Gene Knockout In Murine Embryonic Stem (ES) Cells

Day -3 (Saturday night/early Sunday morning) Thaw out a vial of ES cells (R1, W9.5) and plate on a 6 cm dish which has previously been gelatinised and feeders laid down. Day -1 (Monday) Prepare 10 6 cm dishes with feeders, these will be needed for electroporation tomorrow. If more than one electroporation is to be performed then you will need additional sets of ten feeder dishes. Change the media on the ES cells (5 ml ) Linearise construct overnight (for a promoterless construct approximately 10-25µg DNA or about 10-15µg for a promoter construct. NB more does not mean better in this case as there will be too many G418 resistant colonies to allow you to pick cleanly) Day 0 (Tuesday) EtOH precipitate vectors and resuspend in 750µl MTPBS Before continuing check feeder plates for confluency, if there is something wrong with the feeder layer it is possible to plate them with the ES cells after the electroporation (it is essential to remove the DMSO from the PMEFs as this is a differentiating agent). Also check ES cells, at this point they should be a nice dense monolayer, try to get a feel for the numbers, don't want too many cells (with time doing this it should be possible to have an accurate number) Set up electroporation recovery media, ie. 40 ml ES media in a 50 ml falcon tube/electroporation (10 dishes, 4 ml/dish => 40 ml/electroporation). This step may also be carried out while the ES cells are spinning as describe below. When everything is set up then proceed to trypsinise ES cells as follows. Aspirate off the media, wash twice with MTPBS, add 0.5 ml trypsin versene with 1% chicken serum. Place at 37°C for no more than 3 minutes. At this point one can see that the cells have lifted off, check under microscope and swirl to dislodge. Pipette the cells in trypsin with the plate at an angle to achieve a single cell suspension. Try to avoid getting bubbles. Add 4.5 ml of ES media to neutralise trypsin. Place cell suspension in a 15 ml falcon tube and spin at setting 1 for 5 minutes. While the cells are spinning place DNA and electroporation cuvettes (0.4 cm path length, electrode gap = 50) on ice or an ice slurry to chill. You do not want them to chill for too long as that will effect the frequency of the transformation. Chill pellet for a few minutes after spin then gently resuspend in 750µl DNA suspension with a capillary pipette attached to a pipette boy, no bubbles, transfer to chilled cuvette. At this point timing is critical so be as quick as possible, but don't rush. Electroporation conditions: 250 V (0.25 reading on volt setting)

Once electroporation is finished quickly place in 40 ml recovery media, gently resuspend with a pipette. Aspirate media from the 10 feeder dishes. This needs to be done by tilting dish and aspirating off the side as you do not want to touch the bottom as this will place a hole in the feeder layer and will cause the monolayer to roll quickly when picking. Add 4 ml of electroporation recovery mix to each dish dropwise in the centre, if you add too forcefully you will get a hole in the monolayer. Place in incubator at 37°C over night.

Day 1 (Wednesday) Set up selection in the morning, by changing the media and adding 4 ml of ES media containing 200µg/ml (active portion) G418. NB if you electroporated on Tuesday morning then you will have to come in that night and set up the selection. Day 2 - 10 (Thursday - Friday) Change the media (G418 ES) every other day (remember to aspirate from the side and do not touch the bottom or add media forcefully) to get rid of dead cells, about day 7 cell death should begin to subside and resistant colonies will begin to appear. NOTE : Generally colonies are ready to pick 10 days after electroporation however this is not always the case and you have to learn to judge the culture, it can be anywhere between day 9 and day 14. Day 9 (Thursday) Plate feeders on 10 48 well plates using an eppendorf multi dispensing pipette, pipette off centre or else distribution will uneven, the ES cells will be picked in to these plates. Feeders are set up in G418 media. Also pull needles for picking and place in hood overnight. Day 10 (Friday) Check feeders to make sure they are confluent. Assess ES cells to determine if they are ready for picking. Only remove dishes one at a time from the incubator, one dish must be completely picked before the next one can be trypsinised. Gently: Remove media. Wash with 2-3 ml MTPBS twice, but if lifting only once. Add 2 ml trypsin versene + 1% chicken serum without moving.. Set up dark field microscope in laminar air flow hood. Label 48 well plate as follows

You will finally end up with 3 48 well plates for each of the 10 dishes. The plates you pick into are the tertiary plates and from these you will eventually make DNA to screen with. Wait for five minutes for trypsin to act. Look for white fluffy colonies that do not have a fried egg appearance. Try to pick colonies that are all a similar size. Colonies of different sizes have different growth rates and if a 48 well plate contains a variety of colony sizes you will be in trouble when it comes to splitting the cells as they will not be uniform and some won't be ready. Attach a clean needle to mouth device for each colony. Try to be as quick as possible, as this is better for the cells and also the longer they are in trypsin the stickier they become and are harder to pick. Break into the trypsin with the capillary needle near the colony to break initial capillary action, pick colony from the side in a vacuuming motion and suck gently. Do not attack from the top as this will cause dispersion and holes. If two are close together, turn plate around to be able to get at each from the best angle Blow out into well on feeder dish, blow bubbles to ensure dispersion. Repeat to fill 48 well plate. Return to incubator and repeat with remaining dishes of ES cells. Set up 10 (or however many plates you picked) 48 well plates with feeders for splitting on Monday. The feeders are set up in 200µl ES media. Day 13 (Monday) Check plates for contamination by holding up to the light. Add 2 NaOH pellets to each of the contaminated wells. Mix removed media, add PBS and incubate for 10-15 minutes at 37°C to ensure bacterial/yeast death. Mark contaminated wells with a big X to ensure that you do not carry this over any further. Accesses whether cells are ready to split if you split when too small they may not recover. Check feeders to determine if they are confluent. Gelatinise another set of 48 well plates and add 200µl ES media.

Label the 1° and 2° plates in the same manner as the 3° plates, use a different colour marker for each of the three plates to make them easier to distinguish. Trypsinise all the 3° plates and then split, if this is too many then you can divide into whatever is a manageable number for you. It is probably easier to deal with it in two batches. To ensure cells do not dry out restack after each step ie the first plate aspirated is also the first to get PBS added. Aspirate off media, add 250µl MTPBS with eppendorf multidispense. NB add PBS at an angle rather than straight down so as not to disturb cells, gently swirl, remove add 50µl trypsin versene + 1% chicken serum. Tap gently to get coverage with trypsin. Incubate 5 - 7 minutes at 37°C. Check after a few minutes and bang on side of counter to make a single cell suspension. Add 250µl ES media to bring volume to 300µl. Return to incubator and replicate one at a time. Mark corners of each plate to align for replicating. Using a multichannel pipette take 100µl from the first row of 8 wells on the 3° plate (as marked in diagram) and add to the 200µl feeders + media in 1° plate, repeat with 2° plate. Repeat for the remaining 5 rows. It is a good idea to split into the 1° plate first then if there is any problem with the with volume or wrong wells you may just loose the 2° plate which is a backup. Add 200µl ES media to the 3° plate, return three plates to incubator and repeat procedure for the remaining plates.

Day 14 - 15 Check cells for contamination, also on day 15 if media is yellow, change it. Day 16 (Thursday) Freeze down 1° and 2° plates. Divide into manageable numbers approx 4-5 at one time. Trypsinise in the same way as for splitting. Check under microscope to ensure a single cell suspension this is very important for freezing down. Add 200µl freeze mix (ES media + 10% DMSO, remember its 10% of the final volume (250µl) don't forget to factor in the 50µl of trypsin into the calculation) to each well. Tape with water proof tape and place at -70°C. Day 17 (Friday) Lysis of 3° plate. Aspirate off media and gently add 250µl lysis buffer with proteinase K (100 mM Tris.HCl pH 8.5, 5 mM EDTA, 0.2% SDS, 200 mM NaCl, 100 µg Proteinase K/ml ) place at 55°C overnight, sealed with water proof tape, also place a vessel of water in the incubator to decrease evaporation.. Do not rock. Day 18 (Saturday) This step can also wait until Monday. Add 500µl EtOH dropwise to the wells containing lysis buffer with a multichannel pipette. Wrap in glad wrap and allow to sit at room temperature overnight can go for longer if you want. Day 19/20 (Sunday/Monday) Pour off EtOH gently, the DNA is visible as white spidery substance attached to bottom of wells. (to pour off DNA set up and apparatus as drawn below). Using a multichannel pipette wash twice with 70% EtOH adding dropwise and pouring off carefully. Leave plates with lids on upside down at 37°C to dry out. DNA is digested in a 100µl volume in each well. Each well contains 20U enzyme, 10X buffer and TE/RNaseA (RNaseA @ 1/200 in TE pH8.0). Calculate the total volume of this mix required for all the wells, make a cocktail and add to each well using an eppendorf multi dispensing pipette. Place at 37°C in cell culture incubator overnight (there is too much evaporation in other incubators). Day 21 (Tuesday) Add 10µl sample buffer to each well. Freeze the samples that will not be loaded immediately. Load 30 - 35µl on a TAE gel of appropriate % for 4 hours at 100V (standard running conditions) but depending on the size range that you are looking for separation in this will vary. Blot gels for 4 hrs with the first 20 mins in 0.25M HCl to depurinate the DNA and then the remaining time in 0.4M NaOH. Remove membrane from blotter neutralise in 2XSSC and place in oven to dry. Pre hybridise for at least 15 minutes in your favourite hybridisation solution and hybridise overnight with probe labelled with the Gigaprime kit. Separate unincorporated nucleotides from incorporated by your favourite method. Day 22 (Wednesday) Wash membrane 3X with 0.1% SDS, 0.1XSSC at 68°C and a final wash in 0.5% SDS, 0.1% SSC at 68°C. Preheat all wash solutions before use. These washes should take place over a period of at least one hour or more. Place in sealed plastic bag and place against a phosphor imaging screen for four hours. The gels and subsequent blotting steps take a few days, it is left to your discretion how quickly or slowly you want to push through at this stage. Once all the Southerns are completed DNA from the positive clones needs to be run on gels again to assure that you have not made any mistakes with well labelling. This confirmatory membrane can also be stripped and probed with a neo probe to check for single integration events these are preferable but not essential. Expansion of positive clones Once positive clones have been identified they need to be expanded and stored in liquid nitrogen. Clones are expanded from the 1°48 well plate onto a 12 well plate followed by a 6 cm plate and the a 10 cm plate. Six vials are then frozen from this plate. Plate feeders onto the 12 well plate the day before the expansion is to begin. Pre warm ES cell media to 37°C. Remove a 48 well plate from the freezer add warm media to the tops of wells of interest, this allows the cells to thaw and also dilutes out the DMSO. Act quickly as you may not want the whole plate to thaw unless all positive clones are to be expanded. Mix media well with a pipette to ensure removal of all cells and transfer to the 12 well plate. Change media the next day. Depending on how well the cells have been frozen down it may take a few days before the are ready to expand into the 6 cm dish, generally 2 - 3 days. Once they are reasonably confluent expand them onto a 10 cm plate. When this reaches 70 - 80% confluency with medium size colonies freeze 6 1ml aliquots in 10% DMSO. Store in liquid nitrogen. Supplies Required for Knockout

Gelatinising Plates Make up a 0.1% gelatin solution in PBS using Sigma cell culture grade gelatin from porcine. Autoclave. Plating of Feeders One vial of feeders is approximately 1x106/ml this is enough for a 25cm2 plate. Place enough in a culture dish to cover the bottom. Leave for 30 mins in hood. Do not leave too long as PBS will sart to evaporate making the gelatin solution too concentrated. Aspirate off gelatin solution, the dish is now ready to plate with feeders. for a 25 cm2 flask