Protocols

User guide: Export microscope images from ImageJ/FIJI into Adobe Photoshop and Illustrator

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Here is a step-by-step guide that I made to help people export microscope images from ImageJ/FIJI and then import and alter colours/levels etc in Photoshop. The guide also shows you how to easily move from Photoshop to illustrator to make montage images etc for publication.

PDF Download: Guide to FIJI-Photoshop Image manipulation

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Using Thresholds to Measure and Quantify Cells in Image J

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I often get asked how to uses Thresholds to measure things in Image J.

There are some great guides on the web explaining how to use Thresholds in Image J, and here are a few that are well worth checking out [Link1][Link2].

Below are some of the Basic Steps for using Thresholds:

  1. Open your image and duplicate it (Image>Duplicate)
  2. On the duplicate go to Image>Adjust>Threshold
  3. Play with the sliders until all of your cells are red.
  4. Click ‘Apply’
  5. You should now have a ‘binary’ black and white image
  6. Now go to menu Process>Binary and select ‘fill holes’
  7. You may also want to select erode, dilate, open or close to optimise the binary image so that you have nice solid filling of your cells.
  8. Now go to menu Analyse>Set Measurements. Select all the things you want to measure.
  9. Critical steps: make sure that you select your original image (not the binary) in the ‘Redirect to:’ pull down Menu
  10. Also make sure the ‘Limit to threshold’ checkbox is ticked and also tick the ‘Add to overlay’ and ‘Display label’.
  11. Click ok to close the ‘Set Measurements’ box.
  12. Now go to Analyse>Analyse Particles
  13. Here you will need to play around with the size and circularity settings (bit of trial and error) in order to get accurate identification of your cells or ROIs. I suggest making duplicates before you start so that you can quickly try different things to see which one works best.
  14. Make sure you have the Display results tick box selected.
  15. Once you click ok you should have a the measurements box appear with all your measurements for each cell.
  16. You can copy and paste these into Excel or what ever program you like to use.
  17. Go get a coffee and cake you deserve it!

Good luck!

 

Using ImageJ to Measure Cell Fluorescence

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Image J can be downloaded for free from here .
This guide can also be downloaded as a complete PDF here: Measuring Cell Fluorescence using ImageJ

Here is a very simple guide for determining the level of  fluorescence in a given region (e.g nucleus)

  1. Select the cell of interest using any of the drawing/selection tools (i.e. rectangle, circle, polygon or freeform)
  2. From the Analyze menu select “set measurements”. Make sure you have AREA, INTEGRATED DENSITY and MEAN GRAY VALUE selected (the rest can be ignored).
  3. Now select “Measure” from the analyze menu or hit cmd+m (apple). You should now see a popup box with a stack of values for that first cell.
  4. Now go and select a region next to your cell that has no fluroence, this will be your background.
    NB: the size is not important. If you want to be super accurate here take 3+ selections from around the cell.
  5. Repeat this step for the other cells in the field of view that you want to measure.
  6. Once you have finished, select all the data in the Results window, and copy (cmd+c) and paste (cmd+v) into a new excel worksheet (or similar program)
  7. Use this formula to calculate the corrected total cell fluorescence (CTCF).
    NB: You can use excel to perform this calculation for you.
    CTCF = Integrated Density – (Area of selected cell  X Mean fluorescence of background readings)

     
  8. Make a graph and your done. Notice that in this example that the rounded up mitotic cell appears to have a much higher level of staining, but this is actually due to its smaller size, which concentrates the staining in a smaller space. So if you just used the raw integrated density you would have data suggesting that the flattened cell has less staining then the rounded up one, when in reality they have a similar level of fluorescence.

How to Cite this if you wold like to:

We have used this method in these papers:

McCloy, R. A., Rogers, S., Caldon, C. E., Lorca, T., Castro, A., and Burgess, A. (2014) Partial inhibition of Cdk1 in G 2 phase overrides the SAC and decouples mitotic events. Cell Cycle 13, 1400–1412 [Link]

Burgess A, Vigneron S, Brioudes E, Labbé J-C, Lorca T & Castro A (2010) Loss of human Greatwall results in G2 arrest and multiple mitotic defects due to deregulation of the cyclin B-Cdc2/PP2A balance. Proc Natl Acad Sci USA 107: 12564–12569

But you can also find a similar method published here:

Gavet O & Pines J (2010) Progressive activation of CyclinB1-Cdk1 coordinates entry to mitosis. Dev Cell 18: 533-543

And here:

Potapova TA, Sivakumar S, Flynn JN, Li R & Gorbsky GJ (2011) Mitotic progression becomes irreversible in prometaphase and collapses when Wee1 and Cdc25 are inhibited. Mol Biol Cell 22: 1191–1206

And my apologies to any others that I have not mentioned.

Immunofluorescence Guide

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The Mitchison Lab has an excellent guide on staining and fixing cells for Actin and Microtubules which is worth reading [Link]

Coverslips

Most coverslips come with a fine film coating to stop them sticking to each other. This can reduce the ability of coating agents such as poly-L lysine from working properly, and can thus reduce the ability of cells to properly adhear to the glass. As most mitotic cells ’round’ up and have a much weaker attachment, a poorly coated coverslip can dramatically reduce the numbers of cells you finally have to look at down the microscope. Thus it is always important to first clean the coverslips and then coat them with either Histogrip, Fibronectin, or Poly-L-lysine.

Cleaning Coverslips
1) Boil coverslips in dH2O in a large beaker for several minutes in a microwave
2) Add HCl to a final concentration of about 1M to the hot water. Careful of fumes do in a    fume hood if possible.
3) Cover the beaker with some parafilm, and gently stir/rock the coverslips on for 4-16h or until cool.
4) Rinse the coverslips several times in dH2O.
5) Then rinse 3-5x with 100% Ethanol, leave coverslips in EtOH and go to TC hood
6) In TC hood, separate individual coverslips out onto large piece of clean Kimwipe or similar blotting paper and allow to air dry.
7) They can now be stored (for unto a year) in a 10cm plate or 50ml falcon until coating. Some people like to autoclave them but it is not necessary.

Coating Coverslips
Histogrip (Invitrogen)
1) In a TC hood, make a 1/10 dilution of the histogrip into 100% Acetone in a 50ml Falcon tube. Normally 10-15ml final volume is plenty.
NB: most TC plastic plates will be dissolved by the acetone, but most 50ml Falcons should be ok, but check first.
2) Have a second empty 50ml falcon ready.
3) Drop about 10-20 individual coverslips one by one into the 50ml falcon with the Histogrip solution. Re-cap and invert tube gently several times.
4) Decant the Histogrip solution into the empty 50ml falcon.
5) Place coated coverslips into a 3rd Falcon full of TC clean H2O
6) Repeat steps 3-5 until you have coated enough coverslips
7) Remove H2O and wash coated coverslips 3x with H2O
8) Separate individual coverslips out onto large piece of clean Kimwipe or similar blotting paper and allow to air dry.
9) They can now be stored (for unto a year) in a 10cm plate or 50ml falcon.

General Buffers

PHEM Buffer
25 mM HEPES
1 mM EGTA
60 mM PIPES
2 mM MgCl2
pH = 6.9
(Add in this order.)

Antibody Blocking Solution (ABS)
1X PBS
3% BSA (or 5% Fetal Calf Serum)
0.1% Tween-20

Mix well and filter, aliquot and store at -20°C

Formaldehyde Fixation in PHEM buffer
(Good general use fixation, good for kinetochore proteins, ok for microtubules)

1.    Wash coverslips 2x with 1X PBS.
2 .   If staining a cytoplasmic protein or if you have high background then try a short pre-permeabilize of cells using 0.1-0.5% Triton in PHEM buffer 30 sec -1 min at room temperature (RT).
3.    Carefully fix cells with 3.7% Formaldehyde (fresh is best) diluted in PHEM + 0.5%      TritonX-100 for 10 min at RT.
4.    Wash 4x with 1X PBS at RT. Can be stored at 4°C for 2-3 days at this stage.
5.  Block coverslips for 15-30 min in ABS, then proceed to Antibody Staining

Fixation with -20°C Methanol
(Good for microtubules and most proteins)

1.  Wash coverslips 2x with 1xPBS
2.  Remove all PBS (but do not allow the cells to dry), and immediately add enough -20°C methanol to cover the coverslips. About 2-3ml if using a 6 well plate.
3.  Put the plate in a -20°C freezer for 5 min (NB: can be stored for weeks as long as you keep the coverslips covered in MeOH).
4.  Remove coverslips from MeOH, and rehydrate them in 1xPBS with 0.1-0.5% Triton-X-100 for 10-20 min
5.  Block coverslips for 15-30 min in ABS, then proceed to Antibody Staining.

Antibody Staining

1.    Incubate with ABS  for 15-30 min at RT.
2.    Incubate with 1°Ab diluted in Cell Blocking solution in moist chamber.
3.    Wash 3x 1X PBS-Tween for 5 min at RT.
4.    Incubate in 2°Ab +DAPI in PBS-T in moist chamber.
5.    Wash 3x 1X PBS-T 5 min at RT.
6  .  Mount with Prolong Gold or Mowiol mounting medium on clean glass slide.

Mowiol 4-88 Mounting Medium

Mowiol 4-88 is a high-quality mounting medium with good anti-fade characteristics. It hardens and matches the refractive index of immersion oil, and thus is particularly suited for this form of microscopy. Additional anti-fade (DABCO) is added to further retard photobleaching.

Mowiol 4-88 (Calbiochem; 475904), DABCO (Sigma; D-2522)

1.   Add 2.4g Mowiol to 6g glycerol and stir briefly with a pipette.
2.   Add 12ml dH2O and stir at room temp for several hours or overnight.
3.   Add 12ml 0.2M Tris (pH 8.5) and heat to 50oC for 1-2 hrs while stirring.
4.   When the Mowiol has dissolved, clarify by centrifugation @ 500 x g for 15mins.
5.   Add DABCO to 2.5% (0.72g), aliquot and store at -20oC. Bubbles can be removed by centrifugation. Aliquots can be stored for up to 2 weeks at 4°C or frozen to -20°C for months

How to Synchronise Mammalian Cells in Culture

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General Guidelines

These guides are written primarily for HeLa cells, but it should be possible to extend these to other cell lines with a bit of optimisation.

1) Make sure your cells are happy !
2) Aim for around 75-85% confluence at the time of release.
3) Try and keep everything close to 37ºC, including media, washing media, PBS, TC hood, centrifuges and avoid having the cells out of the incubator for long periods… i.e. work quickly and be organised before you start.
4) Avoid excessive amounts of media on the plate. Cells need to condition the media and the more volume the longer it will take. For a 10cm plate 5-7ml is ideal depending on the timing. Thus short releases (less than 12 hours) use 5ml, longer use 6-7ml.
5) Always try and make sure your cells are very well spread out on the plate.  Clumps and areas of very high or low density will reduce your synchrony.

G1/S Synchronisation
Here you will block cells in late G1 early S phase usually for around 18-28 hours. This time is roughly equivalent to how long the doubling time is for the cell line. For HeLa’s 20-24 hours is normally used. Cells will roughly take around 6 hours to complete S phase, 1-2 hours for G2, and around 1 hour for mitosis. Thus roughly:
S phase = 2-6 hours after release.
G2 = 7-8 hours
Mitosis = 9-10 hours
G1= 11+ hours

Single Thymidine
1) Seed up asynchronously growing cells on your desired plate. Your seeding will depend on when you plan to block the cells. You can block at the time of seeding, once they have resettled or the next day. Generally you will want about 50-60% density.
2) Add 2.5mM (Update we have dropped to using 1mM, best to do a check with your batch of HeLa’s to determine the optimal dose) Thymidine, which depletes the cells of deoxycytidine triphosphate. NB: Thymidine is not very soluble in water so make it up  to a stock [100mM] in PBS, and make sure you use PBS not water will need to add 25µl/ml of media.

3) Wait 20-24h, to arrest the majority of cells in G1/S. NB: you will always still have the odd few cells in mitosis… but thats ok.

4) Wash the cells 3x with pre-warmed PBS or Media. I have found that media sometimes gives slightly better results, although this is the more expensive option.

5) Add back fresh media, with 25µM 2′-Deoxycytidine (Santa Cruz #sc-231247), to replenish the depleted pools, and promote timely entry into S phase. NB some protocols suggest adding 25µM of Thymidine as well, this may help improve the releases slightly.

Double Thymidine
1) After the first release (step 4 above), wait 8-10 hours and then block cells again with 2.5mM Thymidine for an addition 16 hours.
2) Release as before (step 5 above).

NB: a double block is preferred when you require a very tight synchrony and time points from each different cell cycle phase for biochemistry. However, it does add a extra layer of complexity and if not done perfectly can result in a worse synchrony than a single block.

Hydroxyurea (HU)
Similar to thymidine although you use 2mM to block the cells. Also this can only be done as a single block, and no 2′-Deoxycytidine is needed in the release media.
The downside to the method is that some cell lines (e.g. U2OS) will start to over-duplicate their centrosomes if left in Hydroxyurea for too long.

Late G2 Block
This method is excellent for enriching cells in mitosis for either IF or movies.

RO-3306 (Cdk1 inhibitor)
Although a single block can work, best results are achieved by first blocking cells in G1/S with Thymidine or HU.
1) Release cells from G1/S as per instructions above.
2) Add 10µM of RO3306 to cells, ideally 4-6 hours post release to allow cells to get thru S phase.
3) Wait until the majority of cells have reached late G2. This is usually around 12h post release from G1/S.
4) Wash out the drug very well, with at least 3x washes with media or PBS.
5) Add back fresh media. Cells should begin entering Prophase within about 15-30 min of release. Metaphase peaks around 30-60 min and most cells should have completed anaphase/telophase by 90-120 min.

Here is a movie showing a best case synchrony, where around 90% of cells undergo mitosis. Typically, you should expect to see 50-60%.

Pro-Metaphase
This method is excellent for doing biochemistry on mitosis as it allows for highly enriched samples with a tight synchrony. It can also be used for movies if great care is take during the washing stage not to wash the cells off the plate. It is not suitable for IF.

Nocodazole
Like RO3306, you can use just a single  block with Nocodazole for 20-24 hours, but this will result in an increase in the level of death and better results are achieved by doing a pre-sync with thymidine or another G1/S blocker. NB: cells with a functional Chfr/Antephase checkpoint will delay for significant time during late G2 early prophase. HeLa cells do not have a Antephase checkpoint, but you may still notice a 1-2 hour delay in mitotic entry in response to Nocodazole.

1) After release from G1/S add Nocodazole at the desired concentration (25-3000ng/ml) ideally 4-6 hours after release, although you can add it straight away if you’re lazy. If you plan on releasing cells from the Nocodazole arrest then use 25-50ng/ml. If you are only interested in blocking cells use 100ng/ml. If you want to completely depolymerise microtubules then use 1-3µg/ml.
2)  Wait 12-14 hours after release, this should be sufficient for the majority of cells (80-90%) to block in Pro-Metaphase (P-M). You can easily see this by comparing the number of rounded up cells (P-M), to flat attached cells, likely G2.
3) For Biochemistry, recover media and floating cells from plate and put into a falcon tube. Bang the plate several times to help detach mitotic cells. Add a small amount of fresh (warm) media to recover detached cells. Repeat this once more. Check plate under a microscope that you still have interphase cells attached, and have recovered the majority of P-M cells.
4) You now have a nice highly enriched sample of pro-metaphase cells.
5) If you want to release, then gently spin cells down for 2-3 minutes at low speed (usually 1000rpm is sufficient). Remove media, gently resuspend pelleted cells with excess of fresh media (without any Noc in it). Repeat 1-2 times. NB: cells are very fragile at this stage, excessive washing can damage them leading to a poor release. Thus its a balancing act between washing away enough Noc to allow cells to recommence Mitosis. Generally cells will start arresting at doses of around 5ng/ml of Noc.
6) For movies, do not shake off the cells. Be very very very gentle with the plate, remove the media, and slowly add back fresh media. Repeat 2-3 times. Then immediately start filming your cells.

HeLa SILAC conditions

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Grow cells in DMEM (Sigma #D9443) without L-Arg, L-Leu, L-Lys !

This DMEM is low glucose 1000mg/L so you will need to add and extra 3000mg/L to make it unto 4000mg/L in total. Thus 15ml of the 10% Glucose solution is needed for 500ml of DMEM.

Need to add 50ml of Mass Spec Grade Serum for final [conc.] of 10

1X Normal [Conc.] for DMEM media
L-Arg = 398 µM
L-Leu = 802 µM
L-Lys = 798 µM

From testing HeLa cells grow well and have very high incorporation rate at 1/4 to 1/8 of these levels. To play it safe we have been using 1/4. Thus:

L-Arg = 100µM  (Light and Heavy)
L-Leu = 200µM  (Only Light needed)
L-Lys = 200µM  (Light and Heavy)

Grow cells for at least 5 doublings to ensure complete labelling. As a rough guide, if you Start on Monday, you will be ready by the following monday !

 

 

 

Complete Guide to Cell Cycle based Flow Cytometry

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A complete ‘ish’ guide to Flow cytometry or FACS for cell cycle work.

Covers all aspects from tissue culture, through to running and analyzing your samples on a FACScan or FACS Calibur.

It was written by me a few years ago and so is a bit out of date.  For example there is a new “prettier” version of CellQuest (Pro) now in use, but essentially it is still the same.

Antibiotic Concentrations for Bacterial Work

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Taken from Addgene’s guidelines [Link]

Antibiotic Concentrations

The following list shows recommended antibiotic concentrations for LB media or agar plates.

Antibiotic Concentration
Ampicillin 100 μg/mL
Bleocin 5 μg/mL
Carbenicillin 100 μg/mL
Chloramphenicol 25 μg/mL
Coumermycin 25 μg/mL
Gentamycin 10 μg/mL
Kanamycin 50 μg/mL
Spectinomycin 50 μg/mL
Tetracycline 10 μg/mL

Example: To make 100 mL of LB/ampicillin growth media, add 100 μL of a 100 mg/mL ampicillin stock (1000X stock) to 100 mL of LB.

Reagent List

Unless otherwise indicated, the antibiotic powder can be dissolved in H20. Addgene recommends making 1000X stock solutions and storing aliquots at -20oC.

Reagent Catalog Number
Ampicillin EMD Chemicals #171254
Bleocin EMD Chemicals #203408
Carbenicillin EMD Chemicals #205805
Chloramphenicol EMD Chemicals #220551 (dissolve with EtOH)
Coumermycin Sigma #C9270 (dissolve with DMSO)
Gentamycin EMD Chemicals #345814
Kanamycin EMD Chemicals # 420311
Spectinomycin EMD Chemicals # 567570
Tetracycline EMD Chemicals # 583411

Please note that the catalog numbers given in the list above are only examples, and there are many additional companies that supply these reagents.

Adding a Scale Bar to Microscope Images

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With the later versions of Photoshop CS4 and CS5 extended it is now very easy to add a scale bar to your microscope images. But before we go ahead you will need some information first.

1) The actual Pixel size of the camera attached to your microscope (here is a short list of some common cameras).

2) Did you use any binning when acquiring the image?

3) The lens magnification, C mount, and Objective Magnification (NB: normally lens and C mount are 1x, while objectives are e.g. 63X, 100X)

4) A bit of maths.

THE FORMULA:
Actual Pixel size (APS) = CCD Pixel X Binning / Lens Mag x C mount X Objective Mag

Example 1:

Zeiss AxioimagerZ1 with a Coolsnap HQ2 camera,100X objective, 1×1 binning.

APS = 6450*1 / 1*1*100
= 6450/100
= 64.5nm

Ok so now if you want to add a 5µm (5000nm) scale bar to your image you now do this

5000/64.5= 77.8 pixles   So a line that is 77.8 pixels long will be exactly 5µm. Unfortunately the add scale bar feature in Photoshop does not allow for decimal places so its best to round to the closest full integer which in this case is 78 !

So now go to photoshop and from the Analysis menu select “set measurement” then “custom”. Here you can enter 78 for the pixel length and then 5 for the logical length and change the logical units to µm.
Save this as a preset to save time next time you have an image from the same microscope with the same settings.