Vol. 138, No. 2
JOURNAL OF BACTERIOLOGY, May 1979, p. 333-338 0021-9193/79/05-0333/06$02.00/0
Multiple Loci Affecting Photoreactivation in Escherichia coli BETSY M. SUTHERLAND* AND STEPHEN G. HAUSRATH Biology Department, Brookhaven National Laboratory, Upton, New York 11973
Received for publication 6 February 1979
Sutherland et al. mapped a phr gene in Escherichia coli at 17 min and found that induction of an E. coli strain lysogenic for a A phage carrying this gene increased photoreactivating enzyme levels 2,000-fold. Recently, Smith and Youngs and Sancar and Rupert located a phr gene at 15.9 min. We have therefore investigated the properties of photoreactivating enzyme and cellular photoreactivation in cells containing deletions of the gene at 17 min. Cells with this deletion photoreactivated ultraviolet-induced killing at a rate 20% of normal; they also contained approximately 20% of the normal photoreactivating enzyme level. The residual enzyme in these cells was characterized to determine whether the reduced cellular photoreactivation rate and photoreactivating enzyme levels resulted from reduced numbers of normal enzymes or from an altered enzyme. Photoreactivating enzymes from strains carrying a deletion of the region at 17 min had an apparent Km about two- to threefold higher than normal enzyme and showed markedly increased heat lability. The gene at 17 min thus contains information determining the function of the E. coli photoreactivating enzyme rather than the quantity of the enzyme. It is proposed that the gene at 17 min be termed phrA and that located at 15.9 min be termed phrB. UV radiation produces cyclobutane-type pyrimidine dimers in DNA (9). A major pathway for repair of these dimers is photoreactivation (PR), in which a photoreactivating enzyme binds to a dimer-containing region of DNA and, in the presence of light of longer wavelengths (300 to 500 nm), monomerizes the dimer (8). Van de Putte et al. first mapped the phr gene in Escherichia coli to be near gal (14). Subsequently Sutherland et al. located a phr gene between gal and attA by comparing PR of UV-induced killing in strains with known deletions near gal (13). Strains with deletions from gal to attA showed greatly decreased PR of cell survival. They also showed that induction of an E. coli strain lysogenic for a A phage carrying the gal-attA region gave about a 2,000-fold increase in photoreactivating enzyme. Recently, however, Smith and Youngs (Mutat. Res., in press) and Sancar and Rupert (Mutat. Res., in press) were able to detect PR in cells with a deletion of the gal-attA region. They thus termed these strains Phr+ and, in addition, mapped a phr gene at 15.9 min, about 1 min away from the gal-attA region. We have thus reinvestigated cellular PR and photoreactivating enzyme in cells with deletions of the gal-attA region. We found that in these strains PR of UVinduced cell killing occurs at about 20% of the normal rate. Furthermore, these cells contained about 20% of the normal level of photoreactivat-
ing enzyme activity. We have characterized the residual enzyme activity to determine whether the reduced level of biological PR and of photoreactivating enzyme activity resulted from a decreased number of normal molecules (implying that the gene at 17 min is a regulatory gene) or from an altered enzyme (indicating a gene affecting the function of the enzyme). We found that the apparent Km for the enzyme is increased about two- to threefold in cells with the gal-attX deletion, and that the enzyme showed a much greater heat sensitivity than normal. These data indicated that the gene at 17 min contains information affecting the functional properties of the photoreactivating enzyme and, along with the data of Youngs and Smith (Mutat. Res., in press) and Sancar and Rupert (Mutat. Res., in press), indicated that there are (at least) two genes for PR in E. coli. We propose that the gene at 17 min (between gal and attA) be designed phrA, and that the one at 15.9 min be designatedphrB. MATERIALS AND METHODS Strains. E. coli K-12 SA206, SA224, SA244, and SA205, which were derivatives of W3350 obtained by growth on chlorate-containing agar, were gifts from Sankar Adhya, and had been determined by him to contain deletions in the chromosomal region near gal. SA456 and SA446, also the gift of Dr. Adhya, were derivatives of Hfr H, and contain deletions near gal. These deletions are shown in Fig. 1. The strains were checked for the galactose, phosphoglycerolactonase, 333
334
SUTHERLAND AND HAUSRATH Cell
Strain
SA 206
gal
pgl
chl D
att
-
X
bio
uvr B
-
-
-
I-
chi A
~~~~~~~~~~~~~~~~~~~~~~~.......
~~~~~~~~~~~~~~~~~~~~~~......
F
SA 224 SA
J. BACTERIOL.
244
SA 205
+
SA 446
+
SA
+
45 6
4
1
+
-
-
-
-
FIG. 1. Genetic map of E. coli in the region ofgal, adapted from Bachmann et al. (1). Open bars indicate extent of deletions in the various strains as determined by S. Adhya (personal communication); + and symbols indicate results of tests of the indicated markers by us. biotin, and uvrB markers as follows: gal, by the color of colonies on eosin methylene blue galactose plates (27.5 g of EMB agar base [Difco] plus 2% galactose; pgl, by the color of colonies grown on minimal maltose plates and flooded with an 0.1% I2-1.0% KI solution (5); bio, by the ability to grow in the absence of exogenous biotin (1.3 ,ug/ml); and uvrB, by UV survival. UV killing and biological PR. Cells were grown overnight in 10 ml of broth (10 g of tryptone [Difco] plus 8 g of NaCl per liter) at 37°C with shaking and centrifuged at 5,000 rpm for 20 min in an SS34 rotor of a Sorvall RC5. Cells were washed twice by suspension in 10 ml of 0.14 M NaCl and centrifugation as before. Cells were diluted into 0.14 M NaCl to a final concentration of 1 x 107 to 3 x 107 cells per ml, and exposed to 8.2 J/m2 (as measured by a Jagger meter [3] calibrated against a Yellow Springs Instruments radiometer) of UV light from a GE15T8 bulb with most of its emission at 254 nm. Cells were allowed to stand for 30 min (to allow formation of initial enzymedimer complex formation) before exposure to photoreactivating light (192.5 J/m2 per s) from Westinghouse BLB bulbs with their principal emission at 365 nm. Two 0.35-cm layers of window glass plus a glass petri dish cover were placed between the bulbs and the cells to absorb any light of wavelengths less than 300 nm. Samples were taken after 1 to 60 min of photoreactivating light exposure. After dilution in 0.14 M NaCl, samples were plated on modified LC plates (10 g of tryptone [Difco], 5 g of yeast extract, 2 g of sodium citrate, 5.8 g of NaCl, 0.55 g of CaCl2, 15 g of agar [Difco] per liter plus 2% glucose), and the plates were incubated at 37°C for 48 h. All manipulations were carried out by illumination of yellow Sylvania F40G0 fluorescent lamps. Measurement of photoreactivating enzyme. Photoreactivating enzyme activity was measured by the nuclease digestion method of Sutherland and Chamberlin (11). In brief, cell extracts were prepared by grinding 0.5 g of cells with 1.5 g of acid-washed alumina, suspending in 2.0 ml of PR mix (20 mM KPO4 buffer [pH 7.2]-0.1 mM dithiothreitol-0.1 mM EDTA), and centrifuging at 11,000 rpm for 15 min in an SE12 rotor of a Sorvall RC5 centrifuge. Samples
were added to duplicate tubes containing 0.2 ml of a mixture of PR mix containing 10 mM MgCl2, 65 mM NaCl, and highly purified 3P-labeled, T7 DNA containing pyrimidine dimers. One tube was kept in the dark (at 37°C), while the other was exposed to photoreactivating light from a Sylvania 150-W spot lamp. The temperature of the latter samples was kept at 37°C by a circulating water bath. After illumination, samples were digested to a mixture of nucleosides, inorganic phosphate, and dimer-containing oligonucleotides (10) by the sequential addition of 10 ,ug of deoxyribonuclease I, followed by 10 pl of 1 M Tris (pH 8.0), 100 zg of snake venom phosphodiesterase (Sigma V6875), and 1 ,ug of alkaline phosphatase (Sigma P4377), then digestion for 60 min at 37°C; The digestion was terminated by the addition of 10 !d of 0.46 M HCI, and the labeled oligonucleotides were separated from 32P by the addition of 1 ml of a 5% suspension of acid-washed Norit in neutral PPi solution (0.1 M dium PPi-0.1 M sodium phosphate, pH 6). After 15 min of adsorption on ice, samples were filtered over GF/C filters on an ice-cold filter block, washed three times with 3-ml portions of ice-cold PPi solution, and counted in a Baird Atomic planchet counter. Dimercontaining oligonucleotides were retained on the Norit, whereas mononucleosides and Pi were washed through the filter. PR of dimers was calculated by subtracting the amount of radioactivity on Norit in the Aample exposed to photoreactivating light from that on the sample kept in the dark. Protein was measured by the Biuret reaction (6) or by the method of Lowry et al. (7). Determinations of apparent Km values were made by measuring the initial reaction velocities at various DNA concentrations (9.5 x 10-8 M to 4.75 x 10-6 M in nucleoside phosphate); preliminary experiments indicated that similar results were obtained by using the same total nucleoside phosphate concentration and varying dimer content of the DNA. Km data were analyzed by the regression method of Wilkinson (15). Thermal lability of photoreactivating enzymes from the different cell strains was determined both by heating samples of the cell extracts adjusted to equal protein concentrations to 20, 37, 46, and 53°C for 10 min, or by heating to 50°C for various times with so-
PHOTOREACTIVATION IN E. COLI
VOL. 138, 1979
sequential removal of samples. Photoreactivating enzyme activity was then determined as described above.
RESULTS We first compared the rate of PR of UVinduced cell killing in E. coli K-12 strains which contained a deletion of the region gal-chiA (SA206, SA224, and SA244) with similar cells containing deletions from attX-chlA (SA456, SA205, SA446). To allow direct comparisons of photoreactivation rates, all strains tested were similar in dark repair capacity (see reference 2). Figure 2 shows that SA206 and SA244 photoreactivated survival at a rate about 20% of the normal cells. These data indicate that cells with deletions of the region gal-attA have a severe deficiency in their rate of biological PR. This deficiency might result from abnormal levels or structure of photoreactivating enzyme, or from reduced capacity of the celLs for PR, perhaps because of abnornal DNA configuration in which non-photoreactivable damage could be 1000
,
.*s
100
ioo O
0
o
/
0s
Photoreactivotion
Time
(min)
FIG. 2. Relative survival of PhrA+ (circles) or PhrA (triangles) strains of E. coli as a function of photoreactivation time from a black bulb at 192.5 JI m2per s. Relative survivals were calculated by dividing the fraction of the cells surviving after 8.2 J of 254-nm radiation per m2, followed by t minutes of photoreactivation by the fraction of the cells surviving the UV treatment alone. Symbols: PhrA' cells: 0, SA456; *, O), and 0, independent determinations for SA446; e and O, independent determinations of SA205; PhrA cells: A and A, independent determinations of SA206; V and V, independent tests of SA244.
335
produced by UV, or because of factors limiting access of repair enzymes to the lesions. We tested the latter possibility by examining cell survival after very long PR times, at which the reaction is virtually complete. Figure 2 shows that SA206 and SA446 have the same capacity for PR, and that the apparent deficiency in rate of PR results from a change in photoreactivating enzyme level or in its function. We attempted to purify the enzyme present in SA224, SA244, and SA206, but encountered two major problems. First, in Phr+ E. coli there are very few (-20) photoreactivating enzyme molecules (4); in PhrA cells, the activity was reduced to about 20% of that value, making all purifications difficult. Second, the photoreactivating activity in the PhrA cells was unstable. Although the activity could be followed through streptomycin and ammonium sulfate precipitation, after 24 h the activity was no longer detectable. (Activity from Phr+ cells, or from W3350 [Xphr] [12], is stable at this stage of purification for months.) We thus examined photoreactivating enzyme activity in extracts of SA206, SA224, SA244, SA205, and SA446. Table 1 shows the result of five experiments designed to examine photoreactivating enzyme activity in these cells. All data are normnalized to SA446 as 100%; these experiments indicate that strains with deletions of the gal-attA region contain about 20% of the normal level of photoreactivating enzyme. It was possible that the cells with deficiencies in photoreactivating enzyme activities contained an inhibitor which might mask photoreactivating enzyme present in the extracts. We tested this possibility by adding large quantities (0.4 mg) of cell extracts of SA446 or SA244 to about 0.3 to 2 ug of partially purified E. coli photoreactivating enzyme (12). Table 2 shows that neither extract contained inhibitors of photoreactivating enzyme; in fact, the activities in the extracts were additive with the activity in the E. coli photoreactivating enzyme preparation. These data indicate that the depressed photoreactivating enzyme activity seen in vivo and in vitro does not result from the presence of an inhibitor of the enzyme but from a change in the enzyme itself. Is this decrease in biological photoreactivation and in photoreactivating enzyme activity due to a decreased number of normal photoreactivating enzyme molecules (implying that the gene in the gal-attA region is a regul,atory gene, perhaps a positive control elemetit),\ or in an altered photoreactivating enzyme (implying that the gene product of the phr gene located in the gal-attA region affects the function of the enzyme)? We have compared the properties of the enzyme present in SA206, SA224, and SA244 with that
336
J. BACTERIOL.
SUTHERLAND AND HAUSRATH
TABLE 1. Relative specific activities of photoreactivating enzyme in five E. coli strainsa
have similar properties with regard to PR. We have also examined the thermal lability of Relative sp act (%) of photoreactivating enzyme in the enzyme in PhrA+ and PhrA cells in two related experimental series. In the first series, strain: Expt of extract at equal protein concentrasamples SA206 SA244 SA446 SA205 SA224 tions from each cell type were heated to 20, 37, 17.2 1 21.9 100 46, or 53°C for 10 min. Table 4 shows that 19.6 2 100 photoreactivating enzyme from SA206 was con24.6 100 3 more heat labile than that of SA446 siderably 19.2 95 4 100 over all the temperature range tested. The sec22.2 22.0 97 100 5 ond type of experiment was designed to examine a All values are normalized using the specific activity the thermal lability of the enzymes at a single of SA446 as 100%. Strains SA446 and SA205 did not temperature (50°C). Figure 3 shows that the have deletions of the gal-attA region; strains SA224, enzyme from SA206 lost its activity much more SA244, and SA206 had deletions of the gal-attA region. rapidly at 50°C than did that from SA446. A TABLE 2. Photoreactivating activity from purified comparison of Fig. 3 and Table 4 indicates that the data obtained in the first type of experiment E. coli photoreactivating enzyme, cell extracts, or agreed well with that obtained in the second; mixture of enzyme and cell extracts Extract added
Amt (Jlg)
Photoreactivating activitya ob
0.64
1.32b
2.64b
394 633 1,269 0 None 400 1,503 1,819 2,343 2,861 SA206 400 SA446 4,905 5,333 5,365 6,462 a Photoreactivating activities are given in counts per minute of nuclease-resistant, dimer-containing oligonucleotides photoreactivated in a standard 30-min assay. bE. coli photoreactivating enzyme in micrograms.
in SA205 and SA446. Table 3 summarizes the results of determinations of the apparent Km of photoreactivating enzymes from cells with deletions of the gal-attA region, and those in which this chromosomal region is present. The enzyme in the strains with the gal-attA deletion had about a two- to threefold-greater apparent Km than did the enzyme from normal cells. This may indicate that the enzyme in these cells has a reduced affinity for its substrate, pyrimidine dimers in DNA. We have examined photoreactivation in four E. coli K-12 W3350 derivatives (SA205, SA244, SA224 and SA206). In these cells, decreased PR rate and decreased enzyme activity were associated with a deletion of the phr gene at 17 min. Since SA205 is the only one among these to be PhrA+, and might be in some way unusual, we also examined SA446 and SA456, HfrH derivatives with similar deletions to SA205. Tables 1 and 3 indicate that SA446 and SA205 were similar in specific activity and in apparent Km of photoreactivating enzyme. (We consistently observed a higher apparent Km in SA244 than in other PhrA W3350 derivatives.) Figure 2 shows that SA456, SA446, and SA205 carried out biological photoreactivation at about the same rate. We thus conclude that for these cells under our conditions, the W3350 and HfrH derivatives
TABLE 3. Km (apparent) values for photoreactivating enzyme in cell extracts of five E. coli strains Strain
Deletion of gal-att
SA446
No
Apparent Km 0.124
0.122 0.080 0.099
SA205
No
0.084 0.093
SA206
Yes
0.248 0.282
SA224
Yes
0.293 0.350
0.406 0.650 a Values of Km (apparent) are given in micromolar for DNA phosphate in nuclease-resistant sequences. These result from the presence of dimers which render the nucleotide phosphate bond resistant to enzymatic digestion (10, 11); there are approximately 2.3 nuclease-resistant phosphates per dimer-containing oligonucleotide (10).
SA244
Yes
TABLE 4. Thernal lability ofphotoreactivating enzyme activity in two E. coli strains Temp
(OC)
Relative enzyme activity remaining in cell extracts' of:
SA206 100 72 17 0.9 'Extracts of SA446 (PhrA+) and SA206 (PhrA), adjusted to 0.25 mg/ml, were heated to the stated temperatures for 10 min and then assayed as usual. 20 37 46 53
SA446 100 86.4 77.4 16.3
PHOTOREACTIVATION IN E. COLI
VOL. 138, 1979
the data in Table 4 would predict that, after 10 min at 50°C, about 50 and 8% of the starting activities of SA446 and SA206 should have survived, respectively. Figure 3 shows that 10 min at 500C actually left 57 and 12% of the starting activities of SA446 and SA206, respectively. Both the Km and the thermal lability experiments indicate that the decreased photoreactivating enzyme activities and biological PR result from an alteration of the photoreactivating enzyme in cells with deletion of the gal-attA region of their genome. DISCUSSION Our data clearly indicate that E. coli strains carrying deletions of the chromosome from gal to attX suffered about an 80% decrease in rate of photoreactivation, whether measured in vivo or in vitro. This loss was not due to decreased capacity of the cells to be photoreactivated, or to the presence of inhibitors in the cell extract. Measurement of kinetic properties of photoreactivating enzymes from the cells and of thermal lability of the enzymes indicates that there was a functionally altered photoreactivating enzyme in the cells with the gal-attA deletion. Thus the phr gene in the gal-attX region contains structural information affecting the function of the photoreactivating enzyme. Recently, Youngs and Smith (Mutat. Res., in press) have examined PR in cells with deletions of the gal-attA region, and have concluded that they are Phr+. Their method was to carry out UV survival experiments in the presence and absence of 20 min of photoreactivating light. Although they did not state the type of lamp or
337
light flux used in their experiments, if they were similar to those used by us, the data in Fig. 2 indicate that the PR reaction might be nearing completion and, indeed, little if any difference could be detected between cells missing gal-attA region (SA206) and those with that region intact (SA446, SA456). They also state that they did not find a difference in PR rate, but they did not present their data. However, Rupert found a decrease in rate of PR of T4 phage in SA206 relative to normal (personal communication). Sancar and Rupert (Mutat. Res., in press) have used 50 min of photoreactivating light illumination from "daylight" fluorescent lamps in studying PR of SA206 and T4 phage in SA206. They found that SA206, which carried the gal-attA deletion, can photoreactivate both cell and phage killing. Thus, the data obtained by Sutherland et al. (20), our current data, and those presented by Youngs and Smith (Mutat. Res., in press), Rupert (personal communication), and Sancar and Rupert (Mutat. Res., in press) indicate that cells with deletions of the gal-attA region can photoreactivate dimers in their own or in exogenous DNAs, but at a reduced rate. We propose that the simplest explanation of all these data is that there are at least two genes in E. coli for the photoreactivating enzyme. We propose that the gene at 17 min be termedphrA, and that the one at 15.9 min be called phrB. ACKNOWLEDGMENTS We thank Jane Setlow, Robert Rothman, John Sutherland, and S. G. Oliver for helpful discussions, and Sankar Adhya for supplying strains. This research was supported by Public Health Service grant CA 23096 from the National Cancer Institute to B.M.S., by a Public Health Service Research Career Development Award (CA00466) from the National Cancer Institute to B.M.S., and by the U. S. Department of Energy. LITERATURE CIMD 1. Bachmann, B. J., K. B. Low, and A. L. Taylor. 1976. Recalibrated linkage map of Escherichia coli K-12.
0
Time
at
5 50- (ml n)
10
FIG. 3. Percent of initial photoreactivating enremaining in cell extracts of SA446 (PhrA@) and SA206 (PhrA) (both adjusted to 0.24 mg/ml), after treatment at 50°C for varying times.
zyme
Bacteriol. Rev. 40:116-167. 2. Harm, W., C. S. Rupert, and H. Harm. 1971. Photoenzymatic repair of DNA. I. Investigation of the research by flash illumination, p. 53-63. In R. F. Beers, R. M. Herriott, and R. C. Tilghman (ed.), Molecular and cellular repair processes. University Press, Baltimore. 3. Jagger, J. 1961. A small and inexpensive ultraviolet doserate meter useful in biological experiments. Radiat. Res. 14:394-403. 4. Kondo, S., and T. Kato. 1966. Action spectra for photoreactivation of killing and mutation to prototrophy in UV-sensitive strains of Escherichia coli possessing and lacking photoreactivating enzyme. Photochem. Photobiol. 5:827437. 5. Kupor, S. R., and D. G. Fraenkel. 1969. 6-Phosphogluconolactonase mutants of Escherichia coli and a maltose blue gene. J. Bacteriol. 100:1296-1301. 6. Layne, E. 1957. Spectrophotometric and turbidimetric methods for measuring proteins. Methods Enzymol. 3: 447-454.
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SUTHERLAND AND HAUSRATH
7. Lowry, O. H., N. J. Rosebrough, A. L Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. 8. Setlow, J. K. 1966. The molecular basis of biological effects of ultraviolet radiation and photoreactivation, p. 195-248. In M. Ebert and A. Howard (ed.), Topics in radiation research, vol. 2. North Holland, Amsterdam. 9. Setlow, R. B. 1966. Cyclobutane-type pyrimidine dimers in polynucleotides. Science 153:379-386. 10. Setlow, R. B., W. L Carrier, and F. J. Bollum. 1964. Nuclease-resistance sequences in ultraviolet-irradiated deoxyribonucleic acid. Biochim. Biophys. Acta 91:446461. 11. Sutherland, B. M., and M. J. Chamberlin. 1973. A rapid and sensitive assay for pyrimidine dimers in DNA.
J. BACTERIOL. Anal. Biochem. 53:168-176. 12. Sutherland, B. M., M. J. Chamberlin, and J. C. Sutherland. 1973. Deoxyribonucleic acid photoreactivating enzyme from Escherichia coli. J. Biol. Chem. 12:42004205. 13. Sutherland, B. M., D. Court, and M. J. Chamberlin. 1972. Studies on the DNA photoreactivating enzyme from Escherichia coli. 1. Transduction of the phr gene by bacteriophage lambda. Virology 48:87-93. 14. Van de Putte, P., C. A. Van Sluis, J. Van Dillewijn, and A. Rorsch. 1965. The location of genes controlling radiation sensitivity in Escherichia coli. Mutat. Res. 2: 97-110. 15. Wilkinson, G. N. 1961. Statistical estimations in enzyme kinetics. Biochem. J. 80:324-332.
JOURNAL OF BACTERIOLOGY, May 1979, p. 333-338 0021-9193/79/05-0333/06$02.00/0
Multiple Loci Affecting Photoreactivation in Escherichia coli BETSY M. SUTHERLAND* AND STEPHEN G. HAUSRATH Biology Department, Brookhaven National Laboratory, Upton, New York 11973
Received for publication 6 February 1979
Sutherland et al. mapped a phr gene in Escherichia coli at 17 min and found that induction of an E. coli strain lysogenic for a A phage carrying this gene increased photoreactivating enzyme levels 2,000-fold. Recently, Smith and Youngs and Sancar and Rupert located a phr gene at 15.9 min. We have therefore investigated the properties of photoreactivating enzyme and cellular photoreactivation in cells containing deletions of the gene at 17 min. Cells with this deletion photoreactivated ultraviolet-induced killing at a rate 20% of normal; they also contained approximately 20% of the normal photoreactivating enzyme level. The residual enzyme in these cells was characterized to determine whether the reduced cellular photoreactivation rate and photoreactivating enzyme levels resulted from reduced numbers of normal enzymes or from an altered enzyme. Photoreactivating enzymes from strains carrying a deletion of the region at 17 min had an apparent Km about two- to threefold higher than normal enzyme and showed markedly increased heat lability. The gene at 17 min thus contains information determining the function of the E. coli photoreactivating enzyme rather than the quantity of the enzyme. It is proposed that the gene at 17 min be termed phrA and that located at 15.9 min be termed phrB. UV radiation produces cyclobutane-type pyrimidine dimers in DNA (9). A major pathway for repair of these dimers is photoreactivation (PR), in which a photoreactivating enzyme binds to a dimer-containing region of DNA and, in the presence of light of longer wavelengths (300 to 500 nm), monomerizes the dimer (8). Van de Putte et al. first mapped the phr gene in Escherichia coli to be near gal (14). Subsequently Sutherland et al. located a phr gene between gal and attA by comparing PR of UV-induced killing in strains with known deletions near gal (13). Strains with deletions from gal to attA showed greatly decreased PR of cell survival. They also showed that induction of an E. coli strain lysogenic for a A phage carrying the gal-attA region gave about a 2,000-fold increase in photoreactivating enzyme. Recently, however, Smith and Youngs (Mutat. Res., in press) and Sancar and Rupert (Mutat. Res., in press) were able to detect PR in cells with a deletion of the gal-attA region. They thus termed these strains Phr+ and, in addition, mapped a phr gene at 15.9 min, about 1 min away from the gal-attA region. We have thus reinvestigated cellular PR and photoreactivating enzyme in cells with deletions of the gal-attA region. We found that in these strains PR of UVinduced cell killing occurs at about 20% of the normal rate. Furthermore, these cells contained about 20% of the normal level of photoreactivat-
ing enzyme activity. We have characterized the residual enzyme activity to determine whether the reduced level of biological PR and of photoreactivating enzyme activity resulted from a decreased number of normal molecules (implying that the gene at 17 min is a regulatory gene) or from an altered enzyme (indicating a gene affecting the function of the enzyme). We found that the apparent Km for the enzyme is increased about two- to threefold in cells with the gal-attX deletion, and that the enzyme showed a much greater heat sensitivity than normal. These data indicated that the gene at 17 min contains information affecting the functional properties of the photoreactivating enzyme and, along with the data of Youngs and Smith (Mutat. Res., in press) and Sancar and Rupert (Mutat. Res., in press), indicated that there are (at least) two genes for PR in E. coli. We propose that the gene at 17 min (between gal and attA) be designed phrA, and that the one at 15.9 min be designatedphrB. MATERIALS AND METHODS Strains. E. coli K-12 SA206, SA224, SA244, and SA205, which were derivatives of W3350 obtained by growth on chlorate-containing agar, were gifts from Sankar Adhya, and had been determined by him to contain deletions in the chromosomal region near gal. SA456 and SA446, also the gift of Dr. Adhya, were derivatives of Hfr H, and contain deletions near gal. These deletions are shown in Fig. 1. The strains were checked for the galactose, phosphoglycerolactonase, 333
334
SUTHERLAND AND HAUSRATH Cell
Strain
SA 206
gal
pgl
chl D
att
-
X
bio
uvr B
-
-
-
I-
chi A
~~~~~~~~~~~~~~~~~~~~~~~.......
~~~~~~~~~~~~~~~~~~~~~~......
F
SA 224 SA
J. BACTERIOL.
244
SA 205
+
SA 446
+
SA
+
45 6
4
1
+
-
-
-
-
FIG. 1. Genetic map of E. coli in the region ofgal, adapted from Bachmann et al. (1). Open bars indicate extent of deletions in the various strains as determined by S. Adhya (personal communication); + and symbols indicate results of tests of the indicated markers by us. biotin, and uvrB markers as follows: gal, by the color of colonies on eosin methylene blue galactose plates (27.5 g of EMB agar base [Difco] plus 2% galactose; pgl, by the color of colonies grown on minimal maltose plates and flooded with an 0.1% I2-1.0% KI solution (5); bio, by the ability to grow in the absence of exogenous biotin (1.3 ,ug/ml); and uvrB, by UV survival. UV killing and biological PR. Cells were grown overnight in 10 ml of broth (10 g of tryptone [Difco] plus 8 g of NaCl per liter) at 37°C with shaking and centrifuged at 5,000 rpm for 20 min in an SS34 rotor of a Sorvall RC5. Cells were washed twice by suspension in 10 ml of 0.14 M NaCl and centrifugation as before. Cells were diluted into 0.14 M NaCl to a final concentration of 1 x 107 to 3 x 107 cells per ml, and exposed to 8.2 J/m2 (as measured by a Jagger meter [3] calibrated against a Yellow Springs Instruments radiometer) of UV light from a GE15T8 bulb with most of its emission at 254 nm. Cells were allowed to stand for 30 min (to allow formation of initial enzymedimer complex formation) before exposure to photoreactivating light (192.5 J/m2 per s) from Westinghouse BLB bulbs with their principal emission at 365 nm. Two 0.35-cm layers of window glass plus a glass petri dish cover were placed between the bulbs and the cells to absorb any light of wavelengths less than 300 nm. Samples were taken after 1 to 60 min of photoreactivating light exposure. After dilution in 0.14 M NaCl, samples were plated on modified LC plates (10 g of tryptone [Difco], 5 g of yeast extract, 2 g of sodium citrate, 5.8 g of NaCl, 0.55 g of CaCl2, 15 g of agar [Difco] per liter plus 2% glucose), and the plates were incubated at 37°C for 48 h. All manipulations were carried out by illumination of yellow Sylvania F40G0 fluorescent lamps. Measurement of photoreactivating enzyme. Photoreactivating enzyme activity was measured by the nuclease digestion method of Sutherland and Chamberlin (11). In brief, cell extracts were prepared by grinding 0.5 g of cells with 1.5 g of acid-washed alumina, suspending in 2.0 ml of PR mix (20 mM KPO4 buffer [pH 7.2]-0.1 mM dithiothreitol-0.1 mM EDTA), and centrifuging at 11,000 rpm for 15 min in an SE12 rotor of a Sorvall RC5 centrifuge. Samples
were added to duplicate tubes containing 0.2 ml of a mixture of PR mix containing 10 mM MgCl2, 65 mM NaCl, and highly purified 3P-labeled, T7 DNA containing pyrimidine dimers. One tube was kept in the dark (at 37°C), while the other was exposed to photoreactivating light from a Sylvania 150-W spot lamp. The temperature of the latter samples was kept at 37°C by a circulating water bath. After illumination, samples were digested to a mixture of nucleosides, inorganic phosphate, and dimer-containing oligonucleotides (10) by the sequential addition of 10 ,ug of deoxyribonuclease I, followed by 10 pl of 1 M Tris (pH 8.0), 100 zg of snake venom phosphodiesterase (Sigma V6875), and 1 ,ug of alkaline phosphatase (Sigma P4377), then digestion for 60 min at 37°C; The digestion was terminated by the addition of 10 !d of 0.46 M HCI, and the labeled oligonucleotides were separated from 32P by the addition of 1 ml of a 5% suspension of acid-washed Norit in neutral PPi solution (0.1 M dium PPi-0.1 M sodium phosphate, pH 6). After 15 min of adsorption on ice, samples were filtered over GF/C filters on an ice-cold filter block, washed three times with 3-ml portions of ice-cold PPi solution, and counted in a Baird Atomic planchet counter. Dimercontaining oligonucleotides were retained on the Norit, whereas mononucleosides and Pi were washed through the filter. PR of dimers was calculated by subtracting the amount of radioactivity on Norit in the Aample exposed to photoreactivating light from that on the sample kept in the dark. Protein was measured by the Biuret reaction (6) or by the method of Lowry et al. (7). Determinations of apparent Km values were made by measuring the initial reaction velocities at various DNA concentrations (9.5 x 10-8 M to 4.75 x 10-6 M in nucleoside phosphate); preliminary experiments indicated that similar results were obtained by using the same total nucleoside phosphate concentration and varying dimer content of the DNA. Km data were analyzed by the regression method of Wilkinson (15). Thermal lability of photoreactivating enzymes from the different cell strains was determined both by heating samples of the cell extracts adjusted to equal protein concentrations to 20, 37, 46, and 53°C for 10 min, or by heating to 50°C for various times with so-
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VOL. 138, 1979
sequential removal of samples. Photoreactivating enzyme activity was then determined as described above.
RESULTS We first compared the rate of PR of UVinduced cell killing in E. coli K-12 strains which contained a deletion of the region gal-chiA (SA206, SA224, and SA244) with similar cells containing deletions from attX-chlA (SA456, SA205, SA446). To allow direct comparisons of photoreactivation rates, all strains tested were similar in dark repair capacity (see reference 2). Figure 2 shows that SA206 and SA244 photoreactivated survival at a rate about 20% of the normal cells. These data indicate that cells with deletions of the region gal-attA have a severe deficiency in their rate of biological PR. This deficiency might result from abnormal levels or structure of photoreactivating enzyme, or from reduced capacity of the celLs for PR, perhaps because of abnornal DNA configuration in which non-photoreactivable damage could be 1000
,
.*s
100
ioo O
0
o
/
0s
Photoreactivotion
Time
(min)
FIG. 2. Relative survival of PhrA+ (circles) or PhrA (triangles) strains of E. coli as a function of photoreactivation time from a black bulb at 192.5 JI m2per s. Relative survivals were calculated by dividing the fraction of the cells surviving after 8.2 J of 254-nm radiation per m2, followed by t minutes of photoreactivation by the fraction of the cells surviving the UV treatment alone. Symbols: PhrA' cells: 0, SA456; *, O), and 0, independent determinations for SA446; e and O, independent determinations of SA205; PhrA cells: A and A, independent determinations of SA206; V and V, independent tests of SA244.
335
produced by UV, or because of factors limiting access of repair enzymes to the lesions. We tested the latter possibility by examining cell survival after very long PR times, at which the reaction is virtually complete. Figure 2 shows that SA206 and SA446 have the same capacity for PR, and that the apparent deficiency in rate of PR results from a change in photoreactivating enzyme level or in its function. We attempted to purify the enzyme present in SA224, SA244, and SA206, but encountered two major problems. First, in Phr+ E. coli there are very few (-20) photoreactivating enzyme molecules (4); in PhrA cells, the activity was reduced to about 20% of that value, making all purifications difficult. Second, the photoreactivating activity in the PhrA cells was unstable. Although the activity could be followed through streptomycin and ammonium sulfate precipitation, after 24 h the activity was no longer detectable. (Activity from Phr+ cells, or from W3350 [Xphr] [12], is stable at this stage of purification for months.) We thus examined photoreactivating enzyme activity in extracts of SA206, SA224, SA244, SA205, and SA446. Table 1 shows the result of five experiments designed to examine photoreactivating enzyme activity in these cells. All data are normnalized to SA446 as 100%; these experiments indicate that strains with deletions of the gal-attA region contain about 20% of the normal level of photoreactivating enzyme. It was possible that the cells with deficiencies in photoreactivating enzyme activities contained an inhibitor which might mask photoreactivating enzyme present in the extracts. We tested this possibility by adding large quantities (0.4 mg) of cell extracts of SA446 or SA244 to about 0.3 to 2 ug of partially purified E. coli photoreactivating enzyme (12). Table 2 shows that neither extract contained inhibitors of photoreactivating enzyme; in fact, the activities in the extracts were additive with the activity in the E. coli photoreactivating enzyme preparation. These data indicate that the depressed photoreactivating enzyme activity seen in vivo and in vitro does not result from the presence of an inhibitor of the enzyme but from a change in the enzyme itself. Is this decrease in biological photoreactivation and in photoreactivating enzyme activity due to a decreased number of normal photoreactivating enzyme molecules (implying that the gene in the gal-attA region is a regul,atory gene, perhaps a positive control elemetit),\ or in an altered photoreactivating enzyme (implying that the gene product of the phr gene located in the gal-attA region affects the function of the enzyme)? We have compared the properties of the enzyme present in SA206, SA224, and SA244 with that
336
J. BACTERIOL.
SUTHERLAND AND HAUSRATH
TABLE 1. Relative specific activities of photoreactivating enzyme in five E. coli strainsa
have similar properties with regard to PR. We have also examined the thermal lability of Relative sp act (%) of photoreactivating enzyme in the enzyme in PhrA+ and PhrA cells in two related experimental series. In the first series, strain: Expt of extract at equal protein concentrasamples SA206 SA244 SA446 SA205 SA224 tions from each cell type were heated to 20, 37, 17.2 1 21.9 100 46, or 53°C for 10 min. Table 4 shows that 19.6 2 100 photoreactivating enzyme from SA206 was con24.6 100 3 more heat labile than that of SA446 siderably 19.2 95 4 100 over all the temperature range tested. The sec22.2 22.0 97 100 5 ond type of experiment was designed to examine a All values are normalized using the specific activity the thermal lability of the enzymes at a single of SA446 as 100%. Strains SA446 and SA205 did not temperature (50°C). Figure 3 shows that the have deletions of the gal-attA region; strains SA224, enzyme from SA206 lost its activity much more SA244, and SA206 had deletions of the gal-attA region. rapidly at 50°C than did that from SA446. A TABLE 2. Photoreactivating activity from purified comparison of Fig. 3 and Table 4 indicates that the data obtained in the first type of experiment E. coli photoreactivating enzyme, cell extracts, or agreed well with that obtained in the second; mixture of enzyme and cell extracts Extract added
Amt (Jlg)
Photoreactivating activitya ob
0.64
1.32b
2.64b
394 633 1,269 0 None 400 1,503 1,819 2,343 2,861 SA206 400 SA446 4,905 5,333 5,365 6,462 a Photoreactivating activities are given in counts per minute of nuclease-resistant, dimer-containing oligonucleotides photoreactivated in a standard 30-min assay. bE. coli photoreactivating enzyme in micrograms.
in SA205 and SA446. Table 3 summarizes the results of determinations of the apparent Km of photoreactivating enzymes from cells with deletions of the gal-attA region, and those in which this chromosomal region is present. The enzyme in the strains with the gal-attA deletion had about a two- to threefold-greater apparent Km than did the enzyme from normal cells. This may indicate that the enzyme in these cells has a reduced affinity for its substrate, pyrimidine dimers in DNA. We have examined photoreactivation in four E. coli K-12 W3350 derivatives (SA205, SA244, SA224 and SA206). In these cells, decreased PR rate and decreased enzyme activity were associated with a deletion of the phr gene at 17 min. Since SA205 is the only one among these to be PhrA+, and might be in some way unusual, we also examined SA446 and SA456, HfrH derivatives with similar deletions to SA205. Tables 1 and 3 indicate that SA446 and SA205 were similar in specific activity and in apparent Km of photoreactivating enzyme. (We consistently observed a higher apparent Km in SA244 than in other PhrA W3350 derivatives.) Figure 2 shows that SA456, SA446, and SA205 carried out biological photoreactivation at about the same rate. We thus conclude that for these cells under our conditions, the W3350 and HfrH derivatives
TABLE 3. Km (apparent) values for photoreactivating enzyme in cell extracts of five E. coli strains Strain
Deletion of gal-att
SA446
No
Apparent Km 0.124
0.122 0.080 0.099
SA205
No
0.084 0.093
SA206
Yes
0.248 0.282
SA224
Yes
0.293 0.350
0.406 0.650 a Values of Km (apparent) are given in micromolar for DNA phosphate in nuclease-resistant sequences. These result from the presence of dimers which render the nucleotide phosphate bond resistant to enzymatic digestion (10, 11); there are approximately 2.3 nuclease-resistant phosphates per dimer-containing oligonucleotide (10).
SA244
Yes
TABLE 4. Thernal lability ofphotoreactivating enzyme activity in two E. coli strains Temp
(OC)
Relative enzyme activity remaining in cell extracts' of:
SA206 100 72 17 0.9 'Extracts of SA446 (PhrA+) and SA206 (PhrA), adjusted to 0.25 mg/ml, were heated to the stated temperatures for 10 min and then assayed as usual. 20 37 46 53
SA446 100 86.4 77.4 16.3
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VOL. 138, 1979
the data in Table 4 would predict that, after 10 min at 50°C, about 50 and 8% of the starting activities of SA446 and SA206 should have survived, respectively. Figure 3 shows that 10 min at 500C actually left 57 and 12% of the starting activities of SA446 and SA206, respectively. Both the Km and the thermal lability experiments indicate that the decreased photoreactivating enzyme activities and biological PR result from an alteration of the photoreactivating enzyme in cells with deletion of the gal-attA region of their genome. DISCUSSION Our data clearly indicate that E. coli strains carrying deletions of the chromosome from gal to attX suffered about an 80% decrease in rate of photoreactivation, whether measured in vivo or in vitro. This loss was not due to decreased capacity of the cells to be photoreactivated, or to the presence of inhibitors in the cell extract. Measurement of kinetic properties of photoreactivating enzymes from the cells and of thermal lability of the enzymes indicates that there was a functionally altered photoreactivating enzyme in the cells with the gal-attA deletion. Thus the phr gene in the gal-attX region contains structural information affecting the function of the photoreactivating enzyme. Recently, Youngs and Smith (Mutat. Res., in press) have examined PR in cells with deletions of the gal-attA region, and have concluded that they are Phr+. Their method was to carry out UV survival experiments in the presence and absence of 20 min of photoreactivating light. Although they did not state the type of lamp or
337
light flux used in their experiments, if they were similar to those used by us, the data in Fig. 2 indicate that the PR reaction might be nearing completion and, indeed, little if any difference could be detected between cells missing gal-attA region (SA206) and those with that region intact (SA446, SA456). They also state that they did not find a difference in PR rate, but they did not present their data. However, Rupert found a decrease in rate of PR of T4 phage in SA206 relative to normal (personal communication). Sancar and Rupert (Mutat. Res., in press) have used 50 min of photoreactivating light illumination from "daylight" fluorescent lamps in studying PR of SA206 and T4 phage in SA206. They found that SA206, which carried the gal-attA deletion, can photoreactivate both cell and phage killing. Thus, the data obtained by Sutherland et al. (20), our current data, and those presented by Youngs and Smith (Mutat. Res., in press), Rupert (personal communication), and Sancar and Rupert (Mutat. Res., in press) indicate that cells with deletions of the gal-attA region can photoreactivate dimers in their own or in exogenous DNAs, but at a reduced rate. We propose that the simplest explanation of all these data is that there are at least two genes in E. coli for the photoreactivating enzyme. We propose that the gene at 17 min be termedphrA, and that the one at 15.9 min be called phrB. ACKNOWLEDGMENTS We thank Jane Setlow, Robert Rothman, John Sutherland, and S. G. Oliver for helpful discussions, and Sankar Adhya for supplying strains. This research was supported by Public Health Service grant CA 23096 from the National Cancer Institute to B.M.S., by a Public Health Service Research Career Development Award (CA00466) from the National Cancer Institute to B.M.S., and by the U. S. Department of Energy. LITERATURE CIMD 1. Bachmann, B. J., K. B. Low, and A. L. Taylor. 1976. Recalibrated linkage map of Escherichia coli K-12.
0
Time
at
5 50- (ml n)
10
FIG. 3. Percent of initial photoreactivating enremaining in cell extracts of SA446 (PhrA@) and SA206 (PhrA) (both adjusted to 0.24 mg/ml), after treatment at 50°C for varying times.
zyme
Bacteriol. Rev. 40:116-167. 2. Harm, W., C. S. Rupert, and H. Harm. 1971. Photoenzymatic repair of DNA. I. Investigation of the research by flash illumination, p. 53-63. In R. F. Beers, R. M. Herriott, and R. C. Tilghman (ed.), Molecular and cellular repair processes. University Press, Baltimore. 3. Jagger, J. 1961. A small and inexpensive ultraviolet doserate meter useful in biological experiments. Radiat. Res. 14:394-403. 4. Kondo, S., and T. Kato. 1966. Action spectra for photoreactivation of killing and mutation to prototrophy in UV-sensitive strains of Escherichia coli possessing and lacking photoreactivating enzyme. Photochem. Photobiol. 5:827437. 5. Kupor, S. R., and D. G. Fraenkel. 1969. 6-Phosphogluconolactonase mutants of Escherichia coli and a maltose blue gene. J. Bacteriol. 100:1296-1301. 6. Layne, E. 1957. Spectrophotometric and turbidimetric methods for measuring proteins. Methods Enzymol. 3: 447-454.
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7. Lowry, O. H., N. J. Rosebrough, A. L Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. 8. Setlow, J. K. 1966. The molecular basis of biological effects of ultraviolet radiation and photoreactivation, p. 195-248. In M. Ebert and A. Howard (ed.), Topics in radiation research, vol. 2. North Holland, Amsterdam. 9. Setlow, R. B. 1966. Cyclobutane-type pyrimidine dimers in polynucleotides. Science 153:379-386. 10. Setlow, R. B., W. L Carrier, and F. J. Bollum. 1964. Nuclease-resistance sequences in ultraviolet-irradiated deoxyribonucleic acid. Biochim. Biophys. Acta 91:446461. 11. Sutherland, B. M., and M. J. Chamberlin. 1973. A rapid and sensitive assay for pyrimidine dimers in DNA.
J. BACTERIOL. Anal. Biochem. 53:168-176. 12. Sutherland, B. M., M. J. Chamberlin, and J. C. Sutherland. 1973. Deoxyribonucleic acid photoreactivating enzyme from Escherichia coli. J. Biol. Chem. 12:42004205. 13. Sutherland, B. M., D. Court, and M. J. Chamberlin. 1972. Studies on the DNA photoreactivating enzyme from Escherichia coli. 1. Transduction of the phr gene by bacteriophage lambda. Virology 48:87-93. 14. Van de Putte, P., C. A. Van Sluis, J. Van Dillewijn, and A. Rorsch. 1965. The location of genes controlling radiation sensitivity in Escherichia coli. Mutat. Res. 2: 97-110. 15. Wilkinson, G. N. 1961. Statistical estimations in enzyme kinetics. Biochem. J. 80:324-332.
