Screening strains for cyt c expression in 96-well plates

Authors: Heather Jensen, Cheryl Goldbeck, Matt Hepler, Caroline Ajo-Franklin, Yancey Appling
Last updated: 09/17/2013
Goal: To identify Esherichia coli strains that produce high levels cytochrome c</em by measuring their relative red intensity.

Critical Steps:

  • Plan ahead. Since you will be working with many individual samples, you will need to make sure you have all the materials you need, i.e. plates, tips, media, and a well documented plan. This is not experiment that you can figure out as you go along.
  • Since the chromosomal copies of the E. coli cytochrome c biogenesis are turned on by anaerobic conditions (specifically the promoter driving ccm transcription on the genome is Fnr-dependent), factors that affect the aeration of the strains are critical to control. We have found that 37 °C with 275 rpm shaking more accurately replicates growth in 50 mL flasks than 30 °C.
  • We use image processing tools to quantify the redness of the cells, therefore it is also critical to keep the settings on the scanner consistent between the plates on different days. Do not deviate from the settings recommended below, and include the black, white and gray color wheel.

Safety Concerns:

This procedure has involves lots of pipetting into 96 well plates. The procedure has been designed to minimize pipetting motions through the use of multi-channel pipets, but be sure to follow good ergonomic practices. Specifically, take a 5 minute break every 20 minutes and adjust your workspace to minimize stress on your hands. And if you feel any discomfort, please let your supervisor know asap.

Materials needed:

  • Strains containing ccm promoters and the cytochrome c of interest
  • 14 mL round bottom falcon tubes, Falcon 352059 VWR catalog #
  • 2xYT, autoclaved: Note: do not use the “inert binder” type
  • appropriate antibiotics
  • 2.2 mL 96 well plates, presterilized, VWR catalog #7005452
  • P20 and P200 multichannel pipette
  • P20 pre-sterilized tips
  • P200 pre-sterilized tips
  • Repeat pipette
  • 25mL tips for repeat pipette, sterile
  • aminolevulinic acid (ALA), Sigma catalog # A7793-1G, stock solution = 1 M in water
  • IPTG in water, 0.1 M
  • pcr strip tubes
  • 25 mL reservoirs, sterile
  • foil, sterile, VWR catalog #89107
  • V-bottom 96 well plates, Nunc #
  • plate sealers

Software & Scripts Needed:

  • PelletSizeColor2.m, a Matlab function
  • pelletAnalysis_example.m, a Matlab script which will create a 2D heat map of your plate
  • pelletReorder_example.m


Day 1 morning – Gather materials and Map out the plate

  1. How many strains will you be growing up?  This is the number of 5 mL overnight cultures you will be preparing.
  2. What volume of 2xYT media do you need? Can you make one or a few media batches with the appropriate antibiotics and [ALA]?

    1.1 mL of media per well:    __  wells x 1.1 mL media/well x 1.15  = __ mL 2xYT

  3. What, how much antibiotics will you need?

    1.1 uL of antibiotic per well:    __  wells x 1.1 mL media/well x 1.15  = __ uL

    The easiest way to do this is to add your antibiotics to your needed amount of media in an autoclaved Erlenmeyer at a 1:1000 ratio.  The same goes for ALA.

  4. How much ALA will you need?

    ___ mM ALA x ___ wells x 1.1 mL/well x 167.59 g/mol x 1.15 = ____ mg ALA

  5. How many boxes of sterilized tips will you need?

    for p20 tips: ___ plates x 2 boxes/plate = __ boxes of tips

    for p200 tips: ___ plates x 1 boxes/plate + 1 box (to make balance) = __ boxes

  6. Plan out you plate scheme using the excel spreadsheet template here. 96WellPlateSetupTemplate

    Include a negative control strain in the first row of each plate. This strain should be the same bacterial host, i.e. BL21(DE3) or C43(DE3), as the strain you are testing and should have same antibiotic resistance. However, it should express no cytochromes c or cytochrome maturation genes (ccm), so that there are no exogenous or endogenous cytochromes c being expressed in the negative control.  The negative control we most frequently use is MFe540, a C43(DE3) strain which contains the empty vectors pACYC184 and pSB1ET2.

    Include a positive control strain in the second row of each plate. This strain should have a known and fairly reproducible relative red intensity. We frequently use MFe409, a C43(DE3) strain which expresses the ccm genes from the pEC86 plasmid and mtrCAB from the I5023 plasmid.

  7. Day 1, afternoon: Growing overnight cultures in 5 mL tubes

  8. For each strain, use a frozen glycerol stock to incoculate a 5 mL 2xYT culture containing the appropriate antibiotics in a 14 mL Falcon tube. Be sure to clearly label each tube.
  9. Grow the cultures by incubating overnight at 37C with 275 rpm shaking.
  10. Day 2, Induction of back-diluted cultures to express cyt c

  11. In the morning, place the overnight cultures in the 4 °C fridge.
  12. Mix 2xYT with antibiotics and ALA.
  13. Around noon-1 PM, begin back dilution. Gently vortex the 5 mL cultures so that the cells are suspended, not pelleted on the bottom of the tube. Transfer to a labeled, sterile 25 mL basin.
    From the basins, transfer 11 uL per well into a sterile, labeled plate. Repeat for all strains.
  14. Aliquot 1.1 mL of 2xYT with antibiotics and ALA into each well.
  15. Cover each plate with foil and press down with palm to seal, fold and tape. Leave sufficient room to read the well column #. Gently rub the foil over the top of the plate so that the letter row markers are visible as indentations in the foil and the plate label is visible.
  16. Grow for 2.5-3.0 hours at 37 °C and 275 rpm.
  17. Take the OD600 using 100 uL of cells, the OD600 should be between 0.5-0.9
  18. Add the appropriate amount of IPTG

    A simple way to do this is to use a lane of a separate 96 well plate to make a 12x serial dilution of IPTG from 1 M stock, and use a multi-channel pipette for transferring IPTG to multiple wells at a time.

    Note: It’s amazing how easy it is to screw up this step.  Some things that will ensure that you do it properly are:

    • Make sure to label your well plate of IPTG dilutions.  It’s very easy to flip around before filling your plate.
    • Clearly mark the “0 IPTG” well before doing the dilution or else you might end up with a 0.245 uM solution instead.
    • Because the volume of the media you will be adding the IPTG to is ~1.1 mL, you should be adding ~5.5 uL of IPTG solution.
    • You will be piercing the foil with the pipette tip to add IPTG.
  19. Cover foil and press down with palm to seal, fold and tape corners.
  20. Grow overnight for 16 hours at 37 °C and 275 rpm.
  21. Day 3: Imaging cell pellets using the scanner

  22. Gently remove the foil layers from the top of the plate.
  23. Using the multichannel pipet, pipet up and down five times to re-suspend the cells from a row of the deep well plates back into the media. Promptly transfer 150 uL of re-suspended culture to a clear V-bottomed 96 well plate.
  24. Repeat for all rows in the plate.
  25. If you have an odd number of plates, it is helpful to make a duplicate plate of one of your plates so that it provides a balance. This is easier than trying to create an accurate balance using just water.
  26. Spin down the plates for 20 minutes at 3750 rpm on the centrifuge in 5210.
  27. Remove the liquid to isolate the pellets.
  28. Flick the liquid out ONCE, if you flick multiple times you may cause the pellets to become disconnected from the plate.  Flicking is more like burping a baby than flinging supernatant into the sink.  If you do the latter, you will have a blast radius.  It is best to do this into a tray of Wescodyne or bleach solution to kill any escaping bacteria.
  29. Place the plates face down on kimwipes or a diaper and allow the residual media to drain.
  30. Image the plates on the scanner in 5204A alongside the black, white, and gray color wheel
  31. The settings for the scanner are:
    1. Restore Fade Color.
    2. Highlights: 73
    3. Shadows: -42
    4. Midtones: -11
    5. Gain: 2.2
    6. Colors, X: 5, Y: -4
    7. Sharpen: High
    8. Peg: 600
  32. Day 3+: Image Pre-processing using Abode Photoshop

  33. Using Adobe Photoshop, crop the image of interest so that only the wells are visible and save it as a jpg with a distinct name, e.g. 05-10-12 Plate1 Cropped.jpg . It is convenient to crop to round dimensions, e.g. 2500 x 1660 pixels. Write down the width and height of the cropped image.
  34. Make a new image of the same width and height and 600 dpi by selecting file->new.
  35. Set the color range (upper corner) to 195, 140, 119.
  36. Select -> color range. set the fussiness to ~95.
  37. Copy the selected part of the image, and paste it into the new image.
  38. Save the new image as with the date description and selected, e.g. 05-10-12 PL1 Selected.psd. Then save it as a .jpg extension.
  39. Day 3+: Image Processing with Matlab

  40. Reserve time on the Graphics Workstation in advance.
  41. Open Matlab.
  42. Allow Matlab to “see” your .m files and images by adding the folders containing these files to the path with the function addpath. For example, type the following into the command line:addpath(‘\Nano-filerfoundryNanobioSTUDENTS SPACEYancey’)addpath(‘\Nano-filerfoundryNanobioSTUDENTS SPACEYanceyData Expt0(YAA-1-05)’)
  43. Using the Matlab Editor, edit pelletAnalysis_example.m by replace the ‘Selected’ and ‘Cropped’ image names with your image names, and save the altered file.
  44. List any empty wells in [empty]
  45. Attempt to run the program with the default values of high red, and size. If the message,“Number of regions matches number of wells” you can proceed to the next step.
  46. The csv output file contains 6 columns. They contain:
    1. the row number of the pellet
    2. the column number of the pellet
    3. the area (in pixels2) of the pellet
    4. the average intensity in the red channel of the pellet
    5. the average intensity in the red channel divided by the grayscale intensity of the pellet
    6. the average intensity in the red channel divided by the green channel intensity of the pellet

    We use 5 to calculate the relative red intensity. The values in column 6 should be similar to column 5. Do not use 4, because it captures the red intensity without reference to blue, green, or grayscale.

  47. Calculate the relative red intensity by subtracting the red intensity of your negative control with 0 IPTG from all other pellets.
  48. Calculate the change in the biomass by calculating the difference in pellet area between the negative control pellet with 0 IPTG and all other pellets and dividing this difference by the pellet area of the negative control pellet with 0 IPTG.
  49. If you used MFe540 and MFe409 as a negative and positive control, respectively, check the relative red intensity and change in biomass values from your experiment against our published values found here.RelRedandSizeMFe409MFe540
  50. If you used other strains as positive controls, check the relative red intensity and change in biomass values from your experiment against our published values found here.RelRedInt_andSizevsCcmPromoterIPTG_Goldbeck2013

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