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Detection of DNA Fragmentation and Endonucleases in Apoptosis
METHODS: A Companion to Methods in Enzymology 17, 329 –338 (1999) Article ID meth.1999.0747, available online at http://www.idealibrary.com on
Detection of DNA Fragmentation and Endonucleases in Apoptosis P. Roy Walker, Julie Leblanc, Brandon Smith, Siyaram Pandey and Marianna Sikorska Apoptosis Research Group, Institute for Biological Sciences, National Research Council of Canada, Ottawa, Ontario, Canada K1A 0R6
DNA degradation during apoptosis is endonuclease mediated and proceeds through an ordered series of stages commencing with the production of large DNA pieces of 300 kb which are then degraded to fragments of 50 kb. The 50-kb fragments are further degraded, in some but not all cells, to smaller pieces (10 – 40 kb) releasing the small oligonucleosome fragments that are detected as a characteristic DNA ladder on conventional agarose gels. Methodology is presented for the detection of both DNA ladders and the initial stages of DNA fragmentation using pulsed-field gel electrophoresis. We have developed electrophoresis conditions that resolve large fragments of DNA and also retain the smaller fragments on the same gel. Methods for the detection of endonuclease activities responsible for the cleavage of DNA during apoptosis are also presented.
The feature of apoptosis that most characteristically distinguishes it from other forms of cell death is the remarkable change that occurs in the structural organization of the nucleus. The destruction of the nucleus, through the degradation of both specific nuclear proteins and DNA, results in the collapse of chromatin into highly condensed electron-dense masses. This change in nuclear structure was originally observed by electron microscopy (1), but more recently a variety of fluorescence-based techniques have emerged to analyze the nuclear events by microscopy, flow cytometry, or biochemical techniques that detect the underlying DNA fragmentation either in situ or in vitro using agarose gel electrophoresis. These techniques serve as quantitative assays for apoptosis and are also useful for studying
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the actual mechanism of DNA fragmentation, which is still poorly understood. The first technique for the routine analysis of DNA fragmentation in vitro used conventional agarose gel electrophoresis (CAGE) in which a constant electric field at low voltage is used to resolve DNA fragments purified from apoptotic cells (2). This technique, which is straightforward and inexpensive and can resolve fragments up to a maximum of about 30 kb, is ideally suited to observe the ladder of fragments that is generated in many, but not all, apoptotic cells. Although the formation of the DNA ladder has been the hallmark biochemical indicator of apoptosis, it is, in fact, only the end point of DNA degradation and does not reflect the full pattern of DNA fragmentation occurring during apoptosis. Indeed, many cells do not degrade their DNA to this extent. We have recently shown DNA degradation during apoptosis to be a much more complex process that commences with the generation of higher molecular weight (HMW) DNA fragments which are undetectable by CAGE (3–5). These larger fragments are 50 –300 kb in size and reflect endonucleolytic cleavage of interphase chromosomes at the nuclease-sensitive sites that reside in chromatin fibers as a result of their folding into loop (mean size of 50 kb) and rosette (mean size of 300 kb) structures, respectively (3). This suggests that interphase chromosome breakdown in apoptosis is a highly ordered process and is intimately related to the higher order structure of chromatin. However, CAGE cannot resolve fragments greater than approximately 30 kb. Larger fragments either remain in the well or migrate 329
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slowly into the gel as a single band that is an unresolved zone of compression. Pulsed-field gel electrophoresis (PFGE) is a specialized form of agarose gel electrophoresis that permits the ordered separation of larger fragments of DNA (6). The simplest form of PFGE is fieldinversion gel electrophoresis (FIGE) in which the current is periodically reversed between two electrodes oriented 180° with respect to each other. This permits the use of the same gel boxes that are used for CAGE. Since the theory of migration of large DNA molecules during PFGE is not well understood, the frequency at which the current is reversed and the duration of the periods of forward and reverse polarity have been worked out empirically for particular size classes of DNA. We have adapted a form of FIGE called zero-integrated field electrophoresis (ZIFE) to optimize the separation of DNA fragments produced in apoptotic cells. Under appropriate conditions, we can easily extend the range of fragment sizes that can be studied to 1 Mb without losing any of the small fragments from the gel. Since larger DNA molecules are very sensitive to mechanical shear and endonucleolytic attack, radically different sample handling techniques must be used to prepare DNA for PFGE. Another characteristic feature of the degradation of DNA during apoptosis is the appearance of singlestrand breaks in the DNA. Indeed, it is possible that the endonuclease(s) responsible for DNA fragmentation introduces a multitude of single-stand nicks into the DNA, until eventually nicks accumulate sufficiently close together on opposite strands to generate a de facto double-strand break (7–9). We have developed a unique methodology to study the appearance of single-strand nicks in apoptotic DNA. Finally, since the degradation of DNA in apoptosis is endonuclease mediated and there is an intense effort to identify and characterize these proteins (8 –11) we provide methodology for their detection and characterization.
TREATMENT OF CELLS General considerations. The processing of cells or tissues is very dependent on the DNA analysis that is to follow. If only small fragments are to be extracted for “ladder” analysis by CAGE they can easily be solubilized from any cultured cells or tissue with detergent. Cells and tissue must be processed
much more carefully for PFGE analysis of HMW DNA, since a prerequisite for the analysis of high molecular weight DNA is the elimination of mechanical or enzymatic damage to the DNA during the processing of cells or nuclei. We have much more experience working with suspension cultures or thymocyte primary cultures than with either monolayers or tissue (12). A method for the processing of tissue is described by MacManus et al. (13). Processing of cells, tissues, and nuclei. Following the incubation of cells with apoptotic inducers, suspension or primary cultures are easily pelleted from the medium. Monolayer cells are harvested by trypsin treatment [0.15% trypsin in phosphate-buffered saline containing 1mM EDTA] to generate a cell suspension. The suspensions are pelleted and resuspended in nuclear buffer (NB, 15 mM Tris–HCl, pH 7.4, 60 mM KCl, 15 mM NaCl, 1 mM EGTA, 2 mM EDTA, 0.5 mM spermidine, and 0.15 mM spermine). This buffer has a relatively low ionic strength to assist in disruption of the cell and contains EDTA and EGTA to chelate divalent cations, as well as polyamines to bind to chromatin and protect the DNA. It is very difficult to extract DNA from tissue sufficiently quickly to completely prevent endonucleolysis. The best strategy is to isolate nuclei using buffers that are designed to inhibit endonuclease activity and protect chromatin. To achieve this, cultured cells or tissues are homogenized in a buffer consisting of ice-cold 0.25 M sucrose, 15 mM Tris– HCl, pH 7.4, 60 mM KCl, 15 mN NaCl, 2 mM EDTA, 0.5 mM EGTA, 15 mM mercaptoethanol, 0.5 mM spermidine, and 0.15 mM spermine. The nuclei are pelleted at 700g for 10 min at 4°C, resuspended in the same buffer containing 1% Triton X-100 to remove the outer nuclear membrane, pelleted again, and resuspended in an appropriate buffer for the experiment. For PFGE, washed cells or nuclei are usually embedded in agarose to prevent any further degradation. However, to completely eliminate artifactual DNA fragmentation, it is possible, in some circumstances, to carry out entire experiments on embedded cells, since media constituents and inducers of apoptosis can easily diffuse into the agarose plug. To do this, approximately 3 3 107 cells are suspended in 250 ml of medium containing 5% serum and mixed with 250 ml of molten 1.5% (w/v) low-melting-point agarose (prepared in medium without serum and held at 37°C, as indicated below). It is paramount
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not to heat shock the cells by allowing the temperature to rise above 37°C. The mixture is pipetted into 1-ml disposable syringes and allowed to solidify. The plug containing the embedded cells is ejected and incubated in RPMI medium containing 5% fetal bovine serum plus or minus inducers of apoptosis. After the incubation, the plug is processed, as described below, for either CAGE or PFGE analysis of DNA fragmentation. In some experiments we have embedded live cells as described above and then permeabilized them to allow the effects of cations on DNA fragmentation in situ to be studied. To do this the embedded cells are incubated at 37°C in 300 ml of permeabilization buffer (10 mM Tris–HCl, pH 7.0, 60 mM KCl, 0.1 mM EGTA, and 0.5% Triton X-100). Cations can be added and incubations can be carried out at 37°C for various times.
DETECTION OF DNA LADDERS BY CAGE Sample preparation. In this technique cells or nuclei are lysed with detergent and the small, detergent-soluble fragments of DNA are purified using phenol:chloroform. The entire procedure can be carried out in microfuge tubes. The purification steps remove proteins that could retard the migration of DNA in the gel and also RNA which can comigrate with smaller DNA fragments. Approximately 1 3 106 cells, or the equivalent amount of tissue (;10 mg), are resuspended in 400 ml of icecold extraction buffer containing 5 mM Tris–HCl, pH 7.5, 5 mM EDTA, and 0.5% (v/v) Triton X-100. The suspension is centrifuged at 14,000g for 20 min at 4°C in a microfuge, the supernatant is collected, and the DNA is precipitated from the supernatant with 40 ml of 2.5M sodium acetate (pH 5.2) and 2 vol of absolute ethanol at 220°C (usually overnight for convenience, but 2 h is sufficient). The precipitated DNA is collected by centrifugation, resuspended in 400 ml of TE buffer (10 mM Tris–HCl, pH 7.5 1 1 mM EDTA) and treated with RNase A (20 –100 mg) for 60 min at 65°C. Twenty-five microliters (0.5mg) of proteinase K solution (20 mg/ml stock in distilled H2O) is then added along with 40 ml of 103 proteinase K buffer (100 mM Tris–HCl, pH 7.8 1 50 mM EDTA and 5% N-laurylsarcosine) and the incubation continued for 2 h at 37°C. The DNA is then phenol:chloroform extracted by adding 400 ml of phenol:chloroform [saturated phenol mixed 1:1 (v/v)
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with chloroform:isoamyl alcohol, 24:1 (v/v)], gently mixing, and spinning in a microfuge for 5 min. The top layer is transferred to a clean tube and extracted once more. This layer is then precipitated with 40 ml of 2.5 M sodium acetate (pH 5.2) and 1 ml of absolute ethanol at 220°C overnight (or 1 h at 280°C). After precipitation, the samples are spun for 30 min at 4°C in a microfuge and the supernatant is discarded. The pellets are washed with 70% ethanol and centrifuged in a microfuge for 5 min. This wash step is repeated and the pellets are then dried on the bench or in a Speed Vac (Fisher Scientific, Ottawa, Ontario, Canada). The precipitated DNA is dissolved in TE buffer. Preparation of gel. We use 0.8% (w/v) gels for both CAGE and PFGE made with electrophoresis grade agarose from Gibco-BRL (Burlington, Ontario, Canada). The agarose (2.4 g) is suspended in 300 ml of TAE buffer (0.04M Tris–acetate, pH 8.5, containing 2 mM EDTA) and heated for 2 min in a microwave oven, removed, stirred briefly, and heated again for 2 min. The molten mixture is removed from the oven and stirred on a magnetic stirrer until it cools to about 60°C. The agarose is then poured into the gel tray and allowed to cool. When the gel has solidified (in approximately 30 min), it is put in the refrigerator or cold room for 5–10 min to be sure it is hard before removing the comb. If the gel is poured at too high a temperature moisture will condense onto the surface and cause local dilutions of the agarose before it solidifies. Similarly, if the comb is removed too soon moisture will condense in the wells and distort their shape. Electrophoresis. A wide variety of gel boxes, designed for horizontal, submerged gel electrophoresis, including many minisystems, can be used. A basic 200-V power supply is sufficient. The extracted DNA (approximately 5–10 mg) is dissolved in 50 ml of TE buffer, followed by 5 ml of loading solution (30% (w/v) sucrose 1 0.01% (w/v) bromphenol blue in distilled H2O). The sample is then added to each well. We routinely use two size markers for CAGE: the 123-bp ladder and a HindIII digest of l DNA, both from Gibco-BRL. The running buffer is TAE. For regular sized (20 3 25 cm) gels the run is carried out at 30 V for 18 h (i.e., overnight), whereas for minisystems we use 120 V for 1 h. The gel is removed from the tank, still attached to the gel tray, stained, and imaged as described below.
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DETECTION OF BOTH DNA LADDERS AND HMW DNA FRAGMENTS USING PFGE As described earlier, DNA fragments greater than 30 kb cannot be resolved by CAGE. Therefore, PFGE in some form must be used to resolve the larger fragments also produced during DNA degradation in apoptosis. Although fragments as large as 5 Mb can be resolved by PFGE, a disadvantage is that in any given run usually only a fairly narrow size range of fragments can be resolved. Furthermore, in most protocols, the smaller, ladder-sized DNA fragments run off the gel so two gels must be run for each sample—CAGE to detect small fragments and PFGE to detect HMW fragments. To obviate these problems we have devised PFGE conditions that allow fragments from 100 bp to 1 Mb to be resolved on the same gel. This size range spans the sizes of DNA typically seen in apoptotic cells, so it is possible to see the entire spectrum of DNA fragments in a single run. Equipment. There are a number of commercially available PFGE units varying tremendously in capability and cost. For routine detection of low and HMW DNA fragmentation in apoptosis we use the inexpensive FIGE mapper equipment sold by BioRad Laboratories (Mississauga, Ontario, Canada). This unit is programmable with user-defined parameters permitting the use of non-FIGE conditions, such as ZIFE. Sample preparation. Sample preparation is the most critical step in studies on HMW DNA degradation using PFGE. Activation of nucleases often accompanies cell disruption and long DNA fragments are also extremely susceptible to mechanical shear. Therefore, any technique for the analysis of high molecular weight DNA fragmentation in apoptosis is critically dependent on methodology to isolate DNA without further artifactual damage. The technique of embedding cells in agarose plugs evolved to solve this problem. To eliminate mechanical stresses, the whole cell or nucleus is usually embedded in an agarose plug prior to any manipulations on DNA. The proteins, including endonucleases, are then removed enzymatically rather than with solvents. The presence of the denaturing detergent sodium dodecyl sulfate (SDS) and the absence of divalent cations during the incubation with proteinase K ensures the integrity of the DNA. Endogenous RNases are active under these conditions and RNA is usually de-
graded. If necessary, a RNase step can be included at this stage or performed after the gel is run (see below). If cell extracts are to be stored before being processed for PFGE they should be stored at 280°C since DNA degradation occurs during storage at 220°C. Preparation of plugs for PFGE. We use one of two methods for embedding cells in agarose, depending on the number of cells or nuclei available. Usually, DNA from 1–2 3 106 cells must be loaded onto the gel in order to easily detect the DNA by ethidium bromide staining, although DNA from as few as 0.5 3 106 cells or nuclei can still be detected using SYBR gold from Molecular Probes (Eugene, OR). If only 2 3 106 cells, or less, are available then all the cells are encapsulated into a miniplug which is loaded in its entirety onto the gel. If more cells are available, then a larger plug can be made and slices, containing the equivalent of 1–2 3 106 cells or nuclei, can be cut and loaded onto the gel. Preparation of miniplugs. First, low-meltingpoint (LMP) agarose (Gibco-BRL) is weighed into a scintillation vial and an appropriate volume of NB to yield a 1.5% agarose mixture is added. The vial is put into the microwave oven for approximately 20 s, i.e., until it just boils, but does not boil over. Fortymicroliter aliquots are pipetted into Eppendorf tubes arrayed in a heating block (Fisher Scientific) followed by 1 ml of proteinase K (20 mg/ml stock solution in distilled H2O) and the mixture is kept molten at 37°C until ready to use. The sample of cells (or nuclei) is washed in NB, pelleted again, and resuspended in 40 ml of NB. This suspension is mixed with a 40-ml aliquot of the molten agarose–proteinase K mixture, pipetted into the wells of a miniplug casting mold (Bio-Rad), and allowed to set for 10 min at 4°C. When the agarose is set, the plug is ejected into a 1.5-ml microfuge tube containing 300 ml of TEEN buffer (10 mM Tris–HCl, pH 9.5, 25 mM EDTA, 5 mM EGTA, 10 mM NaCl), 15 ml of 10% (w/v in distilled H2O) SDS, and 2 ml of 20 mg/ml proteinase K. The samples are incubated for 1.5 h at 37°C (we use a rotator in an incubator). The objective is to remove protein in as short an incubation time as possible to minimize any further DNA degradation or diffusion from the plug. Complete deproteinization is essential in order to obtain uniform lanes when the gel is run. It is important to realize that proteinase K can often be contaminated with nucleases. We have found the proteinase K
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obtained from Gibco-BRL to be consistently reliable. If nuclease contamination is suspected a plasmid digestion assay (see below) can be carried out to detect any contaminants and the batch can be discarded if necessary. Following the incubation, the supernatant is removed and, if desired, any DNA that may have diffused from the plug can be precipitated from the supernatant at 220°C by adding 0.1 vol of 2.5 M sodium acetate and 2 vol of absolute ethanol. The DNA is washed, redissolved, and run on CAGE. For short deproteinization times, this is not usually necessary. The deproteinized plug is washed in 1 ml of TE at 4°C for 30 min. This step is repeated once and the deproteinized plugs are stored at 4°C in TE buffer until ready to load on a gel. If storing for longer than overnight, remove the TE buffer and store the plug in a sealed microfuge tube.
plugged with Parafilm and allow to set for 20 –30 min at 4°C. Cut the tip off the syringe to leave a blunt end flush with the end of the barrel to permit the solidified plug to be ejected. Eject the plug into a 15-ml plastic tube containing 3 ml of TEEN buffer, 150 ml of 10% (w/v) SDS, and 20 ml of 20 mg/ml proteinase K. The incubation is carried out for 3–16 h at 37°C, as described above, but should be kept to the minimum time required to remove all of the protein. Remove the plug and wash with 10 ml of TE at 4°C for 30 min. Repeat the washing step once and gently suck the plug back into the syringe, cover the end with Parafilm, and store at 4°C until ready to load onto the gel. To load, cut 20- to 30-ml slices, containing 1–2 3 106 cells or nuclei, from the end of the plug and place in the well. The wells are sealed with molten 0.8% (w/v) LMP agarose prepared in 0.53 TBE (45 mM Tris– borate 1 1.25 mM EDTA, pH 8–8.5).
Preparation of larger plugs. Harvest the cells as described above and pellet 20–30 3 106 cells. The plug volume is varied, depending on the number of cells, such that a 30-ml slice (a volume that conveniently fits into a well) gives 1–2 3 106 cells or nuclei. Wash the cells once with 1 ml of NB, resuspend in 250 ml of NB, and add 250 ml of molten 1.5% (w/v) LMP agarose in NB containing 0.1 mg of proteinase K (5 ml of 20mg/ml stock). Pipette the mixture into the barrel of a 1-ml disposable plastic syringe in which the tip has been
PFGE conditions. Gels (0.8%, w/v) are prepared in 0.53 TBE, as described earlier for CAGE. Four sets of standards, with overlapping size ranges, are used: yeast chromosomes (225 kb–1.2 Mb); polymerized l phage DNA (50 –1 Mb), low range markers (0.1–194 kb), all purchased from New England Biolabs (Beverly, MA); and the 123-bp ladder from Gibco-BRL. The quality of markers, particularly the yeast markers, and the extent of polymerization of the l ladder varies considerably. New England Bio-
FIG. 1. Separation of DNA from apoptotic cells by PFGE. (A) EL4 cells were incubated with (lanes 4 and 5) or without (lanes 2 and 3) 10 mM VM26. The cells in lanes 3 and 5 were preincubated for 30 min with 100 mM VAD-fmk, an inhibitor of caspases. Lane 1, control, unincubated cells. Lanes a– e, DNA size markers; 123-bp ladder, low-range PFGE markers (0.1–194 kb); polymerized l phage DNA (50 –1 Mb) and yeast chromosomes (225 kb–1.2 Mb), respectively. The sizes of a selection of these markers are indicated at the right of the gel. (B) Standard curve derived from the positions of the markers shown in lanes a– e of (A), showing ordered separation of the fragments.
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labs consistently produces the most reliable product. Electrophoresis is carried out using a horizontal gel chamber and a FIGE Mapper system (Bio-Rad Laboratories). The buffer, 0.53 TBE prechilled to 14°C, is circulated through a Lauda bath to maintain this temperature. The conventional 13 TBE is not used because the higher ionic strength causes overheating. Electrophoresis is carried out using the following settings: 280 V in the forward direction and 90 V in the reverse direction with an initial switch time of 0.3 s forward and 0.5 s reverse, with nonlinear ramping to 35 s forward and 60 s reverse, for a total run time of 20.5 h. An example of a typical PFGE separation is shown in Fig. 1. In Fig. 1A, the cells were incubated with an apoptotic inducer which results in the degradation of DNA to produce a ladder of small fragments (lane 4). If the cells are also incubated with an inhibitor of caspase activity, then DNA fragmentation is restricted to HMW fragments only (lane 5). Note that the control cells have completely intact DNA which all remains in the well (lane 1). Lanes a– e are standards and were used to produce the standard curve shown in Fig. 1B. This curve shows that, although not linear, there is an ordered separation of fragments over 4 orders of magnitude of sizes (100 bp to 1 Mb).
ANALYSIS OF DNA FRAGMENTATION BY 2D ELECTROPHORESIS Since the DNA strands are not denatured during sample processing in any of the above procedures,
the fragments that are resolved by either CAGE or PFGE are double-stranded molecules. However, these double-stranded molecules may harbor singlestrand breaks along the length of the duplex (7). The presence of these nicks can be detected by denaturing the duplex with sodium hydroxide to release the smaller single-stranded fragments. To study the extent of single-strand cleavage of DNA during apoptosis we developed a two-dimensional separation technique which uses nondenaturing PFGE in the first dimension and denaturing conditions in the second dimension. Conditions for 2D separations. In experiments involving 2D separations, the first dimension PFGE is performed exactly as described above, except that the run is shortened to 19 h from the usual 20.5 h. Only two samples are loaded in order to leave room for the second dimension (Fig. 2A). The gel is stained with ethidium bromide and imaged as described below. SYBR gold cannot be used at this stage since it binds to and severely retards the migration of DNA in the second dimension. The gel is rotated in the gel tray for the second-dimension run and the bottom of the gel is sliced off, if necessary, to fit. The second dimension run is either CAGE, run as described above, or denaturing electrophoresis. For the denaturing run, the gel is soaked in 50 mM NaOH 1 1 mM EDTA for 60 min at 4°C and then run in the same buffer. Both second-dimension runs are carried out at 150 V for either 2.5 h (denaturing conditions) or 2 h (CAGE) with buffer recirculation and cooling to 2°C (the temperature set at the Lauda bath). After the denaturing run, the gel is rinsed
FIG. 2. 2D PFGE-denaturing gel analysis of DNA from apoptotic Jurkat cells. (A) DNA from embedded control (lane 1) and apoptotic (lane 2) cells was resolved by PFGE. Lane a is the polymerized lambda ladder. (B) The gel was rotated and subjected to a second dimension of denaturing electrophoresis. Only the separation for lane 2 is shown. For reference, lane 2 from (A) is rotated and shown on the top of the second-dimension gel.
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twice in distilled H 2 O and then stained with ethidium bromide for 1 h and imaged, as described below. SYBR gold cannot be used for staining denaturing gels. A typical 2D analytical experiment is shown in Fig. 2. Once again the control cells (lane 1) have intact DNA, whereas the apoptotic cells produce primarily fragments in the 10- to 50-kb range with only a small amount of small fragments (lane 2). Lane 2 (i.e., the first dimension of PFGE) is rotated and shown again at the top of Fig. 2B. Underneath is the result of the second-dimension denaturing run. It can be clearly seen that a considerable number of single-strand DNA molecules lie underneath the “curve” of the still intact fragments. This is particularly evident for the 50-kb band which is seen to contain numerous single-strand breaks resulting in the release of many smaller fragments. Digestion of embedded DNA with nucleases. The presence of single-strand nicks and other modifications to DNA from apoptotic cells can be detected enzymatically. Embedded DNA is deproteinized as before and incubated with S1 nuclease (Sigma), mung bean nuclease (Pharmacia), or Bal31 (GibcoBRL). For the S1 nuclease digestions, miniplugs or slices of larger plugs are incubated in a buffer containing 30 mM acetate buffer, pH 4.6, 100 mM NaCl, 0.5 mM zinc chloride, and 450 U of enzyme. Mung bean nuclease digestions are carried out in 30 mM acetate buffer, pH 4.6, containing 50 mM NaCl, 1 mM zinc chloride, 0.01% Triton X-100, and 80 U of enzyme. Bal31 digestions are carried out in 20 mM Tris–HCl, pH 8.0, 200 mM NaCl, 12 mM CaCl2, 12 mM MgCl2, and 5 U of enzyme. All the incubations are typically carried out for 60 min at room temperature and the plugs are subjected to electrophoresis.
VISUALIZATION OF DNA FRAGMENTS AND IMAGE ANALYSIS Gel documentation. There are a number of possibilities for gel documentation and image analysis. The first consideration is the choice of fluorochrome for staining. Routinely, we use ethidium bromide which is inexpensive, easily visualized using a standard transilluminator, and compatible with both Polaroid film and CCD cameras. SYBR gold (Molecular Probes) is more sensitive, particularly when used in conjunction with Polaroid 667 film. However, this
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stain cannot be used in all circumstances (see below). A standard transilluminator with 300-nm lamps is used (such as the one from Ultra-Lum, Inc., Paramount, CA). Although Polaroid 667 film is the most sensitive, acceptable results are also obtained with type 55 positive/negative film for most applications. The image can also be acquired using a CCD camera coupled to a thermal printer and/or a data acquisition board and computer. A variety of software is available for image capture and data analysis. We use an inexpensive Panasonic Model BP312 CCD camera and image capture package from UltraLum. For both Polaroid cameras and CCD cameras it is important to use the most suitable filter for the dye being used. Filters for polaroid film are available from Molecular Probes. For the CCD camera we use a 600 6 25-nm bandpass filter for ethidium bromide and a 560 6 35-nm bandpass filter for SYBR gold (this filter also gives satisfactory results with ethidium bromide for most purposes). The latter filters can be obtained from Melles Groit (Ottawa, Ontario, Canada) in a variety of lens mounts. Following either CAGE or PFGE, the gel is placed in a glass or plastic container and submerged in ethidium bromide (1.5 mg/ml) or in SYBR gold (a 10,000-fold dilution of the stock supplied by the manufacturer in 0.53 TBE). Ethidium bromide needs destaining for at least 30 min, whereas SYBR gold requires no destaining. The gel is placed on the UV transilluminator and either photographed or the image is captured using the CCD imaging system. If there is a lot of nonspecific fluorescence the gels can be soaked in water for several more hours and then photographed again. To minimize the number of manipulations prior to electrophoresis we usually remove RNA after the run by soaking the gel in running buffer containing 1 mg/ml of RNase A for 2 h at room temperature or overnight at 4°C prior to staining and image capture. Data analysis. CCD cameras produce a digitized image of the gel which can subsequently be used to generate scans of pixel intensity vs distance of migration for each lane. A number of software packages (e.g., Sigmagel and Sigmascan from SPSS, Chicago, IL) are available for analysis. Standard curves are generated by regression analysis of the positions of overlapping DNA size markers. The sizes and total amount of fragmented DNA in each sample can then be calculated.
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METHODS FOR DETECTION OF ENDONUCLEASE ACTIVITY Endonuclease activities can be detected by one of two methods. The simplest way is to incubate samples with an intact double-stranded DNA plasmid and use CAGE to assess its integrity. Alternatively, the extracts can be resolved by SDS–PAGE (denaturing polyacrylamide gel electrophoresis) using gels copolymerized with substrate DNA and endonuclease activity bands can be detected after renaturation and digestion of the DNA. The latter technique can be performed using 1D or 2D protein separations and has the advantage of resolving multiple endonucleases in a mixture. In addition, their sizes, isoelectric points, and posttranslational modifications can be determined. As such, it is the tool for the discovery of new endonucleases. Plasmid DNA digestion assay. The protein sample (5–10 mg of total protein) is resuspended in 25 ml of 50 mM Tris–HCl buffer, pH 7.4, 2 mM MgCl2, and 0.2 mM CaCl2 and 1 mg of plasmid DNA is added. Any intact plasmid in the 5- to 20-kb size range can be used. The reaction mixture is incubated for 0.5–2 h at 37°C and the reaction can be stopped by the addition of 4 ml of 103 DNA loading buffer (40% w/v sucrose, 0.01% bromphenol blue, 50 mM Tris–HCl, pH 8.0, and 5 mM EDTA). The integrity of the plasmid DNA is analyzed by loading the reaction mixture on a 0.8% agarose gel. Electrophoresis is carried in TAE at 30 V overnight or 150 V for 2 h, as described above for CAGE. The gel is stained with ethidium bromide and imaged on a UV transilluminator. Usually there are three bands in the undigested plasmid DNA, corresponding to supercoiled (fastest moving band), circular (nicked), and linear forms. Samples containing a low level of endonuclease activity may produce a nick in the supercoiled or circular plasmid resulting in the disappearance of the lower band and appearance of the slower moving relaxed or linear forms. Higher activity samples digest the plasmid completely producing a smear of DNA near the bottom of the gel. It is possible to use this assay to characterize the cation requirements, the pH optima, and the effects of inhibitors. In addition, it is possible to study the nature of the ends of the fragments by attempting to radiolabel them as described below.
Endonuclease activity gels. Endonuclease activities present in cell or nuclear extracts can be identified by SDS–PAGE using a gel in which the DNA substrate is embedded (10, 11, 14, 15). We use SDS– 10% polyacrylamide gels copolymerized with 30 mg/ml of salmon testis DNA (Sigma Chemical Co., St. Louis, MO). The DNA can be either double stranded or heat denatured single stranded. To prepare the DNA, dissolve salmon sperm DNA (Sigma Chemical Co.) in distilled H2O at a concentration of 3 mg/ml. Sonicate the solution for 20 s to produce sheared double-stranded fragments. If required, denature the DNA in a boiling water bath for 5 min and transfer immediately to an ice bath to obtain single-stranded DNA. Use 100 ml of DNA solution (single or double stranded) per 10 ml of acrylamide gel mix. 32P-labeled DNA (106 cpm/ml) can also be added to the DNA solution. A total of 0.2– 0.5 mg of DNA can be radioactively labeled using [a-32P]dATP and [ a - 32 P]dGTP (800 Ci/mmol, NEN DuPont, Oakville, Ontario, Canada) and the multiprime DNA labeling system (Amersham, Oakville, Ontario, Canada). The resolving and stacking gels are then poured using standard procedures. Protein samples are dissolved in a loading buffer of 62.5 mM Tris–HCl buffer, pH 6.7, containing 1% (w/v) SDS, 10% (v/v) glycerol, and 0.01% bromophenol blue, usually prepared as a 43 stock. The samples are not boiled prior to loading. For reducing conditions, 2-mercaptoethanol is added to a final concentration of 1% (v/v). It is important to run the gels, at least initially, under both reducing and nonreducing conditions since some endonucleases are inactive under standard reducing conditions. The samples are separated on SDS–PAGE and the location of the active bands is determined by subsequent reactivation of the endonuclease(s) which remove the embedded DNA from the gel at that location. This is achieved during a protein renaturation step [30 min at room temperature in 50 mM Tris– HCl buffer, pH 7.5, 5 mM MgCl2, and 20% (v/v) 2-propanol with gentle rocking and two changes of the buffer], followed by an overnight reactivation step in 50 mM Tris–HCl buffer, pH 7.5, 5 mM MgCl2, 1 mM CaCl2, and 0.1% Triton X-100 (v/v) at 37°C. Nonradioactive gels are stained in 1 m g/ml ethidium bromide and imaged as before. the ethidium bromide-stained gels show a dark band (no fluorescence) where a nuclease has digested away the DNA in that part of the lane. The gels containing radioactively labeled DNA are dried and exposed to
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Kodak X-Omat film. The autoradiograph from radioactive gels also shows a clear band where a nuclease is present. When searching for new endonuclease activities it is essential to use substrate gels with radioactively labeled DNA since there is a potential artifact in the method that uses unlabeled DNA. On unlabeled gels, DNA-binding proteins, such as histones, can mask the DNA from the ethidium stain giving a “band” on the gel that is indistinguishable from the one produced by an endonuclease. This phenomenon does not occur on autoradiographs, since radioactivity can only be lost due to endonucleolytic degradation of the DNA. DNA–substrate gels also provide a convenient way to establish cation requirements and pH profiles. Equal amounts of the same protein sample are loaded in each lane and separated as described above. After the run, individual lanes are cut separately and subjected to renaturation and activation steps in solutions containing different compositions of cations, cation chelators, or solutions with identical cations but different pH. Two-dimensional activity gel electrophoresis. 2D acrylamide gel electrophoresis can be used to better resolve endonucleases. 2D gels also indicate the isoelectric point of a protein and give some indication to the extent of posttranslational modifications. The first dimension of isoelectrofocusing (IEF) is performed using a Multiphar electrophoresis unit (Pharmacia Biotech, Baie d’Urfe, Quebec, Canada) using the experimental conditions suggested by the manufacturer. Samples of protein extract are mixed with 240 ml of IEF loading buffer (8 M urea, 2% Triton X-100 (v/v), 2% Pharmalytes pH range 3/10) and absorbed onto Immobilin IEF strips, pH 3/10, for 8 h. Reducing agents are omitted if there is reason to believe a redox-sensitive nuclease exists. The proteins are subjected to overnight IEF at 15°C for 20,000 Vh. After the IEF run is complete, the Immobilin strips are loaded onto SDS–10% polyacrylamide gels copolymerized with 30 mg/ml of sheared salmon testis DNA (single stranded, double stranded, and/or radiolabeled) and the proteins are separated in the second dimension. Following the run the endonucleolytic activity is reactivated and visualized as described above. Labeling of 39-OH and 59-OH ends of DNA. DNA fragments can be radioactively labeled at either the 39-OH or 59-OH end using terminal deoxynucleotidyltransferase (TdT) or T4 polynucleotide kinase, re-
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spectively. Since the presence of a phosphate group blocks both of these reactions, this approach can be used to determine the nature of the ends of the DNA fragments in apoptotic DNA, e.g., 59-P/39-OH or 59OH/39-OH or 59-OH/39-P. This has important implications for the mechanism of action of nucleases involved in DNA fragmentation in apoptosis. To assess this, divide the reaction mixture in two tubes, one for TdT and other for the T4 kinase labeling reaction. The TdT labeling reaction is carried out for 1 h at 37°C in 50 ml of reaction buffer containing 25 mM Tris–HCl buffer, pH 6.6, 200 mM potassium cacodylate, 5 mM CoCl2, 0.5 mM dithiothreitol (DTT), 0.25 mg/ml bovine serum albumin, 5 mCi of [a-32P]dATP (NEN DuPont and 60 units of TdT (Gibco-BRL). The T4 kinase labeling is also performed at 37°C for 1 h in 50 ml of reaction buffer containing 50 mM glycine–NaOH buffer, pH 9.2, 5 mM DTT, 10 mM MgCl2, 10 mM Na2HPO4, pH 9.2, 5 mCi[a-32P]ATP, and 50 units of T4 kinase (New England Biolabs). Unincorporated radionucleodides are removed using G-50 Micro columns (Pharmacia Biotech) and labeled DNA is separated by CAGE followed by autoradiography.
CONCLUDING REMARKS There is still a lot to be learned about the mechanism of chromatin degradation in apoptosis and the nature of the endonuclease(s) that catalyze it. To date more than 40 endonuclease activities have been implicated in the process, including both endogenous (i.e., nuclear) and exogenous (extranuclear) activities (8, 9). It is highly likely that more than one activity is required to completely degrade chromatin given that several million single- and double-strand breaks are introduced into chromatin within a matter of minutes once degradation commences. The pattern of fragments generated during this process is primarily a property of the substrate, chromatin. The enormous complexity of chromatin dictates the localization of nuclease-sensitive sites and we can learn about its structural organization from the fragmentation pattern. The challenge is to further develop the technology for analyzing DNA fragmentation, particularly at its earliest stages. Continued improvements in our ability to analyze large DNA fragments by PFGE or 2D electrophoresis will contribute greatly to this effort.
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REFERENCES 1. Kerr, J. F. R., Wyllie, A. H., and Currie, A. R. (1972) Br. J. Cancer 26, 239 –257. 2. Wyllie, A. H. (1980). Nature 284, 555–556. 3. Filipski, J., Leblanc, J., Youdale, T., Sikorska, M., and Walker, P. R. (1990) EMBO J. 9, 1319 –1327. 4. Walker, P. R., Smith, C., Youdale, T., Leblanc, J., Whitfield, J. F., and Sikorska, M. (1991) Cancer Res. 51, 1078 –1085. 5. Oberhammer, F., Wilson, J. W., Dive, C., Morris, I. D., Hickman, J. A., Wakeling, A., Walker, P. R., and Sikorska, M. (1993) EMBO J. 12, 3679 –3684. 6. Schwartz, D. C., and Cantor, C. R. (1984) Cell 37, 67–74. 7. Walker, P. R., LeBlanc, J., and Sikorska, M. (1997) Cell Death Differ. 4, 506 –515. 8. Walker, P. R., and Sikorska, M. (1998) Biochem. Cell. Biol. 75, 287–299.
9. Sikorksa, M., and Walker, P. R. (1998) in When Cells Die: A Comprehensive Evaluation of Apoptosis and Programmed Cell Death (Lockshin, R. A., Zakeri, Z., and Tilly, J. L., Eds.), pp. 211–242, Wiley–Liss, New York. 10. Pandey, S., Walker, P. R., and Sikorska, M. (1997) Biochemistry 36, 711–720. 11. Liu, Q. Y., Pandey, S., Singh, R. K., Lin, W., Ribecco, M., Borowy-Borowski, H., Smith, B., Leblanc, J., Walker, P. R., and Sikorska, M. (1998) Biochemistry 37, 10134 –10143. 12. Walker, P. R., Kokileva, L., Leblanc, J., and Sikorska, M. (1993) Biotechniques 15, 1032–1041. 13. MacManus, J. P., Rasquinha, I., Tuor, U., and Preston, E. (1997) J. Cereb. Blood Flow Metab. 17, 376 –387. 14. Seiliev, A. A., Zvonareva, N. B., Zhivotovsky, B. D., and Hanson, K. P. (1992) Radiat. Environ. Biophys. 31, 123– 128. 15. Heeb, J. M., and Gabriel, O. (1984) Methods Enzymol. 104, 416 – 422.
Detection of DNA Fragmentation and Endonucleases in Apoptosis P. Roy Walker, Julie Leblanc, Brandon Smith, Siyaram Pandey and Marianna Sikorska Apoptosis Research Group, Institute for Biological Sciences, National Research Council of Canada, Ottawa, Ontario, Canada K1A 0R6
DNA degradation during apoptosis is endonuclease mediated and proceeds through an ordered series of stages commencing with the production of large DNA pieces of 300 kb which are then degraded to fragments of 50 kb. The 50-kb fragments are further degraded, in some but not all cells, to smaller pieces (10 – 40 kb) releasing the small oligonucleosome fragments that are detected as a characteristic DNA ladder on conventional agarose gels. Methodology is presented for the detection of both DNA ladders and the initial stages of DNA fragmentation using pulsed-field gel electrophoresis. We have developed electrophoresis conditions that resolve large fragments of DNA and also retain the smaller fragments on the same gel. Methods for the detection of endonuclease activities responsible for the cleavage of DNA during apoptosis are also presented.
The feature of apoptosis that most characteristically distinguishes it from other forms of cell death is the remarkable change that occurs in the structural organization of the nucleus. The destruction of the nucleus, through the degradation of both specific nuclear proteins and DNA, results in the collapse of chromatin into highly condensed electron-dense masses. This change in nuclear structure was originally observed by electron microscopy (1), but more recently a variety of fluorescence-based techniques have emerged to analyze the nuclear events by microscopy, flow cytometry, or biochemical techniques that detect the underlying DNA fragmentation either in situ or in vitro using agarose gel electrophoresis. These techniques serve as quantitative assays for apoptosis and are also useful for studying
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the actual mechanism of DNA fragmentation, which is still poorly understood. The first technique for the routine analysis of DNA fragmentation in vitro used conventional agarose gel electrophoresis (CAGE) in which a constant electric field at low voltage is used to resolve DNA fragments purified from apoptotic cells (2). This technique, which is straightforward and inexpensive and can resolve fragments up to a maximum of about 30 kb, is ideally suited to observe the ladder of fragments that is generated in many, but not all, apoptotic cells. Although the formation of the DNA ladder has been the hallmark biochemical indicator of apoptosis, it is, in fact, only the end point of DNA degradation and does not reflect the full pattern of DNA fragmentation occurring during apoptosis. Indeed, many cells do not degrade their DNA to this extent. We have recently shown DNA degradation during apoptosis to be a much more complex process that commences with the generation of higher molecular weight (HMW) DNA fragments which are undetectable by CAGE (3–5). These larger fragments are 50 –300 kb in size and reflect endonucleolytic cleavage of interphase chromosomes at the nuclease-sensitive sites that reside in chromatin fibers as a result of their folding into loop (mean size of 50 kb) and rosette (mean size of 300 kb) structures, respectively (3). This suggests that interphase chromosome breakdown in apoptosis is a highly ordered process and is intimately related to the higher order structure of chromatin. However, CAGE cannot resolve fragments greater than approximately 30 kb. Larger fragments either remain in the well or migrate 329
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slowly into the gel as a single band that is an unresolved zone of compression. Pulsed-field gel electrophoresis (PFGE) is a specialized form of agarose gel electrophoresis that permits the ordered separation of larger fragments of DNA (6). The simplest form of PFGE is fieldinversion gel electrophoresis (FIGE) in which the current is periodically reversed between two electrodes oriented 180° with respect to each other. This permits the use of the same gel boxes that are used for CAGE. Since the theory of migration of large DNA molecules during PFGE is not well understood, the frequency at which the current is reversed and the duration of the periods of forward and reverse polarity have been worked out empirically for particular size classes of DNA. We have adapted a form of FIGE called zero-integrated field electrophoresis (ZIFE) to optimize the separation of DNA fragments produced in apoptotic cells. Under appropriate conditions, we can easily extend the range of fragment sizes that can be studied to 1 Mb without losing any of the small fragments from the gel. Since larger DNA molecules are very sensitive to mechanical shear and endonucleolytic attack, radically different sample handling techniques must be used to prepare DNA for PFGE. Another characteristic feature of the degradation of DNA during apoptosis is the appearance of singlestrand breaks in the DNA. Indeed, it is possible that the endonuclease(s) responsible for DNA fragmentation introduces a multitude of single-stand nicks into the DNA, until eventually nicks accumulate sufficiently close together on opposite strands to generate a de facto double-strand break (7–9). We have developed a unique methodology to study the appearance of single-strand nicks in apoptotic DNA. Finally, since the degradation of DNA in apoptosis is endonuclease mediated and there is an intense effort to identify and characterize these proteins (8 –11) we provide methodology for their detection and characterization.
TREATMENT OF CELLS General considerations. The processing of cells or tissues is very dependent on the DNA analysis that is to follow. If only small fragments are to be extracted for “ladder” analysis by CAGE they can easily be solubilized from any cultured cells or tissue with detergent. Cells and tissue must be processed
much more carefully for PFGE analysis of HMW DNA, since a prerequisite for the analysis of high molecular weight DNA is the elimination of mechanical or enzymatic damage to the DNA during the processing of cells or nuclei. We have much more experience working with suspension cultures or thymocyte primary cultures than with either monolayers or tissue (12). A method for the processing of tissue is described by MacManus et al. (13). Processing of cells, tissues, and nuclei. Following the incubation of cells with apoptotic inducers, suspension or primary cultures are easily pelleted from the medium. Monolayer cells are harvested by trypsin treatment [0.15% trypsin in phosphate-buffered saline containing 1mM EDTA] to generate a cell suspension. The suspensions are pelleted and resuspended in nuclear buffer (NB, 15 mM Tris–HCl, pH 7.4, 60 mM KCl, 15 mM NaCl, 1 mM EGTA, 2 mM EDTA, 0.5 mM spermidine, and 0.15 mM spermine). This buffer has a relatively low ionic strength to assist in disruption of the cell and contains EDTA and EGTA to chelate divalent cations, as well as polyamines to bind to chromatin and protect the DNA. It is very difficult to extract DNA from tissue sufficiently quickly to completely prevent endonucleolysis. The best strategy is to isolate nuclei using buffers that are designed to inhibit endonuclease activity and protect chromatin. To achieve this, cultured cells or tissues are homogenized in a buffer consisting of ice-cold 0.25 M sucrose, 15 mM Tris– HCl, pH 7.4, 60 mM KCl, 15 mN NaCl, 2 mM EDTA, 0.5 mM EGTA, 15 mM mercaptoethanol, 0.5 mM spermidine, and 0.15 mM spermine. The nuclei are pelleted at 700g for 10 min at 4°C, resuspended in the same buffer containing 1% Triton X-100 to remove the outer nuclear membrane, pelleted again, and resuspended in an appropriate buffer for the experiment. For PFGE, washed cells or nuclei are usually embedded in agarose to prevent any further degradation. However, to completely eliminate artifactual DNA fragmentation, it is possible, in some circumstances, to carry out entire experiments on embedded cells, since media constituents and inducers of apoptosis can easily diffuse into the agarose plug. To do this, approximately 3 3 107 cells are suspended in 250 ml of medium containing 5% serum and mixed with 250 ml of molten 1.5% (w/v) low-melting-point agarose (prepared in medium without serum and held at 37°C, as indicated below). It is paramount
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not to heat shock the cells by allowing the temperature to rise above 37°C. The mixture is pipetted into 1-ml disposable syringes and allowed to solidify. The plug containing the embedded cells is ejected and incubated in RPMI medium containing 5% fetal bovine serum plus or minus inducers of apoptosis. After the incubation, the plug is processed, as described below, for either CAGE or PFGE analysis of DNA fragmentation. In some experiments we have embedded live cells as described above and then permeabilized them to allow the effects of cations on DNA fragmentation in situ to be studied. To do this the embedded cells are incubated at 37°C in 300 ml of permeabilization buffer (10 mM Tris–HCl, pH 7.0, 60 mM KCl, 0.1 mM EGTA, and 0.5% Triton X-100). Cations can be added and incubations can be carried out at 37°C for various times.
DETECTION OF DNA LADDERS BY CAGE Sample preparation. In this technique cells or nuclei are lysed with detergent and the small, detergent-soluble fragments of DNA are purified using phenol:chloroform. The entire procedure can be carried out in microfuge tubes. The purification steps remove proteins that could retard the migration of DNA in the gel and also RNA which can comigrate with smaller DNA fragments. Approximately 1 3 106 cells, or the equivalent amount of tissue (;10 mg), are resuspended in 400 ml of icecold extraction buffer containing 5 mM Tris–HCl, pH 7.5, 5 mM EDTA, and 0.5% (v/v) Triton X-100. The suspension is centrifuged at 14,000g for 20 min at 4°C in a microfuge, the supernatant is collected, and the DNA is precipitated from the supernatant with 40 ml of 2.5M sodium acetate (pH 5.2) and 2 vol of absolute ethanol at 220°C (usually overnight for convenience, but 2 h is sufficient). The precipitated DNA is collected by centrifugation, resuspended in 400 ml of TE buffer (10 mM Tris–HCl, pH 7.5 1 1 mM EDTA) and treated with RNase A (20 –100 mg) for 60 min at 65°C. Twenty-five microliters (0.5mg) of proteinase K solution (20 mg/ml stock in distilled H2O) is then added along with 40 ml of 103 proteinase K buffer (100 mM Tris–HCl, pH 7.8 1 50 mM EDTA and 5% N-laurylsarcosine) and the incubation continued for 2 h at 37°C. The DNA is then phenol:chloroform extracted by adding 400 ml of phenol:chloroform [saturated phenol mixed 1:1 (v/v)
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with chloroform:isoamyl alcohol, 24:1 (v/v)], gently mixing, and spinning in a microfuge for 5 min. The top layer is transferred to a clean tube and extracted once more. This layer is then precipitated with 40 ml of 2.5 M sodium acetate (pH 5.2) and 1 ml of absolute ethanol at 220°C overnight (or 1 h at 280°C). After precipitation, the samples are spun for 30 min at 4°C in a microfuge and the supernatant is discarded. The pellets are washed with 70% ethanol and centrifuged in a microfuge for 5 min. This wash step is repeated and the pellets are then dried on the bench or in a Speed Vac (Fisher Scientific, Ottawa, Ontario, Canada). The precipitated DNA is dissolved in TE buffer. Preparation of gel. We use 0.8% (w/v) gels for both CAGE and PFGE made with electrophoresis grade agarose from Gibco-BRL (Burlington, Ontario, Canada). The agarose (2.4 g) is suspended in 300 ml of TAE buffer (0.04M Tris–acetate, pH 8.5, containing 2 mM EDTA) and heated for 2 min in a microwave oven, removed, stirred briefly, and heated again for 2 min. The molten mixture is removed from the oven and stirred on a magnetic stirrer until it cools to about 60°C. The agarose is then poured into the gel tray and allowed to cool. When the gel has solidified (in approximately 30 min), it is put in the refrigerator or cold room for 5–10 min to be sure it is hard before removing the comb. If the gel is poured at too high a temperature moisture will condense onto the surface and cause local dilutions of the agarose before it solidifies. Similarly, if the comb is removed too soon moisture will condense in the wells and distort their shape. Electrophoresis. A wide variety of gel boxes, designed for horizontal, submerged gel electrophoresis, including many minisystems, can be used. A basic 200-V power supply is sufficient. The extracted DNA (approximately 5–10 mg) is dissolved in 50 ml of TE buffer, followed by 5 ml of loading solution (30% (w/v) sucrose 1 0.01% (w/v) bromphenol blue in distilled H2O). The sample is then added to each well. We routinely use two size markers for CAGE: the 123-bp ladder and a HindIII digest of l DNA, both from Gibco-BRL. The running buffer is TAE. For regular sized (20 3 25 cm) gels the run is carried out at 30 V for 18 h (i.e., overnight), whereas for minisystems we use 120 V for 1 h. The gel is removed from the tank, still attached to the gel tray, stained, and imaged as described below.
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DETECTION OF BOTH DNA LADDERS AND HMW DNA FRAGMENTS USING PFGE As described earlier, DNA fragments greater than 30 kb cannot be resolved by CAGE. Therefore, PFGE in some form must be used to resolve the larger fragments also produced during DNA degradation in apoptosis. Although fragments as large as 5 Mb can be resolved by PFGE, a disadvantage is that in any given run usually only a fairly narrow size range of fragments can be resolved. Furthermore, in most protocols, the smaller, ladder-sized DNA fragments run off the gel so two gels must be run for each sample—CAGE to detect small fragments and PFGE to detect HMW fragments. To obviate these problems we have devised PFGE conditions that allow fragments from 100 bp to 1 Mb to be resolved on the same gel. This size range spans the sizes of DNA typically seen in apoptotic cells, so it is possible to see the entire spectrum of DNA fragments in a single run. Equipment. There are a number of commercially available PFGE units varying tremendously in capability and cost. For routine detection of low and HMW DNA fragmentation in apoptosis we use the inexpensive FIGE mapper equipment sold by BioRad Laboratories (Mississauga, Ontario, Canada). This unit is programmable with user-defined parameters permitting the use of non-FIGE conditions, such as ZIFE. Sample preparation. Sample preparation is the most critical step in studies on HMW DNA degradation using PFGE. Activation of nucleases often accompanies cell disruption and long DNA fragments are also extremely susceptible to mechanical shear. Therefore, any technique for the analysis of high molecular weight DNA fragmentation in apoptosis is critically dependent on methodology to isolate DNA without further artifactual damage. The technique of embedding cells in agarose plugs evolved to solve this problem. To eliminate mechanical stresses, the whole cell or nucleus is usually embedded in an agarose plug prior to any manipulations on DNA. The proteins, including endonucleases, are then removed enzymatically rather than with solvents. The presence of the denaturing detergent sodium dodecyl sulfate (SDS) and the absence of divalent cations during the incubation with proteinase K ensures the integrity of the DNA. Endogenous RNases are active under these conditions and RNA is usually de-
graded. If necessary, a RNase step can be included at this stage or performed after the gel is run (see below). If cell extracts are to be stored before being processed for PFGE they should be stored at 280°C since DNA degradation occurs during storage at 220°C. Preparation of plugs for PFGE. We use one of two methods for embedding cells in agarose, depending on the number of cells or nuclei available. Usually, DNA from 1–2 3 106 cells must be loaded onto the gel in order to easily detect the DNA by ethidium bromide staining, although DNA from as few as 0.5 3 106 cells or nuclei can still be detected using SYBR gold from Molecular Probes (Eugene, OR). If only 2 3 106 cells, or less, are available then all the cells are encapsulated into a miniplug which is loaded in its entirety onto the gel. If more cells are available, then a larger plug can be made and slices, containing the equivalent of 1–2 3 106 cells or nuclei, can be cut and loaded onto the gel. Preparation of miniplugs. First, low-meltingpoint (LMP) agarose (Gibco-BRL) is weighed into a scintillation vial and an appropriate volume of NB to yield a 1.5% agarose mixture is added. The vial is put into the microwave oven for approximately 20 s, i.e., until it just boils, but does not boil over. Fortymicroliter aliquots are pipetted into Eppendorf tubes arrayed in a heating block (Fisher Scientific) followed by 1 ml of proteinase K (20 mg/ml stock solution in distilled H2O) and the mixture is kept molten at 37°C until ready to use. The sample of cells (or nuclei) is washed in NB, pelleted again, and resuspended in 40 ml of NB. This suspension is mixed with a 40-ml aliquot of the molten agarose–proteinase K mixture, pipetted into the wells of a miniplug casting mold (Bio-Rad), and allowed to set for 10 min at 4°C. When the agarose is set, the plug is ejected into a 1.5-ml microfuge tube containing 300 ml of TEEN buffer (10 mM Tris–HCl, pH 9.5, 25 mM EDTA, 5 mM EGTA, 10 mM NaCl), 15 ml of 10% (w/v in distilled H2O) SDS, and 2 ml of 20 mg/ml proteinase K. The samples are incubated for 1.5 h at 37°C (we use a rotator in an incubator). The objective is to remove protein in as short an incubation time as possible to minimize any further DNA degradation or diffusion from the plug. Complete deproteinization is essential in order to obtain uniform lanes when the gel is run. It is important to realize that proteinase K can often be contaminated with nucleases. We have found the proteinase K
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obtained from Gibco-BRL to be consistently reliable. If nuclease contamination is suspected a plasmid digestion assay (see below) can be carried out to detect any contaminants and the batch can be discarded if necessary. Following the incubation, the supernatant is removed and, if desired, any DNA that may have diffused from the plug can be precipitated from the supernatant at 220°C by adding 0.1 vol of 2.5 M sodium acetate and 2 vol of absolute ethanol. The DNA is washed, redissolved, and run on CAGE. For short deproteinization times, this is not usually necessary. The deproteinized plug is washed in 1 ml of TE at 4°C for 30 min. This step is repeated once and the deproteinized plugs are stored at 4°C in TE buffer until ready to load on a gel. If storing for longer than overnight, remove the TE buffer and store the plug in a sealed microfuge tube.
plugged with Parafilm and allow to set for 20 –30 min at 4°C. Cut the tip off the syringe to leave a blunt end flush with the end of the barrel to permit the solidified plug to be ejected. Eject the plug into a 15-ml plastic tube containing 3 ml of TEEN buffer, 150 ml of 10% (w/v) SDS, and 20 ml of 20 mg/ml proteinase K. The incubation is carried out for 3–16 h at 37°C, as described above, but should be kept to the minimum time required to remove all of the protein. Remove the plug and wash with 10 ml of TE at 4°C for 30 min. Repeat the washing step once and gently suck the plug back into the syringe, cover the end with Parafilm, and store at 4°C until ready to load onto the gel. To load, cut 20- to 30-ml slices, containing 1–2 3 106 cells or nuclei, from the end of the plug and place in the well. The wells are sealed with molten 0.8% (w/v) LMP agarose prepared in 0.53 TBE (45 mM Tris– borate 1 1.25 mM EDTA, pH 8–8.5).
Preparation of larger plugs. Harvest the cells as described above and pellet 20–30 3 106 cells. The plug volume is varied, depending on the number of cells, such that a 30-ml slice (a volume that conveniently fits into a well) gives 1–2 3 106 cells or nuclei. Wash the cells once with 1 ml of NB, resuspend in 250 ml of NB, and add 250 ml of molten 1.5% (w/v) LMP agarose in NB containing 0.1 mg of proteinase K (5 ml of 20mg/ml stock). Pipette the mixture into the barrel of a 1-ml disposable plastic syringe in which the tip has been
PFGE conditions. Gels (0.8%, w/v) are prepared in 0.53 TBE, as described earlier for CAGE. Four sets of standards, with overlapping size ranges, are used: yeast chromosomes (225 kb–1.2 Mb); polymerized l phage DNA (50 –1 Mb), low range markers (0.1–194 kb), all purchased from New England Biolabs (Beverly, MA); and the 123-bp ladder from Gibco-BRL. The quality of markers, particularly the yeast markers, and the extent of polymerization of the l ladder varies considerably. New England Bio-
FIG. 1. Separation of DNA from apoptotic cells by PFGE. (A) EL4 cells were incubated with (lanes 4 and 5) or without (lanes 2 and 3) 10 mM VM26. The cells in lanes 3 and 5 were preincubated for 30 min with 100 mM VAD-fmk, an inhibitor of caspases. Lane 1, control, unincubated cells. Lanes a– e, DNA size markers; 123-bp ladder, low-range PFGE markers (0.1–194 kb); polymerized l phage DNA (50 –1 Mb) and yeast chromosomes (225 kb–1.2 Mb), respectively. The sizes of a selection of these markers are indicated at the right of the gel. (B) Standard curve derived from the positions of the markers shown in lanes a– e of (A), showing ordered separation of the fragments.
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labs consistently produces the most reliable product. Electrophoresis is carried out using a horizontal gel chamber and a FIGE Mapper system (Bio-Rad Laboratories). The buffer, 0.53 TBE prechilled to 14°C, is circulated through a Lauda bath to maintain this temperature. The conventional 13 TBE is not used because the higher ionic strength causes overheating. Electrophoresis is carried out using the following settings: 280 V in the forward direction and 90 V in the reverse direction with an initial switch time of 0.3 s forward and 0.5 s reverse, with nonlinear ramping to 35 s forward and 60 s reverse, for a total run time of 20.5 h. An example of a typical PFGE separation is shown in Fig. 1. In Fig. 1A, the cells were incubated with an apoptotic inducer which results in the degradation of DNA to produce a ladder of small fragments (lane 4). If the cells are also incubated with an inhibitor of caspase activity, then DNA fragmentation is restricted to HMW fragments only (lane 5). Note that the control cells have completely intact DNA which all remains in the well (lane 1). Lanes a– e are standards and were used to produce the standard curve shown in Fig. 1B. This curve shows that, although not linear, there is an ordered separation of fragments over 4 orders of magnitude of sizes (100 bp to 1 Mb).
ANALYSIS OF DNA FRAGMENTATION BY 2D ELECTROPHORESIS Since the DNA strands are not denatured during sample processing in any of the above procedures,
the fragments that are resolved by either CAGE or PFGE are double-stranded molecules. However, these double-stranded molecules may harbor singlestrand breaks along the length of the duplex (7). The presence of these nicks can be detected by denaturing the duplex with sodium hydroxide to release the smaller single-stranded fragments. To study the extent of single-strand cleavage of DNA during apoptosis we developed a two-dimensional separation technique which uses nondenaturing PFGE in the first dimension and denaturing conditions in the second dimension. Conditions for 2D separations. In experiments involving 2D separations, the first dimension PFGE is performed exactly as described above, except that the run is shortened to 19 h from the usual 20.5 h. Only two samples are loaded in order to leave room for the second dimension (Fig. 2A). The gel is stained with ethidium bromide and imaged as described below. SYBR gold cannot be used at this stage since it binds to and severely retards the migration of DNA in the second dimension. The gel is rotated in the gel tray for the second-dimension run and the bottom of the gel is sliced off, if necessary, to fit. The second dimension run is either CAGE, run as described above, or denaturing electrophoresis. For the denaturing run, the gel is soaked in 50 mM NaOH 1 1 mM EDTA for 60 min at 4°C and then run in the same buffer. Both second-dimension runs are carried out at 150 V for either 2.5 h (denaturing conditions) or 2 h (CAGE) with buffer recirculation and cooling to 2°C (the temperature set at the Lauda bath). After the denaturing run, the gel is rinsed
FIG. 2. 2D PFGE-denaturing gel analysis of DNA from apoptotic Jurkat cells. (A) DNA from embedded control (lane 1) and apoptotic (lane 2) cells was resolved by PFGE. Lane a is the polymerized lambda ladder. (B) The gel was rotated and subjected to a second dimension of denaturing electrophoresis. Only the separation for lane 2 is shown. For reference, lane 2 from (A) is rotated and shown on the top of the second-dimension gel.
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twice in distilled H 2 O and then stained with ethidium bromide for 1 h and imaged, as described below. SYBR gold cannot be used for staining denaturing gels. A typical 2D analytical experiment is shown in Fig. 2. Once again the control cells (lane 1) have intact DNA, whereas the apoptotic cells produce primarily fragments in the 10- to 50-kb range with only a small amount of small fragments (lane 2). Lane 2 (i.e., the first dimension of PFGE) is rotated and shown again at the top of Fig. 2B. Underneath is the result of the second-dimension denaturing run. It can be clearly seen that a considerable number of single-strand DNA molecules lie underneath the “curve” of the still intact fragments. This is particularly evident for the 50-kb band which is seen to contain numerous single-strand breaks resulting in the release of many smaller fragments. Digestion of embedded DNA with nucleases. The presence of single-strand nicks and other modifications to DNA from apoptotic cells can be detected enzymatically. Embedded DNA is deproteinized as before and incubated with S1 nuclease (Sigma), mung bean nuclease (Pharmacia), or Bal31 (GibcoBRL). For the S1 nuclease digestions, miniplugs or slices of larger plugs are incubated in a buffer containing 30 mM acetate buffer, pH 4.6, 100 mM NaCl, 0.5 mM zinc chloride, and 450 U of enzyme. Mung bean nuclease digestions are carried out in 30 mM acetate buffer, pH 4.6, containing 50 mM NaCl, 1 mM zinc chloride, 0.01% Triton X-100, and 80 U of enzyme. Bal31 digestions are carried out in 20 mM Tris–HCl, pH 8.0, 200 mM NaCl, 12 mM CaCl2, 12 mM MgCl2, and 5 U of enzyme. All the incubations are typically carried out for 60 min at room temperature and the plugs are subjected to electrophoresis.
VISUALIZATION OF DNA FRAGMENTS AND IMAGE ANALYSIS Gel documentation. There are a number of possibilities for gel documentation and image analysis. The first consideration is the choice of fluorochrome for staining. Routinely, we use ethidium bromide which is inexpensive, easily visualized using a standard transilluminator, and compatible with both Polaroid film and CCD cameras. SYBR gold (Molecular Probes) is more sensitive, particularly when used in conjunction with Polaroid 667 film. However, this
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stain cannot be used in all circumstances (see below). A standard transilluminator with 300-nm lamps is used (such as the one from Ultra-Lum, Inc., Paramount, CA). Although Polaroid 667 film is the most sensitive, acceptable results are also obtained with type 55 positive/negative film for most applications. The image can also be acquired using a CCD camera coupled to a thermal printer and/or a data acquisition board and computer. A variety of software is available for image capture and data analysis. We use an inexpensive Panasonic Model BP312 CCD camera and image capture package from UltraLum. For both Polaroid cameras and CCD cameras it is important to use the most suitable filter for the dye being used. Filters for polaroid film are available from Molecular Probes. For the CCD camera we use a 600 6 25-nm bandpass filter for ethidium bromide and a 560 6 35-nm bandpass filter for SYBR gold (this filter also gives satisfactory results with ethidium bromide for most purposes). The latter filters can be obtained from Melles Groit (Ottawa, Ontario, Canada) in a variety of lens mounts. Following either CAGE or PFGE, the gel is placed in a glass or plastic container and submerged in ethidium bromide (1.5 mg/ml) or in SYBR gold (a 10,000-fold dilution of the stock supplied by the manufacturer in 0.53 TBE). Ethidium bromide needs destaining for at least 30 min, whereas SYBR gold requires no destaining. The gel is placed on the UV transilluminator and either photographed or the image is captured using the CCD imaging system. If there is a lot of nonspecific fluorescence the gels can be soaked in water for several more hours and then photographed again. To minimize the number of manipulations prior to electrophoresis we usually remove RNA after the run by soaking the gel in running buffer containing 1 mg/ml of RNase A for 2 h at room temperature or overnight at 4°C prior to staining and image capture. Data analysis. CCD cameras produce a digitized image of the gel which can subsequently be used to generate scans of pixel intensity vs distance of migration for each lane. A number of software packages (e.g., Sigmagel and Sigmascan from SPSS, Chicago, IL) are available for analysis. Standard curves are generated by regression analysis of the positions of overlapping DNA size markers. The sizes and total amount of fragmented DNA in each sample can then be calculated.
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METHODS FOR DETECTION OF ENDONUCLEASE ACTIVITY Endonuclease activities can be detected by one of two methods. The simplest way is to incubate samples with an intact double-stranded DNA plasmid and use CAGE to assess its integrity. Alternatively, the extracts can be resolved by SDS–PAGE (denaturing polyacrylamide gel electrophoresis) using gels copolymerized with substrate DNA and endonuclease activity bands can be detected after renaturation and digestion of the DNA. The latter technique can be performed using 1D or 2D protein separations and has the advantage of resolving multiple endonucleases in a mixture. In addition, their sizes, isoelectric points, and posttranslational modifications can be determined. As such, it is the tool for the discovery of new endonucleases. Plasmid DNA digestion assay. The protein sample (5–10 mg of total protein) is resuspended in 25 ml of 50 mM Tris–HCl buffer, pH 7.4, 2 mM MgCl2, and 0.2 mM CaCl2 and 1 mg of plasmid DNA is added. Any intact plasmid in the 5- to 20-kb size range can be used. The reaction mixture is incubated for 0.5–2 h at 37°C and the reaction can be stopped by the addition of 4 ml of 103 DNA loading buffer (40% w/v sucrose, 0.01% bromphenol blue, 50 mM Tris–HCl, pH 8.0, and 5 mM EDTA). The integrity of the plasmid DNA is analyzed by loading the reaction mixture on a 0.8% agarose gel. Electrophoresis is carried in TAE at 30 V overnight or 150 V for 2 h, as described above for CAGE. The gel is stained with ethidium bromide and imaged on a UV transilluminator. Usually there are three bands in the undigested plasmid DNA, corresponding to supercoiled (fastest moving band), circular (nicked), and linear forms. Samples containing a low level of endonuclease activity may produce a nick in the supercoiled or circular plasmid resulting in the disappearance of the lower band and appearance of the slower moving relaxed or linear forms. Higher activity samples digest the plasmid completely producing a smear of DNA near the bottom of the gel. It is possible to use this assay to characterize the cation requirements, the pH optima, and the effects of inhibitors. In addition, it is possible to study the nature of the ends of the fragments by attempting to radiolabel them as described below.
Endonuclease activity gels. Endonuclease activities present in cell or nuclear extracts can be identified by SDS–PAGE using a gel in which the DNA substrate is embedded (10, 11, 14, 15). We use SDS– 10% polyacrylamide gels copolymerized with 30 mg/ml of salmon testis DNA (Sigma Chemical Co., St. Louis, MO). The DNA can be either double stranded or heat denatured single stranded. To prepare the DNA, dissolve salmon sperm DNA (Sigma Chemical Co.) in distilled H2O at a concentration of 3 mg/ml. Sonicate the solution for 20 s to produce sheared double-stranded fragments. If required, denature the DNA in a boiling water bath for 5 min and transfer immediately to an ice bath to obtain single-stranded DNA. Use 100 ml of DNA solution (single or double stranded) per 10 ml of acrylamide gel mix. 32P-labeled DNA (106 cpm/ml) can also be added to the DNA solution. A total of 0.2– 0.5 mg of DNA can be radioactively labeled using [a-32P]dATP and [ a - 32 P]dGTP (800 Ci/mmol, NEN DuPont, Oakville, Ontario, Canada) and the multiprime DNA labeling system (Amersham, Oakville, Ontario, Canada). The resolving and stacking gels are then poured using standard procedures. Protein samples are dissolved in a loading buffer of 62.5 mM Tris–HCl buffer, pH 6.7, containing 1% (w/v) SDS, 10% (v/v) glycerol, and 0.01% bromophenol blue, usually prepared as a 43 stock. The samples are not boiled prior to loading. For reducing conditions, 2-mercaptoethanol is added to a final concentration of 1% (v/v). It is important to run the gels, at least initially, under both reducing and nonreducing conditions since some endonucleases are inactive under standard reducing conditions. The samples are separated on SDS–PAGE and the location of the active bands is determined by subsequent reactivation of the endonuclease(s) which remove the embedded DNA from the gel at that location. This is achieved during a protein renaturation step [30 min at room temperature in 50 mM Tris– HCl buffer, pH 7.5, 5 mM MgCl2, and 20% (v/v) 2-propanol with gentle rocking and two changes of the buffer], followed by an overnight reactivation step in 50 mM Tris–HCl buffer, pH 7.5, 5 mM MgCl2, 1 mM CaCl2, and 0.1% Triton X-100 (v/v) at 37°C. Nonradioactive gels are stained in 1 m g/ml ethidium bromide and imaged as before. the ethidium bromide-stained gels show a dark band (no fluorescence) where a nuclease has digested away the DNA in that part of the lane. The gels containing radioactively labeled DNA are dried and exposed to
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Kodak X-Omat film. The autoradiograph from radioactive gels also shows a clear band where a nuclease is present. When searching for new endonuclease activities it is essential to use substrate gels with radioactively labeled DNA since there is a potential artifact in the method that uses unlabeled DNA. On unlabeled gels, DNA-binding proteins, such as histones, can mask the DNA from the ethidium stain giving a “band” on the gel that is indistinguishable from the one produced by an endonuclease. This phenomenon does not occur on autoradiographs, since radioactivity can only be lost due to endonucleolytic degradation of the DNA. DNA–substrate gels also provide a convenient way to establish cation requirements and pH profiles. Equal amounts of the same protein sample are loaded in each lane and separated as described above. After the run, individual lanes are cut separately and subjected to renaturation and activation steps in solutions containing different compositions of cations, cation chelators, or solutions with identical cations but different pH. Two-dimensional activity gel electrophoresis. 2D acrylamide gel electrophoresis can be used to better resolve endonucleases. 2D gels also indicate the isoelectric point of a protein and give some indication to the extent of posttranslational modifications. The first dimension of isoelectrofocusing (IEF) is performed using a Multiphar electrophoresis unit (Pharmacia Biotech, Baie d’Urfe, Quebec, Canada) using the experimental conditions suggested by the manufacturer. Samples of protein extract are mixed with 240 ml of IEF loading buffer (8 M urea, 2% Triton X-100 (v/v), 2% Pharmalytes pH range 3/10) and absorbed onto Immobilin IEF strips, pH 3/10, for 8 h. Reducing agents are omitted if there is reason to believe a redox-sensitive nuclease exists. The proteins are subjected to overnight IEF at 15°C for 20,000 Vh. After the IEF run is complete, the Immobilin strips are loaded onto SDS–10% polyacrylamide gels copolymerized with 30 mg/ml of sheared salmon testis DNA (single stranded, double stranded, and/or radiolabeled) and the proteins are separated in the second dimension. Following the run the endonucleolytic activity is reactivated and visualized as described above. Labeling of 39-OH and 59-OH ends of DNA. DNA fragments can be radioactively labeled at either the 39-OH or 59-OH end using terminal deoxynucleotidyltransferase (TdT) or T4 polynucleotide kinase, re-
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spectively. Since the presence of a phosphate group blocks both of these reactions, this approach can be used to determine the nature of the ends of the DNA fragments in apoptotic DNA, e.g., 59-P/39-OH or 59OH/39-OH or 59-OH/39-P. This has important implications for the mechanism of action of nucleases involved in DNA fragmentation in apoptosis. To assess this, divide the reaction mixture in two tubes, one for TdT and other for the T4 kinase labeling reaction. The TdT labeling reaction is carried out for 1 h at 37°C in 50 ml of reaction buffer containing 25 mM Tris–HCl buffer, pH 6.6, 200 mM potassium cacodylate, 5 mM CoCl2, 0.5 mM dithiothreitol (DTT), 0.25 mg/ml bovine serum albumin, 5 mCi of [a-32P]dATP (NEN DuPont and 60 units of TdT (Gibco-BRL). The T4 kinase labeling is also performed at 37°C for 1 h in 50 ml of reaction buffer containing 50 mM glycine–NaOH buffer, pH 9.2, 5 mM DTT, 10 mM MgCl2, 10 mM Na2HPO4, pH 9.2, 5 mCi[a-32P]ATP, and 50 units of T4 kinase (New England Biolabs). Unincorporated radionucleodides are removed using G-50 Micro columns (Pharmacia Biotech) and labeled DNA is separated by CAGE followed by autoradiography.
CONCLUDING REMARKS There is still a lot to be learned about the mechanism of chromatin degradation in apoptosis and the nature of the endonuclease(s) that catalyze it. To date more than 40 endonuclease activities have been implicated in the process, including both endogenous (i.e., nuclear) and exogenous (extranuclear) activities (8, 9). It is highly likely that more than one activity is required to completely degrade chromatin given that several million single- and double-strand breaks are introduced into chromatin within a matter of minutes once degradation commences. The pattern of fragments generated during this process is primarily a property of the substrate, chromatin. The enormous complexity of chromatin dictates the localization of nuclease-sensitive sites and we can learn about its structural organization from the fragmentation pattern. The challenge is to further develop the technology for analyzing DNA fragmentation, particularly at its earliest stages. Continued improvements in our ability to analyze large DNA fragments by PFGE or 2D electrophoresis will contribute greatly to this effort.
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