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
8th Garvan Signalling Symposium
We welcome scientists at all levels, including students, post-docs, research staff and senior lab heads. The intimate nature of the meeting and enjoyable social functions promotes a collegial atmosphere and excellent networking opportunities. A poster session will be held on the Monday afternoon with generous prizes. Slots have been reserved for short (15 minutes) talks to be selected from submitted abstracts.
The meeting is held at the Garvan Institute in the glamorous Darlinghurst region of Sydney, close to the city, Oxford Street, King’s Cross and the harbour.
This years exciting program features state-of-the-art technologies to investigate a wide range of diseases including cancer, immunology, neuroscience and metabolic disorders. Special sessions focus on in vivo/intravital signalling, proteomics, control of gene regulation and the structural basis of signalling.
Click Here for more information and to register
Great news we are currently looking for a new honours student for 2016.
The title of the project is “Developing novel biosensors to monitor DNA damage in cancer cells”.
Its a very exciting new project incorporating cutting edge microscopy and fluorescent biosensors.
If you think you have what it takes and are interested please feel free contact myself, or UNSW SoMS.
For more information on the UNSW honours program please visit: http://medicalsciences.med.unsw.edu.au/students/soms-honours/
Below is an example of the images that will be created during the project.
I often get asked how to uses Thresholds to measure things in Image J.
Below are some of the Basic Steps for using Thresholds:
- Open your image and duplicate it (Image>Duplicate)
- On the duplicate go to Image>Adjust>Threshold
- Play with the sliders until all of your cells are red.
- Click ‘Apply’
- You should now have a ‘binary’ black and white image
- Now go to menu Process>Binary and select ‘fill holes’
- 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.
- Now go to menu Analyse>Set Measurements. Select all the things you want to measure.
- Critical steps: make sure that you select your original image (not the binary) in the ‘Redirect to:’ pull down Menu
- Also make sure the ‘Limit to threshold’ checkbox is ticked and also tick the ‘Add to overlay’ and ‘Display label’.
- Click ok to close the ‘Set Measurements’ box.
- Now go to Analyse>Analyse Particles
- 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.
- Make sure you have the Display results tick box selected.
- Once you click ok you should have a the measurements box appear with all your measurements for each cell.
- You can copy and paste these into Excel or what ever program you like to use.
- Go get a coffee and cake you deserve it!
Here is a very simple guide for determining the level of fluorescence in a given region (e.g nucleus)
- Select the cell of interest using any of the drawing/selection tools (i.e. rectangle, circle, polygon or freeform)
- From the Analyze menu select “set measurements”. Make sure you have AREA, INTEGRATED DENSITY and MEAN GRAY VALUE selected (the rest can be ignored).
- 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.
- 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.
- Repeat this step for the other cells in the field of view that you want to measure.
- 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)
- 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)
- 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
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.
The Mitchison Lab has an excellent guide on staining and fixing cells for Actin and Microtubules which is worth reading [Link]
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.
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.
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.
25 mM HEPES
1 mM EGTA
60 mM PIPES
2 mM MgCl2
pH = 6.9
(Add in this order.)
Antibody Blocking Solution (ABS)
3% BSA (or 5% Fetal Calf Serum)
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.
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