Month: August 2014
Are you interested in Cancer Research and want to do a PhD at the Garvan?
Then come along to the open day and where you can meet with all the supervisors, discover some amazing projects, and get all the information you need to start your career in Cancer Research.
- Date: Wednesday 10th of September 2014
- Time: 1.45-5pm
- Venue: The Kinghorn Cancer Centre, Garvan Institute of Medical Research, 370 Victoria St, Darlinghurst
Registration is free but limited to the first 25 registrants, and closes at 4.00 pm on Tuesday, 09th Sep 2014.
UPDATE: ImpactStory is not free, as I first thought, they currently have a 30 day free trail, then the cost is $45/year.
Its clear that judging a researchers output purely on the impact/quality of the journal they do/don’t publish is not always the best way to accurately judge individual achievement and output. To that end, article level metrics have recently emerged as a potential way to generate a more accurate picture of a individual researchers output.
I hadn’t really taken much notice of article metrics, however a friend at work recently told me about Impactstory, a new website that is….
an open-source, web-based tool that helps researchers explore and share the diverse impacts of all their research products—from traditional ones like journal articles, to emerging products like blog posts, datasets, and software
Its very quick and easy to signup and quickly get a snapshot of your article specific metrics. You can see my results here [link].
Interestingly, my PlosOne paper is ranked 3rd, above the 4th placed EMBO publication, which according to the traditional Impact Factor measurement would be viewed as a much, much, much better publication.
Another player in this space is ResearchGate, which has been around for much longer. It gives you a RG score, which “takes all your research and turns it into a source of reputation”.
How does the RG Score work?
Your RG Score is calculated based on how other researchers interact with your content, how often, and who they are. The higher their score, the more yours will increase.
Here is an example of what a profile looks like:
Its going to be very interesting to see how these new metrics impact on the judging of individual scientists output, and if, when and which metric grant funding bodies will prefer.
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
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.
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
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.
2.5mM (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.
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.
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%.
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.
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.
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 !
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.
Taken from Addgene’s guidelines [Link]
The following list shows recommended antibiotic concentrations for LB media or agar plates.
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.
Unless otherwise indicated, the antibiotic powder can be dissolved in H20. Addgene recommends making 1000X stock solutions and storing aliquots at -20oC.
|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.
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.
Actual Pixel size (APS) = CCD Pixel X Binning / Lens Mag x C mount X Objective Mag
Zeiss AxioimagerZ1 with a Coolsnap HQ2 camera,100X objective, 1×1 binning.
APS = 6450*1 / 1*1*100
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.