Directed Evolution and Identification of Control Regions of ColE1 Plasmid Replication Origins Using...

doi:10.1016/j.jmb.2005.06.051 J. Mol. Biol. (2005) 351, 763–775

Directed Evolution and Identification of Control Regionsof ColE1 Plasmid Replication Origins Using OnlyNucleotide Deletions

Dewey Kim1, Yoon Rhee2, Denise Rhodes2, Vikram Sharma1

Olav Sorenson3, Alan Greener2 and Vaughn Smider1*

1IntegriGen, Inc., 42 DigitalDr. Bldg. 6, Novato, CA 94949USA

2Stratagene, 11011 N. TorreyPines Road, La Jolla, CA 92037USA

3Anderson School of BusinessUniversity of California, LosAngeles, CA 90095, USA

0022-2836/$ - see front matter q 2005 E

Abbreviations used: cfu, colony-fb-galactosidase; LTR, long terminalnuclease-mediated deletional mutagE-mail address of the correspond

[email protected]

Genes can be mutated by altering DNA content (base changes) or DNAlength (insertions or deletions). Most in vitro directed evolution processesutilize nucleotide content changes to produce DNA libraries. We testedwhether gain of function mutations could be identified using a mutagenicprocess that produced only nucleotide deletions. Short nucleotide stretcheswere deleted in a plasmid encoding lacZ, and screened for increasedb-galactosidase activity. Several mutations were found in the origin ofreplication that quantitatively and qualitatively altered plasmid behaviorin vivo. Some mutations allowed co-residence of ColE1 plasmids inEscherichia coli, and implicate hairpin structures II and III of the ColE1RNA primer as determinants of plasmid compatibility. Thus, useful andunexpected mutations can be found from libraries containing onlydeletions.

q 2005 Elsevier Ltd. All rights reserved.

Keywords: molecular evolution; origin of replication; mutagenesis; plasmidreplication
*Corresponding author
Introduction

Molecular evolution techniques have been usedto engineer enhanced or qualitatively different genefunctions across a broad range of biopharmaceu-tical and industrial products. Of paramount import-ance in molecular evolution strategies is the abilityto search the greatest possible functional sequencespace through the creation of diverse libraries.Sequence space is most often described in terms ofthe length and content of a gene or protein,according to SZ4N (for DNA) or SZ20N (forproteins), where 4 and 20 are the number ofdifferent nucleotides or amino acids, and N is thelength of the gene or protein.1 Most methods toevolve genes involve mutagenesis techniques thatalter the DNA content, but not the DNA length.Many related genes and regulatory regions,2

however, differ in length as well as sequencecontent. Additionally, one of the most well-studiedexamples of in vivo molecular evolution, antibody

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orming units; b-gal,repeat; NuDel,enesis.ing author:

gene recombination and affinity maturation, utilizesDNA end-joining mechanisms that result indeletion and insertion events.3 This junctionaldiversity programs the affinity pathway of theantibody.4

A hallmark of any molecular evolution strategy isthe ability to produce gain of function mutations fora defined biological assay. In this regard severalstrategies have been successful. A popular tech-nique is “DNA shuffling”, wherein homologoussequences are recombined to produce DNAlibraries, which are then screened for enhanced oraltered functions.5,6 If homologous sequences differin length, these length differences could be presentin the selected sequences. However, it is difficult toascribe positive functions to the insertions ordeletions due to the concomitant nucleotide contentchanges. We asked whether deletional events alonecould provide a means to enhance genes, and thusexplore an area of sequence space not addressed bycurrent methods.We developed a method termed nuclease-

mediated deletional mutagenesis (NuDel) thatallows a random number of deletions to occur at asingle unspecified position in plasmid DNA. Inorder to enhance expression of a model gene, thistechnique was applied to expression of the

d.

764 Evolution of Novel Plasmid Replication Origins

a-galactosidase (b-gal) a-peptide in Escherichia coli.We used a simple blue/white color screen toidentify several mutants with increased b-galactivity. All of the mutations mapped to the plasmidorigin of replication. Additionally, they alteredpredicted RNA structures known to function inplasmid replication as well as compatibility. In thisregard, we found that some of the mutantsestablished new compatibility groups, and identifyhairpins II and III of the ColE1 RNA primer asrequired structures for plasmid incompatibility.Thus,NuDel provides ameans to enhance expressionby affecting cis-acting regulatory elements, but alsoallows the generation of new and qualitativelydifferent genetic regulatory properties.

Results

Deletional sequence space

A DNA sequence of length N has 4N differentpossible variants if the length is unchanged, and allpositions can be mutated to any of the bases.However, if the length is allowed to vary, newsequences will be present in a theoretically com-plete “library” that are not within the subset of 4N.The main determinants for complexity in a directedevolution library are: (i) the length of the startinggene; (ii) the maximum number of positions thatcan be simultaneously modified; and (iii) thenumber of mutations allowed at any given position.For the last variable, four bases (adenine, thymine,guanine, and cytosine) are allowed at any fixedposition for DNA content changes (without lengthchanges). However, in the case of insertional ordeletional events, any number of additions ordeletions could occur at each position. In Table 1we estimate the reachable sequence space (i.e. thenumber of different sequences that could begenerated from a starting sequence) of deletionalevents versus replacement events for the samenumber of maximum positions altered in a modelgene of 1000 base-pairs. Two points seem worthnoting. First, even with only five deletions perposition, deletional events produce a similar orderof diversity to replacement events. Since these“libraries” do not overlap, deletional events offer anovel source of sequence diversity. Second, the sizeof the sequence space produced by deletional

Table 1. Reachable sequence space from replacement versus d

Deletio

Max number of del

Positions mutated 1 5

1 1.0!103 5.0!12 1.0!106 2.5!13 1.0!109 1.3!14 9.9!1011 6.2!15 9.9!1014 3.1!1

events increases at a faster rate than that generatedby replacement events when the number ofpotential deletions per position grows large. Putmore simply, all of the diversity from replacementmutations comes from the set of sequences of lengthN, whereas the sequences created by deletions allbelong to the set of sequences of length !N. Thus,significant sequence space clearly exists outside ofthat comprised of base changes alone. In order totest whether deletions alone could produce gain offunction mutations, we devised a strategy wheremembers of a library each contained a singlemutation of varying lengths, but at differentpositions. Thus, only the sequence space encom-passed in the first row of Table 1 could be searched.

Strategy

We chose the lacZ a-peptide as a model system totest the feasibility of isolating gain of functionmutations caused by short deletions in geneticregulatory regions. Several regulatory regions thatcould affect lacZ expression were present on aparent vector (Figure 1), including the lac promotor,operator, origin of replication, and untranslatedRNAs. Figure 1 shows the general scheme forNuDel, wherein a library is constructed containingshort deletions at unspecified positions, followedby a screen for enhanced expression.

Generation of short, non-specific deletions

In order to delete nucleotides at non-specificpositions within a nucleic acid, we required amethod to create a single double-strand break at anon-specific position. Some nucleases, like DNase I,can be titrated to produce a nicked or linearizedplasmid,7 however the enzymatic activity continuesto degrade DNA following the linearization event.Other methods, such as chemical cleavage of single-stranded DNA,8 require several steps as well ascareful titration and purification of the linear DNAspecies. Our strategy assumed that a single-stranded nuclease, like S1, P1, or mung beannuclease, would cleave single-stranded regions intightly supercoiled DNA,9 thus producing a nick.Nicked DNA is a natural substrate for theseenzymes,10,11 and would subsequently cleave theDNA to produce a double-stranded break inthe same reaction. Following cleavage, however,

eletion mutations for a typical 1000 bp gene

ns Replacements

etions/position

3 bp/position25

03 2.5!104 3.0!103

07 6.2!108 9.0!106

011 1.6!1013 2.7!1010

014 3.9!1017 8.1!1013

018 9.7!1021 2.4!1017

Figure 2. (a) S1 nuclease can linearize plasmid. Theplasmid pLacZi was untreated (lane 1), treated with thesingle cutting restriction enzyme ClaI (lane 2), orincreasing concentrations of S1 nuclease (lanes 3–6).(b) Cleavage position is non-specific. The linear plasmidfrom lane 6 was run in lane 7, or further cleaved with ClaIand run in lane 8.

Figure 1. General scheme for NuDel. The startingvector pBluescript II KS is subjected to a process whereeach plasmid molecule ultimately contains a singlemutation comprising a short deletion of a non-specificnumber of nucleotides at a random position. This“library” is then screened for enhanced b-gal activity.The lac promotor (lacP), which also contains the lacoperator, a-peptide (a-pep), and ColE1 origin of replica-tion are indicated.

Evolution of Novel Plasmid Replication Origins 765

the single-stranded regions are not present, sincethe plasmid is not supercoiled, and the relaxedplasmid can no longer act as a substrate for theenzyme. Thus, a single double-strand break couldbe placed at non-specific positions in a plasmid,with a population of plasmids containing breaks atdifferent positions.

To test this hypothesis, supercoiled plasmid DNAwas incubated with either a restriction enzymeknown to cut a single time or S1 nuclease (Figure 2).In Figure 2 in the left panel, the DNA plasmid waseither uncleaved (lane 1), cleaved with the singlecutting restriction enzyme ClaI (lane 2), or increas-ing concentrations of S1 nuclease (lanes 3–6). Withhigh concentrations of S1 nuclease virtually all ofthe supercoiled DNAwas converted to linear form,with little effect on relaxed plasmid. In the rightpanel of Figure 2, the linearized plasmid from lane 6was gel purified and run in lane 7, or furtherdigested with ClaI and run in lane 8. This reactionwas done to ensure that S1 did not cleave site-specifically in the plasmid, in which case one ormore bands would be present when digestedwith ClaI. Since ethidium bromide staining isproportional to DNA size, the smear from unit

length downwards in lane 8 indicates that cleavageby S1 was not site-specific or that significant“hot-spots” of cleavage were present. Althoughsingle-stranded regionsmight preferentially form atACT-rich regions, the analysis of our mutantsindicates that cleavage occurred efficiently in non-ACT-rich areas (Table 2).In order to evolve genetic regulatory elements

effectively through deletions, we first determinedthe feasibility of inducing short deletions ofdifferent lengths in DNA fragments. Exo III hasbeen routinely used to generate large deletions ingenes at restriction sites.12 In order to avoid largeand potentially deleterious mutations, we alteredreaction conditions such that Exo III could be usedto generate very small deletions. Using a 232 bpDNA fragment that was fluorescently labeled at one5 0 end as a substrate, we analyzed the 3 0–5 0 activityof Exo III. Deletions were detected on a DNAsequencing apparatus capable of single baseresolution (Figure 3). We were able to demonstratethat Exo III deletes nucleotides in a salt-dependentreaction. More specifically, as salt is increased thenumber of deletions decreases. Nearly 25 nucleo-tides can be removed under conditions of 100 mMNaCl (Figure 3, second panel), up to 15 nucleotideswith 150 mM NaCl (third panel), and a fewnucleotides with 200 mM NaCl (bottom panel).

Library construction

We then tested the feasibility of creating a librarythat contained only deletions. We linearized thepBluescript II KS plasmid with the blunt-cuttingrestriction enzyme SmaI, dephosphorylated theends with calf intestinal phosphatase, and ligatedthe plasmid with phage T4 ligase. As expected,no colonies resulted upon transformation ofEscherichia coli. However, when the same dephos-phorylated plasmid was subjected to furthertreatment with Exo III (as described in Materialsand Methods) prior to ligation, several hundred

Figure 3. Exo III can makecontrolled deletions at a DNAend. The 5-FAM-labeled f232 frag-ment was analyzed on an ABI 373DNA sequencing apparatus in theabsence (top panel) or presence(bottom three panels) of Exo III.The electropherogram in the toppanel is untreated f232, and theother panels contain Exo III andare labeled according to theamount of NaCl added to thereaction.


colonies were formed. Six of six clones from thisligation lost the SmaI restriction site, and sequencedata revealed short deletions at the SmaI site (datanot shown). Additionally, we subjected gel purified,S1 linearized pBluescript to phosphatase treatment

Table 2. Characteristics of deletion mutants

Gain of function (LacC)

Mutant ID No. del. AT:GCFunctionallocationa

2.1 6 3:3 Ori2.2 3 2:1 Ori2.3 7 2:5 Ori3.2 4 2:2 Ori3.3 3 1:2 Ori3.4 11 9:2 Ori4.1 2b 0:2 Ori5.1 11 7:4 Ori5.2 5 5:0 Ori5.3 7 3:4 Ori

a Ori, origin of replication; P, promoter; CRP, cAMP receptor proteistart site. NA, not analyzed.

b Mutant 4.1 also had three sequence changes near the deletion si

prior to a test ligation, and found no colonies uponE. coli transformation. However, several hundredcolonies formed when the plasmid was furthertreated with Exo III prior to ligation. We concludethat removal of the 5 0 phosphate group by

Loss of function (LacK)

Mutant ID No. del. AT:GCFunctionallocationa

2.9 8 4:4 P, ATG5.7 4 3:1 CRP5.8 8 5:3 CRP7.1 15 9:6 ATG7.3 152 NA P, ATG7.7 6 5:1 a-Pep8.7 134 NA P, ATG9.1 10 5:5 a-Pep9.2 13 7:6 P9.3 13 9:4 CRP9.4 8 0:8 a-Pep9.5 12 6:6 CRP9.8 54 NA P

n binding site in the promoter; a-pep, a-peptide; ATG, translation

te, see Figure 5.


phosphatase blocked circularization by T4 ligase,and removal of the dephosphorylated nucleotide inthe following Exo III reaction allowed recirculariza-tion of a deleted product. Thus, phosphatasetreatment eliminated “background” recirculariza-tion of wild-type plasmid that had not undergonedeletion. We expect that all resulting colonies mustcontain a short deletion somewhere in the plasmid.

Library screening

In order to test the ability of deletions to affectgene function, the pBluescript II KS plasmid waslinearized with S1 nuclease, gel purified, dephos-phorylated, and subjected to Exo III digestion asdescribed above, then re-ligated. E. coli weretransformed and incubated at 30 8C on LB platescontaining X-Gal to measure b-galactosidaseactivity. Additionally, unmodified pBluescript plas-mid was plated on X-Gal-containing plates tocontrol for any in vivo mutagenic effects on theplasmid. We screened 4000 colonies from twoindependently constructed deletion “libraries”. Wefound several deep blue colonies in the deletionlibrary arm, representing about 0.5% of the totalcolonies (Figure 4). Among the 10,000 coloniesscreened from the unmutagenized plasmid, nomutants of pBluescript were found with thisphenotype. As a control, we incubated the deletionlibrary on LB plates containing X-Gal and IPTG,where wild-type plasmid will confer a bluephenotype, and found that nearly 30% of thecolonies were white. No white colonies were seenwhen pBluescript was incubated under theseconditions. Thus, loss of function mutations(LacK) can also be identified using this approach.

Figure 4. Phenotype of a LacC mutant colony. A gain offunction mutant colony is seen surrounded by whitebackground colonies on an LB agar plate without IPTGinducer.

Mutation analysis

Plasmids from the LacC mutant cells werepurified and transformed into fresh E. coli andfound to transfer the dark blue phenotype. Whenthe plasmids were sequenced, ten independentmutations were localized to the origin of replication(Figures 5 and 9). The entire lac promotor anda-peptide coding region was sequenced in allmutants, with no alterations detected. The ColE1origin of replication is over 1 kb long; however, ourten mutations clustered within the first 200 nucleo-tides. Several mutations were isolated twice;however, the mutations for 2.1 and 3.4 arose fromindependent libraries, indicating that they werefound from independent mutagenic events. Thedeletions in mutants 2.1, 3.2, 4.1, and 5.3 overlapone another in the ColE1 origin, as do 3.4 and 5.2(Figures 5 and 9). The number of nucleotidesdeleted ranged from two to 11 (Table 2, secondcolumn), with a mean of 5.9. Only three of themutations (on plasmids 3.4, 5.1, and 5.2) occurred inACT-rich areas (Table 2, third column). Onemutant(4.1) also had three nucleotide changes immediatelyadjacent to the deletion (Figure 5).We also analyzed 13 loss of function mutants (i.e.

white colonies in the presence of IPTG) and foundseveral deletions in the lac promoter and a-peptide(Table 2, right). Three of these mutations were over50 nucleotides long and encompassed both thepromoter and a-peptide. Of the ten shortermutations, five were in the promoter, four wereentirely within the coding region, and two deletedthe ATG translation start site. The average deletionlength was 33.6 nucleotides, however exclusion ofthe three large deletions resulted in a mean of 9.7nucleotides. Figure 6 illustrates the location of themutations with relation to known control elementsin the lacZ region.

Functional analysis

Since all of the gain of function mutations werefound in the origin of replication, we tested theplasmid copy number for each mutant at 30 8C(Figure 7). Whole cell nucleic acid analysis revealedthat the mutants had an increase in the amountof mutant plasmid compared to pBluescript(Figure 7(a), compare the first lane to the others).In Figure 7 the chromosomal DNA, and RNAspecies provide internal controls for the nucleicacid extraction. A twofold titration of pBluescriptand 3.4 extract revealed at least a fourfold increasein plasmid for the mutant (Figure 7(b)), despiteslightly more chromosomal DNA found in the wild-type cells. Miniprep plasmid yield was alsoapproximately fivefold higher in the mutants(data not shown). Furthermore, when the originfrom mutant 3.4 replaced the origin of pUC18, theplasmid copy increase as well as b-gal activity wastransferred to pUC18 (Figure 8). The large increasein b-gal activity is possibly due to titration of the lacrepressor in these mutants, allowing an “all or

Figure 5. ColE1 mutant multiple sequence alignment. Mutant sequences were aligned by ClustalW. Uppercase lettersindicate the region transcribed into RNAII. The region transcribed into RNAI is in a black background. Numbering isaccording to the scheme of standard ColE1 origins. Dashes indicate deletions. Mutant 4.1 contained three nucleotidechanges near the deletion, which are underlined.


none” Lac phenotype. The mutants 2.1 and 5.1 wereobserved to confer a moderate slow growthphenotype on the E. coli cells (data not shown).Mutant 5.3 had a very severe growth defect,precluding its analysis in Figure 7. Because someplasmids conferred a growth phenotype, weperformed some stability studies. All of theplasmids were grossly stable as determined byrestriction analysis of minipreps after 100 gener-

Figure 6. Location of loss of function (LacK) mutations. Trepresenting the deletion locations producing white coloniessite, K35 and K10 RNA polymerase binding sites, and lac op

ations of growth (data not shown) and plasmids 3.2,3.3, and 3.4 were resequenced after a year ofpassage, with no sequence changes detected. Themutants 2.1 and 5.1 appeared to have a segregationdefect upon analysis of the copy number of severalindependent colonies; however, we are performingadditional tests to confirm this observation.

In ColE1-type plasmids (such as plasmids in thepBR, pUC, and pBluescript families), both copy

he sequence of the entire lac region is shown, with barsin the presence of IPTG. The positions of the CRP bindingerator are indicated above the sequence.

Figure 7. (a) Plasmid copy number analysis of mutants.Total cell nucleic acid was prepared and analyzed byelectrophoresis and ethidium bromide staining. Threeindependent experiments were done with comparableresults. Wild-type pBluescript (lane 1) and the mutantsare annotated above each lane. The chromosomal band(Chr) and plasmid (Pl) band are noted. The mutant 5.3was not analyzed because of a severe growth phenotype.(b) A twofold titration of wild-type and mutant 3.4 totalcell nucleic acid extract (XT) was performed to estimatethe difference in chromosome to plasmid ratio.

Figure 8. Transfer of mutant phenotype to pUC18.(Left) Whole cell nucleic acid from pUC18 and the originfrom mutant 3.4 subcloned into pUC18 are analyzed.(Right) Relative b-gal activity was performed usingpBluescript KS as 1. Mutant 3.4 (second bar), pUC18,and the 3.4 origin replacing the ColE1 origin from pUC18were analyzed in duplicate. Error bars representC2.5 S.D.


number and plasmid compatibility are under thecontrol of a DNA region within the replicationorigin of the plasmid. A portion of this region istranscribed into RNAs called RNAI and RNAII.13

RNAII serves as a primer for DNA polymerase I,and priming is functionally down-regulated by theinteraction of RNAII with RNAI, which are inantisense orientation to one another (Figure 9(a)).Regulation of RNAII by RNAI is dependent on thelength of RNAII. Full-length RNAII can efficientlyprime DNA synthesis despite the presence ofRNAI. Short RNAII, however, forms a differentsecondary structure that efficiently binds RNAI andinhibits priming. The secondary structures formed

are thought to include a specific hairpin III that onlyoccurs in shorter RNAII transcripts, but that isabsent from longer transcripts. The interactionsbetween RNAI and RNAII are mediated by theunpaired bases at the tips of the hairpins.14–16 Manycommonly used ColE1 cloning vectors are highcopy variants of pBR that have mutations in thesequence that codes RNAI.17,18

Nine of the ten deletion mutants that we isolatedlocalized to the region corresponding to the overlapbetween RNAII and RNAI. The only mutationoutside of this region (2.2) was located four basesupstream from the K35 RNA polymerase bindingsite of the RNAI promotor (Figure 5). Given theimportance of the RNAII/RNAI interaction inregulating plasmid replication, we utilized acomputer program, MFOLD,19 to predict thesecondary structures and free energy of formationof RNAII and RNAI of our mutants. This programhas been used previously to analyze RNAII andRNAI, and its predicted structures were found tocorrelate closely with in vitro data.13 We analyzedthe positions of our mutants with regards to thepredicted structure of RNAII111, which includeshairpin III and is known to interact efficiently withRNAI.13 Mutants 2.1, 3.2, 4.1, and 5.3 were found tolocalize to hairpin I, with each deletion encompass-ing the unpaired bases at the tip (Figure 9(b)).Mutants 3.4 and 5.2 overlapped an unpairedpoly(A) “bubble” at the base of RNAII hairpin I.Mutants 2.3 and 3.3 overlapped with the unpaired

Figure 9. (a) RNA I–RNA II schematic. DNA replication of ColE1 type plasmids utilizes RNA II as a primer. RNAI, ashorter anti-sense transcript, negatively regulates effective primer formation, thus controlling copy number. (b) MFOLDof RNAII111 with mutant positions. The interactions between RNAII and RNAI are mediated by hairpin loops. Thehairpin loops I, II, and III for RNAII are shown and the positions of the deletion mutants are indicated by dark lines.

Table 3. MFOLD analysis of Ori mutations

MutantDG

RNAIDG

RNAII111

DGRNAII555

DG RNAII111–555(diff. from wt)

wt K49.7 K50.0 K230.0 K180.0 (0)2.1 K42.2 K42.5 K226.8 K184.3 (K4.3)2.2 K49.7 K50.0 K230.9 K180.9 (K0.9)2.3 K45.7 K40.9 K225.2 K184.3 (K4.3)3.2 K43.5 K43.4 K223.4 K180.0 (0)3.3 K47 K47.7 K232.0 K184.3 (K4.3)3.4 K34.6 K43.4 K227.7 K184.3 (K4.3)4.1 K47.4 K47.7 K231.5 K183.8 (K3.8)5.1 K45.7 K44.1 K228.4 K184.3 (K4.3)5.2 K43.1 K50.3 K234.6 K184.3 (K4.3)5.3 K39.4 K41.1 K259.3 K218.2 (K38.2)


bases at the tip of hairpin III, whereas mutant 5.1deleted five unpaired residues at the tip of hairpin IIas well as six residues forming the stalk. Weanalyzed the lac RNA by MFOLD to determine ifthe loss of function mutations also occurredpreferentially in unpaired regions; however, wefound no relation of these deletions to unpairedloops (data not shown).

We also compared the DG of formation of theRNAI, RNAII111, and RNAII555 structures for eachof the mutants, and compared them to wild-type(Table 3). In nine of ten mutants the DG of RNAIformation was higher compared to wild-type.Mutant 2.2 was identical with wild-type, due to itslocation outside of RNAI (Table 3, fourth column).Additionally, in eight of ten mutants the DG forformation of RNAII111 was also increased. This isconsistent with lower steady-state RNAI/IIcomplexation, which would result in decreased

inhibition of DNA synthesis primer formation.Mutant 5.2 had a slightly decreased DG forRNAII111, and mutant 2.2 was identical with wild-type, due to the location of the deletion outside of

Table 4. Compatibility of mutant plasmids

Incoming plasmid (KanR)

Harboredplasmid(AmpR) pBS 3.3 5.1

None 100 100 100pBS 4.8 25.5 43.83.3 37.1 4.1 28.25.1 55.8 60.4 4.0

The percentage of KanR plasmid retained after transformation ofE. coli cells that harbor an AmpR ColE1 plasmid.


RNAII111. Five of the mutations decreased DG offormation of RNAII555, the active primer for DNAreplication. This is somewhat surprising, given thefact that a decrease in length would generally beexpected to cause less intramolecular base-pairing,and hence increase the DG. However, this result isconsistent with the idea that these mutants conferan increased copy number, which may be explainedby increased stability of the RNAII555 primer. SinceRNAII111 is a precursor to RNAII555 in vivo, weanalyzed the difference in DG between RNAII111and RNAII555 compared to wild-type. SinceRNAII111 is inhibited by RNAI, this analysis couldgive a surrogate measurement of functional primerformation. Strikingly, all of the mutants had a freeenergy difference between RNAII111 and RNAII555that would favor formation of RNAII555, the activeprimer. Amazingly, six of the mutants had the exactvalue of K4.3. Mutants 2.2, 3.2, 4.1, and 5.3 hadvalues ofK0.9, 0,K3.8 andK38.2, respectively. Thelarge difference in mutant 5.3 is largely due to thefact that this mutation leads to formation of aqualitatively different RNAII555 structure, whichhas a much lower DG than wild-type ColE1.Interestingly, this mutant also had a severe growthphenotype (data not shown).

New compatibility mutants

Besides changing copy number, alterations in theorigin of replication could affect the plasmid’sincompatibility properties. Distinct incompatibilitygroups are known20,21 and regions of the ColE1origin of replication critical for compatibility havebeen identified.22,23 The ColE1-like plasmidRSF1030, for example, is able to be transformedinto and be stably maintained with both pMB1 andp15A-derived plasmids, as well as with non-ColE1vectors such as pSC101.24 However, compatibleColE1 plasmids reside together with at least one ofthe plasmids replicating at a very low copy number.Since we had a panel of mutants of the ColE1 originof replication, we tested their compatibility with awild-type ColE1 origin residing on a kanR vector.We performed an initial screen of the ten mutantsby attempting to cotransform them into an E. colihost (XL1-Blue) along with a kanR pBluescriptderivative that expressed an orange fluorescentprotein. Selection was done for ampR only andthe percentage of fluorescent colonies wereevaluated. For the wild-type pBluescript and mostof the copy number mutants, 10% or less of theampR colonies were fluorescent. However, formutants 3.3, 5.1, and 2.3, greater than 50% of theampR colonies were fluorescent. Thus, it waspossible that these mutants also had alteredincompatibility properties compared with thewild-type counterpart.

To quantify more specifically the compatibilityproperties of the 3.3 and 5.1 mutants, a kanR

(ampicillin-sensitive) variant was constructed ofthese two origin mutants and the wild-typeplasmid. Competent cells harboring the ampR

version of the wild-type or mutant origins wereprepared and transformed with the kanR version ofeach. Selection was for kanR only. We controlled forpotential differences in DNA concentration byusing a plasmid-free host (labeled None inTable 4) and controlled for the relative efficiencyof the competent cells by using a plasmid R6Kderivative that is compatible with pBluescript (pBS;see Materials and Methods). This experiment wasperformed in triplicate and the total number oftransformants (minus those kanamycin-resistanttransformants that became ampicillin-sensitive,and hence, were not co-transformants) recorded.After normalizing for the DNA concentration andhost cell transformation efficiency, the data arepresented in Table 4. In each case, DNA of the KanR

plasmid transformed E. coli containing the identicalampR version at a low level. Less than 5% of theexpected colony-forming units (cfu) were observed.In all other combinations, cotransformation efficien-cies ranged from 25–60%, indicating a quantitativereduction in the incompatibility expressed.

Discussion

Standard mutagenesis procedures generally relyon altering the nucleotide content of genes. EvenDNA shuffling, which allows a combinatorialrearrangement of members of gene families, isconstrained by the fixed lengths of the familymembers. The real “sequence space” of a givengene should also consider length alterations alongwith nucleotide content changes. Calculation of thereachable “deletional space” of a model generevealed more possible combinations of sequencesthan nucleotide changes at the same number ofpositions. Thus, a technique that could alter genelength could allow access to unexplored areas ofsequence space.25

From a practical standpoint, gene and proteinoptimization would benefit from a technique thatcould rapidly identify and remove genetic elementsthat negatively impact a desirable function.Removal of the calcium binding property ofsubtilisin was useful for engineering this proteinto function more efficiently in laundry detergent.26

Removal of protease sites could enhance proteindrug half-life.27 Additionally, regulatory regions of


genes such as operators, silencers, or siRNAencoding regions are known to affect geneexpression. Deletional mutations in such regionswould be unlikely to be found using techniques likerandom mutagenesis or DNA shuffling.

We have developed a simple procedure to createa single double-strand break in a supercoiledplasmid using a single-strand nuclease. Althoughwe did not observe any hot-spots for cleavage, wecannot conclude that the S1 method is completelyrandom. Although gain of function mutationsoccurred in predicted hairpin structures, no clearcorrelation with hairpin structures was found forthe loss of function mutations. Supercoiled plasmidis susceptible to hydroxide attack preferentially atcruciform structures,28 so it is possible that these orother structural features may be preferential sub-strates for S1. It is promising, however, that tenindependent gain of function mutations wereisolated, and that 13 loss of function mutationswere also independent. This indicates that the S1cleavage event is not strongly biased towardspecific sequences. Additionally, no proclivitytowards ACT-rich regions was found in themutants, which might be expected since ACT-richareas might preferentially form single-strandedregions in plasmids, and be preferred cleavagesites for S1.

The mean deletion length of the gain of function(LacC) mutations was 5.9, and for the loss offunction (LacK) was 33.6. Three of the LacK

mutations, however, were large (54, 134, and 152bases). Exclusion of these three large mutationsrevealed a mean of 9.7 bases deleted in theremaining LacK mutants. We speculate that largerdeletions are more likely to result in loss of b-Galactivity, which were chosen in our screen to becompletely white colonies in the presence of the lacinducer IPTG. Additionally, the LacK mutantsclustered near the 5 0 end of the lac region. Onlyseven of 13 mutations were in the coding region,and three of these also overlapped the promoter.Thus, only four of the ten shorter deletion mutantshad deletions entirely within the coding region,even though the coding region makes up approxi-mately 75% of the lac region. This is somewhatsurprising, since NuDel should produce 66% frame-shift mutations, which would inactivate lac in thecoding region. This could reveal a bias in themutagenesis procedure to the promoter, or a moredramatic phenotypic effect of deleting expressionregulatory elements. In regards to the latter, fourmutations remove the ATG start site, and sixoverlap the CRP binding site in the promoter(Figure 6). Perhaps deletions in these regionsproduce the most severe LacK phenotype. Onlyone mutation (7.7) was both in the a-peptide codingregion and in-frame. Thus, all of the deletions affectan area of lac known to be involved in b-gal activity.

NuDel allows short stretches of nucleotides to bedeleted at a non-specific position. Insertion ordeletion of a fixed number of nucleotides into arandom position induced by chemical cleavage of

single-stranded DNA was capable of altering orenhancing the spectral characteristics of greenfluorescent protein.8 Although this method couldtheoretically be used to allow up to 16 insertions ordeletions, a large number of specific oligonucleo-tides would be required in this method to coversuch a vast sequence space. The enzymatic nature ofNuDel allows it to achieve significant diversitywithout the use of multiple oligonucleotides ofdifferent lengths. Other methods can incorporateoligonucleotides in a site-specific manner to achievedeletional mutagenesis; however, they do not havethe advantage of the positional and deletional sizedistributions of NuDel.29 Because a double-stranded plasmid can be opened without positionalspecificity using single-strand nucleases like S1, weanticipate that it will be straightforward to generatefurther diversity by inserting sequences at the breakusing standard ligation reactions. Other methods,like linker-scanning mutagenesis,12 could alsolikely be modified to produce libraries similar toNuDel. In this regard, deletions and insertionscould be combined to perform sequence replace-ments, or specific sequences, such as immunogenicepitopes, could be inserted into DNA vaccines. Theutility of this approach is also apparent in that onlyone mutagenic event can occur per molecule, whichprecludes deleterious mutations from occurring incis to advantageous ones. In the case of DNAinsertions, large “jumps” in sequence space couldoccur with only one mutagenic event. This methodcould also be used iteratively, such that additionalpositions could be subject to deletion/insertion insuccessive rounds of mutation and screening. Thiswould allow a much larger area of sequence spaceto be sampled (Table 1).

We have used NuDel to identify unique plasmidorigins of replication that increased b-gal activityin vivo. The mutations cluster in the regionsencoding RNAII and RNAI of ColE1, which areknown to control plasmid copy number as well ascompatibility. Confirmation that the mutationsconferred the phenotype was accomplished by:(i) sequencing the entire lac region and origin of theplasmid and finding no other mutations;(ii) transferring the copy number and b-galphenotype to a new plasmid; and (iii) subcloningthe compatibility mutants to new plasmids andtransferring compatibility. The fact that no gain offunction mutations were found in the lac promoteror operator may indicate that deletions in theseareas may not enhance function, or may bedeleterious to other plasmid functions. Alterna-tively, mutations in the origin may allow a moredramatic b-gal enhancement resulting in the bluestcolonies. The increase in b-gal activity could be dueto: (i) an increase in plasmid copy number, and thusgene dosage; (ii) increased levels of functionalRNAII that could conceivably read-through to thea-peptide, resulting in increased transcription; or(iii) a combination of (i) and (ii).

At a molecular level, mutations in the RNAI/RNAII region could have several consequences.


First, they could allow a more stable version ofRNAII555, the active primer for DNA replication,resulting in a higher plasmid copy number. Second,the mutations could result in decreased RNAI/RNAII complexation, resulting in more activeRNAII555 and an increased copy number. Third,the mutations could decrease the half-life of RNAI,leading to an increase in copy number. In thisregard, the half-life could be affected directly bydestabilizing the RNAI structure, or indirectly byaltering RNA processing such as RNaseE cleavagesites30 or polyadenylation,31 which are known toaffect copy number. The fact that the changebetween the DG for RNAII555 and RNAII111 isidentical for six of the ten mutants argues for thefirst explanation. However, the location ofmutations in unpaired regions at the tips ofhairpins, as well as the compatibility mutants,suggest the second mechanism. Crystallographicstudies of the RNAI and RNAII interaction foundthat they occur through “kissing hairpins”, theunpaired regions at the tip. All of our mutants occurin, or overlap, unpaired areas. In fact, some mutantsmay utilize different, and even more than onemechanism, to achieve higher copy number. Kissinghairpin interactions of plasmid replication originsare strikingly similar to certain siRNA interactions,so it is not unreasonable to speculate that NuDelcould be used to identify naturally occurring smallinhibitory RNAs. For example, RNAi-based mech-anisms that control long terminal repeat (LTR)activity32 might be identified using NuDel deletionlibraries and LTR-dependent gene expressionscreens in eukaryotic cells.

Interestingly, the mutations that confer a compat-ibility phenotype occur in the tips of hairpins II andIII. Neither the four mutations in hairpin I, nor thetwo mutations in the poly(A) bubble, were com-patible with wild-type ColE1. These results suggestthat hairpins II and III are more important in theRNA interactions affecting compatibility comparedto hairpin I. Interestingly, mutations in all threehairpins appear to increase the copy numberphenotype. Thus, functional roles of the differenthairpin structures may be overlapping but in somecases separable. Alignment of several ColE2 origins,which differ in both compatibility and copy numberproperties, reveals insertion and/or deletionevents.2 However, functional attributes cannot beassigned to the length changes because there arealso several concomitant sequence content changesin these replication origins.

Until now, it has been difficult to transform andstably maintain two different plasmids containingColE1 origins. For researchers who wished to co-express multiple proteins in the same E. coli cell, theonly option was to use plasmids of differentincompatibility groups that were present at varyingcopy numbers and lacked the cloning andexpression features of the ColE1 vectors. Thus,new compatibility groups of the ColE1 plasmidfamily will have considerable interest for research-ers in any field where regulation between two genes

or proteins could be analyzed in E. coli. Such areasinclude functional proteomics and genomics.

Materials and Methods

Sequence space calculations

For replacement mutations (SR), a given gene of lengthN could be mutated to any of three nucleotides (differentfrom the original nucleotide) at a maximum number ofpositions (P), we calculated the total number of possiblevariants according to:

SR Z 3PN!

P!ðNKPÞ!

For deletional mutations (SD) a gene of starting lengthN could be mutated at a maximum number of positions(P), and each position could have up to a maximumnumber of deletions (D), we calculated the total numberof possible variants according to:

SD ZDP N!

P!ðNKPÞ!

The formula for deletional diversity is an approxi-mation for N[D!P. As D!P approaches N, increasingnumbers of potential combinations result in deletions ofthe same sequences (overlapping deletions). However, inthe case of up to 25 deletions at five positions, wecalculate the overestimation to be only 0.06%.

DNA and strains

Supercoiled plasmid DNA was purified by a Qiagenmidiprep kit according to the manufacturer’s instruc-tions. Plasmid pLacZi was from Clontech and pBluescriptII KS was from Stratagene. The f232 PCR fragmentwas amplified from pLacZi using the primers F1111(ATCGGCATAACCACCACGCTCATC) and R882(ACGTCTCGTTGCTGCATAAACC), wherein a 5 0 amineend of primer R882 was fluorescently labeled with 5-FAM(Molecular Probes) according to the manufacturer’sinstructions. Electrophoresis was on an ABI373 DNAsequencer and electropherogram analysis was performedby Genescan software v2.0. E. coli strains XL1-Blue orTop10F were used in all experiments.

S1 digestion

Supercoiled plasmid DNA at 200 ng/ml was incubatedwith 0.1 to 1.0 unit/ml of S1 nuclease (Promega) in 100 mlof the manufacturer’s buffer for ten minutes at roomtemperature. The reaction was stopped with 5 ml of 0.5 MEDTA, twice extracted with phenol/chloroform/isoamylalcohol (25:24:1, by vol.), and precipitated with sodiumacetate and ethanol. Plasmid DNA was purified from1.5% (w/v) agarose gels using Qiaex beads (Qiagen)according to the manufacturer’s instructions.

Exo III digestion

In a 10 ml reaction, ten units of Exo III (New EnglandBiolabs) were incubated with 100 ng of linear plasmid orPCR product at 158C for five minutes in a buffercomprising 66 mM Tris (pH 8.0), 6.6 mM MgCl2 and theindicated amount of NaCl. The reaction was stoppedwith100 ml of S1 buffer containing five units of S1 nuclease. This

† http://mfold.burnet.edu.au/


reaction, which eliminated exonuclease III-generatedoverhangs, was for 15 minutes at room temperature.The reaction was stopped with 5 ml of 0.5 M EDTA, andincubated with 1 mg of protease K at 70 8C for fiveminutes, then extracted with phenol/chloroform/isoamyl alcohol (25:24:1), and precipitated with sodiumacetate and ethanol.

Library preparation and screening

Two independent libraries were produced using pBlue-script II KS as a template for S1 cleavage as describedabove. Following S1 cleavage, plasmid was depho-sphorylated using calf intestinal phosphatase (NewEngland Biolabs) according to the manufacturer’sinstructions. Gel-pure linear dephosphorylated DNAwas incubated with Exo III as described above in twotubes with either 100 mM or 150 mM NaCl. DNA wasligated with T4 DNA ligase (New England Biolabs), andelectroporated into E. coli. Tubes containing the differentsalt concentrations were combined in a single tube“library”. Two independently constructed libraries wereproduced with this approach. Libraries were screened onLB agar plates containing 40 mg/ml of X-Gal at 30 8C.Under these conditions the lac repressor is produced,resulting in a white colony phenotype. Plasmids weresequenced through the entire lac region using T3 and T7primers, and in the origin using primersGGTGCCTCACTGATTAAGCATTG and CGGTAACTATCGTCTTGAGTCCAAC. Multiple sequence analysiswas performed using ClustalW.

Total cell nucleic acid preparation

Total cell nucleic acid was prepared by a modificationof the method described by Lin-Chao & Bremer.33 Cellswere grown overnight at 30 8C and four A600 units(approximately 1 ml) were pelleted and resuspended in100 ml of 10 mM Tris, 1 mM EDTA, 150 mM NaCl. Anequal volume of phenol/chloroform/isoamyl acohol(25:24:1) was added and vigorously vortex mixed. Theaqueous phase was saved on ice and 10 ml was analyzedby electrophoresis on a 1.2% agarose gel.

b-Galactosidase activity

Relative b-gal activities of E. coli extracts were deter-mined using a kit according to the manufacturer’sinstructions (Stratagene).

Compatibility testing

Plasmid incompatibility was evaluated as follows.Kanamycin-resistant versions of pBluescript (wild-type)and mutants 3.3 and 5.1 were constructed by inserting aPCR product (Pfu, Stratagene) of the kanamycin gene intoXmnI and ScaI sites of the plasmids, replacing part of theopen reading frame of b-lactamase. This renders themampicillin-sensitive. Competent cells of XL1-Blue(Stratagene) and hosts harboring the ampR version ofpBluescript, 3.3 and 5.1 were prepared. Each host strainwas transformed with one of the kanR plasmids describedabove or an R6K control plasmid (KanR, compatible withColE1) as a reference for plasmid transformationefficiency. The number of kanR transformants into theplasmid-free XL1-Blue served as a reference for DNAconcentration. The number of kanR colonies wasrecorded. Of these, approximately 40 kanR transformants

for each group were then patched onto ampicillin platesto determine whether the resident plasmid was retained.If no growth was observed on the ampicillin plate, arelative reduction in the overall compatible percentagewas made.

RNA folding analysis

Predictions of RNA secondary structures for wild-typeand mutant RNA II of varying lengths were performedusing the MFOLD server at the MacFarlane Burnet Center(MFOLD v3.0)†.19 All calculations were performed usingthe default settings except for the folding temperature,which was set at 30 8C.

Acknowledgements

We thank Stanley N. Cohen for helpful commentson the manuscript, and Bill Heriot for technicalsupport.

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Edited by J. Karn

(Received 2 May 2005; received in revised form 6 June 2005; accepted 9 June 2005)Available online 5 July 2005

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