Enhanced Deletion Formation by Aberrant DNA Replication in Escherichia coli

(1)

Copyright 0 1997 by the Genetics Society of America

Enhanced Deletion Formation by Aberrant DNA Replication

in

Escherichia coli

Catherine

J. Saveson

and

Susan T.

Lovett

Department of Biology and Rosenstiel Basic Medical Sciences Research Center, Brandeis University,

Waltham, Massachusetts 02254-91 10

Manuscript received September 26, 1996 Accepted for publication February 21, 1997

ABSTRACT

Repeated genes and sequences are prone to genetic rearrangements including deletions. We have

investigated deletion formation in Eschm'chia coli strains mutant for various replication functions. Deletion

was selected between 787 base pair tandem repeats carried either on a ColElderived plasmid or on the

E. coli chromosome. Only mutations in functions associated with DNA Polymerase I11 elevated deletion

rates in our assays. Especially large increases were observed in strains mutant in dnaQ the E editing

subunit of Pol 111, and dnuB, the replication fork helicase. Mutations in several other functions also

altered deletion formation: the a polymerase ( d n a , the y clamp loader complex (holC, dnaX), and

the

p clamp

(dnaN) subunits of Pol I11 and the primosomal proteins, dnaCand pui. Aberrant replication

stimulated deletions through several pathways. Whereas the elevation in dnaB strains was mostly recA-

and Zeddependent, that in dnaQstrains was mostly recA- and Zed-independent. Deletion product analysis

suggested that slipped mispairing, producing monomeric replicon products, may be preferentially in-

creased in a dnaQ mutant and sister-strand exchange, producing dimeric replicon products, may be

elevated in dnaE mutants. We conclude that aberrant Polymerase I11 replication can stimulate deletion events through several mechanisms of deletion and via both recA-dependent and independent pathways.

T

ANDEMLY repeated DNA sequences, common in

the genomes of many organisms, are vulnerable to deletions and amplifications. Such rearrangements may occur between repeated genes, kilobases in length, or between short segments, of as little as a few nucleotides in length. These tandem repeat rearrangements are common sources of genetic mutation and human

genetic disease (MEUTH 1989; KRAWCZAK and COOPER

1991; Hu and WORTON 1992; NELSON 1993).

Genetic analysis of repeated sequence rearrange-

ments in Escherichia coli has revealed that multiple mech-

anisms contribute to the process. In particular, we have

previously investigated the genetic dependence of spon-

taneous deletion events between 787 base pair (bp)

repeated sequences within the tetA gene of E. coli (Fig-

ure 1 ) (LOVETT et al. 1993). The majority of recovered deletions occur by a homologous recombination pathway requiring the RecA strand transfer protein. How-

ever, a substantial proportion of these events (30%)

occur independently of RecA.

Both RecA-independent and -dependent deletion

formation occurs between homologous target se-

quences. However, there is a minimal homology re-

quirement for the Red-dependent pathway and no

such requirement for the Red-independent pathway. The RecA-dependent mechanism does not appear to

function on homologies less than -200 bp in length

(DIANOV et al. 1991; WIN et al. 1991; BI and LIU 1994).

Corresponding author: Susan T. Lovett, Rosenstiel Basic Medical Sci-

ences Research Center MSO29, Brandeis University, Waltham, MA

0225491 10. E-mail: lovett@hydra.rose.brandeis.edu

Genetics 146: 457-470 (June, 1997)

The RecA-independent mechanism can mediate deletion between homologies of several nucleotides, although the efficiency of deletion is improved dramati-

cally with increased homology (ALBERTINI et al. 1982;

DIANOV et al. 1991; WIN et al. 1991; PIERCE et al. 1991;

BI and LIU 1994).

DNA replication may play a role in both mechanisms

of deletion formation. The events that may trigger

RecA-dependent homologous recombination between

tandem repeats are not known but could include a

blocked replication fork. Our genetic studies implicated the RecF, RecO and RecR homologous recombi-

nation proteins of E. coli in the RecAdependent dele-

tion pathway

LOVE^

et d . 1993; M. Bzymek and S. T.

Lovett, unpublished results). These functions are also

required for recombinational repair of DNA lesions

that block replication in the cell (TSENG et al. 1994).

Therefore, some tandem rearrangements could result from recombinational gap-filling reactions in a blocked

replication fork as depicted in Figure 2. Unequal cross-

ing-over between sister chromosomes or recombination between the two repeats on the same chromosome dur-

ing the course of this repair may lead to deletion of

one of the repeated segments. Alternatively, as shown

in Figure 3, a stalled replication fork may be broken;

recombination of the broken chromosome with its sister to restore the fork could result in deletion of tandem

repeats (ASAI et al. 1994; KUZMINOV 1995a,b).

(2)

458 C. J. Saveson and S. T. Lovett

fetA+

.:.:.:.:.:.:.:.:.:

:.:.:.:.:.:.;.:.:. .:+:+:.:.:.: :.:.:.:.:.:.:.:.

tetAdu

EmRV Nrul

:.:.:.:.:.:.:.: _... ...

... ... :::::::::I

FIGLIRE 1.-Diagram of the k t A locus and the 787-bp dupli- cation used in these experiments. Duplication of the segment from the EcoRV to the NmI sites of the gene disrupts the t d A

gene. Deletion of one copy of the tandem repeat restores

the IdA' gene structure and can be selected by tetracycline-

resistance.

cative mechanism for the RecA-independent deletion of both short (TRINH and SINDEN 1991) and long (LO-

VETT and FESCHENKO 1996) tandem repeats. Exposure

of singlestrand DNA during the replication process may provide an opportunity for tandem sequence homologies to interact. A simple slipmispairing model (Figure 4A) proposes that slipped realignment between the nascent strand and its template leads to deletions or amplifications (STREISINCER et al. 1967; ALBERTINI et al. 1982). Additional models propose that deletions may arise by strand misalignment within a replication fork concomitant with sisterchromosome exchange (Figure 4B) (LOVEIT et al. 1993; BI and LIU 1996). The location of these slipped mispairing events within closely juxta- posed and singlestranded regions of the replication fork would explain why a strand transfer protein such as R e d is not needed for such reactions.

Because DNA replication is implicated in both RecA- dependent and -independent deletion mechanisms, we have investigated the genetic relationship between DNA replication components and deletion formation. There are three DNA polymerases in E. coli: Polymerase I11 bears the major responsibility for chromosomal replication, Polymerases I and I1 are both believed to be in- volved in repair pathways. Polymerase I is required for some plasmid replication initiation and may also play a role in Okazaki fragment joining (KORNBERG and BAKER 1992).

Polymerase I11 itself is a complex enzyme with multiple subunits encoded by different genes (BAKER and WICKNER 1992; MARIANS 1992; KELMAN and O'DONNELL

1995; MARIANS 1996). The polymerase activity is pro- vided by the a subunit, encoded by dnaE.

Its

proofread- ing e subunit, possessing exonuclease activity, is encoded by dnaQ. The function of the final subunit of the core polymerase, 8, encoded by hoE, is apparently nonessential (STUDWELL-VAUGHAN and O'DONNELL 1993; SLATER et al. 1994). It is believed that both leading and lagging strands are coordinately synthesized by an asymmetric polymerase dimer (JOHANSON and MCHENRY 1984). The T subunit, encoded by dnuX, facil- itates the interaction between the two core polymerase molecules (MCHENRY 1982; STUDWELL-VAUGHAN and

C.

D.

E.

Branch migration

J/

J.

Resynthesis

*rn??%?*?A?Y*?* ₃_:_;_I

:=

_K._~ _:..,. .,,,,, _I ! R x * ~ : ? ~ w $ $ ~ ? ? x < x x < > ~ _I _~ _~_<;

:=

_~ _~ _....:.:.. _..:s.,_~ _: _* _~ _.

::::., .:+ _..._...n.. 6:: .... .... .... .... _...

.... ... _..._....

... _... ...

... ....

..:.:.>. ... x&

+??

.... ...

.... ... >:.:+>:.:.:.:.:.:.>>:.:.>:.:.:.

... ... ...

<*+-

... ... < ...

... ....

... ... ... ...

... .:.:..

...

.:.:.. .:.:., .*.:.. &.

-< >.*.Mm$$: fi:::..<+w >y* a &% w ->

* *

k.& e g< s:; .:$;*??x%K&Y

*;

ss:*...

Junction resolution

c

...

FIGURE 2.-Recombinational gapfilling (post-replication) repair. (A) DNA synthesis cannot proceed past a replication- blocking lesion, giving rise to a daughterstrand gap. (B) The gap can be filled by recombination with the sister chromo- some. (C) Branch migration of the Holliday junction can bypass the lesion and cause the incomplete nascent strand to

switch templates. (D) Replication is completed on the sister template. (E) The Holliday junction is resolved. This particu- lar resolution produces a cross-over between sister chromo- somes.

O'DONNELL 1991). The subunit, encoded by dnaN, is a processivity factor that forms a clamp around the DNA to anchor the polymerase (STUKENBERG et al. 1991). This clamp is loaded by a polymerase-associated complex (O'DONNELL 1987), which includes the y

(also encoded by dnaX), 6 (holA),

6'

(holB),

x

(holCj,

and t,b ( h o D ) proteins.

(3)

Hyperdeletion in Replication Mutants 459

A.

B.

C. J,

JI

Degradation and recombination with sister chromosome dqradatQn

J.

FIGURE 3.-Replication fork collapse and repair (Kuzminov

1995b). (A) Inhibition of polymerization stalls the replication fork. (B) Nuclease attack of single-strand DNA breaks the fork. (C) The broken end invades i t s sister chromosome. Dur- ing this step unequal recombination can occur at a repeated

sequence (dark hatching). (D) The fork is reestablished with

a deletion on one chromosome.

association is mediated by interactions with the dnaB protein, also the replication fork helicase (TOUGU et al.

1994). The primase, like the

P

clamp, is required at the start of each Okazaki fragment, placing a potential greater burden on these functions for lagging strand synthesis. Other proteins that are required for primosome assembly and may also be components of the primosome include the p i A , @‘B, pzC, dnaC and dnaT gene products

(ARAI

and KORNBERG 1981). An alternative primosome assembly occurs at the E. coli origin of replication, using the d n d protein in addition to dnaB and dnaC (FUNNELL et al. 1987; M A S N et al. 1990).

In this work, we have investigated whether aberrant replication promotes deletions at repeated sequences. We constructed a series of isogenic strains containing mutations that impair selected components of the replication machinery. Because there are three DNA polymerases in E. coli, we have tested the genetic dependence of deletion on each polymerase gene as well as

other functions associated with DNA replication. E. coli mutant strains were surveyed for effects on deletion formation using both plasmid and chromosomal deletion assays. To gain insight into the mechanisms of deletion during aberrant replication, we also analyzed the

deletion products from selected strains. We present evidence that aberrant or incomplete replication via DNA polymerase I11 stimulates deletion at tandem repeats via several mechanisms.

MATERIALS AND METHODS

Bacterial strains, media and antibiotics: Strains were grown

on LB media: 1 % Bacto-tryptone, 0.5% yeast extract, 0.5%

sodium chloride and 0.005% thymine, and, for plates, 1.5%

agar (WIl.LETTS et dl. 1969). Growth temperatures are indi-

cated in the text. Minimal complete media consisted of 56/

2 salts (WILLETTS et al. 1969), 0.2% glucose, 1 pg/ml thiamine,

50 pg/ml each of required amino acids and, for plates, 2% agar. LCG media for preparation of P1 lysates and transduc- tions consisted of LB media supplemented with 1% glucose

and 2 mM calcium chloride, and, for plates, 1 % agar. A 0.7%

agar concentration was used for LB and LCG top agar. Media

were supplemented with antibiotics: kanamycin ( K m ) at 60

pg/ml, tetracycline (Tc) at 8-15 pg/ml, chloramphenicol

(Cm) at 15 pg/ml and ampicillin (Ap) at 100 pg/ml (for

plasmid resistance genes) and 30 pg/ml (for chromosomal genes).

Isogenic strains used for deletion assays were constructed

by P1 transduction and are described in Table 1 . These strains

include derivatives of AB1 157 for plasmid deletion assays (A)

and STL695 for chromosomal deletion assays (B). P1 u i r A

phage lysates and transduction were performed as described (MILLER 1992). For many strains, a TnlQkan element linked

to the mutation of interest was used for selection in strain

constructions. In these cases, control strains were constructed

carrying only the Tn IOkun element to ensure that this marker

did not contribute to the deletion phenotype. Deletion assays confirmed that deletion rates in these control strains were within 50% of the wild-type value (data not shown). Mutants in dnuQ were assayed for mutator phenotype on rifampin plates (at 100 pg/ml).

Deletion assays: Deletion formation was measured using

plasmid pSTL55 (LOVE= et al. 1993). This plasmid is a deriva-

tive of pBR322, conferring ampicillin resistance, and contains

a tandem repeat within the tetA gene (Figure 1). The dupli-

cated region spans 787 bp between the EcoRV and NruI sites

of tetA. Plasmid DNA was introduced into strains via electro-

poration (DOWER et ul. 1988). Chromosomal deletion fre-

quencies were measured in strains carrying an insertion of

bla and tetAdup787 from pSTL55 on the E. coli chromosome

at lacZ (LOVE= et al. 1993). Deletion of one tandem repeat

results in recovery of a functional tetA gene and confers resis-

tance to tetracycline.

Strains containing either the chromosomal or plasmid con-

struct were grown at their permissive temperatures on LB plates containing ampicillin. Individual colonies were then picked whole, and grown in LB+Ap broth for 2 hr. The cul-

tures were diluted in 56/2 buffer and the number of Ap‘ and

Tc‘ cells in the population was determined by plating cell dilutions on LB+Ap and LB+Ap+Tc media. Because of poor

growth in rich medium, polA and

pnA

mutant strains were

grown on minimal-complete agar plates and in minimal-liquid media with the appropriate antibiotics. Deletion assays were performed for 2 1 0 independent cultures in parallel with the wild-type control strain. Deletion rates were calculated by the

method of the median (Lw and COULSON 1949) for the en-

tire set of assays using the formula: deletion rate = M / N ,

where M is the calculated number of deletion events and N

is the final average number of Ap‘ cells in the 1-ml cultures. Mis solved by interpolation from experimental determination

(4)

460 C. J. Saveson and S. T. Lovett

A. Simple slipped mispairing B. Sister-strand slipped mispairing

f Nascent strand

<Template strand

wk::~s7~::;*;;~x ... ..". ...

:.:.:.:. '.' 1 II ...~.:.~.~.:.:.~.:.:~.. ... ... ...

E?%dR???%?

...????>% ... ".::.:.A _.,.!?... ... ... ... _...

_:*

x.:.:. ... ...

*:<

*

_.....:.&$:-

.:.:.x;:.:.:.:-.:~<~<~:~:~~: :.:,.:.:.> ... .... :.:.:..

Sister strands anneal

+

Slipped alignment .., >,>..:.:.:.>:<.:.:.: _...W? VS

...

w.s-<:- ... ... ...

I

*

Monomer deletion product

+

Dimer deletion product

FIGURE 4.-Slipped realignment mechanisms for deletion formation. (A) Simple slippage. Displacement and realignment of

a nascent strand with its template can delete one copy of a tandem repeat. On a circular chromosome, such deletion products should be replicon monomers. (B) Sister-strand slipped mispairing. Nascent strands are displaced and mispair with each other, causing a deletion to be formed. [Although we have illustrated leading strand synthesis impeded relative to the lagging strand,

the ends of the nascent strands may be created by nucleolytic cleavage rather than representing the point at which DNA synthesis

has stopped. See LOVEIT et al. (1993) for further discussion of this model.] With pairing of the parental strands, a recombinational

intermediate is produced. Resolution o r replication through the Holliday junction intermediate causes sisterchromosome

exchange. Crossing-over between circular sister chromosomes produces a dimeric deletion product. Both mechanisms are

presumed to operate independently of RecA.

cultures, by the formula r,, = M( 1.24

+

In

M).

The observed

numbers of Tc' cells for a given data set were ranked and a 95% confidence limit for the median was established using

Table A-25a of DIXON and MASSEV (1969); this was converted

to a 95% confidence interval for deletion rates as suggested

by Wierdl et al. (1996) using the formula above.

Analysis of plasmid deletion products T e d ' plasmid deletion products were examined by electrophoresis to determine

monomer and dimer product distribution. These deleted

products were readily distinguishable from each other and from the parental plasmid. Each product examined was iso-

lated from an independent culture. Dimeric plasmid DNA was

never detected from nondeleted Apresistant (Tc-sensitive) colonies, confirming that dimerization occurred concurrent

with deletion formation. Plasmid DNA was prepared from

each culture by alkaline lysis (SAMBROOK et al. 1989) or by

phenolchloroform extraction and subjected to agarose gel

electrophoresis. Phenolchloroform DNA extraction was performed as follows: single colonies were randomly selected and

suspended in 25 pl of TE (10 mM Tris-HCI, pH 8.0, 1 mM

EDTA). For slow-growing strains, individual colonies were

streaked to LB+Tc and incubated overnight, producing a

larger streak, which was then suspended in 25 p1 of TE. This

suspension was extracted with 25 pl of phenolchloroform-

isoamyl alcohol (25:24:1 by volume), mixed vigorously for 1 min and cleared by microcentrifugation for 2 min. The entire

supernatant was then subjected to agarose gel electrophoresis.

RESULTS

Effects of replication mutations on deletion forma-

tion: We constructed a series of isogenic strains in the

AB1157 strain background that carry various replication mutations (Table 1). Deletion of tandem direct repeats was measured using

both

a plasmid-based and chrome somal assay. Plasmid pSTL55 (LOVEIT et aZ. 1993), containing a 787-bp internal tandem repeat disrupting the

tea

gene on plasmid pBR322, was introduced into each strain. Precise deletion of one repeat restores a functional

tea

gene and tetracyline resistance to the cell. The c h r e mosomal assay used the identical 787-bp ,!&A duplication construct integrated into the chromosomal

lac

gene (Lo-

VEIT et al. 1993). In the case of

pnA

and pol4 mutants,

deletion rates could be measured only for the chrome somal construct because of poor plasmid maintenance. Deletion rates were calculated from the number of cells in the population that acquired tetracycline resistance (as

described in MATERIAIS AND METHODS).

(5)

Hyperdeletion in Replication Mutants 46 1

1994) on rich medium, these strains were grown and

assayed on minimal medium at

37”

(Table

2B).

Strains

with temperature-sensitive lethal phenotypes (Table

2,

C and D) were measured at their permissive tempera-

ture for growth, either 25 or 30”. In each case, we show absolute deletion rates for the mutant strain and the rate relative to the wild-type control strain under the same growth conditions (Table 2). For the wild-type strains, the various growth conditions had little effect on the efficiency of deletion formation.

We found that many replication mutants exhibited higher deletion rates in both plasmid and chromosomal

deletion assays (Table

2).

In particular, some mutants

with defects in DNA polymerase I11 core enzyme exhib-

ited high deletion rates. This includes a sevenfold eleva-

tion of plasmid deletion rates seen for the temperature- sensitive dnaE486 (poZC486) mutant, which is defective

in the a polymerase subunit itself. However, a minor

twofold effect on chromosomal deletion rates was seen. Deletion assays of strains carrying the antimutator allele

of dnaE, dnaE925, were normal. Mutants carrying

dnaQ49, affecting the 3’ to 5‘ editing exonuclease E

subunit, exhibited dramatic increases in deletion formation for both assays, with ninefold and 100-fold stimulation of plasmid and chromosomal deletion rates, re-

spectively. Likewise, another allele of dnuQ, mutD5,

stimulated deletion rates threefold in the plasmid assay and 600-fold in the chromosomal assay. Null mutants in hoE, the third 8 subunit of the polymerase core enzyme complex, were unaffected for deletion formation in either assay. In contrast to these mutations affecting DNA polymerase 111, deletions of the genes for polymerase I

(poZA) and polymerase I1 (poZB) had no influence on deletion rates.

Mutants in the P-processivity “clamp” associated with polymerase 111, dnaN, gave seemingly contradictory results in the two assays: 14fold higher deletion rates in the plasmid assay and eightfold lowered deletion rates

in the chromosomal assay as compared to wild-type

strains. This latter result is of special interest because it implicates replication as an essential component of deletion formation and suggests that when the polymer-

ase clamp is impaired, formation or recovery of deletion

events on the E, coli chromosome is reduced. The dis-

crepancy between the two assays will be addressed in

the DISCUSSION below.

Other mutants affecting the “clamp loader” (the y

complex of the DNA polymerase I11 holoenzyme) were analyzed. Deletion formation in temperature-sensitive

mutants of the dnaXgene, which encodes both T and

6 subunits of polymerase 111, was at normal levels for the

plasmid assay but 30-fold reduced in the chromosomal assay, reminiscent of the dnaN results above. A null

mutant in hoZC, encoding the

x

subunit of the y com-

plex, exhibited a fivefold elevated deletion rate in the plasmid assay and a 16-fold elevated rate in the chromosomal deletion assay.

Other mutants affecting primosome assembly and

the replication fork were also assayed. Elevated deletion

rates were seen for a temperature-sensitive mutant of

the dnaB helicase and primosome mutants in dnaC and dnaT, assayed at their permissive temperatures. These strains showed a substantial increase in the plasmid deletion formation as compared to wild-type strains: 20- fold for dnaB107, 11-fold for dnaCl and 15-fold for the double dnaC2 dnaTl2 mutants. The dnaBmutant exhibited a large effect in the chromosomal assay with a 130- fold stimulation of deletion rate. Unfortunately, the chromosomal deletion assay could not be used for dnaC

and dnaT mutants due to their extremely poor growth.

Null mutants in the primosome helicase gene PriA

showed a 10-fold increase in the chromosomal deletion

assay and could not be measured in the plasmid assay because of plasmid instability.

We conclude from these studies that aberrant DNA replication via polymerase I11 often results in high rates of deletion between tandem repeated sequences. DNA polymerases I and I1 are not required for deletion for-

mation in E. coli. Recovery of deletions on the chromo-

some is impaired in dnaNand dnaxmutants with defective processivity clamp and clamp loader complex.

Is elevated deletion formation in d m mutants r e d - dependent or -independent? Both recAdependent and -independent pathways contribute to deletion forma-

tion between repeats of the 787-bp size assayed. For

wild-type strains, approximately 70% of the observed deletion is recA-dependent and 30% is recA-independent (LOVETI et al. 1993). The recA-dependent pathway also

uses the recombination genes, recF, recR and rec0 and

may reflect a normal recombinational pathway known

as the RecF pathway (reviewed in SMITH 1989; CLARK

and SANDLER 1994). The genetic basis of the recA-independent pathway is unknown but has been proposed to involve a replication-slippage (or realignment) mechanism.

We chose three mutants with elevated deletion rates (dnaE486, dnaQ49and dnuBlO7) to determine whether the increased deletion formation was due to recA-dependent or -independent processes (Table 3). Plasmid deletion rates for dnaE486 mutants were independent of recA, with an 11-fold stimulation in recA- derivatives compared to sevenfold in recA+ derivatives. (The chromosomal assay had shown no significant elevation and

so was not tested). Assays of the dna449 mutant strains

showed only a minor effect of recA (twofold elevation

in recA- dnaQ49mutants vs. ninefold elevation in recA+

dnaQ49 mutants for the plasmid assay; 52-fold stimula-

tion in recA- dnaQ49 us. 98-fold in recA+ dnaQ49deriva-

tives for the chromosomal assay). In contrast to the

above, assays of the dnaB107 derivatives showed that most of the large increase seen in the chromosomal

assay was dependent on the recA gene: deletions rates

were 130-fold elevated in recAf but only twofold elevated

(6)

TABLE 1

Escherichia coli strains

Strain Relevant genotype Derivation

AB1157" DM49 JC10287

RM4193 STL1270 STL1316 STLl336 STL1818 STL1845 STL1861 STL1866 STL1880 STL1909 STL2107 STLZ 134 STLZ 182 STL2234 STL2314 STL2545 STL'2674 STL2702 STL2706 STL3000

STL695 STL753 STLl324 STL1353 STL1885 STL1887 STL1902 STL1957 STL2077 STL2082 STLZ 109 STL2125 STL2 129 STL2131 STL2 149 STL2181 STL2 186

STL2332 STL2385

STL3022

RM4848 STL672 STL700 STL774 STL1277 STL2128

STL2133

STLZ 174

A. Strains used in plasmid deletion assays

rec+ _BACHMANN₍₁₉₉₆₎

lexA3 A. J. CLARK, MOUNT et al. (1972)

A(srlR-recA)304 CSONKA and CIARK (1979)

holE202: : cat R. MAURER, SIATER et al. (1994)

dnaBl07(ts) malE::TnlOkan Km' Ts transductant P1 STLll52 X AB1157

dnaB107(ts) malE: :TnlOkan A(srlR-recA)304 W Cys' transductant P1 JC10287 X STL1277

ApolBl: : spc str ESCARCELLER et al. (1994)

dnuE486( ts) zae-3095: : Tn 1 Okan Km' Ts transductant P1 STL1155 X AB1157

dnaN159(ts) rid-3162::TnIOkan Km' Ts transductant P1 STL1205 X AB1157

dnaQ49( ts) zae-3095: : Tn 1 Okan K m ' Mut transductant P1 STL1368 X AB1157

priA2: : kan Km' transductant P1 PN103 X AB1157

dnaX2016( t s ) zba-3101: : Tn lOkan Km' Ts transductant P1 STL1157 X AB1157

dnaT12(ts) dnaC2(ts) zji-3188::TnlOkan Km' Ts transductant P1 STL1147 X AB1157

holCl02:: cat Cm' transductant P1 RM4848 X AB1157

sulAI I led71 : : T n 5 Km' transductant P1 RDKl397 X STL2181

dnaE486( ts) zae-3095 : : Tn 1 Okan A(srlR-recA)304 uv' Cys+ transductant P1 JC10287 X STL2174 dnaQ49( ts) zae-3095: : Tn I Okan A(srlR-recA)304 U V Cys+ transductant P1 JC10287 X STL2267

dnaCl (ts) zj-3188: : Tn 1 Okan Km' Ts transductant P1 STL2519 X AB1157

dnaE925 zae: : TnlOd-cam Cm' Km'transductant P1 NR11258 X STL1819

priA2: : kan s u o l l l K m ' transductant P1 STL1866 X STL'2134

dnaE925 zae: : T n l Od-cam A(srlR-recA)304 uv' Cys+ transductant P1 JC10287 X STL2704

mutD5 Pro' Mut transductant P1 RDK1276 X AB1157

S U l A l l Pyr+ Tc' transductant P1 RDK1397 X STL774

B. Strains used in chromosomal deletion assays

l a d : : bla' tetAdup787

A(srlR-recA)304 lacZ: : bla' tetAdup787

dnaBIO7( ts) maE: : Tn 1Okan lacZ: : bla+ tetAdup787 ApolBl : : spc str lacZ: : bla+ tetAdup787

dn&486( ts) zae-3095 : : Tn 1 Okan lacZ: : bla' tetAdup 787 dnaNI59(ts) zid-3162:: TnlOkan lacZ:: bla+ tetAdup787 dnaX2016(ts) zba-3101 ::TnlOkan lacZ:: bla+ tetAdup787 dnaQ49(ts) zae-3095: : Tn 1 Okan lacZ: : bh' tetAdup787 holE202:: cat lacZ:: bla' tetAdup787

priA2:: kan lacZ:: bla' tetAdup787 holClO2: : cat LacZ: : bla+ tetAdup787 lexA3 l a d : : bla' tetAdup787

dnaBlO7(ts) malE: :TnlOkan led3 lacZ:: bla' tetAdup787 dnaQ49(ts) led3 zae-3095::TnlOkan lacZ:: bla+ tetAdup787 ApoL4:: kan lacZ: : bla' tetAdup787

sulAll lacZ: : bla+ tetAdup787

dnaQ49(ts) zae-3095: : TnlOkan A(srlR-recA)304 lacZ: : bla+

s u o l l l lexA71::Tnj lacZ::bla+ tetAdup787

dnaBlO7( ts) malE: : T n l Okan A(srlR-recA)304 lacZ: : bla+ mutD5 lacZ: : bla+ tetAdup787

tetAdup 787

tetAdup787

LOVETT et al. (1993) LOVETT et al. (1993)

Km' Ts transductant P1 STL1152 X STL695 Ap' transductant P1 STL695 X STL1336 Km' Ts transductant P1 STL1155 X STL695 Km' Ts transductant P1 STL1205 X STL695 Km' Ts transductant P1 STL1157 X STL695 Km' Mut transductant P1 STL1368 X STL695 Cm' transductant P1 RM4193 X STL695 Km'transductant P1 PN103 X STL695 Cm' transductant P1 RM4848 X STL695 Ap' transductant P1 STL695 X DM49 Km'Ts transductant P1 STL1152 X STL2125 Km' Mut transductant P1 STL1368 X STL2125 Km' transductant P1 SClOl X STL695 Ap' transductant P1 STL695 X STL2134 uv' Cys+ transductant P1 JC10287 X STL2133

Ap' transductant P1 STL695 X STL2182

U V Cys' transductant P1 JC10287 X STL2128

Pro+ Mut transductant P1 RDK1276 X STL695

C. Intermediate strains derived from AB1157

holCI02:: cat recA430 sr1::TnlO zjg-2086:: kan R. MAURER

cysC95 : : Tn I O Tc' transductant P1 CAG12173 X AB1157

cysC95 : : Tn I O lacZ: : bla+ letAdup 78 7 Tc' transductant P1 CAG12173 X STL695

pyrD34 zcc-282: : Tn I O Tc' Pyr- transductant P1 DC305 X AB1157

dnaB107(ts) malE::TnlOkan cysC95::TnlO Tc' transductant P1 CAG12173 X STL1270

dnaB107(ts) malE::TnIOkan cysC95::TnlO lacZ:: bla+ Km' Ts transductant P1 STLll52 X STL700 dnaQ49( ts) zae-3095 : : Tn IOkan cysC95: : Tn 10 lacZ: : bla' Km' Mut transductant P1 STL1368 X STL700

tetAdup787

tetAdub787

(7)

Hyperdeletion in Replication Mutants

TABLE 1

Continued

463

Strain Relevant genotype Derivation

STL2267 STL2704

AX727" CAG12107' CAG12119' CAG12173' CAG18558' CAG18580' CAG18619' DC305" E107' E48d HC 1 94" KH1366" MElO1' NR1125W PC1

PN103' RDKl276" RDK1397" SC101" STLl 147k STLl152' STL1155/ STL1157" STL1205" STL1368" STL251gk PC2k

dna449( ts) zae3095: : Tn 1 Okan cysC95: : Tn I O

dnaE925 zae: ; Tn 1 Od-cam cysC95 : : Tn 10

D. Other strains

dnaX20I 6( ts) = dnaZ2016

zba-3101: : Tn 1 Okan

malE: : Tn 1 Okan

cysC95: : T n 10

zid-3162: : Tn 1 Okan

zae-3095 : : Tn 1 Okan

y'j-3188: : Tn 1 Okan

zcc-282 : Tn 10 pyrD34

dnaB107(ts)

dnaE486( ts) = polC486( ts)

dnaN159(ts) dna449(ts)

ApolBl : : spc str

dnaE925 zae: : Tn I Od-cam

dnaCl

dnaTl2( ts) dnaC2( ts)

priA2: : kan

mum5

lexA 71 ; : T n 5 sul4l I

ApolA: : kan ApolBl : : spc str

dnaT12(ts) dnaC2(ts) zjj"jr188::TnlOkan

dnaBlO7( ts) malE: : Tn 1 Okan

dnaE486( ts) = polC486( ts) zae-3095: : Tn I Okan

dnaX2016( ts) zba-3101: : Tn 1 Okan

dnaN159( ts) zid-3162: : Tn I Okan

dnaQ49( ts) zae-3095: : Tn 1 Okan

dnaCl (ts) zjj-3188: : Tn I Okan

Km'Mut transductant P1 STL1368 X STL672

Tc' transductant P1 CAG12173 X STL2674

B. BACHMANN, FILIP et al. (1974)

SINGER et al. (1989)

B. BACHMANN

B. BACHMANN, WECHSLER and GROSS (1971)

B. BACHMANN, SAKAKIBARA and MIZUKAMI (1980)

B. BACHMANN, HORIUCHI et al. (1981)

ESCARCELLER et al. (1994)

R. SCXAAPER, FIJALKOWSKA and SCHAAPER (1995)

B. BACHMANN, CARL (1970)

K. MARIANS, NURSE et al. (1991)

R. KOLODNER R. KOLODNER M. GOODMAN

Km'Ts transductant P1 CAG18619 X PC2

Km'Ts transductant P1 CAG12119 X E107

Km'Ts transductant P1 CAG18580 X E486

Km'Ts transductant P1 CAG12107 X AX727

Km'Ts transductant P1 CAG18558 X HC194

Km'Mut transductant P1 CAG18580 X KH1366

Km'Ts transductant P1 CAG18619 X PC1

" Genotype of AB1 157, STL695 and their derived strains, unless otherwise indicated, includes F- X- hisG4 argE2 leuB6 A(gpt-

I' Genotype of AX727 and its derivative includes F- thi lac rpsL rpsL.

'All CAG strains are derived from MG1655 and their genotype includes F- X-.

"Genotype of DC305 includes F- galK2 mal41 xyl-7 mtl-2 rpsL118.

'Genotype of E107 and derivative includes F- thr-1 leuBGJhwl21 supE44 X- @Dl thyA6 rspL67 thildeoC1.

/Genotype of E486 and its derivative includes F- thr-1 leuB6 tonA2l lacy1 supE44 X- @Dl thyA6 rpsL67 thil deoC1 met89.

"Genotype of HC194 and its derivative includes Hfr tow422 ompF627 reoll thyA148 metBl T T .

'Genotype of ME102 includes F- X- tonA2l thrl leuB6 lacy1 supE44 @Dl thil.

I Genotype of NR11258 includes F- aru thi A(pro-lac) mutL:: Tn5.

'

Genotype of PN103 includes HfrC X-.

proA)62 thr-1 thi-1 rpsL31 galK2 lacy1 ara-14 xyl-5 mtl-1 kdgK5l supE44 tsx-33 @Dl mtl-1 rac-.

Genotype of KH1366 and its derivative includes Hfr metD88 Alac-6 tsx-7 X- srl-8 reUl spoT1 metB1.

Genotype of PC1, PC2 and their derivatives includes F- leuB6 X- thyA47 qsL153 deoC3.

Genotype of RDK1276 includes lacZ trpA metE.

Genotype of RDKl397 includes F- leuB6 thr-1 p o met x y l mtl gal ara.

"Genotype of SClOl includes F+ A(ga1-bio) thi-1 rel4l spoT1.

rates were also somewhat recA-dependent, with 20-fold elevation in recA' dnaB107strains and 4.5-fold elevation

i n recA- dnaB107 strains. We conclude that both recA-

d e p e n d e n t and -inde pe ndent d el e ti o n form a t i on can

be induced by aberrant DNA replication and that indi-

vidual replication mutants differ in the mechanism by

which deletion formation is stimulated.

Effect of the SOS response on deletion forma- tion: The SOS response in E. coli is the coordinated

induction of a set of genes whose function is to p r o m o t e

survival after production of replication-blocking lesions

i n DNA. Some replication mutant strains such as dnuQ

and PriA exhibit constitutive expression of the SOS re-

sponse (NURSE et al. 1991; SLATER et al. 1994) and it

is u n k n o w n w h a t t h e c o n s e q u e n c e o f t h i s m a y be o n observed tandem deletion rates.

Induction of the SOS response was neither necessary

nor sufficient to induce deletion formation (Table 4).

(8)

TABLE 2

Deletion rates in red' background

Plasmid assay Chromosomal assay

Temperature Deletion rate Relative Deletion rate Relative

Genotype (deg) ( ~ 1 0 ~ ) rate n ( X

lo6)

rate n

A.

+

37 1.5 (0.85-2.0) = I 16 5.1 (3.4-7.9) = I 15

dnaE925 ₃₇ _{1.0 (0.82-1.9)} 0.66 12 ND

dnaQ49 (ts) 37 14 (7.8-19) 9.3 16 500 (200-810) 98 16

dnaQ (mutD5) 37 3.9 (2.8-6.5) 2.6 12 3100 (2200-11,000) 610 22

holCl02 : : kan ₃₇ _{8.1 (6.3-14)} 5.4 21 80 (53- 110) 16 16

holE202: : cat _{37 1.8} _(1.3-2.7) 1.2 19 3.4 (3.0-6.0) 0.67 16

ApolBl 37 1.8 (1.3-2.0) 1.2 24 5.3 (3.1-7.8) 1

.o

16

B.

+

37 ND _{6.7 (4.7-12)} =1 32

ApolA 37 ND 3.4 (2.3-4.5) 0.51 30

priA2: : kan 37 ND 66 (55-97) 9.9 22

c.

+

30 1.1 (0.59-1.6) =1 24 4.8 (3.8-7.2) =1 16

dnaBl07 (ts) 30 22 (18-36) 20 16 600 (470-750) 130 16

dnaE486 (ts) 30 8.0 (7.3-12) 7.3 25 9.8 (5.6-18) 2.0 16

dnaN159 (ts) 30 15 (13-21) 14 11 _{0.64 (0.60-1.1)} 0.13 24

dnaX2016 (ts) 30 0.84 (0.39-1.0) 0.76 16 _{0.17 (0.11-0.62)} 0.035 11

D.

+

₂₅ _{0.95 (0.74-1.2)} = I 37 ND

dnaCl ₂₅ _{10 (4.3-19)} 11 24 ND

dnaTl2 dnaC2 (ts) 25 14 (9.8-17) 15 17 ND

All strains are derivatives of AB1157 and are described in Table 1. The strains in A and B were assayed at 37" and the strains in C and D were assayed at permissive temperatures of 30 and 25", respectively. The strains in A, C and D were grown on LB medium, while the strains in B were grown on minimal complete medium. All mutant strains were assayed in parallel with the wild-type control strains AB1157 or STL695. Rates were calculated by the method of the median (LEA and COULSON 1949) and represent an estimation of the number of deletion events that occurred in the population, calculated from the observed median number of Tc' colonies for the number of independent cultures ( n ) , as described in the MATERIALS AND METHODS. Numbers in parentheses correspond to 95% confidence values for the measured rates. Relative rates were determined by comparison to the control strain assayed under the same growth conditions. ND, not determined.

causes constitutive expression of SOSregulated genes, However, the SOS response may augment the deletion

showed wild-type levels of deletion formation in both rate in this strain somewhat, which accounts for the

the plasmid or chromosomal assays. Furthermore, the twofold decrease in rates seen in both recA and ZexA

large increase in deletion formation promoted by E mu- Ind- dnuQmutant derivatives relative to rec+ kx+ dnuQ

tations does not require the SOS response with the strains. In contrast to dnaQ a substantial proportion of

double mutant dnaQ49 kxA3 (Ind-) showing a 47-fold increased deletion formation in

dnuBlO7

strains was

increase of deletion formation in chromosomal assays. ZexA-dependent. Deletion rates on the chromosome

TABLE 3

Deletion rates in red derivatives

~~~ ~

GenotvDe (deg) ( ~ 1 0 ~ ) rate n (X106) rate n

A. A recA304 37 0.69 (0.38-1.3) =1 16 5.2 (2.7-5.7) =1 16

ArecA304 dnaQ49 (ts) 37 1.2 (0.48-2.3) 1.7 19 270 (60-610) 52 12

B. ArecA304 30 0.51 (0.28-0.56) =1 23 5.3 (2.9-7.6) =1 16

ArecA304 dnaB107 (ts) 30 2.3 (1.1-5.7) 4.5 19 9.5 (6.3-1.7) 1.8 16

ArecA304 dnaE486 (ts) 30 5.4 (2.3-7.8) 11 23 ND

All strains are derivatives of AB1157 and are described in Table 1. The strains in A were assayed at 37" and the strains in B were assayed at a permissive temperature of 30°, in parallel with ArecA304 control strains JC10287 or STL753. Rates were calculated for the number of independent cultures ( n ) as described in MATERIALS AND METHODS. Values in parentheses correspond

(9)

465 Hyperdeletion in Replication Mutants

TABLE 4

Deletion rates for kxA derivatives

Genotype (deg) ( ~ 1 0 ' ) rate n ( X 106) rate n

Zed3 (Ind-) 37 ND 4.3 (2.2-8.6) 0.84 16

Zed3 (Ind-) dnaQ49 37 ND 240 (91

-

280) 47 12

Zed3 (Ind-) dnuBl07 (ts) 30 ND 140 (47-350) 29 15

sulAll Zed71 (De0 37 1.7 (1.1-2.8) 1.1 20 12 (8.3-22) 2.4 16

All strains are derivatives of AB1157 and are described in Table 1. The strains were assayed in parallel with the wild-type

control strains AI31157 or STL695. Rates were calculated for the number of independent cultures (n) as described in MATERIAIS

AND METHODS. Values in parentheses correspond to the 95% confidence limits for the rates. Relative rates were determined by

comparison to the control strain assayed under the same growth conditions. ND, not determined.

S U l A l l 37 1.1 (0.76-1.6) 0.73 20 9.2 (5.7-14) 1.8 12

were 29-fold elevated over wild-type strains for Zed3

dnaBlO7 compared to 130-fold for lex+ dnaBlO7 derivatives. Mutations in the DnaB fork helicase and primosome protein therefore promote deletions, in part, via

a mechanism requiring induction of the SOS response.

Analysis of plasmid deletion products reflects two

types of Red-independent deletion formation: There are several mechanisms that may give rise to deletion events between tandem duplications. Unequal sisterchromosome exchange can lead to deletion formation; such events fuse circular replicons and can be detected in the plasmid assay by the formation of dimeric plasmid deletion products. Other mechanisms of deletion formation, such as slipped mispairing, are expected to produce only monomeric products (see Figure 4).

Examination of the plasmid deletion products for their monomeric or dimeric structure can therefore be informative about the particular deletion mechanism

in play. However, product analysis in recA+ strains is

confounded because of the occurrence of recombination between plasmids before and after the deletion event. For this reason, we have examined products in the recA strain background, where there is no detectable plasmid recombination (and plasmid dimerization) in

the absence of the deletion event. Among deletion

products in recA strains, 40% were dimeric plasmid

products that represent intramolecular recA-indepen-

dent sister-chromosome exchange. The remaining 60%

of products were monomeric, formed presumably by

slipped mispairing ( LOVETT et al. 1993).

To determine whether mutations in the replication machinery affect one of these mechanisms specifically, we analyzed the plasmid deletion products of selected

replication mutants (Table 5). Among dnaE486 ArecA

mutants assayed at their permissive temperature, where an 11-fold stimulation of recA-independent deletion is observed, proportionately more products are found in

the dimeric form. Conversely, in dnaQ49 ArecA strains,

where recA-independent deletion events are stimulated

twofold relative to dnaQ+ ArecA strains, more products

are found in the monomeric form. The transformation

efficiency of monomer and dimer forms into these mu-

tants, with and without a coresident monomer plasmid (data not shown), confirmed that the skewed product

distribution is not merely due to a change in the estab-

lishment or maintenance of various plasmid forms.

Rather, these results suggest that polymerase mutation

dnaE486 increases the likelihood of a dimer-producing deletion event, such as sister-chromosome exchange

(see Figure 4). The editing subunit mutation dnaQ49,

on the other hand, increases the likelihood of monomer-producing mechanism such as slipped mispairing on the same strand (as shown in Figure 4). The antimu-

tator polymerase mutation dnaE925 and helicase muta-

tion dnaBl07have no effect on the distribution of dele-

tion products. Because dnaB107 caused a 4.5-fold

stimulation of recA-independent deletion, both dimer- and monomer-producing pathways must be equally enhanced.

DISCUSSION

We have demonstrated that mutations in many replication genes affect deletion events at tandem repeats. For most of the mutant strains examined, we observed increased deletion of repeats both carried on a

multicopy plasmid and on the E. colichromosome. How-

ever, for dnaQ and dnaB mutants there were much

larger increases in chromosomal as compared to plasmid deletion rates. This discrepancy may be due to

several reasons. (1) The differences between plasmid

and chromosomal data may reflect a real difference in

the way that replication occurs on the plasmid us. the

chromosome. Both the chromosome and the plasmid

depend on Pol I11 for normal replication although the

structure of the replication fork could differ in some way [there are differences in the genetic requirements

for initiation and primosome assembly (KORNBERG and

BAKER 1992)l. (2) The chromosome may be subject to

(10)

TABLE 5

Deletion product analysis for dnuB, dnaE and dnaQ mutants

Growth

temperature Dimer

Genotype products

A. A recA304 ₃₇ _{32/82 (39)}

ArecA304 dnaQ49 (ts) 37 15/93 (16)

ArecA304 dnaE925 ₃₇ _{32/82 (39)}

B. ArecA304 ₃₀ _{48/140 (34)}

ArecA304 dnaBlO7 (ts) 30 39/107 (36)

ArecA304 dnaE486 (ts) 30 54/87 (62)

All strains are derivatives of AB1157 and are described in

Table 1. The strains in A were grown at 37" and in B at

30". Dimer product ratios represent the number of deletion products that were dimeric plasmids (sister-chromosome ex-

change events) out of the total number of plasmid deletion

products examined for each strain, with the percentage in

parentheses. The alternative product was monomeric plas-

mid, equivalent to pBR322.

x'

analyses showed that the distri-

butions of products in strains dnaE486 and dnaQ49 were sig-

nificantly different from control strain ArecA304.

on plasmids. For example, the plasmid lacks

x

se-

quences that could stimulate recombinational reactions on the chromosome. (3) Differences observed may reflect relative recovery of products. For multicopy plasmids, deletion intermediates may be selectively lost. In- deed, plasmid loss may be magnified in mutants that impair replication, causing reduced recovery of plasmid

deletion products. Determination of plasmid copy num-

ber (data not shown) suggested that dnaB and dnaC

dnaT mutants, in particular, have two- to fivefold low-

ered levels of plasmid DNA, which could lead to an

underestimate of plasmid deletion rates in these strains.

The other replication mutants tested here were not

significantly affected in this analysis (data not shown).

In two cases, the dnaN and dnaX mutant strains, we

observed seemingly contradictory results: no effect or an increase in deletion events for repeats on the plasmid but a large decrease in deletions of the same repeats on the chromosome. These mutations affecting the processivity clamp may truly affect tandem repeat

deletion in opposite manners on a plasmid us. a chro-

mosome replicon. An alternative explanation, which we prefer, is that, in these mutant strains, chromosomal deletion products may be less likely to be recovered. For instance, aberrant or incomplete replication may

initiate deletion formation on both the plasmid and

the chromosome. However, on the chromosome, the deletion intermediate may fail to be processed into a viable deletion product. Because the plasmid presents a shorter replication template, it is possible for ineffi- cient Pol I11 replication or uncoupled leading and lagging strand replication to be completed or replaced by repair replication via Pol I or 11. On the much larger chromosome, the same situation may lead to failure to

complete replication and loss of the potential deletion product.

Of E. coli's three known DNA polymerases, only func-

tions associated with Polymerase 111, responsible for

chromosomal replication, affected deletion at tandem repeats in our experiments. This includes mutations in

the a polymerase (dnaE gene) and E exonuclease

(dnaQ) subunits of the core enzyme as well as mutations

in subunits of the y complex (dnaX, hoZC) and the ,B

clamp (dnaN). Mutants in priA, dnaB, dnaC and dnaT

with defects in the primosome or primosome assembly also exhibited high deletion rates. Another mutant of

Polymerase 111, the antimutator dnaE (FIJALKOWSKA and

SCHAAPER 1995) did not show altered deletion rates. Deletions of genes encoding Polymerases I and I1 had no effect on deletion rates in otherwise wild-type strains. This result does not, however, exclude the possibility that these polymerases also contribute in a minor way

to deletion formation. The ssb-113 mutation of single-

strand DNA binding protein had been previously shown

to elevate deletion formation fourfold in the same plas-

mid assay as used here (LOVETT et al. 1993). This mutant

protein has normal DNA binding properties and is be-

lieved to be defective in protein-protein interactions

(CHASE et al. 1984) with components of the replication complex.

We believe that deletion formation is stimulated in these mutants by aberrant DNA replication rather than other indirect effects. The phenotype of elevated deletion frequency is not correlated with slow growth. Dele- tion rates in wild-type were found to be largely independent of growth medium and temperature. Whereas some slow-growing replication mutants showed elevated deletion rates (dnaB), others showed essentially wild-

type deletion rates despite poor growth (poZA)

.

In addi-

tion, our results suggest that induction of the SOS re-

sponse does not itself stimulate deletion formation nor

is it required for the elevated rate seen in some hyperde-

letion mutants, as in the case of the dnaQ mutant. Al-

though this suggests that aberrant replication in a vari- ety of replication mutant strains can lead to enhanced deletion formation at tandem repeated sequences, this does not necessarily mean that deletion events in wild- type strains occur strictly during the process of replication. Recently, we have presented genetic evidence that most deletions formed between 101-bp tandem repeats occur during, or shortly after, DNA replication because they can be subject to correction dependent on hemi-

methylation (LOVETT and FESCHENKO 1996). Moreover,

the reduced recovery of chromosomal deletion products in dnaNand dnaxmutants also implies that replication is an essential component of the deletion process.

More detailed analysis of several selected mutants

(dnaB, dnuE and dna4) suggested that aberrant replication can stimulate deletion through different pathways. The hyperdeletion phenotypes for polymerase mutants

(11)

Hyperdeletion in Replication Mutants

whereas that for dnaR helicase mutant was primarily rPcAdependent. Also, a functional SOS response was necessary to see the full stimulatory effect of dnaR mutations on deletion formation, in contrast to dnuQ. The mutant polymerase must therefore stimulate a recA and Zed-independent deletion pathway whereas the mutant DnaB helicase induces primarily a recA and Zeddepen- dent recombinational deletion pathway.

Analysis of the structure of plasmid deletion products has revealed that there are two mechanisms of RecA- independent deletion formation. Deletion formation on circular replicons results in both monomeric and dimeric forms. According to replicational models of deletion, these products reflect distinct mechanisms: simple slippairing, resulting in monomer replicon

products, and sisterchromosome slippairing, resulting in dimeric replicon products (Figure 4). The distribution of products in the dnaQ49 strain was skewed to monomers and that in dnaE486 strains was skewed to dimers, relative to the proportion observed for wild- type strains. Product analysis in a dnaB strain revealed a distribution similar to wild type. We conclude that aberrant replication can stimulate deletions in both a RecAdependent and -independent manner, and via both simple slippage and the sisterchromosome exchange mechanisms. Different replication mutants differ in the mechanism by which these deletions occur.

The correlation of hyperdeletion phenotypes with specific biochemical defects in replication fork propa- gation may shed light on the mechanisms of deletion formation. However, more exhaustive genetic and biochemical analysis will be necessary before clear conclu- sions can be drawn. For many of the mutants examined here, the specific consequences of the mutation on the structure or function of the replication complex in vivo is not well understood. The generality of phenotypic effects is also not clear because only one mutant allele was examined in most cases. Moreover, many of our assays were performed at “permissive” temperatures, where the function of temperature-sensitive components of the replication machinery may be impaired in subtle ways. It is not known whether observed effects on deletion formation are genetically recessive or domi- nant, and therefore due to a missing or an altered activity. Deletion events occur rarely, in a small fraction of cells, and therefore may not necessarily correlate to the biochemical defects determined for the cell population

as a whole. Nevertheless, in theory, the biochemical functions affected by these hyperdeletion replication mutants allow us to speculate on several ways by which deletion formation may be stimulated.

Some mutants may encourage deletion by promoting the persistence of single-strand DNA in the replication fork. Mutations that adversely affect replication on only one strand may cause extended stretches of single- stranded DNA to accumulate during replication. Muta- tions affecting the primosome complex (e.g., dnaB,

467

Replication gap

“:::z&Rw>>>>>.

A,. ,. :.:*, %... ..x.< ,..% <Target for slippage

is unpaired

\

Slipped realignment

.:,:.s.:.z<. Ey >>y X?> :*

. . . c<:sz.*:::::

Normal replication

y

.

:

#

,

I

,

.

I

.

<Target for slippage is already paired

FIGURE 5.-Persistence of singlestrand DNA promotes slipped realignment. If a replication gap encompasses a copy of the tandem repeat, the nascent strand is free to realign at the second repeat. This slipped realignment, if unrepaired, will give rise to deletion. During normal replication, no such singlestrand DNA is available for alternative pairing and slipped alignments are discouraged.

dnaC, dnaTand p i A ) could be expected to disturb replication specifically on the lagging strand, either by un- coupling leading and lagging strand replication or by leaving gaps of unreplicated ssDNA on the lagging strand.

Persistence of single-strand gaps could promote a RecAdependent mechanism of deletion formation. Daughter-strand gaps caused by replication-blocking lesions in DNA, such as UV-induced pyrimidine dimers, are filled by recombinational postreplication repair (Figure 2 ) . This repair process requires RecA protein in order to recombine nascent strands across the replication fork (RUPP et dl. 1971). Alternatively, a stalled replication fork may be broken and then reestablished by RecAdependent recombination with its sister chromosome ( h A I et al. 1994; KUZMINOV 1995a,b) (Figure 3). One or both of these mechanisms may be induced by the dnaR mutation which stimulated deletion formation in a recA- and lexAdependent fashion. The dnaR mutation may block unwinding of the fork or leave daughter-strand gaps by disrupting primosome function, thereby inducing the recombinational repair processes that culminate in deletion.

RecA-independent deletion formation may also be induced by the accumulation of single-strand DNA in the replication fork (Figure 5). Strand realignment would be disfavored if the secondary pairing site had already been replicated to duplex DNA; available tar- gets of single-stranded DNA template would promote slippage and mispairing either on the same strand or across sister strands. A minor component of the stimulation by dnuR on deletion formation was recA-independent and may occur by this mechanism.

(12)

468 C. J. Saveson and S. T. Lovett

Displacement

of

nascent strand during replication

4

.:.:.:.:.:.:.:.~.x.:.:.:.:.: k??: ::::::: ::;::::

=

*e

:C*S.X.h >x+x.:.:.:.x.:.:.:.:.>

J

Slipped realignment

1 ₁

FIGURE 6"Prevention of deletion by the exo- nuclease activity of e ( d m @

protein. Simple slipped re- alignment may be initiated by the displacement of the nascent strand, which s u b

sequently mispairs with its t e m p l a t e s t r a n d . Exo- nuclease degradation of the displaced single strand o r

of the mispaired strand by the 3' exonuclease activity of DNA polymerase subunit

E will abort the deletion

event.

Deletion on nascent strand

ciation of the polymerase could free the nascent DNA strands and allow their translocation across the fork or on their template strands to produce slipped mispairing. Mutations in the a subunit of the polymerase may stimulate Red-independent recombination via this mechanism.

Other mutants in the

p

clamp and clamploading/ recycling machinery y complex may stimulate deletion formation by decreasing processivity of the polymerase. We would expect mutations in the clamp or mutations to the polymerase that hinder its interactions with the clamp, to inhibit the processivity of the enzyme complex, thereby increasing the incidence of deletions. As with the primosome, one might expect defects in the clamp loader to affect lagging-strand synthesis more than leading-strand synthesis, slowing replication or leaving single-stranded gaps. However, in some cases, deletions on the chromosome may not be recoverable because of failure to reinitiate and complete replication.

Replication mutations may also stimulate deletion by allowing slipped mispairing intermediates to persist. We observed that both the dnaQ49 and mutD5 mutation increased deletion events. At least in the case of dnuQ49, this increase was via a recA-independent mechanism. This may be via simple slipped mispairing, as we observed proportionately more monomer deletion products in dnaQ49mutants. Figure 6 illustrates that, during normal replication, the nascent strand may be displaced from its template. The E subunit of Pol

I11

may degrade

this displaced single strand from its 3' end and thereby prevent slipped pairing events from occurring. Alterna- tively, E may degrade the 3' end of the nascent strain

in the slipped alignment with its template and abort the deletion process. Although these scenarios postu-

late that it is the loss of the exonuclease activity that is responsible for the dnaQeffects, E has also been shown

to modulate a and the processivity of the polymerase (STUDWELL and O'DONNELI. 1990) and this property may influence the deletion process, as described above. Our observations that mutations in the replication machinery may greatly stimulate genetic rearrangements may have implications for human genetic disease. Rearrangements between tandem repeated sequences or genes are a source of genetic mutation in humans (MEUTH 1989; K R A W C ~ A K and COOPER 1991;

HU and WORTON

1992; NEISON 1993). It is likely that aberrant replication in humans, as in other organisms, can also promote deletion events. Mutations in the anal- ogous replication functions of human cells to those we have investigated here may well predispose certain individuals for more frequent genetic rearrangements and genetic disease.

We thank PAOIA DRAPKIN and ~ . E R GI.UCKMAN for preliminary

work o n this project and VIADIMIR FESFIENKO for construction o f the

poL4 mutant strain. We are indebted to the following individuals for

providing strains: B. BACHMANN of the P;. coli Genetic Stock Center,

A. J. CIARK, M. GOODMAN, C. GROSS, R. KOI.ODNF,R, K. MARIANS, R.

MAURER and R. SCIIAAPER. We thank the anonymous reviewer for

suggesting the statistical method for setting 95% confidence limits for

rate determinations. This work was supported by U.S. Public Health

Service grants R 0 1 GM-51753 to S.T.L. and T32 GM-07122 to C:J.S.

LITERATURE CITED

AIBERTINI, A. M., M. HOFER, M. P. C h o s and J. H. MII.I.ER, 1982 O n the formation o f spontaneous deletion: the importance of short sequence homologies in the generation of large deletions.

Cell 2 9 319-328.

ARAI, K., and A. KORNRERG, 1981 Unique primed start of phage

4x174 DNA replication and mobility of the primosome in a

direction opposite chain synthesis. Proc. Natl. Acad. Sci. USA