The choice of solvent systems for multidimensional chromatography of postlabeled DNA adducts is dependent on the type of adducts to be chromatographed and the removal of interfering ³²P-labeled compounds (i.e., normal nucleotides) that can contribute to the radioactive background on the chromatograms. Urea (used to reduce the interaction of hydrophobic molecules with the PEI-cellulose sheets), pH, and salt concentrations all influence the rate at which adducts migrate. Increasing the solvent strengths of the salts or urea, or both, will generally increase migration. However, as the solubility limits of the solvent components are approached, the risk of these components precipitating out of solution onto the PEI-cellulose sheets increases rapidly. A consequence of this precipitation is the tailing of radioactivity on the chromatograms which can hamper quantitation. Concentrations of urea greater than 8.5 M should be avoided because chromatography times increase significantly without any appreciable improvement in adduct migration. Generally, we dip the edge of the PEI-cellulose sheet in water or the buffer component of the urea/buffer solvents before placing the sheet in the solvent to reduce the problem of supersaturation and precipitation of salts in the solvent front. For large hydrophobic structures derived from PAHs a 1 M sodium phosphate, pH 6.0, solvent is generally used for D1 to migrate normal nucleotides and nonspecific radioactivity off the PEI-cellulose sheets and onto paper wicks. However, for faster migrating and less hydrophobic structures, the phosphate concentration may have to be raised to 2.3 M sodium phosphate, pH 5.5, to get the DNA adducts to salt out of solution and onto the sheets. The lower pH (5.5) is necessary to minimize the problem of sodium phosphate crystallizing on the PEI-cellulose sheets when using phosphate solutions that are 1.7 M or greater. Also, raising the laboratory temperature and sealing the tanks will help to reduce this problem. However, increasing the ionic strength of the phosphate buffer will also cause some interfering ³²P-labeled structures to salt out onto the sheets, and the resulting autoradiograms may not be as clean (Beach and Gupta 1992). An optional clean-up chromatography step can be done in the D2 direction (Fig. 5) using 2.5 M ammonium formate, pH 3.5, although this step is not necessary when chromatographing bulky, hydrophobic DNA adducts like those derived from PAHs (>2 rings). The solvent systems for D3 and D4 are used to separate the large, hydrophobic, xenobiotic DNA adducts. The most commonly used D3 solvent system for large, hydrophobic adducts on commercial sheets is 3.5 to 4.5 M lithium formate, 8.5 M urea, pH 3.5 (LFU). If the DNA adducts in D3 are extremely slow migrating then consider 3.5 to 4.5 M pyridinium formate, 8 M urea, and pH 3.4 (made by neutralizing formic acid with pyridine to desired pH). The traditional solvents for D4 are 1.6 M lithium chloride, 0.5 M Tris, 8.5 M urea, pH 8.0 (LTU) or 0.8 M sodium phosphate, 0.5 M Tris, 8.5 M urea, pH 8.2 (PTU) for commercial PEI-cellulose sheets. When using laboratory prepared sheets, dilute the solvents to 80 to 90% of the strength used for commercial sheets.

The PAH-DNA adducts are generally found within a diagonally shaped region when LFU is used for D3 and LTU is used for D4. If the diagonal radioactive zone (DRZ) formed on the chromatogram (as visualized in the autoradiogram) is strong with many intense overlapping spots, then the isopropanol:4N ammonia system is an excellent alternative for D4, because it will disperse the adducts over a larger area of the chromatogram (Fig. 6).

Another promising D4 solvent system is 0.08 to 0.4 M NH₄OH that gives a zone of adducts more disperse than LTU but less than the isopropanol/ammonia system (Spencer et al. 1993). In addition, this system develops rapidly, and chromatography for D4 is complete in approximately 0.5 hours. However, if the DRZ is weak, then the LTU system would be superior because the DNA adducts are compressed into a zone of overlapping spots that can be easily quantitated. Sometimes when autoradiograms are hazy, a final development (D5) in 1 M sodium phosphate, pH 6.8, can improve autoradiogram quality by removing some of the nonspecific radioactivity from the chromatogram.

Some of the solvents used for chromatography are based on formic acid or acetic acid. The concentration of these solutions is based on anion concentration, for example 4 M lithium or ammonium formate, pH 3.5 would have 4 moles per liter of formic acid and sufficient lithium hydroxide (or ammonia) would be added to bring the pH to 3.5. Another important consideration is the type of pH electrode used. With Tris solutions, a calomel electrode is essential because Tris interferes with the standard silver/silver chloride electrode in alkaline media. Also, pH must be set at the temperature of chromatography because some buffers, such as Tris, have strong temperature dependent pH profiles. Small changes in pH above 8.0 can have a substantial effect on the migration of some DNA adducts.

Note: Some researchers report that direct exposure to fluorescent light or stray sunlight can cause photodecomposition of benzo[a]pyrene-DNA adducts on PEI-cellulose sheets (R. Roggeband and M. J. Steenwinkel, pers. commun.). These findings indicate the need to use filtered light (400 nm cutoff) in the laboratory during chromatography.

The following protocol is for large hydrophobic DNA adducts.

1. Spot 5 to 20 µl of each ³²P-labeled sample slowly on the origin of a premarked PEI-cellulose sheet that has an 11-cm filter paper wick (Whatman No. 17 CHR) stapled to it (see Fig. 5) and allow the PEI-cellulose to draw the moisture from the pipette tip. If the sample is spotted too quickly, a puddle will form around the pipette tip giving a larger initial spot size. The spots will expand during chromatography. Slow, careful spotting will give better resolution of spots on the autoradiograms and this will help appreciably with quantitation. After the spot has been applied, the sheet is placed in a multisheet TLC chamber (see Reddy and Blackburn 1990 for specifications) containing 1.0 M sodium phosphate, pH 6.0, and developed in the D1 direction overnight. The wick should be saturated with the D1 solvent at the completion of this step.
2. Remove the sheet from the TLC chamber and touch the bottom edge to a piece of absorbent paper to remove any chromatography solvent adhering to the bottom edge of the PEI-cellulose sheet. Cut the sheet at Line 1 (see Fig. 5) using a paper cutter. Handle the PEI-cellulose piece with the attached wick carefully, it will have greater than 98% of the radioactivity initially spotted on the chromatogram.
3. Hold the PEI-cellulose sheet on the side opposite from the cut edge and rinse it in a slow stream of cool tap water to remove loose specks of PEI-cellulose containing radioactivity. These specks are caused by the cutting process and may give false images on the autoradiograms. They are usually easy to detect on the autoradiograms because they are intense and have a sharp border.
4. The sheet is then soaked in a tank (a 2 gallon aquarium works well with the multisheet holders) with either distilled water or soft tap water to remove the salts from the chromatography solvent. Agitation of the water in the tanks with a stirrer will enhance the removal of salts from the PEI-cellulose sheet. Faint light refraction differences next to the sheet can be seen as the salts leach off. Usually a 10 minute soaking is sufficient.
5. The sheet is placed on a drying rack and dried under a gentle flow of warm air using hair blowers.
   Note: Do not use extremely hot air to dry the sheet. We have observed cracking and peeling of the PEI-cellulose. It takes approximately 10 to 20 minutes for the sheet to dry. Be sure that the sheet is completely dry, otherwise solvent migration up the sheet may not be satisfactory in the next chromatography step.
6. Dip the bottom edge (approximately 1 cm) of the sheet in 0.45 M lithium formate, pH 3.5, or water, and the excess solvent on the bottom edge is wicked away with a paper towel. Develop the sheet in the D3 direction (Fig. 5) using a lithium formate-urea solvent system (8.5 M urea, 4.5 M lithium formate, pH 3.5, for commercial sheets or 85% strength for laboratory-prepared sheets). It usually takes 2 - 3 hours for the solvent front to reach the top of the sheet.
   Caution: The level of the liquid in the chromatography chamber must be below the sample application site on the PEI-cellulose sheets.
7. Remove the sheet from the TLC chamber and touch the bottom edge to absorbent paper to remove the solvent. The PEI-cellulose sheet must be held vertical until the solvent bead has been removed from the bottom of the sheet; holding the sheet in any other position allows the solvent bead to run across the sheet, and a displaced band of radioactivity will appear on the autoradiogram. Also, the solvent bead along the bottom of the sheet must not be allowed to wet the PEI-cellulose side of any other sheet in the chromatography tank during removal.
8. Rinse with a slow stream of tap water then place the sheet in a tank containing distilled or soft water and rinse in the same manner as D1. Remove the sheet from the water tank and cut along Line 2 (see Fig. 5). Rinse the sheet with tap water to remove any loose cellulose flecks.
9. Place the sheet on a drying rack and dry under a gentle flow of warm air.
10. Dip approximately 1 cm of the bottom edge of the sheet in a 0.05 M Tris, 0.8 M lithium chloride buffer, pH 8.0, and wick the excess solvent with absorbent paper. Develop the sheet in the D4 direction (see Fig. 5) using 0.5 M Tris, 1.6 M lithium chloride, 8.5 M urea, pH 8.0 (85% strength for laboratory prepared sheets).
11. Remove the sheet from the TLC chamber and touch the bottom edge to a tissue. Rinse in the same manner as for the D3 step.
    Optional cleanup step. Staple a 5 to 7 cm filter paper wick (Whatman No. 17 CHR) to the PEI-cellulose sheet according to Fig. 7. Dip the edge of the sheet in distilled water and wick away excess water with paper towel. Develop the sheet in the D5 direction (see Fig. 7) using a 1.0 M sodium phosphate buffer (pH 6.8) overnight. This sodium phosphate development step may improve the autoradiogram by removing some of the residual nonspecific radioactivity. After removing the wick, place sheet in a tank containing water and rinse them in the same manner as D1.
12. Trim the sheet for placement in the cassette while it is still wet (see Fig. 7). Excise the origin to remove an intense spot of radioactivity that may interfere with quantitation. Rinse the sheet gently with tap water to remove any flecks of radioactive PEI-cellulose left by the cutting process. Place the sheet on a drying rack and dry under a gentle flow of warm air. Tape the dry chromatograms face down to a piece of 14" by 17" paper and put in a cassette for autoradiography.

Please see section on autoradiography for further information.

Isopropanol/4 N ammonia is an excellent alternative D4 solvent system that will breakup the strong diagonal radioactive zones that the LTU solvent system gives and disperse the individual adduct spots over a larger area of the chromatogram (see Fig. 6).
NOTE: The isopropanol/4 N ammonia solvent system may cause some cracking to occur in Plexiglas boxes and equipment. Use glass tanks for this solvent system. The ratio of isopropanol/4 N ammonia (v/v) can vary from 0.8:1 to 2:1; 1.2:1 works well for large hydrophobic adducts on laboratory prepared sheets and 1:1 works for commercial sheets. When using the isopropanol/4 N ammonia solvent system for D4, the sheet is placed in 10 mM Tris base after the D3 water rinse step to soak for 3 to 5 minutes (this resets the pH of the sheet). The sheet is then rinsed under the tap and dried. A 2 cm filter paper wick (Whatman No.1) is then stapled to the top edge. A longer wick stapled to the top of the PEI-cellulose sheet will increase the distance that the adducts migrate and this can be used as an additional variable along with the isopropanol/4 N ammonia ratio to control adduct migration on the chromatogram. Place the sheet in a TLC tank containing the isopropanol/4N ammonia, seal it with plastic wrap, and develop in the D4 direction. Do not move the tank during development. Development time is usually 2 to 3 hr. When the solvent has reached the top of the wick, remove the sheet from the tank and let the sheet with the attached wick air dry in a fume hood. Place the dry chromatogram with the attached wick in 1 M sodium phosphate (pH 6.0) for D5. The D5 is a necessary step for the isopropanol/4 N ammonia solvent system, otherwise a strong dark band may appear in the middle of the autoradiogram. Trim sheet (see Fig. 7) for autoradiography.

Dilute ammonia (0.08 to 0.4 M) is also an excellent solvent system for D4, and the dispersion of adducts on the chromatograms falls in between the LTU and isopropanol/4 N ammonia solvent systems. The dilute ammonia solvent systems also require the Tris base pretreatment and the attachment of a 2 cm wick as described for the isopropanol/4 N ammonia system. The development time for the dilute ammonia solvent system is 20 to 40 minutes. After D4, develop in the D5 direction in the same manner as was done for the isopropanol/4N ammonia system.

At present, there are several ways to quantitate radioactivity on chromatograms. The traditional "cut and count method" aligns the autoradiogram with the corresponding chromatogram and uses a marking pen to outline the radioactive regions on the chromatogram. Radioactive regions are then cut from the chromatogram, placed in a liquid scintillation vial containing liquid scintillation cocktail, and counted by liquid scintillation spectrometry (LSS). Cerenkov counting, which measures the light emitted when a charged particle passes through water, can also be used; however, a correction for counting efficiency is necessary. In addition, areas near radioactive spots are excised and counted to obtain background corrections. However, background corrections based on this approach can be somewhat subjective and one must exercise careful judgment when the background correction is substantial relative to the spot or region of interest. Another analytical approach is the use of computer-aided imaging systems that will locate and directly measure the radioactivity on the chromatogram.

Autoradiography is used to locate the position of ³²P-derived radioactivity on the chromatograms; and it provides information on the relative intensity of each individual adduct. However, there are restrictions on the interpretation of autoradiograms. First, film has a limited range of response (less than 300) and the range of linear response is less than 100. Multiple exposures are usually required for a complete set of autoradiograms to show both strong and weak regions of radioactivity present on the chromatograms. Moreover, the autoradiographic images of faint spots may be only 20 to 30% of their expected densities based on the level of radioactivity present. Because of these limitations, caution is required when making assumptions on the relative intensity of faint spots and the levels of background radioactivity present based on a visual inspection of the autoradiogram. Autoradiography of the chromatograms either can be run at room temperature or film sensitivity can be enhanced by a factor of up to 18 with the use of intensifying screens at -80°C (Swanstrom and Shank 1978).

Handle the chromatograms carefully to avoid any flaking or loose particles being generated that are radioactive and could leave confusing images or spots on the autoradiograms.

1. Tape the chromatograms to a 14" x 17" piece of paper. The chromatograms can be taped to the paper with the vinyl side up if the samples are labeled with ³²P. If the chromatograms are placed on the paper with the cellulose side up, then cover with a thin plastic wrap. Be sure to make a map of the chromatogram placement on the paper sheets for future reference. The chromatograms should be marked with fluorescent ink so that they can be aligned with the autoradiograms later to be marked for cutting. If the intensifying screen surface in the film cassette is dirty, clean with Kodak screen cleaner.
2. The chromatograms are placed in a cassette and taken to the darkroom. All darkroom operations must be done using a photographic safelight. Add autoradiography film to each cassette, marking each film with a different scissor cut. For most samples, film exposure in the presence of intensifying screens for 2 to 72 hours at -80°C is satisfactory. For extremely radioactive samples, autoradiography at room temperature for up to several hours is often sufficient.
3. The cassette is removed from the -80°C freezer and brought to room temperature; this is done to prevent the film and PEI-cellulose sheets from cracking when the cassettes are opened.
4. In the darkroom remove the film from the cassette and place it in a tray containing Kodak GBX developer. The aqueous concentration of the developer should be about 5%. Development time varies from 2 to 4 minutes. The film is removed from the developer with tongs and rinsed in a flowing waterbath for at least 30 seconds. The film is then transferred to a tray containing Kodak GBX fixer at a concentration of 5%. Total time for fixing is 5 minutes. Do not allow fixer to accidentally get in the developer tray as it will ruin the developing solution. Remove film from the fixer and rinse it in a flowing waterbath for at least 5 minutes to remove all chemicals. To obtain high quality autoradiograms, it is important to use developer and fixer that have been made recently. As these solutions get older, the rate at which the film is developed or fixed slows down and the background "grayness" of the autoradiograms gets darker. The lights can be turned on now.
5. The autoradiograms are hung to dry and then labeled. Each autoradiogram should have the following information on it: date of autoradiography, length of exposure, temperature of film exposure, and identification of each individual chromatogram.

Photography of Autoradiograms
We use a 4 by 5 inch format Polaroid camera mounted on a stand and Polaroid type 55 positive/negative film for making prints of individual autoradiograms. Satisfactory prints can be made using a 35 mm camera and Kodak TX200 film. However, the larger format of the 4 by 5 inch Polaroid type 55 film yields a sharper print.

1. Place the autoradiogram on a lightbox that has a uniform light distribution.
2. The image is brought into focus through the Polaroid viewfinder. The viewfinder on the Polaroid camera has a centimeter scale which helps to define the print image size and is useful when more prints need to be made. The camera settings are normally f-4.5 for aperture and the shutter speed varies from 1/30 to 1/8 second. These settings can vary because of differences in lightbox intensity, film speed, the distance between the lens and the autoradiogram, and the quality of the autoradiogram.
3. After the picture has been taken, remove the film from the camera and develop for 20 to 30 seconds. Peel the paper apart to get the print and immediately wipe the surface of the print with fixer to set the image. The film pack also contains a negative which needs to be treated in a sodium thiosulfate bath immediately if you want a negative for making enlargements (see directions in the filmpack for preparing negatives).

Quantitation of Radioactivity by the "Cut and Count" Method
Place the autoradiogram on the chromatogram and align it with the fluorescent ink marks. If you do not have fluorescent ink marks to align the chromatogram, then use the solvent front edges on 2D autoradiograms to align the autoradiogram with chromatogram. Carefully outline the regions of interest including background areas on the chromatogram with a marking pen.

A map is drawn of the radioactive regions that are to be excised and each region is numbered. This will give you a permanent record of areas quantitated and their respective positions on the autoradiograms. The regions are then excised using a sharp razor blade or a scalpel. Each of the excised chromatogram pieces is weighed (this allows a background correction to be made per unit weight of chromatogram) and placed in a 20 mL scintillation vial. Add 10 mL of scintillation cocktail to each vial and count.

Liquid Scintillation Spectrometry
Liquid scintillation counting has some restrictions. Samples containing low levels of radioactivity (i.e., faint spots and samples for estimation of background corrections) must be counted for longer time periods for adequate counting statistics. Counting statistics are based on total counts measured and not the counting rate (i.e., cpm or dpm). The following numbers give a comparison between total counts and % error (i.e., total counts / % error): 200 / ± 14%, 500 / ± 9%, 1000 / ± 6%. Also, the liquid scintillation spectrometer has an inherent background of 20 to 25 cpm. A small variation of 3 to 10 cpm in the estimation of background per cm² of surface (easily achievable because of poor counting statistics for samples with low radioactivity levels) can greatly affect adduct level computations. For single, well-resolved spots, one can use a clean, adjacent area to define background. However, for diagonal radioactive zones, which often have a faint hazy area around them (the haze may be due to adducts), the question of where to sample for defining background corrections becomes highly subjective. The subjective aspect of defining background from a remote part of the chromatogram points to the need to do the chromatography and measurement of radioactivity carefully. Moreover, proper handling and storage of tissue and purified samples is important, because DNA breakdown products will raise the overall background.

The liquid scintillation counting efficiency for ³²P in weakly quenched solutions is 98 to 99% which means cpm is essentially the same as dpm.

Correction for background is as follows:
corrected cpm (region of interest) = cpm (region of interest) - [wt (mg) of region of interest x (background cpm/wt of background region in mg)].

Cerenkov Counting
An alternative to counting samples by LSS is Cerenkov counting which is cheaper because it does not require the use of scintillation cocktail and eliminates disposal problems associated with some organic-based cocktails. However, the counting efficiency for radioactivity can be substantially lower. A correction factor for counting efficiency can be made by counting aliquots of ³²P in both liquid scintillation cocktail and water. Since the counting efficiency in the cocktail is approximately 99%, the ratio of cpm-cocktail to cpm-water will give a correction factor for converting Cerenkov counts to dpm.

We are currently using storage phosphor imaging technology to locate and measure radioactivity associated with the DNA adducts present on the chromatograms. This technology offers high sensitivity (approximately 0.2 nmol adduct/mol nucleotides), extremely low counting background, a large linear range of response to radioactivity (10⁵ to 1), and sufficient data points to allow accurate mapping and quantitation of radioactivity using computer image analysis methods. (For an in-depth discussion on storage phosphor imaging see Reichert et al. 1992). An additional benefit of this imaging system is that, unlike radioactivity measurements by liquid scintillation spectrometry, all raw data for the distribution and intensity of radioactivity on the chromatograms is available for review and reprocessing at any time in the future.

Scanning the Chromatograms
The imaging screens are first erased using an Image Eraser (Molecular Dynamics, Sunnyvale, CA). Storage phosphor imaging is performed at ambient temperature by placing storage phosphor screens (Molecular Dynamics, Sunnyvale, CA) over the radioactive chromatograms and exposing for a specified time period. A factor to convert the screen signal to dpm is generated by imaging a test strip containing a serial dilution of a ³²P solution concurrently with the samples. The signal response from the screen is then divided by the ³²P dpm in the test spots to generate a factor for converting the imaging screen signal response directly to ³²P dpm. The length of screen exposure to a chromatogram is dependent on the maximum levels of radioactivity present and is usually about one-tenth the time required for autoradiography. Generally, for a 10 µg DNA sample a 6 to 24 hr exposure is sufficient. The chromatograms for DNA bases and the specific activity analyses require only a 10-minute exposure. Saturation of the screens occurs as the pixel values approach 80,000 and is evident when the radioactivity profiles of lines drawn through radioactive areas on the chromatogram images have a truncated appearance. However, this is a problem only with extremely "hot" samples where the radioactivity is concentrated in a small area. After exposure of the chromatograms to the screens, the latent radioactivity image is read by scanning the screen with a PhosphorImager (Molecular Dynamics, Sunnyvale, CA).

Data Storage and Computer Requirements
Image data from the PhosphorImager is stored on a 620 MB optical disk (Pioneer DEC-702 rewritable optical disk, Upper Saddle River, NJ) using a Pioneer rewritable optical disk drive (Upper Saddle River, NJ). Data are then processed on a 486-33 MHz computer equipped with a 330 MB hard drive, 32 bit local bus and 48 MB of RAM. A Diamond Stealth VRAM video accelerator card is used to increase the rate of screen repainting. Because the file sizes generated by the imaging process are up to 41 MB, it is desirable to place the files in a virtual RAM drive where processing is 3 to 6 times faster. This computer, as configured, processes large data files quickly and efficiently. The files can be processed on a 386-33 MHz; however, there is a substantial increase in data processing time. Software used to process the data is ImageQuant (version 3.15) from Molecular Dynamics.

Processing of Image Analysis Data

1. Restore the image file from the tape cartridge or other data storage device to the hard disk. If the file is on an optical disk, you can work directly from this drive, although processing will be slower.
2. If your computer has sufficient RAM, then use ImageQuant to copy the image file into a virtual RAM drive.
3. Adjust the size of the image using the "magnification" (footnote 8) command.
4. Choose the desired color scheme. If the resolution of the color image is poor, then try the gray scale.
5. The "color range" of the image can be optimized by drawing a "line object" through the strongest and weakest radioactive regions on the chromatogram and creating a profile of radioactive intensity along this line. From this graph, determine both the pixel values of the strongest spot that you are interested in and the background region. Set the color range using these values as a starting point for your maximum and minimum color values.
6. Draw an "object" around the region of radioactivity you wish to measure and give it a "name."
7. Determine the background value to be subtracted from your "object" by drawing "segments" through the area that surrounds the "object" that represent background. The "define background" command is used to obtain an average pixel value along each segment. From this information determine a value to be used as background.
8. "Integrate volume" for your region. (The background value will automatically be subtracted from each pixel in your "object".)
9. Transfer the table of values to a spreadsheet program. See Table 1 for a typical Excel spreadsheet after the data have been transferred from ImageQuant software.

The ³²P-postlabeling results are usually presented as a ratio of the number of adducts detected divided by the amount of DNA used in the assay (e.g., nmol DNA adducts/mol DNA, amol DNA adducts/µg DNA, fmol DNA adducts/µg DNA). Presenting the data in this fashion avoids the problem of not being able to extract DNA quantitatively from tissues.

It is necessary to keep track of all numbers used in the final calculations. It is also important that your laboratory protocols assure that the necessary information regarding aliquot sizes and dates when samples are counted is recorded. See Table 1 for sample calculations that include the raw data from the imaging system (in Microsoft Excel 4.0 format).

Sample calculations
Calculation for total DNA adducts measured:

Calculation for total DNA analyzed for adducts:

Calculation of DNA damage level:

nmol DNA adducts/mol DNA	= amount of DNA adducts measured in a sample/amount of DNA used.
	= 0.968 fmol DNA adducts/29 nmol DNA
	= 968 amol DNA adducts/29 nmol DNA x (109/109)
nmol DNA adducts/mol DNA	= 33 nmol DNA adducts/mol DNA

After the calculations are completed, the results are reviewed for accuracy. An important check for computational or data errors is to compare the final numbers with the autoradiograms. The numbers should vary directly with the intensity of the autoradiograms provided that comparable amounts of DNA were used in the assay.

The PPL method is an involved procedure that requires numerous determinations to be made that are dependent on enzyme efficiencies and radioactivity measurements. The method should be considered semiquantitative when working with tissue samples from animals exposed to complex mixtures, because the identities of the adducts formed from complex mixtures are unknown, and it is not possible to simultaneously optimize the conditions for all adducts present. However, if sets of samples are run under uniform conditions, then meaningful comparisons between samples collected from different sites can be made. To have confidence in the results, appropriate controls must be used to signal the presence of significant errors. The following are quality assurance procedures that should be included in the ³²P-postlabeling assay:

1. Salmon testes DNA is used for measuring the efficiency of DNA hydrolysis and as a sample blank throughout the assay (chromatography and reference standard).
2. The compound 7R,8S,9S,10R-(N²-deoxyguanosyl-3'-phosphate)-7,8,9,10-tetrahydrobenzo[a]pyrene (BaPDE-dG-3'p) is used as an external standard to monitor the efficiency of enzyme-mediated transfer of the ³²P-phosphate from [gamma-³²P]ATP to polycyclic aromatic hydrocarbon derived DNA adducts. The concentration of the stock BaPDE-dG-3'p from the manufacturer should be verified by postlabeling the BaPDE-dG-3'p with ³²P and using the specific activity of the [gamma-³²P]ATP to determine concentration. The concentration of the BaPDE-dG-3'p determined from the ³²P labeling should agree with the manufacturers stated concentration. The radioactivity measurement of the chromatographed spot will give information on the efficiency of ³²P labeling and can be used to verify the specific activity of the [gamma-³²P]ATP used in the assay (specific activity of [gamma-³²P]ATP = dpm of BaP spot/amount of BaP labeled). In addition, the postlabeled BaPDE-dG-3'p can be used as a chromatography standard.
3. In the butanol adduct enrichment method an aliquot of BaPDE-dG-3'p is used as an extraction efficiency standard for recovery of PAH derived DNA adducts. However, when a specific DNA adduct is being targeted then a standard for that compound, if available, should be used to determine extraction efficiency.
4. An aliquot of contaminant-modified DNA from fish injected with a contaminated sediment extract is also used to monitor labeling efficiency of complex mixtures between assays and as an additional chromatographic standard.
5. A 2'-deoxyguanosine-3'-monophosphate standard is used to monitor the efficiency of enzyme-dependent labeling of the normal nucleotides by ³²P-phosphate from [gamma-³²P]ATP.
6. To assess reproducibility, every 10th tissue sample is analyzed in duplicate for DNA adducts.