A new method of assay for polynucleotide ligase

452 Biochimica et Biophysica A cta, 340 (1974) 452--462 Q Elsevier Scientific Publishing Company, Amsterdam -- Printed in The Netherlands

BBA 97966 A NEW METHOD OF ASSAY F O R POLYNUCLEOTIDE LIGASE

JOHN D. KARKAS Cell Chemistry Laboratory, Department of Biochemistry, College of Physicians and Surgeons, Columbia University, New York, N.Y. 10032 (U.S.A.)

(Received October 29th, 1973)

Summary A new m e t h o d for the assay of polynucleotide ligase is described. It consists of two parallel DNA polymerase reactions with poly(dA) • oligo(dT) as template-primer and d[ 3 HI TTP as precursor; ligase is added to both, but one is supplemented with NMN and the other with NAD. In the presence of NAD, ligase joins several oligo(dT) molecules, thus reducing the number of primer sites available to the polymerase, while the NMN-supplemented reaction serves as a control. The difference in dTMP incorporation between the two assays provides a measure of the activity of the ligase. The m e t h o d is simple, rapid, and sensitive and can be used for the comparison of ligase levels in crude cell extracts or for monitoring ligase activity in the course of the purification of the enzyme.

Introduction The joining of discontinuous or broken DNA chains by polynucleotide ligases plays a central role in a number of important biological processes, including chromosome replication, genetic recombination and repair of radiation damage. Although several techniques are available for the assay of ligase activity [1--11], almost all present some serious difficulties, such as elaborate preparation of the substrates, time consuming determination of the products, dependence on the purity of the enzymes used as tools, etc. This communication describes a new and rather simple m e t h o d for the assay of polynucleotide ligase. All of the required substrates and enzymes are commercially available; the m e t h o d is fast and sensitive; and it includes a built-in control which renders it rather specific even in crude extracts. Materials P o l y n u c l e o t i d e s . Poly(dA) and poly(dT) were synthesized with terminal

453 deoxynucleotidyl transferase (EC 2.7.7.31) [12--13] according to Bollum [14]. Oligo(dT). Oligo(dT) was prepared by treatment of poly(dT) with deoxyribonuclease (EC 3.1.4.5): 240 #g of poly(dT) were incubated with 200 pg of pancreatic deoxyribonuclease (Worthington, DPFF, electrophoretically purified) in 1.2 ml of 0.05 M Tris--HC1 (pH 7.5)--4 mM MgC12 ; after 20 min at 37°C, the deoxyribonuclease was inactivated by heating at 100°C for 10 min. The solution was made up to 2 ml with water. DNA polymerase I. DNA polymerase I (EC 2.7.7.7) was isolated from Escherichia coli according to published procedures [15,16]. The preparation had a specific activity (defined and determined as in ref. 16) of 15600 units/ mg. Polynucleotide ligase. Polynucleotide ligase (EC 6.5.1.1) was isolated by the procedure described by Olivera [17] from frozen E. coil cells of the ligase overproducer mutant N1625 lop8 [18], kindly provided by Dr Richard Moyer and coming originally from Dr M. Gellert. In some experiments, indicated in the text, the enzyme used was prepared by the same procedure from commercial frozen E. coli B cells (Grain Processing Co., Muscatine, Iowa). Chemicals. NAD, NMN, and dTTP were products of Sigma. d[ 3 H/TTP was purchased from New England Nuclear. Methods

Principle Poly(dA) • oligo(dT) is a very efficient template-primer for E. coli DNA polymerase I [19]. Exposure of this template-primer to ligase reduces this efficiency, presumably by joining several adjacent oligo(dT) molecules [20] and thereby eliminating potential primer sites for the polymerase. The proposed method consists in performing two parallel DNA polymerase assays with poly(dA), oligo(dT) as the template-primer and labelled dTTP as the precursor; the ligase preparation is added to both assays, but one is supplemented with NMN, an inhibitor of the ligase reaction, and the other with NAD, a required cofactor. The difference in dTMP incorporation between the two assays, provides a measure of ligase activity. Procedure The procedure described below is for ten assay tubes, which correspond to five sets of determinations (one with NMN and one with NAD per set). It can be scaled up or down according to need. 100 #1 of a solution of poly(dA) (200 #g or 6 A260 ,m units per ml) are mixed with 200 #1 of oligo(dT) solution (120 gg/ml, prepared as described in Materials), 650 pl of buffer (0.1 M Tris--HC1 (pH 8.3)--0.01 M MgC12-0.1 M KC1), and 100 #1 of a 2.5 mM solution of d[ 3 H/TTP (20 cpm/pmol). After 15 min at room temperature, the mixture is distributed to 10 small test tubes (100 #1 per tube). Five of the tubes are supplemented with l 0 pl of 10 mM NMN and five with 10/zl of 10 mM NAD. 10 #1 of the ligase preparation are added to each tube and incubation is started immediately at 32°C. After 5 min, 10 pl of DNA polymerase solution (1 unit) are added to each tube and incubation

454 continued for another 30 min. The reaction is terminated by addition of 1.5 ml of cold 10% trichloroacetic acid. After 15 min at 0°C, the precipitates are collected by filtration through membrane filters (Schleicher and Schuell, Type B-6), washed five times with cold 5% trichloroacetic acid, and counted in a dioxane--naphthalene scintillation mixture. The difference between dTMP incorporation of the NMN-supplemented assay and t h a t of the assay containing NAD, AL, is the measure of ligase activity. Comments on the parameters of the assay The conditions of the assay, as described above, are optimal for the determination of bacterial ligase. In other situations, modifications may be necessary. The following comments on the various parameters of the assay and the suggested calibration experiment are offered as guidelines for possible modifications. Template. Since the assay is, in essence, a demonstration of a reduction in the number of primer sites available to the polymerase, it is obvious that it must be performed under conditions where this number is the limiting factor, i.e., with an excess of polymerase with respect to the template-primer. The a m o u n t of the latter can be increased only as long as it remains far from saturating the polymerase. This must be ascertained whenever a new polymerase preparation is to be used, or a change in conditions is contemplated. The calibration experiment described below can serve for this purpose. There is also another consideration with regard to the a m o u n t of template-primer: the measure of ligase activity is the difference in incorporation between the assay with NMN and that with NAD; increasing the amounts of template-primer, or polymerase, or both relative to the ligase will increase the absolute incorporation figures for both assays but not their difference. In fact, the accuracy of the determination will be reduced if A L becomes the small difference between two large numbers. Primer. Both the a m o u n t and the size of the oligo(dT) primer are important. The a m o u n t of oligo(dT) must exceed that of poly(dA), in order to achieve a close alignment of the oligonucleotides on the polynucleotide strand, with as few gaps between them as possible; such gaps are primer sites for the polymerase but cannot be repaired by the ligase. The size of the oligo(dT) is the most crucial factor. Commercial oligo{dT) preparations of specified length are too short for this purpose. A very satisfactory product can be obtained by controlled deoxyribonuclease digestion of poly(dT). The conditions for this digestion described under Materials were found to give the best results in our hands, but because of the importance of the size of the oligo{dT) for the success of the assay, they must be established by the calibration experiment described below when setting up the assay for the first time or whenever new batches of polynucleotides or of polymerase are to be used. Calibration experiment. An example of this experiment is presented in Fig. 1. Poly(dT) is exposed to deoxyribonuclease and at various time intervals samples are withdrawn, added to poly(dA), and the mixtures used as templateprimers for DNA polymerase. As indicated in Fig. 1, dTMP incorporation in-

455 4

E

3

1

to

I

310

I

I

I

I

M Jnutes of i n c u b a t i o n w i t h n u c l e a s e

Fig. 1. Calibration experiment. 60 /~g of poly(dT) w e r e i n c u b a t e d at 37°C w i t h 50 #g of p a n c r e a t i c deoxyribonuelease in 0.3 ml of 0.05 M Tris--HCl (pH 7.5)---4 mM MgC12. At the indicated times 40 /~1 samples were removed, diluted with 40 /~1 of water, and h e a t e d at 100°C for 10 rain to inactivate t h e n u c l e a s e . After cooling, 20/~1 of each sample were added to 10 pl of p o l y ( d A ) solution (200/~g or 6 A 2 6 0 nm units per ml), 65 #1 of buffer (0.1 M Tris--HC1 (pH 8.3)---0.01 M MgC12--0.1 M KC1), and 10/~1 of 2.5 mM d [ 3 H ] TTP (20 cpm/pmole). After 15 rain at room temperature, 25 pl of DNA polymerase I s ol ut i on (1 unit) were added and the samples i n c u b a t e d at 32°C for 30 min. T h e r e a c t i o n w a s t e r m i n a t e d and incor po ratio n was d etermined as d e s c r i b e d for the regular assay in Methods.

creases with increasing time of incubation of the poly(dT) with deoxyribonuclease; the increase is linear in the beginning and then a plateau is reached. It is apparent that in the linear part of the curve the efficiency of dTMP incorporation is determined by the size of the oligo(dT); it is, therefore, in this region that the joining of adjacent oligo(dT) pieces into larger molecules would be expected to ~have a most pronounced effect on dTMP incorporation. The plateau, on the other hand, indicates that there are more primer sites available than the polymerase can utilize; ligation, in this region, would have no apparent effect on incorporation. The object, then, of the experiment is to find the linear part of the curve. This type of experiment, with proper modifications, can be used in order to calibrate the polymerase assay in terms n o t only of the proper conditions of exposure of the poly(dT) to deoxyribonuclease, b u t also of other parameters, such as the amount of template-primer, the amount of polymerase, etc. Whenever a change in any of the parameters of the assay is to be made, it should be established, by this calibration, that one works in the linear part of the curve where the size of the oligo(dT) is critical. Time sequence. The time interval between the addition of the ligase and that of the polymerase does n o t seem to be critical, at least with the purified ligase. One can actually perform the assay in two discrete steps, exposing first the template-primer to the action of the ligase and then use the "repaired" template for a separate polymerase assay. On the other hand, one can demonstrate ligase action even if the two enzymes are added to the template-primer simultaneously, in mixture. Neither of these 2 extremes is practical when dealing with less purified enzyme preparations, especially if several different samples are to be compared. It is preferable to add first the ligase to a series of assays and then the polymerase, keeping the interval constant from tube to tube.

456

TABLE

I

SPECIFICITY The

assays

of the

OF were

omitted

the averages

THE

LIGASE

performed component.

of two

ASSAY

as described

in Methods

1 ~tg of purified

(Stage

with V)

the indicated

additions;

ligase was used

per assay.

water The

was added figures

in place

reported

are

determinations.

Additions

dTMP incorporation (pmoles)

NMN

NAD

Ligase

-

-

-

+

+

+

2190

-

Polymerase

2205

+

-

-

+

2268

-

+

-

+

2154

+

-

+

+

2223

-

+

+

+

1286

Specificity The specificity of the assay is demonstrated by the experiment presented in Table I. It can be seen that neither NAD, nor NMN, nor ligase alone have any significant effect on the extent of dTMP incorporation in the DNA polymerase reaction and the same is true for the combination of ligase and NMN. Only when ligase and NAD are present together, is a drop in dTMP incorporation observed. These results were obtained with purified ligase. With cruder enzyme preparations, a drop in dTMP incorporation can be seen even when ligase is added to the polymerase reaction alone, w i t h o u t NAD. This can be attributed to the presence in the crude ligase preparation of (a) some endogenous NAD or some AMP-charged ligase and (b) factors inhibiting the polymerase reaction, the most obvious being deoxyribonucleases acting on either the template or the product, or both. The NMN-supplemented parallel assay provides a valuable control in both of these cases. In the first place, NMN "discharges" ligase from any bound AMP [21]. That part of the drop in dTMP incorporation which is due to the presence of NAD or of AMP-charged ligase in the preparation is restored in the presence of NMN. An example of this is presented in the experiment of Table II, which was performed with a less purified ligase preparation. TABLE

II

EFFECT

OF

NMN

ON

CRUDE

LIGASE

PREPARATIONS

Conditions s a m e as in Table I, except that 4 p g of a less purified (Stage IV) ligase preparation w e r e used per assay.

Additions NMN

d T M P incorporation (pmoles) NAD

-

-

+ -

+

Ligase

Polymerase

-

+

1925

+

+

1551

+

+

1821

+

+

1319

457 On the other hand, the drop in dTMP incorporation due to factors inhibiting the DNA polymerase reaction is not restored by NMN but can be accurately measured in its presence. The NMN-supplemented assay permits the determination of the level of polymerase activity in the presence of all contaminants of the ligase preparation but without the action of the ligase itself; thus, a reliable baseline is provided from which to measure ligase activity, when NAD replaces NMN in the parallel assay. What is being measured, in other words, by the dual assay is an activity inhibited by NMN and requiring NAD, which can affect DNA polymerase action; of the known enzymes, ligase alone fulfills these requirements; none of the known nucleases falls into this pattern. One cannot, of course, exclude the possibility that an enzyme with such characteristics does exist but has not yet been discovered.

Sensitivity A measure of the sensitivity of the assay is provided by the experiment of Fig. 2 in which the standard assay was performed with increasing amounts of ligase. It can be seen that under these conditions as little as 0.1 pg of enzyme protein can be detected. The difference in dTMP incorporation, AL, increases linearly with the a m o u n t of enzyme up to 1 #g of protein per assay. This upper limit of linearity could be increased by increasing the amounts of templateprimer and polymerase. The ligase used in this experiment was purified about 600-fold {stage V of the procedure of Olivera [17] ) and is far from being a single protein; therefore, the actual figures for both the sensitivity and the limit of linearity of the assay must be considerably lower. Examples Most of the results presented above were obtained with purified polynucleotide ligase. In order to demonstrate the usefulness of the proposed assay in more demanding situations, we present below experiments in which the

20

o

x "~ ~C E Q.

Q5 ug

1 ligase

15

I 2

I 25

Fig. 2. S e n s i t i v i t y a n d l i n e a r i t y o f t h e ilgase a s s a y . T h e a s s a y s w e r e p e r f o r m e d u n d e r t h e s t a n d a r d c o n ditions described in Methods with increasing amounts of liga~. Stage V enzyme was used and the dilutions were made with 1 mg/ml serum albumin just before the assay.

458 TABLE III LIGASE CONTENT OF THE CRUDE O V E R P R O D U C E R M U T A N T lop8

EXTRACTS

O F W I L D - T Y P E E. coli B A N D O F T H E L I G A S E

To p r e p a r e t h e c r u d e e x t r a c t s , 5 g of f r o z e n cells o f e a c h s t r a i n w e r e s u s p e n d e d in 1 2 . 5 m l o f b u f f e r (0.1 M g l y c y l g l y c i n e , p H 7 . 0 - 1 m M E D T A ) a n d s o n i c a t e d f o r 10 r a i n in a M o d e l W 1 8 5 S o n i f i e r Cell D i s r u p t o r ( o u t p u t s e t t i n g 10). T h e t e m p e r a t u r e w a s k e p t b e l o w 1 5 ° C . T h e p r o t e i n c o n t e n t o f t h e c r u d e e x t r a c t s was determined and appropriate dilutions were made with the same buffer containing 1 mg/ml serum a l b u m i n in o r d e r t o o b t a i n t h e i n d i c a t e d ( C o l u m n 3) a m o u n t o f e n z y m e p r o t e i n p e r 10 ttl o f e x t r a c t . T h e a s s a y s w e r e p e r f o r m e d as d e s c r i b e d in M e t h o d s .

Expt No.

S o u r c e o f ligase

#g protein per assay

dTMP incorporation (pmoles)

AL

with NMN

with NAD

1 2 3 4 5

E, coli B w i l d t y p e

2.5 5 10 20 40

2108 1734 1538 1206 823

2113 1775 1450 1037 562

--88 169 261

6 7 8 9 10

E, coli N 1 6 2 5 1 o p 8

2.5 5 I0 20 50

2206 1964 1561 1303 797

2041 1780 1339 733 285

165 184 222 570 513

assay was used (1) to detect quantitative differences in ligase content between crude extracts of different strains of E. coli, and (2) to m o n i t o r ligase among the proteins eluted drom DEAE-cellulose columns in the course of the purification of the enzyme.

Crude extracts Table III presents the results of a comparison of ligase levels, determined by the proposed assay, between the crude extracts of two types of E. coli: commercial, wild t y p e B, and N1625 lop8, a ligase overproducing m u t a n t [18]. The comparison is performed at several levels of protein concentration. An examination of the results in Table III leads to a number of conclusions: (1) The DNA polymerase reaction is inhibited by the extracts, even in the presence of NMN (column 4); (2) The degree of this inhibition is a b o u t the same for the two extracts when the same amount of protein is used; (3) In spite of this inhibition, it is possible to detect the presence of ligase in the two extracts by comparing the NMN- and the NAD-supplemented assays (columns 4 and 5, respectively); (4) At all protein levels, the amount of ligase, measured by AL, (last column) is higher for the lop8 m u t a n t than for the wild type cells. An exact quantitative comparison of the ligase levels of the two types of cells is n o t easy with extracts as crude as these because there is obviously no linear relationship between the amount of protein used and A L (columns 3 and 6, respectively). If one compares the A L values for the same a m o u n t of protein (for instance, lines 3 and 8, or 4 and 9) the ligase content of the lop8 extract seems to be 2.5--3 times higher than that of the wild-type extract. On the other hand, if one compares the amounts of protein that would give the same A L

459

. . . . . . . A 28Onto - Z~,L

~ 500

10

Q. _.1 c t <

250 o

<

B • *NAD o *NMN 125( "5 E

.~ 100(

X b 75o

I

0

I 10

I Fraction

I 20

i

I 30

i

40

number

Fig. 3. D e t e c t i o n o f ligase in c o l u m n e f f l u e n t s . 5 g o f c o m m e r c i a l f r o z e n E. coli B cells w e r e carried t h r o u g h S t a g e I V o f the i s o l a t i o n procedttre o f Olivera [ 1 7 ] involving s o n i c a t i o n , s t r e p t o m y c i n precipitat i o n , a m m o n i u m s u l f a t e f r a c t i o n a t i o n , and D E A E - c e l l u l o s e c h r o m a t o g r a p h y . T h e e f f l u e n t s f r o m the D E A E c o l u m n s w e r e t e s t e d f o r ligase w i t h the p r o p o s e d assay. Ligase a c t i v i t y ( A L ) and a b s o r h a n c e at 2 8 0 n m o f the f r a c t i o n s are p l o t t e d in A and the individual d T M P i n c o r p o r a t i o n data, f r o m w h i c h A L w a s c o m p u t e d , in B.

values (lines 4 and 6), one comes to the conclusion that the lop8 extract contains 8 times more ligase than that of the wild type cells. Gellert and Bullock who isolated and characterized the lop8 mutant, estimated its ligase content as being 4--5 times higher than that of the wild type cells [ 1 8 ] .

Detection of ligase in column effluents Figs 3 and 4 illustrate the elution profiles of two ligase preparations from DEAE-cellulose columns. This fractionation is the fourth step in the ligase isolation procedure described by Olivera [ 1 7 ] . Again the sources of the two enzyme preparations are the commercial wild type E. coli B (Fig. 3) and the lop8 mutant (Fig. 4). Figs 3A and 4A present the absorbance of the eluates at 280 nm and their ligase activity, determined by the proposed assay and expressed as the A L . The activity curves indicate clearly the location of the ligase in the effluents. The individual incorporation figures in the presence of NMN and of N A D from which the A L was computed, are plotted in Figs 3B and 4B. They are included in order to illustrate the usefulness of the dual assay; indeed, it is interesting to note that incorporation in the presence of NMN does fluctuate

460

!

........ A28Qnm AL

15

c o

500

o0 10 c

OL "~ 250

0.5 ~

1000

e*NAD o*NMN

g 75o a_

I[

t--o 5 0 0

0

10

20

30

40

Frection number F i g . 4 . S a m e as Fig. 3 b u t w i t h 5 g o f E. coli N 1 6 2 5

lop8 cells as t h e s t a r t i n g m a t e r i a l .

considerably throughout the elution profiles, probably in response to endogenous polymerases, nucleases, or other factors, but incorporation in the presence of NAD follows these fluctuations except in the area of the ligase peak. Also, the possibility that the minor peak that precedes the major one in Fig. 3A is a second ligase fraction can be dismissed upon examination of the two curves of Fig. 3B: it is evident that A L is not due here to a decrease in dTMP incorporation in the presence of NAD but to an increase in the presence of NMN and does not, therefore, represent ligase. Thus, it is advantageous, when monitoring column effluents for ligase activity, to plot not only A L but also the individual incorporation data (as it was done in Figs 3B and 4B) in order to eliminate possible artifacts. Figs 3 and 4 are drawn on the same scale and, as indicated in the legends, the same amount of cells were used in the 2 experiments. It is obvious that there is more ligase in the lop8 cells than in the wild type. It is not possible, however, to perform a direct quantitative comparison of the ligase content of the two types of cells by measuring the areas under the respective activity peaks because the amount of ligase used in the assays exceeds the limits of linearity of the method, at least in the case of the enzyme from the lop8 cells. The peak fraction (tube No. 19 in Fig. 4) had to be diluted at least 20-fold before a linear relationship between enzyme protein and AT. was established.

461

Discussion

The proposed assay for polynucleotide ligase presents several advantages over the techniques presently employed for the same purpose. One of the most important advantages is the simplicity of the substrate. In most existing methods the preparation of the substrate is a laborious operation involving several chemical or enzymatic steps and often unstable isotopic markers. In contrast, poly(dA) and poly(dT) are easily prepared polymers and they can also be obtained from commercial sources; they are not labeled and they are very stable. A simple deoxyribonuclease treatment of the poly(dT) is the only required operation. Another advantage of the proposed method is its procedural simplicity. It involves a single 35 min incubation and the results are immediately available upon filtration and counting. Moreover, the technique is applicable even to crude extracts, unlike some of the simplest of the existing methods, such as the adenylylation of the ligase with labeled NAD [9] or the ATP-PPi exchange [8]. Finally, an advantage of the proposed method over what seems to be the simplest and most reliable of the existing assays, the measurement of circle formation from linear poly(dA-dT) [ 1 0 ] , is that it does not depend on the purity of the enzyme used as tool. In the latter assay, the purity of the exonuclease used for measuring circle formation is of paramount importance. For the proposed assay, even commercial preparations of DNA polymerase are suitable because the absolute activity of this enzyme is n o t a crucial factor. The optimal amount of polymerase can be determined by a calibration experiment, and the NMN control provides a baseline for the measurement of ligase activity alone*. The proposed assay should be applicable also to the viral and mammalian ligases by substituting ATP for NAD. We have n o t yet tested this possibility. One complication that might be expected in this case, when working with crude extracts, is the presence of ATP-stimulated nucleases. Work is n o w in progress on the quantitative aspects of the assay, especially with crude extracts, and on the definition of a unit comparable to those n o w in use. But even in its present form, the assay can be very useful for comparing ligase levels in cell extracts and for monitoring the purification of the enzyme. Acknowledgements I wish to express m y gratitude to Dr Erwin Chargaff for his support and encouragement and to acknowledge the excellent assistance of Mr Richard Liou. This work was supported by research grant CA-12210--13 from the National Institutes of Health, U.S. Public Health Service, and grant NP-56P from the American Cancer Society. The author is the recipient of an Irma T. Hirschl Career Scientist Award. *

P r e p a r a t i o n s o f D N A p o l y m e r a s e I a r e n o t a l w a y s free o f traces o f ligase a c t i v i t y . A set o f assays ( o n e w i t h N M N a n d o n e w i t h N A D ) o m i t t i n g t h e Ugase c o u l d d e t e c t t h e p r e s e n c e o f this a c t i v i t y in t h e p o l y m e r a s e and p r o v i d e t h e a p p r o p r i a t e b l a n k i f a c o r r e c t i o n is n e e d e d .

462

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