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Appendix C
Protocols

C.1  Reverse transcription

C.1.1  Superscript II/III

C.1.2  How much?

Use 200 U for the first 1 mg , then use 200 U for each additional mg .

C.1.3  Heat stability

To Do!!!  Heat superscript II and III to 95C for N minutes (try 2, 5). Cool and try reverse transcription. See if either still works.

C.2  Cloning

C.2.1  Buying chemically competent cells

For routine cloning, I use the DH5a Subcloning competent cells [Invitrogen 18265-017]. They don't recommend these for cDNA cloning. For cDNA cloning they recommend using MAX Efficiency DH5a Competent Cells or better. They cost 2x as much. The subcloning cells are so cheap, I hesistate to ever make competent cells for cloning again, since theirs are better than mine. I'll use my own cells only when I need to clone into a wierd strain.

C.2.2  Cloning vector into purchased competent cells

This protocol is for when you have intact (non-ligated) DNA. It is designed for speed not efficiency.
  1. thaw a 50 ml aliquot of competent cells on ice
  2. add 250 pg - 5 ng of plasmid
  3. flick tube to mix plasmid with vector
  4. leave on ice 15 minutes (I should try making this time even smaller)
  5. heat shock 20 seconds at 37 C
  6. place tubes back on ice for 2 minutes; during the 2 minutes add 350 ml of SOC (or LB) to the tube
  7. incubate at 37 C with shaking for 45 min
  8. spread 30 ml onto an agar plate with proper antibiotic resistance

C.2.3  Cloning vector into purchased competent cells

This protocol is for tranforming ligated DNA. It is designed for efficiency not speed. The protocol is similar to the Invitrogen protocol that comes with the cells, except the dilution with SOC is much less.
Use 50 ng of vector, 1 ml of Ligase, and a 3:1 insert to vector ratio in a 20 ml reaction volume.
  1. thaw a 50 ml aliquot of competent cells on ice
  2. add 2 ml of the ligation mixture
  3. flick the tube a few times to gently mix the cells / ligated plasmid
  4. incubate tubes on ice for 30 minutes (Ilaria uses just 15 minutes)
  5. heat shock for 20 seconds in a 42° water bath
  6. place tubes on ice for 2 minutes
  7. add 250 ml of SOC
  8. incubate the cells at 37° C for 1 hr at 225 rpm (Ilaria grows 45 min at 300 rpm)
  9. plate 100-150 ml on a pre-warmed agar plate44 with appropriate antibiotics
  10. wait 12-24 hrs (typically overnite) for colonies to grow




C.2.4  Chemically competent cell preparation

C.3  ChIP Protocols

This protocol is a mixture of three protocols from abcam, upstate, upstate Tips/Protocol, and palsson et al 2005.

C.3.1  Lyse, Crosslink, Shear

The following protocol should be started in the morning and will finish later on in the evening (allocate  10hrs). The morning will be only sporatic work but as the day goes on and on, full time must be devoted to the experiment (from the lysis step forward).
Growth
  1. Grow stain(s) in 50ml of liquid media in a 250ml flask with appropriate antibiotics
  2. add IPTG or other inducer as needed for induction of any cloned constructs (if you have a specific non-cloned gene or tagged gene in the genome, this step is not necessary)
  3. take M, Nml samples as replicates from the same culture (replicates in different cultures are important too; typically I use M=1 and N=15). I put this in a 15ml corning centrifuge tube.
Crosslinking
  1. add 37% formaldehyde to 1% total concentration for each (400ml for 15ml sample) and mix by inversion (invert tube 10 times). Incubate 10 min at room temp. Do NOT crosslink too long!
  2. pellet cells by centrifugation for 10 min at 3200g (this doesn't pellet ALL of them but enough for downstream purposes; if OD is low ( < 0.3) all of sample will pellet)
  3. decant supernant in formaldehyde/media hazardous waste bin
  4. wash cells 2x in ice-cold PBS (spin 3200g for 8 min) (from here on, keep samples on ice to prevent proteases from messing up the experiment; I use 2/3 initial volume for washing: e.g. with 15ml of cells I use 10ml of PBS to wash)
  5. prior to lysis remove any remaining drops of PBS that wouldn't go away by decanting with a P1000 pipettor
Lysis
  1. add 0.4ml Ready-Lyse per ml of lysis buffer into a lysis buffer master mix (just before using)
  2. begin lysis by adding 500 ml ChIP lysis buffer and incubate for 37C for 30 min (no shaking)
  3. add 10ml of 100mM PMSF and 2ml RNAse cocktail per ml of 2x Palsson buffer (just before using)
  4. add 500 ml 2x Pallson IP buffer with 1 ml fresh RNAse cocktail [Ambion] and incubate at 37C with shaking at 300rpm
  5. lysate should be clear like water (if it's not, I don't know maybe you did something wrong. For me it's always looked like slightly-soapy water)
  6. transfer the 1ml lysate to a 1.5ml eppy tube with a P1000
Shearing
  1. sonicate samples using a Branson 250 Sonifier for 30 secs at 20% power (more consistent results can be obtained by using a digital Branson where the exact time can be set. also, it helps to rig up the stand holding the sonifier tip to also hold the eppy tube so the tip stays in the same part of every eppy tube sample.
  2. keep samples on ice for at least 1 min between sonifications
  3. sonicate each sample 4 times for shearing range of between 1000bp and 100bp with an average around 500bp (See Figure 1.24 for an example)
12 samples takes around an hour; dreadfully slow and boring; I play my ipod under the protective sonicator earphones. After half-an-hour this becomes painful if you have the ipod earbuds, so adjust them until it doesn't hurt anymore. Shearing is much less tedious with some tunes.
Quantification Part 1
  1. remove 100 ml from each sample to quantify the amount of DNA in each lysate and to verify the shearing range
  2. place the remaining 900 ml in the -80C freezer for use in immunoprecipitation (it'll be more than enough)
  3. add 350 ml of water to the 100 ml sample (450 ml total volume); add 5 ml of 20mg/ml proteinase K added;
  4. reverse crosslinks overnite in a 65C heat block

C.3.2  Sheared DNA yield / Beginning of immunoprecipation

This protocol also takes a day but with lots of breaks.
Quantification Part 2
Although the DNA yield from this step is pretty high, I can never see a DNA pellet (even though when doing precipitations with much smaller amounts I have seen one; perhaps cause it's sheared?). This used to worry me; now I just trust that it'll work and make sure to suck out the liquid on the side of the eppy tube where the DNA shouldn't be when doing my ethanol precipitation.
  1. separate DNA from proteins by phenol:chloroform extraction using gel phase lock tubes; I use the gel-phase tube only for the second half (the chloroform only step) and I get cleaner DNA than when I use the gel phase lock for both steps. Plus, it seems to smell less like phenol this way.
  2. add 1/10th volume NaAcetate and 2 ml of N mg/ml glycogen as a DNA carrier (not sure this does anything at all, but doesn't hurt)
  3. add 1ml ethanol
  4. place in -80C for 30min
  5. spin at 0C at maximum rpm for 20min
  6. remove supernant with vacuum at 50 mbar, place tip on/near side opposite the g-force (i.e. the side the DNA is not stuck to)
  7. add 1ml of cold 70% ethanol (some people say to resuspend DNA pellet here by vortexing, I don't).
  8. spin at 0C for 5 min
  9. remove supernant with vacuum at 50 mbar, place tip on/near side opposite the g-force (i.e. the side the DNA is not stuck to)
  10. air dry in fume-hood to allow ethanol to evaporate
  11. resuspend DNA in 100 ml of TE
  12. quantify using spectrophotometer (e.g. a Nanodrop)
The cleaned up DNA should be frozen at -20C and saved, as it will serve as the positive control for all downstream qPCR reactions. 500-600ng of each sheared DNA sample should be run an a 1.5% agarose gel to verify shearing range. Examples of yields vs OD can be seen on pages pageref and pageref. For the next step it is helpful to have the DNA concentration over 250ng/ml .
Immunoprecipitation
Immunoprecipitations are begun with equal starting DNA (25 mg). So DNA is first quantified using the protocol above. The following steps I do in 2ml eppy tubes.
  1. divide the 900 ml left of the sheared lysate into N replicates where N is divisible by 3. Each sample should contain 25 mg of DNA. Label these A, B, and C (I normally only use N=1).
  2. dilute all samples 1:10 in dilution buffer (if your DNA isn't concentrated, dilute as much as possible, should still work).
  3. preclear solution by adding 40ml of agarose beads and rotate at 4C for 90 minutes
  4. spin at 1000g for 1 min and transfer supernant to a new tube
  5. add 2ug of the correct antibody to sample A (e.g. if your tag is myc put anti-myc here)
  6. add 2ug of an incorrect antibody to B (a negative control) (e.g. if your tag is myc put anti-Xpress here)
  7. C is a negative control with no antibody
  8. rotate all samples overnite at 4C, preferably in 2ml eppy tubes (these allow better mixing during rotation because the liquid doesn't get stuck as poorly in the bottom; they do however make the later bead washing more difficult because it is hard to see the beads in the 2ml tube)
By pulling all three of these from the same lysate you get nice samples for detecting the different between A, B, and C. Independent sample replicates are important as well. Just try to start with the same concentration of DNA in every immunoprecipitation.
Last, I'm not convinced the C negative control is useful. In the end, I might dump this control as it is expensive to have three controls (one positive and two negatives), takes up valuable space on the qPCR plate, and adds only a small sanity check.

C.3.3  Immunoprecipitate Bead washing

This protocol takes 3-5hrs. Be careful not to suck out the beads by mistake. I use a vacuum with a vacuum regulator adjusted to make the suction very weak (I use approx 50mbar). I'm also conservative with my washing steps and leave  100 ml or so in the tube each step except the final elution step which I work hard to remove all liquids and no beads before starting.
Centrifuge at 4C for 1 min at 1000g; be careful not to disturb the bead pellet when moving samples. Keep samples on ice, unless specified otherwise. Rotate each wash 5 min in cold room, except for TE wash, which is rotated at room temp.
Add beads add 60ml of salmon sperm / agarose beads (I buy the mixture from upstate). Rotate at 4C for 2hr.
Bead Washing
  1. wash beads 1x with low salt wash
  2. wash beads 1x with high salt wash
  3. wash beads 1x with LiCl wash
  4. wash beads 2x with TE (steps at room temp from now on)
  5. elute by adding 225 ml fresh elution buffer and rotating for 15min. Keep the supernant in a 1.5ml eppy tube.
  6. repeat elution with 225 ml more elution buffer and combine supernant with previous supernant.
Crosslink reversal
  1. add 10 ml 5M NaCl solution to the 450 ml elution and incubate overnite in a 65C heat block

C.3.4  Final DNA cleanup

Takes a morning. Make sure to let the ethanol precipitation sit for a long time and spin for a LONG time to pellet/precipitate the short DNA fragments. I proceed similar to the section Quantification Part 2 above, except I add an extra 10 min to each centrifuge time, just to be safe.
  1. add 10 ml EDTA, 20 ml Tris [pH 8] then add 1 ml of proteinase K and incubate 1hr at 45C.
  2. separate DNA from proteins by phenol:chloroform extraction using gel phase lock tubes
  3. add 1/10th volume NaAcetate and 2 ml of N mg/ml glycogen as a DNA carrier and ethanol precipitate
  4. resuspend DNA in 100 ml of TE
Quantification of the product from the above protocol doesn't seem to be useful and just wastes a little sample. There's not enough DNA to quantify.

C.3.5  qPCR to determine TF binding site enrichment

Even with everything in plates, setting up the entire 384-well plate (i.e. using most or all of the wells) takes 1hr 30min.
qPCRs are run with an ABI Sybr Green master mix in a 384-well plate using an ABI 7900HT qPCR machine. The qPCR is a 2-cycle PCR with 60C melting temp so primers should be designed accordingly (e.g. with Primer3 software).
Use 1.5 ml of template (from the total 100 ml ), 150 nM primer, 10 ml 2x master mix and add water to a final volume of 20 ml . I dilute the primer and the template quite a bit in water to make it easier to pipette with the multichannel (I use 1.5ml as the smallest pipetting amount). Primers are typically prearranged in 96-well plates so everything (primers, template, master mix) is added 12 wells at a time with a 12-channel pipettor (Finnpipette). Make sure to have more than enough of everything or else it is hard to get consistent volumes from the multichannel. Also making sure every tip is well attached to the pipettor is important for consistency reasons, otherwise they show up half-full.

C.4  ChIP Protocol Post 1st Round Factorial Optimization

I took the ChIP protocol above and optimized it through one fractional factorial experiment with eight factors (see section 2.2 on page pageref for details). This isn't the final protocol, but it will probably be used to verify some predictions from Vwani's lab at UCLA, because this protocol is much faster and less taxing than the original protocol above that I used for the PLoS paper.

C.4.1  Lyse, Crosslink, Shear

The following protocol should be started in the morning and will finish later on in the evening (allocate  10hrs). The morning will be only sporatic work but as the day goes on and on, full time must be devoted to the experiment (from the lysis step forward). Little has changed in this part of the protocol except the additional glycine quenching step.
Growth
  1. Grow stain(s) in 50ml of liquid media in a 250ml flask with appropriate antibiotics
  2. add 0.01 mM IPTG or other inducer as needed for induction of any cloned constructs (if you have a specific non-cloned gene or tagged gene in the genome, this step is not necessary)
  3. put a 15 ml sample from the flask into a 15ml corning centrifuge tube.
Crosslinking
  1. add 37% formaldehyde to 1% total concentration for each (400ml for 15ml sample) and mix by inversion (invert tube 10 times). Incubate 10 min at room temp. Do NOT crosslink too long!
  2. quench the crosslinker with 1/20th volume of 2.5M glycine (750 ml for a 15 ml sample)
  3. pellet cells by centrifugation for 10 min at 3200g (this doesn't pellet ALL of them but enough for downstream purposes; if OD is low ( < 0.3) all of sample will pellet)
  4. decant supernant in formaldehyde/media hazardous waste bin
  5. wash cells 2x in ice-cold PBS (spin 3200g for 8 min) (from here on, keep samples on ice to prevent proteases from messing up the experiment; I use 2/3 initial volume for washing: e.g. with 15ml of cells I use 10ml of PBS to wash)
  6. prior to lysis remove any remaining drops of PBS that wouldn't go away by decanting with a P1000 pipettor
Lysis and Shearing Do these the same as in the previous protocol.
Quantification Part 1
  1. remove 100 ml from each sample to quantify the amount of DNA in each lysate and to verify the shearing range
  2. place the remaining 900 ml in the -80C freezer for use in immunoprecipitation (it'll be more than enough)
  3. add 25 ml of water to the 100 ml sample (125 ml total volume); add 5 ml of 20mg/ml proteinase K;
  4. reverse crosslinks overnite in a 65C heat block or water bath

C.4.2  Sheared DNA yield, immunoprecipation, and bead washing

This section is the largest change to the previous protocol. These steps used to take at least 2 days, now you can easily finish them in one - if you work hard you can even reverse the crosslinks and have the DNA cleaned up at the end of this day.
Quantification Part 2
I found that a Qiagen PCR purification actually seems to be a more consistent way of determining the relative yields of your sheared DNA. Depending on how robust the ChIP proceedure is to the concentration of DNA, I might eliminate this step altogether, and use it only as a sanity check on the shearing range.
  1. clean up sheared DNA with a Qiagen DNA purification kit
  2. resuspend DNA in 30 ml of EB buffer
  3. quantify using spectrophotometer (e.g. a Nanodrop)
500-600ng of each sheared DNA sample should be run an a 1.5% agarose gel to verify shearing range. Examples of yields vs OD can be seen on pages pageref and pageref. For the next step it is helpful to have the DNA concentration over 250 ng/ml . I no longer use this DNA as a positive control in my qPCR rxns, because I never used that data and it was just an unnecessary cost.
Immunoprecipitation and bead washing
This step is much faster in the new version. Rather than doing an overnite antibody incubation + 2 hrs bead incubation, I do a 10 min antibody and a 10 min bead incubation. I also removed the preclear step, because it had little to no effect (if anything it was deleterious). I switched from agarose beads to dynal beads, because the performance was similar, and the dynal beads require less brain power and attention to wash.
Immunoprecipitations are begun with equal starting DNA (25 mg). So DNA is first quantified using the protocol above. The following steps I do in 1.5 ml eppy tubes (I used to use a 2.0 ml eppy tube, but the 1.5 works better with the dynal beads).
  1. divide the 900 ml left of the sheared lysate into N replicates where N is divisible by 2. Each sample should contain 25 mg of DNA. Label these B, and C (I normally only use N=1).
  2. dilute all samples 1:10 in dilution buffer (if your DNA isn't concentrated, dilute as much as possible, should still work).
  3. add 2 ug of the correct antibody to sample B (e.g. if your tag is myc put anti-myc here)
  4. I no longer use the incorrect antibody negative control. sample C is just a no antibody negative control
  5. rotate all samples 10 minutes at 4C (in the cold room), preferably in 1.5 ml eppy tubes
By pulling both of your immunoprecipitations from the same lysate you get nice samples for estimating the enrichment between B, and C. Independent sample replicates are important as well. Just try to start with the same concentration of DNA in every immunoprecipitation.
Wash beads
With the switch to dynal beads, it is much more difficult to remove the beads by mistake. I do each wash with 1 ml. And I remove the washes with a P1000 pipettor. I do all of this work in the cold room to save time whilst still keeping the samples cold.
Rotate each wash 5 min in cold room, except for TE washes, which are rotated at room temp.
I prepare the dynal beads using a similar strategy to the Young Lab protocol in Nature Protocols.
  1. add N ml of beads
  2. collect with magnet;
  3. add 15 x N ml of block solution (0.5% BSA in PBS)
  4. repeat steps 2 and 3;
  5. resuspend beads in N ml of block solution
This can be done during the first antibody incubation. It doesn't hurt if that incubation goes too long (I saw no difference between 2 hrs and 10 minutes).
Add beads
add 60ml of the prepared dynal protein G magnetic beads [invitrogen]. Rotate at 4C for 10 minutes.
Bead Washing
  1. wash beads 1x with low salt wash
  2. wash beads 1x with high salt wash
  3. wash beads 1x with LiCl wash
  4. wash beads 2x with TE (steps at room temp from now on)
  5. For the elution, I used the Young protocol (which used something similar to TE + 1% SDS) for the dynal elution. Crosslinks were reversed overnight in a water bath at 65C (but the crosslinks are reversed after 6hr if you want to continue in the same day)
Crosslink reversal
  1. add 1 ml of proteinase K samples the next morning after taking them out of the water bath To the dynal samples I added H2O to 450 ml total volume for phenol chloroform extraction
DNA cleanup and qPCR This steps are still done the old way with phenol/chloroform and 20 ml qPCR rxns. Instead of having four samples (+ control, correct antibody, 2 x negative controls) per tested edge, I now only use two, which cuts the expensive qPCR rxns in half and still yields the same results.

C.5  Preparation of E. coli genomic DNA

The following protocol is modified from this website, which modified the protocol in Experimental Techniques in Bacterial Genetics, Jones and Bartlet 1990. It will produce purified DNA suitable for PCR of genes, promoters, etc... (much better than just tossing in some E. coli cells into your PCR), but the DNA will be fairly sheared. If you want contiguous chromosomal DNA it is better to use other protocols.




  1. Grow 5ml E. coli overnight in rich broth.
  2. Transfer 2ml to a 2ml eppy tube (or a 1.5 if that's the biggest you have)
  3. pellet cells by centrifugation for 60 sec at 5600g
  4. vacuum off supernatant
  5. resuspend in 482 ml TE45. Add 15 ml of 20% SDS and 3 ml 20 mg/ml proteinase K and incubate 1 hr at 37C.
  6. optional: sonicate the DNA to make it easier to handle a Branson 250 Sonifier for 30 secs at 10% power (higher power than this in the 2 ml tube results in excessive foaming and not very much shearing e.g. Figure 5.5, page pageref)
  7. add 500 ml phenol/chloroform and mix well
  8. prepare a phase-lock gel (light) tube by spinning at max speed for 30 sec
  9. transfer the mixed solution to the prepared phase-gel lock tube
  10. spin at max speed for 10 min
  11. transfer the aqueous layer (the part above the gel) to a fresh 1.5 ml tube
  12. add 5 ml RNAse Cocktail
  13. incubate at 37° C for 25 min
  14. add 500 ml phenol/chloroform and mix well
  15. prepare a phase-lock gel (light) tube by spinning at max speed for 30 sec
  16. transfer the mixed solution to the prepared phase-gel lock tube
  17. spin at max speed for 10 min
  18. transfer the aqueous layer (the part above the gel) to a fresh 1.5 ml tube
  19. add 50 ml sodium acetate
  20. add 500 ml isopropanol and mix gently (you should see the genomic DNA in 1-2 minutes if you didn't shear it, wait 3-5 minutes total before preceeding)
  21. spin at max speed for 4 minutes
  22. remove supernatant with a weak vacuum (I use -200 mbar)
  23. add 1 ml of 70% ethanol and incubate at RT 1 minute
  24. spin at max speed for 2 minutes (you want the DNA pellet to stick to the tube, unfortunately with genomic DNA it often is more of a gooey ball that won't stick so be careful. If I can get the DNA to stick, I suck as much liquid as I can without getting close to the pellet and add more ethanol (filling the tube) and mix well. This seems to remove some of the gooeyness and allow the pellet to stick.
  25. remove supernatant with a weak vacuum (I use -200 mbar)
  26. resuspend in 200-500 ml TE

C.6  Preparation of E. coli plasmid DNA

C.6.1  Standard Method

I almost always use the Qiagen Miniprep kit. I've also used the Eppendorf Miniprep kit but don't like it as much.

C.6.2  Old School Method

This method is very similar to the Qiagen procedure except you have to make all the solutions for yourself. It is quite a bit slower. Yield???
  1. Grow 5ml E. coli overnight in rich broth.
  2. Transfer 2ml to a 2ml eppy tube (or a 1.5 if that's the biggest you have)
  3. pellet cells by centrifugation for 30 sec at 7500g
  4. vacuum off supernant
  5. repeat above two steps with an additional 2ml of cells in the same tube (only necessary if you want a lot of DNA)
  6. resuspend in 100 ml of GTE buffer (50 mM Glucose, 25 mM Tris-Cl, 10 mM EDTA, ph8). Vortex gently if necessary.
  7. Add 200 ml of NaOH/SDS lysis solution (0.2 M NaOH, 1% SDS). Invert tube 6-8 times (solution should become very quickly)
  8. Immediately add 150 ml of 5 M potassium acetate solution (pH 4.8) to neutralze the NaOH from the previous step and precipate the genomic DNA and SDS into a white goopy mass. Spin at max spin for 1 minute.
  9. Transfer supernant to a new tube; don't transfer any of the white junk.
  10. precipitate with 500 ml isopropanol on ice for 10 minutes and centrifuge at 4C for 2 minutes
  11. aspirate all of the isopropanol supernant. Dissolve the pellet in 400 ml TE.
  12. phenol/chloroform
  13. add 50 ml NaAcetate
  14. ethanol precipitate
  15. resuspend in 50 ml TE

C.6.3  RNA-free Midiprep

I found this protocol on the internet, and I've added my own personal comments about the tricky steps.
  1. Grow 60 ml of cells in rich media (e.g. LB) with appropriate antibiotics to maintain the plasmid. Grow them to late stationary (high cell density).
  2. transfer 50 ml of cells to a 50 ml falcon tube
  3. spin at 4000 rpm46 for 15 min at 4° C 47.
  4. Add 20 ml of H2O and mix a little (I don't resuspend the entire pellet; not a good wash, but it removes most of the LB which can mess things up). Spin 4 min at 4000 rpm48. Remove all of the solution (suck out most of if with a vacuum and get the last little bit with a pipettor).
  5. Resuspend the pellet in 5 ml of Solution I (see B.5.1)
  6. Add 10 ml of Solution II (see B.5.2) and mix by inverting the tube 10 times (solution should become clear)
  7. Add 7.5 ml of Solution III (see B.5.3) and mix by inverting the tube 10 times (solution should fill with chunky white stuff)
  8. Spin at 4000 rpm for 15 minutes49. Since the centrifuge I use doesn't reach the ideal speed, not all of the white stuff (genomic DNA and cell wall) pellets to the bottom. However, most of the precipitate does go to the bottom and the remaining part sits as a firm layer on the top of the tube and the following step is still pretty easy to accomplish.
  9. Transfer the supernatant to a new 50 ml falcon tube (avoid taking the white stuff).
  10. Add 15 ml of isopropanol, mix well, and store at room temperature for 10 minutes.
  11. Spin at 4000 rpm for 15 minutes at 4 C. Discard the supernatant. Remove any remaining fluid with a pipettor.
  12. Dissolve the pellet in 600 ml TE. Transfer to a 1.5 ml eppy tube.
  13. add 200 ml of 8M LiCL. Mix well and spin at 14,000 rpm for 5 min at 4° C (this precipitates the larger RNAs so you can get rid of them).
  14. transfer the supernatant containing the plasmid DNA to a new 1.5 ml eppy tube. Add 600 ml isopropanol. Mix well and incubate 2 minutes at room temperature. Spin at 14,000 rpm for 5 minutes at 4° C.
  15. discard the supernatant. rinse the pellet and the wall of the tube (by inverting it a few times) with 1 ml of cold 70% ethanol.
  16. add 400 ml of TE with RNase A (20 ug/ml) (I actually use Ambion RNAse cocktail; just add 2ml of the cocktail). Incubate 30 minutes at 37° C (this chops up the remaining RNA into little bits)
  17. After 30 min, if a nucleic acid pellet is visible at the bottom of the tube, vortex well to dissolve and incubate another 30 minutes
  18. add 240 ml 2M NaCl, 20% PEG8000 (10 g PEG 8000 and 5.844 g NaCl in 50 ml H2O) (I think this part removes small bits of RNA, but I'm not totally sure)
  19. spin at 14,000 rpm for 5 minutes




  20. Discard the supernatant. Rinse the pellet with 300 ml of cold 70% ethanol (pellet will become white but is much smaller than before due to the absense of the RNA).
  21. resuspend the pellet in 400 ml of TE
  22. add 400 ml of phenol:chloroform and mix well
  23. spin 5 minutes at max rpm and keep the aqueous phase
  24. add 400 ml of chloroform, mix
  25. tranfer the solution to a gel phase lock (Light) tube. spin 5 min max rpm, decant the aqueuous phase to a 1.5 ml eppy tube
  26. add 40 ml 3 M sodium acetate and 880 ml of 95% ethanol. Mix well. Spin at max rpm for 5 minutes at 4° C.
  27. Discard supernatant. Wash with 1 ml 70% ethanol.
  28. Discard supernatant, allow ethanol to evaporate
  29. resuspend in 200-500 ml TE (I usually use 250 ml )

C.6.4  Midiprep





For 20-50 ml, resulting in quite a lot of plasmid DNA. Grow cells in LB with appropriate antibiotics. I typically grow 50 ml in a 250 ml flask. This protocol is from the Sambrook molecular cloning manual.
  1. add 15 ml of overnite to a 15 ml centrifuge tube
  2. spin at max speed (4000 rpm) in the bucket centrifuge for 10 min at 4 C
  3. aspirate the media (according to Sambrook it is very important to get the pellet very dry to prevent DNA that is hard to cleave with restriction enzymes)
  4. resuspend pellet 200 ml Alkaline Lysis I (see B.5.1) by vortexing
  5. tranfer the 200 ml to a 1.5 ml eppy tube
  6. add 400 ml of Alkaline Lysis Solution II (see B.5.2 make fresh each time)
  7. invert tube rapidly 5 times 50
  8. immediately add 300 ml of ice-cold Alkaline Lysis Solution III (see B.5.3)
  9. centrifuge at max speed for 5 min
  10. transfer 600 ml of the supernatant to a fresh tube
  11. add 5 ml of RNAse cocktail and incubate at 37° C for 25 min
  12. add an equal volume of phenol:chloroform to the tube and mix by vortexing.
  13. centrifuge at max speed for 2 min
  14. add 600 ml isopropanol at RT and 2 min
  15. centrifuge at max speed for 5 min
  16. remove supernatant
  17. add 1 ml 70 % ethanol
  18. remove supernatant and dry
  19. resuspend in 100 ml TE

C.7  Preparation of E. coli RNA

C.7.1  RNAeasy preps

How much sample to use

This is what I do with E. coli.
OD culture (ml) RNA protect (ml) RNAlater (ml)
0.5 2 4
0.25 4 8

C.8  Size-separation / exclusion of DNA

C.8.1  Size-exclusion of DNA using microcon filters

Microcon filters are ultrafiltration columns that can be used to remove salts, concentrate DNA, and remove DNA less than a particular size (125 bp maximum double-stranded and 300 bp maximum single-stranded). This isn't as complete a removal as gel filtration. Each of the nucleotide cutoffs below indicates which lengths retain at least 90% of their molecular species. So it is almost always better to go with bigger sizes if the pieces you want to retain are much larger than the smallest cutoff.
The following table is copied from the millipore website: http://www.millipore.com/publications.nsf/docs/6dkp6d
NMWL single-strand cutoff (bp) double-strand cutoff (bp)
3K 10 10
10K 30 20
30K 60 50
50K 125 100
100K 300 125
Here are their recommended g-force and spin times for the microcon columns:
NMWL max g-force spin time (min) at 4° C spin time (min) at 25° C
3K 14,000 185 95
10K 14,00050 35
30K 14,000 15 8
50K 14,000 10 6
100K 500 25 15
Note that these columns also retain proteins which are larger than the pores in the filter, so this isn't a good way to remove proteins from your reaction.

C.8.2  Size-exclusion using Qiagen spin-columns

The Qiagen columns use the glass fiber to catch the DNA and wash the salts and proteins off. The sample loss is much higher than with an EtOH or a microcon filter. But it denatures proteins and should remove them better.
From the Qiagen website:
Specifications: PCR purification kit nucleotide removal kit gel extraction kit
Recovery:
Oligonucleotides - 17-40mers -
dsDNA 100bp - 10kb 40bp - 10kb 70bp - 10kb
Removal:
< mers Y Y Y
17-40mers Y N N

C.8.3  Size-separation of DNA using Sephacryl 500

I've taken a

C.9  Preparation of PET libraries

C.9.1  Growing cells

You want to grow your cells to almost max out the RNAeasy column in the RNA step below (max is 100 mg ). If you don't, you will have problems getting enough cDNA downstream, particularly because the rRNA removal step removes most of the RNA.
For E. coli, I grow the cells to around 0.5 (OD600) and add 2.5 ml of this to 5 ml of RNAprotect, vortex 5 sec, incubate at RT for 5 minutes, and spin at 4000 rpm in a bucket centrifuge for 12 minutes. Then I decant the RNAprotect and get the residual off by tapping the tube on a paper towel. RNAprotected RNA is safe in the -20C for two weeks (I try to use it ASAP though).

C.9.2  RNA extraction

When making PET libraries, I really want to remove every trace of DNA possible from the RNA before making cDNA. Otherwise, if I unknowingly have DNA in there it would create lots of false positives when defining genes with the sequenced PETs. Therefore, I use the RNAeasy kit (preferentially selects RNA over DNA, but still has a lot of DNA left over; also removes short RNAs). I then precipitate the RNA with LiCl, which does not precipitate DNA (this removes the bulk of the remaining DNA and short RNAs). Finally, I digest the trace remaining DNA with the Ambion DNA-free kit.
The entire RNA extraction process takes 4-5 hours (?)
  1. Lyse cells in 100 ml of TE with 1 mg/ml lysozyme. Incubate 2 min, vortex every minute. Add 10 ml Proteinase K. Incubate 3 more minutes, vortex every minute.
  2. add 350 ml RLT (with b-ME added) and follow the RNAeasy kit; elute with 50 ml 2 times (100 ml total)
  3. SAMPLE POINT A: measure yield with Nanodrop, save 750 ng for a gel (takes approximately 1 hr to reach SAMPLE POINT A)
  4. add 50 ml (1/2 volume) of 7.5 LiCl [ambion] to the 100 ml of RNA; place at -20C for 30 minutes
  5. centrifuge at max rpm for 15 minutes
  6. wash RNA pellet in 1 ml of 70% ethanol; incubate at RT 2 minutes, spin 5 minutes, dry pellet 7 minutes
  7. resuspend in 35 ml of RNAse free TE [Ambion] 51 (Note: It takes 1 hr to get to this point from SAMPLE POINT A)
  8. follow DNA-free TURBO kit instructions for high-conc DNA. Briefly: add 3.5 ml Buffer, add 1 ml DNAse, incubate 30 min, add additional 1 ml DNAse, incubate 30 more minutes. Deactivate with 7 ml of deactivation buffer and keep supernatant.
  9. transfer the upper, aqueous phase to a new eppy tube
  10. SAMPLE POINT B: spec DNA free RNA and save 750 ng for a gel (it takes 2 hr 30 minutes to get here from SAMPLE POINT A)
  11. use MICROBExpress to remove 16S and 23S from 10 mg of total RNA (max volume 15 ml ).
  12. resuspend in 16 ml (not the recommended 25 ml )
  13. SAMPLE POINT C: spec
  14. save 200 ng to run on gel (more if possible?)

C.9.3  1st strand synthesis of cDNA

This step takes about 2 hrs
Use Superscript III and the following protocol:
Do in PCR tubes:
  1. add 1 ml of random hexamers (100 ng)
  2. add 1 ml of dNTP (10 mM each)
  3. add 1.5-3mg of mRNA 52
  4. add H2O to 13 ml
  5. heat to 65° C for 5 minutes, chill on ice, brief centrifuge
  6. add 4 ml First-strand buffer, 1 ml DTT
  7. add 1-3 ml of SuperScript II, mix by flicking tube a few times 53
  8. incubate at 25° C for 5 minutes to bind random primers
  9. incubate at 50° C for 60 minutes
  10. heat-inactivate at 70° C for 15 min

C.9.4  2nd strand synthesis of cDNA

This step take 3 hours
Do this in the same tube as first strand. Keep on ice while preparing.
  1. add 66.15 ml of H2O
  2. add 10 ml of NEBuffer 2
  3. add 3 ml dNTP mix (10 mM each)
  4. add 5 ml E. coliDNA polymerase I (40 Units)
  5. add 0.25 ml RNAse H (1 Unit)
  6. incubate 2 hours at 16 C
  7. add 5 ml E. coli DNA ligase buffer (NOT T4 ligase buffer)
  8. add 1 ml E. coli DNA ligase (NOT T4 ligase) and add another 0.25 ml of RNAse H (1 Unit)
  9. incubate 15 minutes at 16 C
  10. heat inactivate both enzymes 20 min at 75 C (it takes about 11 hrs from the beginning to reach here)
  11. (I go home for the day after starting the previous step. I set the thermocycler to keep the tubes at 4C until the next morning)
  12. cleaned up with Qiagen PCR clean up; eluted into 35 ml EB buffer 54
  13. end repair with epicenter kit using 34 ml cDNA (all of it; just keep the same tube); incubated at RT 45 min
  14. heat deactivated enzymes 70 C for 10 min
  15. clean up the end-repaired DNA with a phenol:chloroform cleanup; and elute into 30 ml TE buffer
  16. SAMPLE POINT D: spec 1 ml

C.9.5  Add adaptors to double-stranded cDNA

Use 2 ml of N mM adaptor pairs in each reaction (appx 4.2 mg ). Anneal them first in TE+salt.
  1. to the 29 ml of cleaned up, end-repaired DNA (1 ml was used to spec), add 3.6 ml T4 DNA ligase buffer
  2. add 2 ml (appx 4.2 mg ) of BamISH adaptor (see section 6.6.1 page pageref for details on BamISH)
  3. add 1 ml of T4 DNA ligase
  4. mix by flicking the tube a few times
  5. incubate for 12 hrs at 16° C
  6. heat inactivate T4 ligase at 65 C for 10 min
  7. add 1 ml of T4 DNA ligase buffer55
  8. add 1 ml of T4 polynucleotide kinase (no need to add ATP because it is in the ligase buffer) 56
  9. incubate at 37° C for 30 minutes
  10. heat inactivate for 20 minutes at 65° C
  11. clean up with Qiagen PCR purification kit, elute into 30 ml
clean up now or just run on gel?

C.9.6  Size-selection of cDNA

We need to remove the primers and to only grab the longer cDNA as the short pieces are preferentially amplified by RCA. We'd prefer to have longer cDNA as they should be closer to full length genes/operons. I select two sizes from the gel 500-1500bp and > 1500 bp.