CHAPTER 7

A BAC vector can carry an insert greater than 100,000 base pairs in length. However, standard DNA isolation methods tend to shear DNA into fragments too small to take advantage of the carrying capacity of BACs. To limit DNA shearing, purified nuclei are embedded in agarose plugs. The agarose provides a physical support for the DNA preventing significant shearing. Plugs are incubated in buffer containing proteinase(s) to digest proteins and detergent to emulsify nuclear lipids, respectively.

Preparing nuclei suitable for BAC library construction can be one of the most difficult steps in making a plant BAC library. The predominant problems involved in trying to isolate plant nuclear DNA are ones that animal researchers do not typically encounter. For example, (a) plant cell walls must be physically broken or enzymatically digested without damaging nuclei, (b) chloroplasts must be separated from nuclei and/or preferentially destroyed, an important process since copies of the chloroplast genome may comprise the majority of DNA within a plant cell, (c) volatile secondary compounds such as polyphenols must be prevented from interacting with the nuclear DNA, and (d) carbohydrate matrices that often form after tissue homogenization must be prevented from trapping nuclei. While it would be ideal if there were a nuclear DNA isolation protocol that worked for all plant species, the biochemical and morphological diversity within the plant kingdom make development of such a protocol unlikely (Loomis 1974, Peterson et al. 1997). Below we present two quite different nuclear DNA isolation protocols that we have used to construct BAC libraries from plants. OPTION X is a promising technique that has only recently been used in BAC library construction. OPTION Y (or prototypes of this protocol) has been used in the construction of BAC libraries for several years.

OPTION X is an adaptation of a procedure originally designed for isolating milligram quantities of highly pure nuclear DNA from tomato (Peterson et al. 1997, 1998). It has several features that make it well suited for use in BAC library construction as well as other molecular biology applications: (a) Prior to homogenization, tissues are treated with ether to make nuclei more friable. Ether treatment markedly increases the yield of nuclei (Watson and Thompson 1986; our observations). (b) Homogenization is performed using a simple kitchen blender. (c) The nuclear isolation buffer (MEB) is designed to deal with several common problems in plant nuclear DNA extraction. First of all, the buffer contains 2-methyl-2,4-pentanediol (MPD), a compound that helps stabilize nuclei and prevents their premature lysis. Nuclear yield using MEB is > 10 times that obtained using sucrose-based buffers (Peterson et al. 1997 and our observations). The buffer also contains the antioxidants ß-mercaptoethanol, sodium diethyldithiocarbamate, and sodium metabisulfite. These compounds limit the oxidation of polyphenols. In their oxidized forms, polyphenols covalently bind to DNA turning it brown and making it useless (Katterman and Shattuck 1983; Guillemaut and Maréchal-Drouard 1992). Polyvinylpyrrolidone in the buffer adsorbs polyphenolic compounds preventing them from interacting with DNA (Loomis 1974). (d) The low pH of the buffer (pH 6.0) serves to inhibit polyphenol oxidation. (e) After homogenization, addition of Triton X-100 to a concentration of 0.5% results in preferential lysis of chloroplasts and mitochondria. The presence of divalent cations (Mg²⁺) in the MEB prevents nuclei from being lysed by the Triton X-100 (Watson and Thompson 1986). (e) Nuclei are separated from most debris by centrifugation through a Percoll gradient. Further low-speed centrifugation steps are used to remove some, if not most, of the starch grains that typically pellet with nuclei. (f) Throughout the protocol, nuclear preparations are examined using a light microscope. This allows the investigator to visually assess nuclear concentration and purity.

We have used OPTION X to isolate nuclei and nuclear DNA from numerous plant species (e.g., sorghum, sugarcane, grape, cotton, loblolly pine, prickly-pear cactus, fern, peanut, Leyland cypress). The protocol's usefulness in BAC library construction was first demonstrated in August 1999 when it was used to isolate DNA that subsequently was used to construct a 9X library for Gossypium raimondii (Peterson et al., in preparation). This success quickly was followed by construction of a 16X library for grape (Vitis vinifera) (Tomkins et al., in preparation) and a 9.2X library for Gossypium hirsutum (Acala Maxxa) (Tomkins et al., in preparation). Using OPTION X, agarose plugs containing nuclear DNA of suitable size and restrictability for BAC library construction have been generated for peanut (Arachis hypogaea 'Florunner') and tomato (Solanum lycopersicum). The peanut plugs are currently being used in BAC library construction.

OPTION X has produced megabase-sized, restrictable DNA from all dicots in which it has been tested, but whether it will be useful in isolating megabase-sized DNA from monocots is still uncertain. Several attempts at isolating high molecular weight DNA from Sorghum bicolor using OPTION X yielded fragments too small (< 100 kb) for BAC cloning, presumably due to partial digestion of the DNA by endogenous nucleases. Attempts to limit nuclease activity by removing Mg²⁺ (a DNase cofactor) from the nuclear isolation buffers (either directly or indirectly) resulted in premature nuclear lysis. However, addition of the nuclease inhibitors EGTA (6 mM) and L-lysine-HCl (200 mM) (see Liu and Wu 1999) to the nuclear isolation buffers permitted isolation of sorghum DNA fragments about 800 kb in length without a noticeable decrease in nuclear yield. Further research on the use of OPTION X to isolate megabase-sized DNA from monocots is in progress.

Another potential drawback of OPTION X is that it is designed for extraction of nuclei from relatively large quantities (ca. 500 g) of fresh tissue. Whether the protocol can be scaled-down to accommodate smaller quantities of tissue and/or used to isolate high molecular weight DNA from frozen tissue has yet to be tested.

OPTION Y is relatively simple and can be used to isolate nuclear DNA from somewhat smaller quantities of fresh or frozen tissues. It has been used to construct DNA libraries from numerous species including rice, sorghum, wheat, sugarcane, cotton, soybean, barley, and Arabidopsis. In OPTION Y, plant tissue is ground in liquid nitrogen, the resulting homogenate is suspended in a sucrose-based buffer (SEB = sucrose extraction buffer), Triton X-100 is added to destroy chloroplasts and mitochondria, the homogenate is filtered several times to remove most cellular debris, and nuclei are pelleted by centrifugation. The relatively high pH of the SEB (pH 9.1) inhibits the activity of endogenous nucleases. The presence of ß-mercaptoethanol and PVP in the SEB counteract some of the negative effects of oxidized polyphenols. OPTION Y has been used to prepare megabase-sized DNA from monocots and some dicots. It is particularly well suited for species/tissues that contain little in the way of secondary compounds or carbohydrates. Disadvantages of OPTION Y include relatively low nuclear yields (and subsequently production of agarose plugs with lower DNA concentrations), significant contamination of nuclear preps with debris and carbohydrates, and limited control of polyphenols.

For dicots and non-angiosperms, we recommend that investigators start by trying OPTION X. At present, we suggest that investigators working on monocots try OPTION Y first. If neither OPTION X nor OPTION Y work for a species of interest, there are a number of other nuclear isolation protocols that can be tried (e.g., see Katterman and Shattuck 1983; Guillemaut and Maréchal-Drouard 1992; Watson and Thompson 1996; Hamilton et al. 1972; Couch and Fritz 1990).

SUPPLIES, EQUIPMENT, AND REAGENTS (see CHAPTER 2 for details): MEB; MPDB; cheesecloth; 1X TE slurry; 1X TE (non-sterile); Percoll; diethyl ether; 1% methylene blue; kitchen blender; light microscope

METHODS:

1. This nuclear DNA isolation method can be used to isolate DNA from adult plants and seedlings.

1. For mature plants, young leaves are gently picked from the plant of interest. After removing a leaf, it is immediately submerged in a 1X TE slurry. Approximately 200-500 g of leaves are collected for one isolation.
2. For DNA isolation from seedlings, plant seeds at high density in soil in greenhouse trays, cover seeds with 0.5 cm of soil, water, and place trays in the greenhouse. At the time of harvest, cut the tops from the plants and submerge the tops in a 1X TE slurry. The optimum time of harvest depends on the plant. For monocots, allow seedlings to grow to a height of approximately 20-30 cm before harvesting. For dicots, wait until true leaves appear and flourish before harvesting as cotyledons are senescent tissue and do not yield good quality DNA. Approximately 500 g of seedlings are collected for one isolation.

2. Remove leaves/seedlings from the 1X TE slurry and place them in 1-3 L of ice-cold diethyl ether (i.e., enough to submerge all tissue) for three minutes.

­ Note 7.1: Perform this step in a fume hood!

Ether removes waxes and makes cells more friable. Remove leaves/seedlings from the ether and then wash them three times (1 min each wash) in 4 L of 4°C 1X TE (prepared from the 100X TE stock). Place the leaves/seedlings in 3 L of ice cold MEB. Homogenize leaves/seedlings in the MEB using a kitchen blender (highest speed attainable for 30 seconds). Squeeze the homogenate through six layers of cheesecloth, and then filter the resulting filtrate through 32 layers of cheesecloth. If possible, let this second filtration occur by gravity only (i.e., no squeezing).

3. Add Triton X-100 to the filtrate to a final concentration of 0.5% v/v. Fill four 500 ml centrifuge bottles with the mixture and spin at 800 x g for 20 min at 4°C. Decant supernatants, place the remaining homogenate in the bottles, and centrifuge as described above (this reduces the number of bottles one has to wash at the end of the experiment).
4. Resuspend each pellet in 4 ml of MPDB, and pool suspensions. Mix a small drop of the suspension with an equal volume of 1.0% methylene blue on a microscope slide. Add a coverglass and examine the slide by phase-contrast and/or bright-field microscopy. If all has gone well, the mixture should contain numerous dark-blue stained nuclei, no intact cells, and no mitochondria or chloroplasts. Starch grains (visible by phase-contrast microscopy) should stain little, if at all, with methylene blue (see FIGURE 7.1).
5. Place 7.5 ml of Percoll in a 50 ml polypropylene centrifuge tube. Add MPDB to a final volume of 20 ml to produce "37.5% Percoll". Layer the suspension of nuclei onto the 37.5% Percoll bed. Centrifuge the gradient in a swinging bucket rotor at 650 x g for 60 minutes (4°C). Nuclei are relatively dense structures and normally form a pellet (along with large starch grains) at the bottom of the centrifuge tube. Less dense materials (e.g., most cell debris, organelles, plastid DNA) remain in the supernatant.

­ Note 7.2: On occasion, some or all of the nuclei will not pellet. This can occur if the batch of Percoll is more viscous than normal, and/or cell debris has formed a semi-solid barrier within the gradient preventing passage of nuclei. Thus it is important not to discard the supernatant unless you are confident that the nuclei are in the pellet. It is prudent to look at a drop of the supernatant under a microscope (see step 4) before discarding it. If the supernatant has several distinct layers, examine a drop from each layer before discarding that layer. If nuclei are found in the supernatant (no matter what the cause), the simplest solution is to add 5-10 ml of MPD buffer, vortex the mixture, and spin the tube at 650 x g for an additional 30 minutes. If the nuclei still have not pelleted, dilute the contents further and spin again (though, in our experience, this has never been necessary).

6. Decant the supernatant, and gently resuspend the pellet in 0.5 ml of MPDB. Examine a drop of the suspension under the microscope as described in step 4 (see FIGURE 7.1). Add 10 ml of MPDB, and transfer the nuclear suspension to a 15 ml polypropylene centrifuge tube.
7. Centrifuge at 300 x g for 10 min, and then turn the centrifuge up to a speed equivalent to 650 x g. Allow the tube to spin for an additional ten minutes. Discard the supernatant.
8. The resulting pellet may or may not contain two or more distinct layers. If there is a single layer, proceed to step 11. If not, do steps 9 and 10.
9. If more than one layer is present, add two ml of MPDB and gently vortex the tube until the upper-most layer is resuspended. Pour this suspension into its own 15 ml centrifuge tube leaving the rest of the pellet in the first tube.
10. Repeat step 9 until all of the original pellet layers are segregated into different tubes. Prepare a slide from the contents of each tube (see step 4). Based on the microscopy results, pick the tube(s)/layer(s) that contains the most nuclei and the least debris. Discard all other tubes.
11. Add 10 ml of MPDB to the nuclei and mix by gentle inversion. Centrifuge the suspension at 650 x g for 10 minutes, and carefully decant the supernatant.

­ Note 7.3: Highly pure nuclei do not form a very hard pellet and may slide out of the tube if the supernatant is discarded too forcefully.

12. Gently tap the tube so that the nuclei become resuspended in the residual MPDB left in the tube. Place the tube containing the nuclei on ice.
13. Proceed to II.

SUPPLIES, EQUIPMENT, AND REAGENTS (see CHAPTER 2 for details): liquid nitrogen; SEB; SEB^+BME; SEB^+BME/Triton; mortar and pestle; cheesecloth; Miracloth

METHODS:

1. Freeze 15-100 g of fresh plant tissue in liquid nitrogen or take 15-100 g of frozen tissue out of the freezer.
2. Place the frozen tissue in a mortar containing liquid nitrogen. Grind the tissue to a powder with the mortar and pestle.
3. Transfer the powder into ice-cold SEB^+BME in an appropriate container(s). For every gram of tissue, use approximately 10 ml of SEB^+BME (e.g., place 20 g of frozen tissue in 200 ml of ice-cold SEB^+BME).
4. Place the mixture on ice for 12 minutes. During this time, swirl the contents of the container every two minutes (each swirl time = 20 sec).

­ Note 7.4: Nuclei in sucrose-based buffers must be handled with extreme care. The absence of divalent cations coupled with the extreme osmotic conditions in the SEB^+BME make the nuclei extremely fragile. If roughly agitated, the nuclei will break.

5. Filter the homogenate through 2 layers of cheesecloth and 2 layers of Miracloth into a clean flask(s).
6. Add ¹/₂₀ volume of SEB^+BME/Triton to the flask(s), and leave it on ice for 10 min. During this time, swirl the contents of the flask every two minutes.
7. Transfer the mixture into several 50 ml polypropylene centrifuge tubes, and spin the tubes in a centrifuge at 650 x g for 15 min (4ºC).
8. Very gently decant the supernatants. Add 10 ml of SEB^+BME to each pellet, and gently resuspend nuclei with a small paintbrush. Consolidate the nuclear suspensions into as few 50 ml centrifuge tubes as possible. To each tube add SEB^+BME to 50 ml, and centrifuge tubes at 650 x g for 15 min (4ºC).
9. Repeat step 8 until all of the nuclear suspensions have been consolidated into a single tube.
10. Decant the supernatant, and resuspend the nuclei in 20 ml of SEB (without ß-mercaptoethanol). Centrifuge as described above, and very gently remove all but 1-2 ml of the supernatant with a large pipettor.
11. Gently resuspend the pellet in the residual SEB with a paintbrush. Examine a drop of the suspension by light microscopy (see step 4 in OPTION X).
12. Proceed to II.

SUPPLIES, EQUIPMENT, AND REAGENTS (see CHAPTER 2 for details): MPDB or SEB (depending on whether OPTION X or OPTION Y was used to isolate nuclei); Lysis Buffer; LMP agarose; plug molds

METHODS:

1. In a 50 ml flask, mix 0.15 g of LMP agarose with 10 ml of MPDB (if OPTION X was used to isolate nuclei) or 10 ml of SEB (if OPTION Y was used to isolate nuclei). Place the flask on a balance. Tare the balance. Transfer the flask into a microwave and heat the LMP agarose/buffer mixture until all of the LMP agarose has gone into solution (this may require some boiling). Place the flask back on the scale. Add MBG water to the flask until the scale reads "0.00 g". Place aluminum foil over the top of the flask, and place the flask in a 45°C water bath.
2. Place an agarose plug mold(s) on ice. If plug molds are being re-used, make sure that they are clean (thoroughly washed and rinsed with ethanol or washed and UV-sterilized) and that fresh tape has been placed on the bottom of the wells.
3. Place the tube containing nuclei in the 45°C water bath for 10 minutes.
4. Mix an equal volume of the LMP agarose solution with the pre-warmed nuclei. Using a plastic pipet with a relatively large-bore tip, place the nuclei/agarose mixture into the wells of the pre-chilled plug mold (FIGURE 7.2). Place the ice bucket containing the plug mold and plugs in a refrigerator. Allow plugs to solidify for 30 minutes.
5. Push plugs out of plug molds into 50 ml of Lysis Buffer. BioRad plug molds come with a small plastic tab designed for pushing plugs out of the molds (see FIGURE 7.2). However, a clean spatula will suffice. If the nuclear isolation was performed as described above, the plugs should be white to light yellow in color.
6. Incubate the plugs in Lysis Buffer at 50°C for 24 hours. Drain the Lysis Buffer from the plugs, and add 50 ml of fresh Lysis Buffer. Incubate the plugs at 50°C for an additional 24 hours.
7. Transfer plugs into 50 ml of fresh Lysis Buffer and store at 4°C overnight.

   ­ Note 7.5: The length of time plugs can be stored in Lysis Buffer at 4°C without significant DNA degradation seems to vary from species to species. For optimal results, store plugs in Lysis Buffer for no more than a few days. If a longer storage time is required, transfer plugs (or other agarose pieces containing DNA) to a 50 ml polypropylene tube containing aqueous 70% ethanol (20°C). Allow plugs to equilibrate in the 70% ethanol at room temperature for at least four hours. Place the tube containing the plugs at -20°C. Plugs can be stored in this fashion for at least 6 months (if not considerably longer) without noticeable DNA degradation. Before performing digestions, etc., on plugs that have been stored in 70% ethanol, transfer the plugs into 50 ml of an appropriate aqueous buffer at 4°C. Initially, the plugs will float on top of the buffer. However, once the ethanol has diffused out of the plugs they will sink to the bottom of the tube. At this point, the plugs are ready for further use.

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