume of the chamber is known. These features enable researchers to count the number of cells within a specific volume of liquid. Furthermore, because the spacing of the gridlines is also defined it is possible to estimate the size of the cells within the chamber of the hemocytometer.

Calculating cell culture density
Using a hemocytometer, you can determine the density (cells per mL) of cell culture.

Counting cells using a hemocytometer.
  1. Carry the tube with your 90 μL cell suspension aliquot to the center microscope bench and add 10 μL of trypan blue cell stain. Mix by pipetting up and down.
  2. Carefully pipet 10 μL of the stained cells between the hemocytometer and (weighted) glass cover slip.
  3. Count the cells that fall within the four corner squares (with a 4x4 etched grid pattern), average (i.e. divide by 4), and then multiply by 10,000 to determine the number of cells/mL.
    • What was the density of the original culture you used? How many cells did you seed to inoculate your fresh culture?

Measuring cell size

Hemocytometer grid (see table)

The gridded area of the hemocytometer consists of nine 1 x 1 mm (1 mm2) squares.

  • These are subdivided in 3 directions; 0.25 x 0.25 mm (0.0625 mm2), 0.25 x 0.20 mm (0.05 mm2) and 0.20 x 0.20 mm (0.04 mm2).
  • The central square is further subdivided into 0.05 x 0.05 mm (0.0025 mm2) squares.
  • The raised edges of the hemocytometer hold the coverslip 0.1 mm off the marked grid, giving each square a defined volume (see figure on the right).
    • To calculate volume, use the conversion 1 cm3 = 1 mL.

Compare the size of a typical cell seen on your hemocytometer to a characteristic dimension of the grid (see color code below).

  • What is the order of magnitude of the diameter of these cell lines?
Dimensions for Area Area Dimensions for Volume Volume
1 x 1 mm 1 mm2 1 mm2 x 0.1 mm 0.1 mm3 = 100 nL
0.25 x 0.25 mm 0.0625 mm2 0.0625 mm2 x 0.1 mm 0.00625 mm3 = 6.25 nL
0.20 x 0.20 mm 0.04 mm2 0.04 mm2 x 0.1 mm 0.004 mm3 = 4 nL
0.05 x 0.05 mm 0.0025 mm2 0.0025 mm2 x 0.1 mm 0.00025 mm3 = 0.25 nL

Reagents list

CometChip:

  • agar, normal melting point (Invitrogen)
  • phosphate buffered saline (VWR)
  • GelBond film (Lonza)
  • 1 well dish (VWR)

Cell culture:

  • Human glial fibroblasts from malignant glioblastoma: M059J (+DNAPKcs) and M059K (-DNAPKcs)
  • 1:1 Dulbecco's Modified Eagle's Medium (DMEM) : Ham's F12 medium (Gibco)
    • with 10% fetal bovine serum (Atlanta Biologicals)
    • 0.05% Non-essential amino acids
    • 100X antibiotic solution (Gibco)
      • 10,000 U/mL Penicillin
      • 10,000 U/mL Streptomycin
  • Trypsin (Gibco)
  • Trypan Blue
  • Incubator maintains 37°C, 5% CO2 and 95% relative humidity

Navigation links

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