Western blots

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• Please dilute all your antibodies in 2% BSA/PBS + phenol red. You will find this in the fridge. If it runs out, please notify whoever is in charge to make more.

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• All diluted primary antibodies that we have in the lab are kept in one fridge – please check which. Diluted antibodies should be kept until the red solution turns yellow. ‘In use’ secondary antibodies are in the common fridge. Diluted antibodies located in the fridge will be discarded after 1 year/turn yellow, if they are commercially available. Antibodies that we get from other labs/discontinued are stored either way.

• When diluting primary antibodies, you MUST write the information on the side – Serial number, dilution, date (If you want to add more - enjoy) and at least the serial antibody number on the cap.

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• A list of antibodies available in the lab can be found in the shared Dropbox folder along with indications where to find it in the antibody freezer (‘working stock’ and secondary boxes for respective aliquots in use, ‘DO NOT TOUCH’ boxes for stored aliquots). The labeling on the aliquoted tubes refers to the fluorescent channel used for detection with our LICOR (700 or 800). For example, an antibody originally called after its emission maximum of 680, then is listed as 700 for simplification. If info about the specific identity of an antibody & manufacturer is missing in the list, please ask Olga (guardian of our antibodies).

• When an antibody arrives, it needs to be aliquoted (usually to 10ul aliquots but it depends on the antibody), labeled properly, the aliquots stored in ‘working stock’, the original vial stored in ‘DO NOT TOUCH’ box with some indication on it that it’s empty. This is the responsibility of whoever ordered it/used it first. If you have any problems or questions – talk to Olga.

When you take an aliquot, keep that for yourself and use it until it’s finished. The antibody vial from the company/collaborator (which contains a few microliters) will be stored in the main box (‘DO NOT TOUCH’) and should not be used unless in an emergency (aka – someone used all the aliquots in the ‘working stock’ box and the re-ordered Ab has not yet arrived). It is important to keep the company vial for reference in case anything was entered incorrectly onto our spreadsheet. – do not throw away any vial without checking with Olga first.

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• When you decide on a primary antibody, be aware that it might not/only work against the yeast or mammalian protein. This does not apply to secondary antibodies which only need to be matched to the primary antibody (raised in mouse or rabbit) and not to the organism (yeast or mammalian cells). You can check in this website https://www.citeab.com/ if the antibody has been used before and for which applications.

• Milk contains lots of biotinylated and phosphorylated proteins, so do not use this for streptavidin blots or anti-phospho antibodies. Instead, we have Fish Serum blocking solution in the lab (white bottle, 1:5 in PBS, 4oC fridge in the hall).

• The anti-histone H3 antibody (#59) is commonly used in the lab as a loading control. It is a rabbit polyclonal, and the detected band runs at ~17kD, so rarely interferes with your protein of interest. If it does, or you do need an anti-mouse loading control, then you can try our Hsp70 (#72) antibody. A new anti-Actin antibody can also be used (#191). Or the mouse anti GAPDH (#217). Fr the GAPDH it is better to use in RT for 1 hour, since it doesn’t do well in the cold room.

• To reduce the size of the membrane to fit the box better, you can cut the borders of the membrane after Ponceau staining.

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-It is recommended not to cut your membrane in your first trial of running the samples, you might miss some bands that you did not expect.

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• It is essential to run a control sample (e.g. BY4741) to check specificity of the antibody. There might be times that a band is not specific to your samples and can also be found in the control lane.

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• Pre-cast gels are to be used ONLY for publication quality images or if gradient gels are required. For all other uses, please pour your own gels or ask special permission from Maya.

• Pre-cast gels do not always work with RIPA buffer. Avoid RIPA if possible or test it before doing a big experiment.

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• Be aware that some buffers interfere with accurate, quantitative running of BioRad pre-cast gels. Bands may (or may not) appear at the right molecular weight but look heterogeneous, without really being different, so it is a tricky artefact – not quantifiable and not pretty. (BioRad does not determine which buffers but be cautious with RIPA buffer, commonly used mammalian cell lysis.)

• Do not forget to remove the green tape from the bottom of pre-cast gels before loading your samples.

• To look at proteins of small molecular weight (10kDa and smaller) that would normally run at the front of a regular 15% gel, you can prepare instead a 19% “UREA gel”. 10 kDa will appear in the center of the gel! And everything above 70kDa will most probably not look good and be squeezed. The Prestained protein ladder works fine, although it reveals an additional band between 20-40kDa (don’t be surprised!) The recipe includes Urea and a higher ion concentration so a few things to consider: i) heat generation above 0.3A, means running at 80V works fine, ii) precipitation in the cold, so store not in fridge but in a wet chamber to prevent drying (lasts a week or two, then let the gel soak and rehydrate in the blot chamber), iii) Weigh Urea powder according to the new recipe and wait until completely dissolved, iv) use 2M Tris buffer stock for the separation gel. Other reagents stay the same, just used in different amounts, see recipe.

• Antibodies in sodium azide can be discarded in the sink. In case the azide concentration is higher than 0.05%, ask safety for guidance - Safety Unit (phone: 2950 or 3457). The antibodies that are diluted into our ‘red solution’ are safe to be discarded into the sink (the azide in the solution is diluted enough).

• If you need to, you can leave your membrane in blocking solution or primary antibody (if you know it produced a clean signal after overnight incubation) over the weekend, shaking at 4deg. However, never do this with the secondary antibody.

• If you are doing an IP followed by WB, then avoid using the same species Abs for IP and WB as you will only ever detect the heavy and light chain bands. If there is no option and you have to use the same species, then if your protein of interest is significantly above 25kD, you can run the gel and transfer as normal, then cut the lower part of the membrane (just above 25kD) and detect your band using a light-chain-specific secondary Ab.

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• For western blot quantification you should always check the total protein amount in your sample using BCA (or another method). The loading control, Ponceau or the pre-lysis OD is not enough.

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• When performing BCA – dilute your samples by 4/5 and always make 2ul extra - in the protocol it’s 25ul per well but works great with a 1:5 dilution and 15ul per well. Whatever you choose, remember to add the same amount of the standards as your samples.

• If you need to quantify your bands - in the protocol folder there is a protocol for how to best quantify and which program to use

• The NaOH method can be used only for qualitative assessment because you cannot quantify the total protein level.

• The NaOH method can only be used for soluble proteins as it requires boiling which is incompatible with membrane proteins.

• Do not boil membrane proteins, they tend to aggregate. Instead put them at 45oC (or up to 75 if needed) for 15m.

• For proteins above 175kDa, do not use the semi-dry transfer but only the wet transfer. The semi-dry will work but will not be quantitative.

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• Bromophenol blue (in the standard sample buffer) is highly fluorescent in the 680 channel, orange G is a recommended alternative to reduce background.

• If you have many proteins to look at and you decide to cut your membrane for incubation with antibodies, you may like using 50ml falcons (swirl the membrane on the falcon wall if the panel is very long) and our two new tube rollers (one in the cold room and one inside the lab at room temperature). It saves resources (less than 10ml of antibody solution required), saves you from looking for tons of other boxes and allows you to conveniently store your antibody solution afterwards in the falcon.

• Before starting the protocol – make sure you know the relevant type of the first antibody (mouse/rabbit) and make sure to use the correct secondary antibody.

• If needed, instead of the Extra Thick Blot Filter Paper that we use to transfer the gels, you can use 3-5 layers of regular Whatman paper (which you need to cut to the relevant size).

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• If you want to be extra resourceful in saving antibody or you prefer to store your Western Blot wet (re-decoration of dried membranes can be tricky), you may like using home-made plastic bags for incubation and storage of your Western Blot (complete or cut). To do so, we use our little heating machine, next to the laminating machine in front of Maya’s office. It melts and seals the plastic foil into a bag of every size you need, matching the size of your membrane. Usage of 3ml antibody solution is sufficient. You may go even lower.