Note: this is a search engine friendly version of my lab notebook, please see the pdf version of this document for a more human friendly, printer friendly version.

Chapter 3
Serial Analysis of Promoter Enrichment

THIS CHAPTER/PROJECT IS DEAD
See the first paragraph of this chapter for an intro to what I was trying to do. See the last paragraph of this chapter for info about why I stopped trying.
Wed Apr 19 11:49:18 EDT 2006
Chromatin Immunoprecipitation has become the standard in vivo method for determining whether a particular transcription factor binds a particular piece of DNA. ChIP-Chip, a techinique that combines ChIP with a tiling array, performs a similar task except the location doesn't have to be known in advance as all locations are tiled on a microarray. The sensitivity of ChIP-Chip in determining the binding site of a transcription factor is around +/- 500bp, and the accuracy of ChIP-Chip with noisy microarrays is unclear. Here, I propose to develop a new technique building on an earlier idea for quantifying gene expression termed: Serial Analysis of Gene Expression (SAGE). The new technique, Serial Analysis of Promoter Enrichment, is more similar to genetic footprinting approaches (Fig. ).
DNA is enriched in vivo or in vitro using a tagged transcription factor (as in ChIP). The bound DNA is digested with restriction enzymes or endonucleases. As with footprinting, the protein protects the binding site from digestion. Like in SAGE the fragments are ligated together with a separating linker, amplified, (size selected), and sequenced. Unlike SAGE the fragements are variable in length and the linker is connected in a different way.
Please see the pdf version for figures

3.1  Initial Steps

The linkers and primers have been chosen (Fig. 3.1). I'm going to try initially to digest a known plasmid (ignoring the ChIP part), glue the pieces back together, clone, sequence. This allows me to verify that the last steps are working when using a quantifiable amount of DNA and ratios (since ChIP DNA is not abundant enough to be quantified).
Here is some potentially useful info from IDT:
Annealing Protocol
It is sometimes necessary to make double-stranded DNA from single-stranded oligos. While the annealing procedure is very simple, attention to a few details can greatly reduce the presence of undesired single stranded material.
Method:
Things to consider: If the product will be used in a ligation reaction, the addition of 5' -phosphate may be needed. This can be done at the time of oligo synthesis (chemical phosphorylation) or at any time thereafter using PNK (enzymatic phosphorylation). If the oligos are relatively long or to be used in cloning, starting with PAGE purified oligos is recommended.
from the FAQ
Can you tell me the minimum number of bases that can be annealed?
You can anneal an oligo of any size to its target, but longer sequences will lead to more stable duplexes. A minimum of 10 bases is needed for a PCR primer to find and remain annealed to its target long enough for extension to occur. This may be a good minimum threshold to consider if you plan to make dsDNA.
The initial linker has a melting temperature around 20C, slightly less than room temperature, but above the 16C optimum for ligation. It is also 8bp long (minus the 3' T hanging off each end), so it is shorter than the minimum needed for PCR primers and hopefully this will prevent any unligated linkers from making primer dimers during the amplification step.

3.1.1  First SAPE Protocol

Initially, I am removing the ChIP enrichment step. I know I can do that part. The tricky part is putting all the transcription factor binding sites into plasmids separated each with a linker. I'll start with a piece of known DNA (a plasmid), digest it, ligate it to the linkers, amplify it, and finally clone it. The BOLD capital letters for each step indicate the corresponding step in Figure 3.1.
  1. digest 100 ng and 1000 ng of plasmid DNA with 0.5 and 5 units of EACH restriction enzyme in a 50 ml reaction for 45 min at 37C (MspI, MseI, HinP1 I) [5 units if 0.5 ml this is 5x the recommended amount]
  2. ethanol precipitate
  3. (C) add 20ml PCR master mix (do we really need to clean up the buffer after this?) incubate 20min at RT to add A's to the ends of each digestion product
  4. (D) do either step 5, 6, 7, or 8
  5. (D)
    1. heat deactivate the restriction enzymes by incubation at 65C for 25 minutes
    2. add ATP to a final conc of 1 mM
    3. ligate to linker for different time lengths using 1 ml T4 ligase and linker at 16C
    4. heat deactivate T4 at 65C for 10 min
  6. (D)
    1. clean up products with a Qiagen nucleotide removal or PCR clean up kit (NOTE: this removes the products below 40 and 100 bp respectively)
    2. add ligase buffer
    3. ligate to linker for different time lengths using 1 ml T4 ligase and linker at 16C
    4. heat deactivate T4 at 65C for 10 min
  7. (D)
    1. clean up products by ethanol precipitation
    2. add ligase buffer
    3. ligate to linker for different time lengths using 1 ml T4 ligase and linker at 16C
    4. heat deactivate T4 at 65C for 10 min
  8. ethanol precipitate (we have to get rid of the ATP from the ligation)
  9. (E) (for 10-20ng DNA) add 5 ml NEBuffer 5, 5 ml 2.5 mM CoCl2 soln, 0.5 ml Terminal Transferase enzyme, 1 ml 10mM dGTP incubate at 37C for 30 min
  10. heat deactivate at 70C for 10 min
  11. ethanol precipitate
  12. (F) D amplify with the two PCR primers try a range of temps, lean on the high temp side to prevent the primer dimer problem; Min temp is 47C use easy-A kit
  13. size select and gel purify 1-2 kb band
  14. (G) TOPO clone
  15. (H) sequence clones that have an insert

Finally getting started...

Fri May 12 12:25:40 EDT 2006
Unlike the above plan this is what I actually did.
  1. miniprep 4 ml of lrpA and lrpB
    Sample DNA (ng/ul) 260/280 260/230
    lrp A 110
    lrp B 109.3
  2. digest 2.5 ug plasmid DNA minipreped from lrpA and lrpB (in TOP10 cells); master mix of enzyme cocktail (0.25 ml MspI, 0.5 ml Mse, and HinPI; 30 ml H20, 10 ml BSA); digests were run in 50ml rxns for 1 hr at 37C followed by 20 min at 65C to heat deactivate the enzymes.
  3. the ethanol precipitation was not done (step 2 above). rather 10 ml of water and 10ml NEB PCR master mix were added and the samples were incubated for 10 min at 72C to add the A's to the end. The ethanol precipitation was skipped because NEB Buffer 2 and the PCR buffer are fairly similar and I don't need an efficient amplification, I only need 2-3 bp filled in plus the A on the end.
  4. ethanol precipitate to remove (hopefully) the Taq, dNTPs and the buffer; 7 ml NaAcetate (into the 70 ml ), mix, add 160 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum out ethanol, add 750 ml COLD 70% ethanol, spin 4 min, (dry), resuspend in 40 ml TE
  5. quantify DNA with the nanodrop
    Sample DNA (ng/ul) 260/280 260/230
    lrp A 11.9
    lrp B 24.8
    16 ml of each were run on a 2% agarose gel (see Figure )
  6. ligate; aliquoted 16 ml of lrpA and B (approx 190 ng and 396 ng of DNA respectively) and 2x 1.6 ml for both digests for a total of 6 ligation reactions; 3 of the tubes were ligated for 2 hr at 16C, the other 3 were ligated for 5 hrs. all six reactions were halted by 65C for 10 min.
  7. ligations were cleaned up with a PCR purification kit. This kit removes DNA less than 100 bp which is ok because we don't want to clone or sequence those anyways, so we might as well not amplify them with the PCR. Ligated DNA was quantified
    Sample DNA (ng/ul) 260/280 260/230
    lrp A 2hr L 10.2
    lrp A 5hr L 9.9
    lrp A 2hr H 11.8
    lrp B 2hr L 11.0
    lrp B 5hr L 10.1
    lrp B 5hr H 12.5
    *** Some strange things here *** No matter what my starting conc of DNA the amount of DNA in the ligation product is the same. All are very dirty too. ***
  8. add GGGGGGGGGG tag with terminal transferase (TdT); 2 ml ligated DNA (post PCR purification), 5 ml Buffer 4, 5 ml CoCl2, 1 ml 100 mM dGTP, 0.5 ml TdT, 36.5 ml H2O.
  9. amplified each sample via PCR; 12.5 ml NEB master mix, 1 ml 200 nM SAPE primer1 Forward and Reverse mix, 1 ml mix from TdT rxn above (not cleaned up. could this cause a problem? particularly the CoCl2?), 10.5 ml H2O; 10 ml of each PCR product was run on a 2% gel (see Figure ). PCR annealing was at 48C. Maybe I should drop it, and raise the extension time (which as only 30sec at 72C, should make that much of a difference though).
Please see the pdf version for figures
Figure 3.1: Diagnostic 2% agarose gels from first attempt at SAPE. The plasmid was digested for 1hr with three 4-mer cutters. The digested plasmid (A) seems so chopped up that I can't see any DNA on the gel (or perhaps it was lost in the ethanol precipitation?). The amplified PCR product didn't fair any better. There is a little something around 30bp but probably either primer dimer or just the primers.
Brief Conclusions:   As is clear in Figure 3.1, the first round didn't work. Lower temp on PCR? Use less enzymes during digestion? (why the 10% DNA loss? why can't see it on gel?) Run ligation longer? ***Run ligated DNA on gel. This should help localize the problem (like a print statement in debugging computer scripts). *** Try Tim's idea of attaching a linker with phosphorylated ends? Idea from Josh, use exonuclease to chop up single stranded stuff.

Trying to narrow down what went wrong in SAPE round one

Mon May 15 15:26:12 EDT 2006
Ran/running the ligated fragments for the three lrpA samples from step 7 above on a 2% agarose gel to see if there was anything at that stage. Ran 27ml of each (that's all I had). If the table for step 7 is accurate (and I don't believe it is) this should be about 275 ng of DNA. I initially pre-stained the gels with agarose. There appeared to be a very large band only visible in the tube that should presumably have the most DNA (based on starting material not based on the nanodrop reading in which is was the most abundant but only by a small margin). The band wasn't bright enough to concretely state that something was there. If it is correct, it appears the ligation step is definitely too long (that was only a 2hr ligation). I am not post staining the gel with sybr gold to see if it brightens the band a little. (I tried to remove a little of the EtBr by rotating the gel with H2O for 15 min with a water change after the first 10 min). The sybr gold is being rotated with the gel for 25 min.
There does seem to be a very faint LARGE band (see Figure ) at around 10000bp. Unfortunately the agarose gel percent chosen is horrible for separating at this range, so its hard to really say what size it is. The band is slightly more visible with Sybr Gold than EtBr (I think our Sybr Gold is getting old). You really have to zoom in a lot to see the band (700% or so).
Please see the pdf version for figures
Figure 3.2: Diagnostic 2% agarose gels from first attempt at SAPE. 27ml of the qiagen PCR cleaned-up ligation reaction was run on the gel. The gel was originally pre-stained with EtBr (A). Subsequently it was washed 10 min and 5 min with water and finally dyed with 1x Sybr Gold for 25 min and washed for 10min in H2O (B). The faint band shows what seems to be an appx 10kb band of the ligated product.
Brief Conclusions:   Figure 3.2 seems to indicate the ligation was too long. I'm not sure what to conclude about why this PCR didn't work based on this. It could either be 1) the ligation products were too long to allow such a long amplification (particularly given that my amplification time was only 30 seconds) 2) the end labeling with poly-G didn't work 3) both

Trying to further narrow down what went wrong in SAPE round one

Tue May 16 12:25:13 EDT 2006
I'm running out the lrpB ligations on a gel as I did for lrpA yesterday. I am only going to stain with Sybr Gold, so hopefully I'll get a better signal. Used 7.5 ml of ladder instead of the standard 10 ml so it doesn't overwhelm the weaker signals. I also included 110 ng and 109 ng of lrpA and lrpB miniprep product respectively for size comparison and a sanity check just to make sure the minipreps all of these results are dependent on really worked.
I really think the Sybr gold is going bad. I used the solution from yesterday and the gel was completely blank. I tried switching to EtBr and could barely see the bands. Now I increased the concentration of Sybr Gold (after so much watching I don't know if I have any DNA left in the gel anymore).
Please see the pdf version for figures
Figure 3.3: Diagnostic 2% agarose gels from first attempt at SAPE. 27ml of the qiagen PCR cleaned-up ligation reaction was run on the gel. The gel was originally pre-stained with Sybr Gold. Subsequently it was stained with EtBr. Subsequently it was stained with Sybr Gold again (but much more concentrated). I think the old Sybr Gold killed this gel. The two lanes of plasmid DNA are the correct size
Brief Conclusions:   This didn't work terribly well (see Figure 3.3). Mainly a problem with the gel staining. Glad to at least see the two plasmids were there.

SAPE1 try 2 (probably without going to the PCR step)

Tue May 16 12:28:03 EDT 2006
I'm trying the initial steps again, this time with shorter ligation times. I'm also using fewer restriction enzymes (only MspI) so hopefully I'll be able to see the digest DNA better (and I'll use Sybr Gold to help that problem too). Below is the sizes of the pieces I should theoretically get (based on an in silico digestion with NebCutter2).
Fragment Cutter Ends Coordinates Length (bp)
1 MspI-MspI 4048-208 551
2 MspI-MspI 2531-3057 527
3 MspI-MspI 334-834 501
4 MspI-MspI 869-1310 442
5 MspI-MspI 1764-2167 404
6 MspI-MspI 3307-3698 392
7 MspI-MspI 1311-1552 242
8 MspI-MspI 3699-3939 241
9 MspI-MspI 2168-2357 190
10 MspI-MspI 3092-3239 148
11 MspI-MspI 2384-2530 147
12 MspI-MspI 209-333 125
13 MspI-MspI 1553-1662 110
14 MspI-MspI 3940-4047 108
15 MspI-MspI 1663-1729 67
16 MspI-MspI 3240-3306 67
17 MspI-MspI 835-868 34
18 MspI-MspI 1730-1763 34
19 MspI-MspI 3058-3091 34
20 MspI-MspI 2358-2383 26
For the digests, all of the remaining DNA was used (around 2.5 mg ). The digestions were done with MspI only for 1 hr followed by addition of 10 ml H2O and 10 ml PCR Master mix [NEB]. After ethanol precipitation, the samples were resuspended in 20 ml TE. lrpA was immediately run on a 2% gel (Figure A). The lrpB digestion was ligated for 30 min and then the entire ligation reaction was run on a gel (Figure B).
DNA was quantified after the digestion was cleaned up via ethanol precipitation:
Sample DNA (ng/ul) 260/280 260/230
lrp A digestion 71.2
lrp B digestion 31.1
Please see the pdf version for figures
Figure 3.4: (A) is 1% (B) is 2%
Brief Conclusions:   The DNA yield post-digestion cleanup were much better though this may be because the previous elution volume was 40 ml which was too dilute for reliable measurement.
I think the problem with SAPE1 is clear now. Tomorrow, I'll run the digestion again with out the addition of the Taq at 72° C for 10 min. If the digestion looks correct, then the puzzle is solved. It appears that all the little pieces from the digestion bind to each other when in the solution with the Taq at 72° C and prime the PCR reaction resulting in all the pieces being turned into a series of long giant pieces (see Figure 3.4). It's kinda puzzling to me why NO smaller pieces appear though.

SAPE1: solving the puzzle

Wed May 17 16:12:35 EDT 2006
Three replicates of lrpB were miniprepped with the following yields:
Sample DNA (ng/ul) 260/280 260/230
lrp B 1 82.5
lrp B 2 70.4
lrp B 3 80.6
I took a 24 ml sample from lrpB1 and lrpB2 and digested them with MspI (0.25 ml ) and HinP1 I (0.5 ml ) respectively. I ran the entire digestion on a gel (unfortunately I forgot to run a little of each plasmid out as the same time Figure ).
Please see the pdf version for figures
Figure 3.5: in silico and in vitro digests of the lrpB plasmid with 4-mer cutters MspI and Hin1PI without the addition of PCR master mix which messes everything up.
Brief Conclusions:   So the problem is now found (but not solved at least not experimentally yet). Figure 3.5 shows that the digestions are working correctly this leaves only the Taq based addition of A's (step 3 of SAPE First SAPE Protocol) as the culprit for the huge pieces of DNA found in Figures 3.2 and 3.4. TA based SAPE will not work without something clever. I plan to abandon TA based SAPE in SAPE2.

3.1.2  SAPE1: the return of SAPE1

Fri May 26 11:31:32 EDT 2006
Jay Shendure did a technique almost identical to what I want to try in George Church's lab in Accurate Multiplex Polony Sequencing of an Evolved Bacterial Genome a recentish Science paper. His situation was sheared DNA with messy ends. He used the End-It DNA Repair Kit [Epicenter] to make the ends blunt. Then he added taq with ONLY dATP to put the A on the end. I'm going to try this and see if it fixes the huge product problem (see Figure 3.4)
  1. digest 24 ml plasmid DNA minipreped from lrpB1 and lrpB2; master mix of enzyme cocktail (0.25 ml MspI, 5ml Buffer 2, 21 ml H2O ); digests were run in 50ml rxns for 1 hr at 37C followed by 20 min at 65C to heat deactivate the enzymes.
  2. ethanol precipitate; 5 ml NaAcetate (into the 50 ml ), mix, add 110 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum out ethanol, add 750 ml COLD 70% ethanol, spin 3 min, (dry) 4
  3. resuspend in 34 ml TE
  4. add 5 ml End-Repair Buffer, 5 ml dNTP Mix, 5 ml 10mM ATP, 1 ml End-Repair Enzyme Mix
  5. incubate RT for 45 min, deactivate at 70 C for 15 min
  6. run all of lrpB3 on a gel to make sure it is ok, post End-Repair (put lrpB3 in freezer, so all could be run on the same gel)
  7. ethanol precipitate; 5 ml NaAcetate (into the 50 ml ), mix, add 110 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum out ethanol, add 750 ml COLD 70% ethanol, spin 3 min, (dry) 5
  8. resuspend in 25 ml TE
  9. A-tail DNA: 5 ml standard taq (mg-free) rxn buffer [NEB], 6 ml 25 mM MgCl2 (3 mM final), 0.25 ml dATP (0.5 mM final), 0.5 ml Taq (2.5 units), 25 ml sample
  10. run all of lrpB2 on a gel to make sure it is ok, post A-tail (put in freezer, so all could be run on the same gel)
  11. ethanol precipitate; 5 ml NaAcetate (into the 50 ml ), mix, add 110 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum out ethanol, add 750 ml COLD 70% ethanol, spin 3 min, (dry)
  12. resuspend in 16 ml TE
  13. ligate; 16 ml of lrpB3 for 2 hr at 16C. halted by incubation 65C for 10 min. (1 ml SAPE1 linker, 16 ml sample, 2 ml buffer, 1 ml T4 DNA ligase [NEB])
  14. run lrpB3 on a gel (again put in freezer)
Sun May 28 18:35:09 EDT 2006
All three samples (lrpB1,2,3) are being run on a 1.5% agarose. In addition, I made two midipreps using the on page 6, so I should have lots of plasmid and not have to waste a bunch of money and time on the Qiagen preps. One of the samples was also run on the gel. Yields were:
Sample DNA (ng/ul) 260/280 260/230
lrp B 1 Midi 1613.4
lrp B 2 Midi 1779
Please see the pdf version for figures
Figure 3.6:
Brief Conclusions:   I wish I started with a little more DNA. It would be nice if the DNA were concentrated enough that I could track a single sample through all the steps (rather than assuming that lrpB1 is like B2 is like B3). I shouldn't have switched to a 1.5% gel (see Figure 3.6 right pane). The lower wt bands are hard to see. Even the ladder is hard to see as far as that goes. My gels seem to be getting crappier recently. However, I think that ligation is working. Unfortunately it is hard to see the lrpB3 lane (again I need to start with more DNA). The lrpB2 post A-tail where I used to have problems looks a lot better. It does start to get a little smeary towards the top, but not the giant piece like I was getting previously (e.g. Figure 3.4). And the best thing is the DNA starts to get larger in size after the ligation step which is just right. I think adding a phosphatase step before the A-tailing would be good. I don't think I can conclude if lrpB3 sucks because the end-repair messed it up or because I spilled the tube. Ideally I'd run one sample straight through starting with about 5x my normal starting amount of DNA (e.g. 12.5 mg ) and pull out a sample to run after: 1) digestion, 2) end-repair, 3) A-tail, 4) ligation. And maybe 200 ng of uncut plasmid just to be safe. The MIDI prep using the new protocol does not seem to have worked (Figure 3.6). I need this to work if I'm going to have enough DNA.

Midipreps

Mon May 29 2006
I need more DNA if I'm going to trace one sample all the way through this process (I think I need around 10 mg of DNA). I tried to do a midiprep using the protocol in Molecular Cloning. My version of the protocol (little to no modifications just clarified things based on my previous attempt above) can be found on page . I ran 4 preps using 15 ml of culture for each. The results of each of these four and the one from the previous day that I didn't run on a gel are in Figure . One microliter was run in each lane (combined with 9 ml H2O and 1.6 ml 6x loading dye). Yields were very high:
Sample DNA (ng/ul) 260/280 260/230
lrp B 1 Midi 2073
lrp B 2 Midi 1679.6
lrp B 3 Midi 1720.9
lrp B 4 Midi 2215.6
Please see the pdf version for figures
Figure 3.7:
Brief Conclusions:   This sucks. I seem to have only gotten a large amount of tiny fragments in both of my attempts at midipreps (see Figures 3.6 last column and 3.7). I'm giving up on this protocol for now. My only guess as to what went wrong is that the phenol:chloroform is old, but I don't know if that stuff goes old. The bottle is almost empty so I ordered new and will just toss the remaining 3 ml or so. For now I ordered the Qiagen HiSpeed Midi kit. Hopefully that will give me more success than the old school Sambrook method.

3.1.3  SAPE1 returns: following the same sample all the way (including PCR)

Wed May 31 14:45:54 EDT 2006
I won't get started until Mon cause sis and mom are in town, but want to build my gameplan for the week. Gameplan broadly:
  1. inoculate 3 x 50 ml cultures for monday (Sunday)
  2. make a ton of DNA with the midiprep kit for 3 samples (Monday)
  3. test the 58 operon promoters alone and perhaps in couples with NEB master mix (Monday)
  4. run SAPE1 to completion test samples at each step include G tail and PCR (Tuesday)
  5. try SAPE2 (Wednesday)

The plan for SAPE1

Wed May 31 15:32:25 EDT 2006
Modify below based on conc of DNA at each step.
  1. digest 24 ml plasmid DNA midipreped from lrpB1 and lrpB2; master mix of enzyme cocktail (ml MspI, 5ml Buffer 2, 21 ml H2O ); digests were run in 50ml rxns for 1 hr at 37C followed by 20 min at 65C to heat deactivate the enzymes.
  2. run 1/5 of sample on gel
  3. ethanol precipitate; 5 ml NaAcetate (into the 50 ml ), mix, add 110 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum out ethanol, add 750 ml COLD 70% ethanol, spin 3 min, (dry)
  4. resuspend in 34 ml TE
  5. add 5 ml End-Repair Buffer, 5 ml dNTP Mix, 5 ml 10mM ATP, 1 ml End-Repair Enzyme Mix
  6. incubate RT for 45 min, deactivate at 70 C for 15 min
  7. is Antarctic phosphatase step possible. try adding antarctic phosphatase buffer to 1x. calculate units based on conc of DNA. incubate 37 C for 30 min, heat deactivate 65C for 5 minutes
  8. run 1/4 of sample on gel
  9. ethanol precipitate; 5 ml NaAcetate (into the 50 ml ), mix, add 110 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum out ethanol, add 750 ml COLD 70% ethanol, spin 3 min, (dry) 7
  10. resuspend in 25 ml TE
  11. A-tail DNA: 5 ml standard taq (mg-free) rxn buffer [NEB], 6 ml 25 mM MgCl2 (3 mM final), 0.25 ml dATP (0.5 mM final), 0.5 ml Taq (2.5 units), 25 ml sample
  12. run 1/3 of sample on gel
  13. ethanol precipitate; 5 ml NaAcetate (into the 50 ml ), mix, add 110 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum out ethanol, add 750 ml COLD 70% ethanol, spin 3 min, (dry)
  14. resuspend in 16 ml TE
  15. ligate try a different ligation time for each of the samples; 16 ml of lrpB3 for 2 hr at 16C. halted by incubation 65C for 10 min. (1 ml SAPE1 linker, 16 ml sample, 2 ml buffer, 1 ml T4 DNA ligase [NEB])
  16. run 1/2 of sample on gel
  17. end label with dGTP
  18. PCR amplify with some of the G-labeled DNA
  19. run PCR product to see if it worked

3.1.4  More DNA: midipreps, midipreps, midipreps

Wed Jun 7 17:42:23 EDT 2006
I've been having a heck of a time getting midipreps to work. I tried an old-fashioned approach and a the new HiSpeed Qiagen midiprep. The Qiagen kit gave me clean DNA but not very much of it (see and table below). I'm not sure why the yield was so low. I haven't yet tried Qiagen's debugging process. Meanwhile I figure out the problem with the earlier midipreps and the huge amount of crap right around 50 bp (see Figures 3.7 and A). The problem was that adding RNA too the 1st lysis buffer was insufficient to get rid of all the RNA it seem to chop it into this small fragment size. I read on the web a good place for the RNA digestion is right after you precipitate the cellular wall and genomic DNA (right before the phenol chloroform step). In Figure , I digested the midiprep with a large amount of RNAse cocktail and RNAse A (sample on the right only) and the RNA the samples (before RNAse Figure A). The RNA was also drastically messing up my measurements of DNA concentration.
Yields from Qiagen midipreps (500 ml total volume; run on Mon Jun 5, 2006):
Sample DNA (ng/ul) 260/280 260/230 total yield (ug)
1 32.6 16.3
2 77.1 38.6
Yields from old-school midipreps (100 ml total volume; run on Tue Jun 6, 2006):
Sample DNA (ng/ul) 260/280 260/230 total yield (ug)
1 3838.2 383.8
2 3727.8 372.8
Yields from old-school midipreps post RNA digestion (100 ml total volume; quantified Thur Jun 8, 2006):
Sample DNA (ng/ul) 260/280 260/230 total yield (ug)
1 72.3 7.2
2 63.3 6.3
Please see the pdf version for figures
Figure 3.8: Qiagen midipreps didn't produce nearly as high a yield as they claim I can get in the manual. Genomic DNA in the first lane is for use in the operon study.
Please see the pdf version for figures
Figure 3.9: 0.75 ml of midiprep DNA (total volume) from the old-school method on a 1% agarose gel. Initially RNA was high and yield was high. Then I digested with RNAse cocktail and the RNA went away (as did the yield which I guess was mostly RNA).
Mon June 12, 2006 Ran six more Sambrook mini-preps with only slightly more luck than before. Here are the yields before RNAse digestion.
Sample DNA (ng/ul) 260/280 260/230
lrp B 1 Midi (10 ml starting culture) 670
lrp B 2 Midi (10 ml starting culture) 1160.8
lrp B 3 Midi (15 ml starting culture) 950.9
lrp B 4 Midi (15 ml starting culture) 1246.6
lrp B 5 Midi (25 ml starting culture) 1935.0
lrp B 6 Midi (25 ml starting culture) 1519.8
and after RNAse digestion.
Sample DNA (ng/ul) 260/280 260/230
lrp B 1 Midi (10 ml starting culture) 186.7
lrp B 2 Midi (10 ml starting culture) 284.5
lrp B 3 Midi (15 ml starting culture) 246.1
lrp B 4 Midi (15 ml starting culture) 237.2
lrp B 5 Midi (25 ml starting culture) 246.2
lrp B 6 Midi (25 ml starting culture) 152.9

3.1.5  Midipreps that work and are RNA free!

Sun Jun 18, 2006
I was just about to give up on the midi idea altogether. I grew cells to make 6 Qiagen minipreps (with 5 ml of culture apiece to get maximal yield). I also included a new protocol for midipreps (see section page ). I planned to ethanol precipitate the 6 preps into one more concentrated prep if the midi didn't work.
Yields were as follows:
Sample DNA (ng/ul) 260/280 260/230
lrp B 1 Mini (5 ml starting culture) 163.9
lrp B 2 Mini (5 ml starting culture) 155.5
lrp B 3 Mini (5 ml starting culture) 143.2
lrp B 4 Mini (5 ml starting culture) 150.5
lrp B 5 Mini (5 ml starting culture) 146.7
lrp B 6 Mini (5 ml starting culture) 159.1
lrp B 1 Midi (50 ml starting culture) 395.5
Please see the pdf version for figures
Figure 3.10: 0.75 ml of miniprep (6 samples) and midiprep (1 sample) DNA (total volume) from on a 1% agarose gel. RNA is not visible. DNA is not nicked for the Midiprep as it has been with the Sambrook Midiprep.
Brief Conclusions:   After much frustration getting a simple technique to work, I finally found a protocol that works. And it seems to work well (see Figure 3.10). Unlike with the Sambrook protocol midipreps, this new method doesn't result in nicked DNA (not sure why). Unfortunately, I only had two midiprep samples and one of them was spilled before I completed the protocol. But for now, I'll assume this annoying search for a plasmid prep with high yield and RNA free DNA is over.

3.1.6  SAPE2 a modified protocol that hopefully works (original SAPE did not)

Quick thought on SAPE1... Maybe it didn't work because I didn't clean up the reaction before adding the Taq? I'd like to check that out using the method from Shendure in the Church lab to add ends (actually I have see section 3.1.2).
Please see the pdf version for figures
Figure 3.11: If one primer PCR is a problem, we can try the end labeling trick from SAPE1

3.1.7  stab at an initial SAPE2 detailed protocol

Wed May 31 15:32:57 EDT 2006
Stab at a first protocol.
  1. test Sss I by running on a smaller sample and verifying that it blocks MspI and AatII digestion (next step)
  2. digest 24 ml plasmid DNA midipreped from lrpB1 and lrpB2; master mix of enzyme cocktail (ml MspI, 5ml Buffer 2, 21 ml H2O ); digests were run in 50ml rxns for 1 hr at 37C followed by 20 min at 65C to heat deactivate the enzymes.
  3. SssI protection of CG sites; heat inactivation 20 min at 65 C
  4. add Antarctic phosphatase buffer and enzyme to remove phosphates from the end 15 min 37 C (is 15 min enough?), deactivate 65 C for 5 min
  5. run 1/5 of sample on gel
  6. ethanol precipitate; 5 ml NaAcetate (into the 50 ml ), mix, add 110 ml 95% ethanol; -85C 20 min, spin 4C 10 min, vacuum ou t ethanol, add 750 ml COLD 70% ethanol, spin 3 min, (dry)
  7. resuspend in ml TE
  8. ligate phosphorylated linker in NEB buffer for with 1 mM ATP added. heat inactivate ligase
  9. digest with AatII (do I need to ethanol precipitate?). or digest with SmaI then with AatII
  10. ligate linked promoters (maybe add more dATP in)
  11. (stop for now run on a gel) don't have primers for the rest of this. make sure this works first
  12. remove phosphates with Antarctic Phosphatase
  13. finally get to use a freaking kit. before PCR amplification, clean up DNA with Qiagen PCR cleanup kit

3.2  The end of SAPE

Wed Aug 23 15:19:15 EDT 2006
I think I'm going to abandon SAPE. The paper: Multiplex sequencing of paired-end ditags (MS-PET) a strategy for the ultra-high-throughput analysis of transcriptomes and genomes. pretty much does what I wanted to do, but maybe even better (though there protocol is no easier than mine and perhaps harder). I think it would be more effective to make PETs like I'm doing with my experimental transcript determination study. Then I could just sequence them to determine binding sites.