20.109(S16):DNA repair assays (Day6)

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20.109(S16): Laboratory Fundamentals of Biological Engineering

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Introduction

The flow cytometry machine has revolutionized biology by allowing researchers to analyze and isolate cells based on their spectral qualities. Both genetic reporters and physical tags may be used to introduce fluorescent signal into cells. For example, if you have a fluorescently tagged antibody that preferentially binds to a unique cell receptor – or more likely, a set of antibodies that taken together identify a unique cell type – you can count the number of this cell type within a mixed sample using flow cytometry. Furthermore, you can isolate a pure sample of this cell type from a complex mixture by using a modified flow cytometry equipped to perfrom FACS: Fluorescence Activated Cell Sorting. Technically, if the machine is only able to count, and not sort subpopulations of cells, then the procedure is called flow cytometry, while cell sorting or isolation is called FACS. Because FACS is shorter to say than flow cytometry, many people call anything to do with a flow cytometer FACS.

Schematic diagram depicting principle of cell counting with flow cytometer.
Principle of FACS analysis.
Before researchers had flow cytometers, there were Coulter counters. Coulter counters are automated cell counting machines developed in the 1950s that count cells as they flow in a liquid stream. In an ingenious conceptual leap, Mack Fulwyler combined the technology of ink jet printers with that of Coulter counters to develop the first flow cytometer. The ink jet printer head works by vibrating a nozzle so that a spray of discrete droplets is formed. Similarly, in a flow cytometer, a liquid suspension of cells is forced at high pressure through a vibrating nozzle to create tiny charged droplets, each containing a single cell. The stream of droplets passes in front of a laser beam, and the scattered light is analyzed by a series of filters and photomultiplier tubes that convert the light signal into electrical impulses. Thus, each cell is "interrogated." In FACS, the spectral qualities of the cell are analyzed nearly instantaneously and compared to user-specified spectral qualities. For example, if you have a mixture of green fluorescent cells and non-fluorescent cells, you can ask the machine to isolate the green cells. If a cell registers as green, an electrical charge deflects the cell causing it to fall into a collection chamber. In flow cytometry, each cell is interrogated, and documented as fluorescent or not, but then all the cells go into the same waste stream.

Cell sorting, or FACS, is technically challenging and most FACS machines are only run by experts. In contrast, regular old biologists and biological engineers are often trained to perform flow cytometry as graduate students. In preparation for an FC experiment, the user must appropriately set voltage, compensation, and gating using control samples; failure to set these parameters correctly will result in GIGO (garbage in, garbage out). The next step of an FC is experiment is to measure the experimental samples on the machine. The final step is to document and analyze the statistics output by the flow cytometer. In our case, this last step involves making an additional set of analysis gates. Although these three steps will primarily be done by the teaching faculty, we will describe them briefly below.

How does the machine "know" which cells are fluorescent or not? There is no magic here; the user must tell the flow cytometer which cells are which. The first key control to include is a mock/negative control: these cells undergo the lipofection treatment, but without any DNA. The reason to mock transfect these cells is in case their viability or flow cytometry profile is affected by the lipofectamine. We view these cells using side scatter versus forward scatter. Forward scatter (FSC) is proportional to size, while side scatter (SSC) tells us something about shape/roughness of the cell. The cells should be the largest objects in the solution, while dust and cell debris will tend to fall at very low FSC. Note that in the gates boxed below we do exclude very high FSC and very high SSC events, as these are likely aggregates of cells and dead cells, respectively. The voltages should be set such that few or no cells are lost on the SSC axis. Raising the voltage shifts cells up or right, while lowering the voltage shifts the cells down or left.

Next we look at red fluorescence (from pMAX_mCherry) versus green fluorescence (from pMAX_EGFP). We set the voltage to keep these negative cells in the lower left corner, while still keeping all of the cells visible, and set the gating such that 0-0.1% of negative cells show up as fluorescent. Then similar adjustments are performed using single-color controls: cells that have been transfected only with pMAX_mCherry or only with pMAX_EGFP.

Flow cytometry plots of control samples used to establish gates. Cells tranfected with only pMAX_mCherry (left plot) and only pMAX_EGFP (center plot) were used to assess the number of cells that were co-transfected with both fluorescent reporters (right plot).

Finally, the experimental samples, which are co-transfected with pMAX_mCherry and either intact or damaged pMAX_EGFP_MCS, may be examined, and will look similar to the co-transfection plot above. By measuring the percentage of cells that fluoresce green, you will have some measure of the frequency of non-homologous end-joining within the M059K cells.

Flow cytometers have been around since the 1970s, and while the details have gotten fancier (more colors, faster measurements, etc), the general principles have remained the same. It is truly a testament to the versatility and utility of flow cytometry that the technology does not look to be supplanted by another approach any time soon.

After preparing your samples to be measured by flow cytometry, you will prepare one final experiment for this module: a dose response curve to further examine the NHEJ inhibitor you selected.

Protocols

Both experimental exercises will be completed in the tissue culture room. Unfortunately, not all of the teams can be accommodated in TC at one time...you are strongly encouraged to use this time to read the paper to be discussed with Prof. Samson on M2D7.

Part 1: Prepare cells for flow cytometry demonstration

The cells you transfected during the previous laboratory section were analyzed using the flow cytometer 24 h post-transfection by Dr. Alex Chaim, who just defended his PhD thesis from the Samson Laboratory. Today you will prepare cells for a flow cytometry demonstration to ensure you are familiar with how the data are collected. The samples you will work with today are the controls used to designate the gates that enable the flow cytometer to count fluorescent cells (see plate map below). On M2D7 you will learn how to normalize and interpret the data that were collected for your samples for your Systems engineering research article...then on M2D9 we will discuss how you can use statistics to give meaning to your data.

Plate map for flow cytometry demonstration.
  1. Begin by briefly looking at the cells under the microscope.
    • Record your observations. Specifically, do the cells in any wells appear less dense or less healthy than in others?
  2. Aspirate the media from each well according to the following steps:
    • Tip the plate such that it is at a 45 degree angle to pool the media at the bottom of each well.
    • Place the tip of the Pasteur pipet on your aspirator at the bottom of the well to remove the media. Aspirate all the liquid, but remove the aspirator promptly to avoid damaging/aspirating the cells.
    • Before moving to the next (non-duplicate) well, dip the Paster pipet on your aspirator briefly (less than a second!) in 70% ethanol.
    • Then hold the Pasteur pipet up to dry and count to 3.
    • Alternatively, you may use a fresh yellow tip on the end of the Pasteur pipet, changing between non-duplicate wells.
  3. Gently distribute 0.5 mL of warm PBS to each well, using a 5 mL serological pipet.
    • Be careful not to blast the liquid right at your cells.
  4. Repeat the aspiration step, again using the ethanol to clear the Pasteur pipet or changing yellow tips between samples.
  5. Add 150 μL of phenol-red-free trypsin/EDTA to each well, using a P200.
    • Incubate for 2 min at 37 °C using a timer.
  6. Meanwhile, label 6 eppendorf tubes according to the plate map above.
  7. Retrieve your plate from the incubator and add 150 μL of warmed media to each well.
    • You may use the same pipet tip for each well, as long as you don't touch the tip down into the trypsin or well!
  8. One well at a time, follow the protocol below to transfer a uniform cell suspension to the appropriately labeled eppendorf tube.
    • Pipet the ~300 μL of solution up and down about four times. Make sure to vary where your tip is in the well, concentrating on the four "edges" of the circle (if the circle were, you know, a square) to more fully detach cells that are adhered to the well.
    • Check the first well or two under the microscope to be confident of your technique. You should see rounded, freely floating cells.
    • When you are confident your cells are resuspended, move the suspensions to the labeled eppendorf tubes.
  9. Alert the teaching faculty when your samples are ready for the flow cytometer.

Part 2: Drug treat cells for inhibitor dose response assay

Today you will further examine the effect of your selected drug on DNA repair via NHEJ. This will serve as the final experiment for Module 2. Yesterday the teaching faculty plated 250 cells per well in a 6-well plate -- yes, only 250 cells! The goal of this experiment is to determine how many single cells survive 4 Gy of ionizing radiation after drug treatment. Remember, NHEJ is key to repairing double strand DNA breaks due to irradiation, so if the NHEJ pathway is inhibited the cells will be unable to repair their DNA and, therefore, more less likely to survive exposure to irradiation. You will test 4 drug concentrations, leaving one well of the M059K cells without drugs as a positive control...one well of M059J cells was also included on your plate as a negative control.

Plate map for irradiation assay.

For one of the concentrations you will use the reported IC50 for your selected drug. This value corresponds to the concentration of inhibitor that causes 50% inhibition of a response. For DMNB the IC50 suggests the concentration where 50% of the activity of DNA-PKcs is lost. For Loperamide the IC50 values for NHEJ inhibition were reported in the literature. You will select three additional 'doses' of drug that surround the IC50 value. Use the following information to calculate the volume of drug you will add to reach the concentrations you select:

Drug Reported IC50 Stock concentration
Loperamide 2.8 μM 12 mM
DMNB 15 μM 24 mM
  1. Calculate the volume of drug needed for the concentrations you will test in the irradiation assay.
    • For each well you will prepare 3 mL of media.
    • DMNB and Loperamide are soluble in DMSO, therefore you will need to add DMSO to the 'no drug' wells to account for any DMSO-specific effects on the cells. It is important that all wells receive the same amount of DMSO.
      • To achieve this, determine the largest volume of drug you will add (i.e. the volume needed to reach the highest concentration).
      • Let's say this volume is 10 μL. Then you will add 10 μL of DMSO to the no-drug wells.
      • You also need to ensure the exposure to DMSO is the same between samples receiving different concentrations of drug. Therefore, in drug wells you will add the appropriate volume of drug then add additional DMSO to bring the final volume up to 10 μL.
    • It is best not to pipette less than 2 μL into a large volume -- instead make intermediate dilutions if necessary.
  2. Label six 15 mL conical tubes and aliquot 3 mL of media into each.
  3. Add the appropriate volume of drug, drug + DMSO, or DMSO to each conical tube.
  4. Collect one 6-well plate from the incubator and aspirate the media.
  5. Transfer 3 mL from each conical tube you prepared to the appropriate well, according to the plate map above.
    • Be sure to label the top of the wells with your drug and concentration (and team information!).
  6. Return your plate to the incubator. This evening the plate will be irradiated with 4 Gy using the gamma-irradiator in building 76 (Koch Institute).
    • Thank you to Marcus Parrish, a graduate student in Prof. Bevin Engelward's laboratory for helping with this step!
  7. In ~24 h the teaching faculty will replace the media with fresh (drug-free) cell culture media.
  8. Next week you will fix the cells and stain the colonies that survive.
    • Because of the very low cell seeding density that was used, each colony represents one cell and you will use the colony count as an indication of inhibitor efficacy.

Reagent list

  • 0.5% Trypsin/4.8 mM EDTA from Life Technologies
  • BD LSR II flow cytometer (BD Biosciences)
    • Green excitation: 488 nm, Green emission: ~530 nm
    • Red excitation: 561 nm, Red emission: ~640 nm
  • Gammacell 40Exactor (Best Theratronics)
    • 137-Cesium irradiation source

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