Genomic Lambda Library Methods

Written by Michael Frohlich
Based on methods from the Promega Protocols and Applications Guide, second edition
Elliot Meyerowitz Lab, Cal Tech

This is a description of methods I've used to make genomic DNA libraries. Most of this discussion deals with the partial digestion, because that is the step which caused me the most difficulty. These methods are based on the instructions in the Promega Protocols and Applications Guide, second edition, for use with the "Lambda Gem 11 Xho partial-filled-site kit", but I have made modifications to save DNA and enzyme, and to get better reproducibility between small-scale and large-scale digests. The instructions in the third edition of the guide are similar to those in the second edition.
These instructions assume that you have a 2 µl pipettor (e.g., Gilson P-2). The small volumes called for here could not be measured accurately with a standard P-20 pipettor. I think the methods (to my mind wasteful) recommended by Promega are designed to avoid measuring small volumes, which makes waste inevitable.

First isolate the DNA. It must be high molecular weight, over 40 Kb in length. This is not difficult; the methods I use are described in another text ("CTAB DNA isolation"). I band the DNA in CsCl, to make sure it is pure. While this may not be strictly necessary, it does remove enzyme inhibitors that could interfere with digestion, and cause different DNA samples to require very different enzyme concentrations for partial digestion. The DNA must be quantified by spectrophotometry or by comparison on a gel to samples of known amount.

The first and most troublesome step is the partial digestion. If one has DNA to waste then one can do a series of large-scale partial digestions (using a total of a few hundred µg of DNA) and then choose the best for further processing. More often the DNA is precious, so one does a series of small-scale partial digestions to determine optimal conditions, and then one does a single large-scale digestion at these conditions. This requires that the digestions be reproducible, so that the large-scale digestion results in the same size spectrum of fragments as the optimal small-scale trial. The enzyme and DNA concentrations must be the same in both large- and small-scale digestions. The volume of the large-scale digestion is limited by the need to precipitate it conveniently, so the trial digestion, done at the same concentrations, uses very small volumes. Of course, the temperatures and times must also be the same, but this is easy to achieve.

There should be a vial of enzyme for use in partial digestions only, so variation in enzyme activity is minimized. One vial will last through many digestions. I use the Bohringer-Manheim enzyme. I used the buffer they supplied until it ran out; then I made my own buffer, which worked well (recipes are at end of this file).

Detailed instruction for partial digestion:

1. Check the calibration on the p-200 pipettor, by weighing ca 80 µl of water. Put it into a small eppindorf to prevent evaporation. (This will be important at step 29.)
2. Make the gel for visualizing the trial digestions. It should be 0.3% agarose in TBE buffer (TAE should also be OK). These gels are very fragile. I always keep them on the carrier when visualizing the bands on the UV box. One must have a UV-transmitting gel carrier. I use 10 µl of 3mg/ml Ethidium Bromide per 40 ml of gel, and I put double this amount into the downstream well of the gel box. Teeth on the comb should be narrow- ca. 2.5 mm wide (smaller amounts of DNA can be visualized in narrower lanes - I use ca 200 ng per lane). The teeth and the gel should each be thick enough so 10 µl of material can be loaded into each well. My gels are ca 10 cm long.
3. Label tubes. I use one (or two) 1.5 ml eppindorfs for the enzyme dilution buffer, and 1.5 ml eppindorfs for the diluted enzyme solutions (see table of dilutions on page 5 to determine how many and what tubes are needed). I do the actual digestions in 0.5 ml eppindorfs. These should be labeled on their tops.
4. Put labeled eppindorfs into ice in two (or one) ice buckets. The tubes should be cold before enzyme is placed into them.
5. Put tube of oil (for covering test digestions) on ice.
6. Weigh (on a good balance) the small eppindorfs in which the DNA mix will be prepared. Record the weight.
7. Prepare the DNA mix. Concentration of DNA in the mix should be 100 ng/µl. I typically use a total of 10 µg of DNA, so the final volume is 100 µl. (Table headings on page 5 give a template for recording calculations). I dilute the DNA with TE, not with water, so that the amounts of tris and EDTA are constant in the mix, no matter the concentration of the original DNA solution (which is surely in TE). I also put 1/10 volume of Sau3A buffer in the mix, so the DNA has everything except BSA needed for digestion. Keep the DNA mix on ice at all times. If there are nucleases in the DNA solution it may become degraded; one can check for this by warming an aliquot of DNA mix to 37° and comparing it to material kept on ice. No change should be apparent. Any nucleases would defeat the partial digestion, so if problems are encountered, then presence of nucleases should be checked.
8. Mix enzyme dilution buffer (recipe on page 5). I make up enough for both the large scale digestion, and for the trials, to eliminate a possible difference between trials and large scale digestions.
9. Put indicated amounts of enzyme dilution buffer in the tubes for diluted enzyme. Close tops.
10. Get Sau3A enzyme from the freezer, and put 1 µl of enzyme into dilution tube one, using the P-2 pipettor. I wash the tip by sucking in-and-out a couple of times; I do not do that in subsequent dilution steps. It is especially important to just touch the top surface of liquid in the enzyme vial when measuring with these small tips; large amounts of liquid can stick to the outside if the tip is stuck into the liquid.
11. Mix the contents of dilution tube number one by sucking it in-and-out with a 20 µl (yellow) pipette tip. Discard this tip and use a dry one to measure the 15 µl to put into dilution tube 2.
12. Mix dilution tube two by vigorous finger vortexing.
13. Put 10 µl into the other dilution tubes (up to tube nine; mix and aliquot from tube seven if further dilutions are needed). Use a fresh dry tip for each transfer; wet tips may yield slightly different volumes.
14. Mix each dilution tube by vigorous finger vortexing. If tubes are frothy, suck air out of bubbles with a 20 µl pipette tip, so bubble-free surface is available for aliquoting small volumes.
15. Put exactly 2 µl of the appropriately-diluted enzyme into each small eppindorf. Return small tube to ice and cap immediately. Also keep the large tubes of diluted enzyme on ice (these same dilutions will be used for the large-scale digestion.)
16. With fingers only, gently "vortex" the DNA mix solution for 30 sec. High molecular weight DNA will probably not be completely dissolved, so clumps of DNA will settle out of the mix to the bottom part of the tube, causing aliquots removed from the top to have too-little DNA, which will allow them to get more digested than DNA in the supposedly comparable large-scale. This is a dangerous source of non-reproducibility between small- and large-scale digestions.
17. Put exactly 2 µl of DNA mix into each small eppindorf. Cap and return to ice immediately.
18. Bring ice bucket containing small eppindorfs to a refrigerated microfuge (most likely in a cold room). Also bring a box of 20 µl tips, the cold oil, and TWO pipettors (one 20 µl; the other 20 or 200 µl).
19. Put sleeves into the microfuge so small tubes can be spun safely. (Topless 1.5 µl eppindorfs work fine, but they will eventually fail, so decapitate new ones to replace any ancient ones lying around.)
20. Spin the small eppindorfs briefly to get all liquid to the bottom.
21. As tubes are removed from the centrifuge, open each one and mix contents by sucking in-and-out with a fresh yellow tip. Then put ca 10 µl oil over each one, and put on ice immediately.
22. Put tubes in 37° water bath or heat block.
23. After 30 minutes remove from heat block and put on ice.
24. Open tubes and put in 6 µl of EDTA-bromophenol blue loading buffer; leave the 20 µl tip in the tube.
25. Load test digestions onto gel. Use ca 50 or 100 ng lambda Hind III DNA as migration standard.
26. Run gel at low voltage for 2 to 5 hours. I use 24 volts for a 10 cm gel. Degree of digestion is easier to evaluate after a longer run, and enzyme does seem to remain stable for 5 hours, but gel can be evaluated after 3 or, with difficulty, after 2 hours.
27. Photograph and evaluate gel. Because fluorescence measures the mass distribution, rather than fragment number, and because smaller fragments are more spread-out than larger ones, the brightest part of a lane is not the region with the highest number of fragments- it is at a much smaller size. Standard wisdom says one should observe what enzyme dilution gives a maximum of fluorescence in the 23 to 9 kb region, and then use one-half this enzyme concentration for the large scale digestion. In practice I look for the lane with maximum fluorescence on the larger side of 27 (or 23) Kb as the optimal enzyme concentration. This gives about the same dilution as the other method.
28. Weigh the eppindorf containing what is left of the DNA mix. Subtract the weight of the empty tube (determined in step 6). This indicates how much DNA mix remains, and how much enzyme solution should be added.
29. Check that enough diluted enzyme remains to do the large scale digestion. If not, then make more of this dilution, using the left-over enzyme dilution mix and dilution number two (or seven). (The recipe on p. 5 calls for lots of dilution number two [and seven] to be made, so there will be enough to dilute the enzyme for the large-scale digestion.)
30. Add precisely the same weight of the appropriate diluted enzyme to the DNA mix. This is most easily done by adding ca 10% too little enzyme mix, then weighing the tube again, and adding the few additional microliters needed to get the correct amount. Most pipettors are off a little bit, so the calibration in step 1 is important here.
31. Mix by finger vortexing, and return to ice for about the time that the test digestions were on ice between mixing and putting at 37°
32. Put at 37° for the same length of time as the small-scale digestions.
33. After removing from 37° immediately add an equal volume of phenol-chloroform. Mix, Spin and remove top layer to a large eppindorf.
34. Back-extract by putting a little TE into the small eppindorf, mixing the top layer, and putting the top layer into the large eppindorf.
35. Add an equal volume of chloroform. Mix, Spin and remove top layer to another large eppindorf. Be careful not to touch the plunger on the cap with fingers, as the DNA will be stored in this tube, and fingers can leave DNAases behind.
36. Back-extract with a little TE. Top off the large eppindorf with TE to a convienent volume for precipitation.
37. Precipitate with 1/2 volume of 7.5 M ammonium acetate and 2.0 total volume of abs ETOH. Store in freezer for 30 minutes or more. To pellet DNA I first put it in the microfuge with hinge facing inward, then I spin again with hinge facing outward. This gets DNA into a pellet at the bottom of tube, rather than along the side of the tube.
38. Decant, remove droplets with a Q-tip, add 0.5 ml 80% ETOH and keep in freezer for 5 min, then spin, decant the ETOH, remove droplets with a Q-tip, dry in air, and dissolve in 20 µl TE.

Steps after the Partial Digestion:

• From this point I follow instructions in the Promega Protocols and Applications book, edition 2, page 182, part A (Partial fill-in reaction for cloning into Xho I half-site Arms).
• I do the partial fill-in as recommended by Promega. The fill-in buffer can go bad in a -20 degree freezer, especially if the freezer is not working well, and maybe if people keep the door open too long too often. I destroyed lots of partial digests when a batch of this buffer went bad. Both the DTT and the dNTPs can go bad. I mixed my own buffer, according to Promega's recipe, with fresh DTT and dNTPs; I store it in small aliquots at -80. I thaw a new aliquot each time I do reactions; I don't re-freeze it.
• After the fill-in I do the phenol-chloroform extraction and precipitation. With care one can keep losses of DNA to a minimum, but that is not necessarily desirable. I've had some of my best luck in library making when I lost half of the DNA in the two precipitations. I suspect the losses were mostly in the shorter pieces, which eat up lamdba arms in the ligation step.

I mix my own ligation buffer from fresh ingredients and also store that in small alliquots at -80. I do the ligation optimization, as described in part E (p. 183), and when very lucky I've even got enough clones from these optimization reactions for a whole library. The only major problem I've had in the ligation step is getting the lambda arms to dissolve, which they are often reluctant to do, even after sitting at room temperature for an hour. (I've seen tiny bubbles in the solution that move in concert, proving that a DNA gel is still present.) I add all the water that will be used in the reaction directly to the vial of lambda arms, mix vigorously by finger-vortexing, and let it sit for a while before use. If possible, I let the ligations proceed for several days in the refrigerator, both due to the slowness of the reaction itself and to let the DNA dissolve so it can react.

When amplifying I plate ca 10,000 clones on a 15 cm petri plate. I wash the clones off with SM (Sambrook, et al. 1989 p. A7), rather than by scraping off the top agar. It is worth keeping the library divided into several sub-libraries, so one can be sure phages in the different sublibraries came from different original fragments of partially digested genomic DNA.

Recipes:

MF Sau 3A buffer (10x):

NaCl	1 M
Tris HCl pH 7.5	100 mM
MgCl2 100	100 mM
DTT	5 mM

Enzyme dilution buffer: [Promega calls for BSA in both the DNA solution and the diluted enzyme. I put it all in the enzyme dilution, so I use twice the recommended amount for the enzyme dilution buffer.]

MF sau 3A buffer (10x)	100 µl
Acetylated BSA 10 mg/ml	20
H2O	880
Total	1000 µl

Enzyme Dilutions: Note that dilution number TWO has a lower concentration than in Promega’s instructions. Generally I use only the concentrations in bold face for trial digestions. Most often numbers eight or nine are the optimal concentrations for large-scale digestion.

Tube #	Amnt enz to add	Which enz dil to use	Amnt of dil buffer to add	Relative enzyme concentration
1	1.0	Sau3A stock	19	1
2	14.3	# 1	200	0.0667
3	10	2	10	0.0334
4	10	2	30	0.0167
5	10	2	50	0.011
6	10	2	70	0.0083
7	10	2	90	0.00667
8	10	2	140	0.0044
9	10	2	190	0.0033
10	10	7	20	0.0022
11	10	7	30	0.00166
12	10	7	50	0.0011
13	10	7	70	0.00083
14	10	7	90	0.00066

EDTA loading buffer (use no Xylene cyanol).

15% ficol		200 µl
Bromphenyl blue (11 mg dye in 500 µl Tris 1 M pH 8)		ca 5
EDTA (0.25 M)		75
Ethidium Bromide solution (3 mg/ml)		5
Total		280 µl

Table for recording DNA amounts used in test partial digestions of several DNA samples for library making. (I assume 10 µg of DNA is being put into a mix of 100 µl total volume.)

DNA isolation number	DNA Name	ng/µl in stock DNA soln [DNA]	µl DNA stock per tube 200/[DNA] (gives 200 ng)	Total volume of stock soln for 10 µg of DNA 10000/[DNA]	Vol 10x MF Sau3A buffer	Additional TE needed (100 - 10000/[DNA])
					10 µl