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Will this strategy work for verifying successful cloning of a DNA fragment into a plasmid vector?

Will this strategy work for verifying successful cloning of a DNA fragment into a plasmid vector?


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So I am in a bit of a time constraint. Essentially, I inserted a DNA fragment via molecular cloning which contains a unique RE site. I need to confirm whether my colony has or does not have the fragment. I need to do this in one day so would this work as a confirmation strategy:

  1. Pick colony and perform colony PCR with primers around RE site.
  2. Cleanup the PCR product and digest using the enzyme which was inserted.
  3. Run on a gel. If a cut is noticed, this confirms that the colony had the inserted fragment (containing the RE site).

Would this strategy work? Are there any notable issues people can for see in this strategy?


Yes, your strategy would work, with the caveats already mentioned by @Cell.

Depending on which restriction enzyme and PCR buffer you use, you might even be able to skip step 2. NEB has a good list over their restriction enzymes and their activities in some common PCR buffers. With some reasoning that information is often applicable also to restriction enzymes of other brands.

Depending on your primer availability, another quick strategy is to simply use a primer pair such that one primer binds inside your fragment, and the other binds to the backbone. This should result in a PCR product only if your cloning was successful, and also verifies correct direction (in case you used blunt-end or single-site cloning). Two possible problems with this approach is:

  1. If you get no bands at all you may not know if what failed was your cloning or the PCR reaction. The latter can be easily controlled for with a well-thought positive PCR control.
  2. You may detect cases in which more than one copy of your fragment have been inserted. Although such cases are rare, it sometimes happens that inverted repeats of 3, 5, 7,… etc copies get inserted in tandem.

One issue is if your fragment amplified is too small you may not be able to see the positive result of two bands because they run off the gel. Also the RE may not cut a small fragment efficiently.

I would recommend extracting and purifying plasmid, then do a double digest using the unique fragment cutting RE and a RE that only cuts the backbone. You should either see linearized plasmid DNA or two bands that are probably both >1 kb and easily resolvable.


A restriction-free method for gene reconstitution using two single-primer PCRs in parallel to generate compatible cohesive ends

Restriction-free (RF) cloning, a PCR-based method for the creation of custom DNA plasmids, allows for the insertion of any sequence into any plasmid vector at any desired position, independent of restriction sites and/or ligation. Here, we describe a simple and fast method for performing gene reconstitution by modified RF cloning.

Results

Double-stranded inserts and acceptors were first amplified by regular PCR. The amplified fragments were then used as the templates in two separate linear amplification reactions containing either forward or reverse primer to generate two single-strand reverse-complement counterparts, which could anneal to each other. The annealed inserts and acceptors with 5’ and 3’ cohesive ends were sealed by ligation reaction. Using this method, we made 46 constructs containing insertions of up to 20 kb. The average cloning efficiency was higher than 85%, as confirmed by colony PCR and sequencing of the inserts.

Conclusions

Our method provides an alternative cloning method capable of inserting any DNA fragment of up to at least 20 kb into a plasmid, with high efficiency. This new method does not require restriction sites or alterations of the plasmid or the gene of interest, or additional treatments. The simplicity of both primer design and the procedure itself makes the method suitable for high-throughput cloning and structural genomics.


Introduction

Recombinant DNA techniques are indispensable in any modern laboratory that relies on molecular biology methods. Traditionally, molecular cloning depends on restriction endonucleases to produce linear vectors and inserts that are fused by the DNA ligase [1]. Recombinant DNA technology has also been greatly facilitated by the use of Polymerase Chain Reaction (PCR). A standard DNA cloning practice is to amplify a DNA of interest with oligonucleotides containing 5′ tails specifying endonuclease recognition sites. These allow subsequent cleavage and insertion of PCR fragments into any desired cloning vector. Notwithstanding the universality of these traditional cloning approaches, the preparation of reactant molecules with restriction enzymes and their combination by DNA ligase involves many steps, being not only laborious, but often rendering meager or frustrating results. To overcome such limitations, several PCR-based cloning protocols have proposed to skip the use of DNA ligase and also dispense the need for restriction endonucleases [1]. Among them, homology-based methods take advantage of PCR products flanked on both sides by 15 to 60 bp long sequences that perfectly match the ends of a linear vector [1]. This is facilitated by PCR amplification of fragments with oligonucleotides containing 5′ appendages homologous to the cloning plasmid. After fragment preparation, the cloning reaction can be driven in vitro by enzymatically-assisted recombination between the vector and the PCR inserts (e.g., SLiCE [2] and Gateway [3] methods). Alternatively, the PCR fragments and linear plasmid can be enzymatically treated to generate single-stranded DNA (ssDNA) on their termini (e.g., SLIC [4] and USER [5] methods). The resulting vector and insert overhangs are complementary and hybridize in a double-stranded DNA (dsDNA) formation containing a nick. During transformation in E. coli, the nick is repaired in vivo, sealing a perfect circular recombinant plasmid. Other homology-based cloning procedures rely on a typical PCR setting in which the primers have been omitted (e.g., OEC [6] and CPEC [7] methods). Following the heat denaturation step, overlapping free termini of vector and PCR fragments can anneal by base pair complementarity. The hybrid regions then serve as “megaprimers” in the extension step catalyzed by the DNA polymerase. As a result, a nicked recombinant plasmid is generated that is repaired in vivo in E. coli. Besides those recent innovations, traditional gap-repair in yeast stands as one of the most effective homology-based cloning approaches [8,9]. This procedure consists of merely co-transforming yeast cells with a mixture of the linear vector and PCR fragments, both having common flanking sequences. Following transformation, the yeast DNA repair machinery is very effective to recombine vector and insert homologous termini, generating a closed plasmid.

Interestingly, an analogous in vivo gap-repair cloning method in E. coli was described by two groups as early as 1993 [10,11]. The principle of this cloning approach in E. coli is equally as simple as in yeast. By co-transforming a linear vector and PCR fragments containing homologous ends, several E. coli laboratorial strains are capable of recombining the reactants in vivo, generating a closed propagative plasmid. Although this technique is simpler than traditional ligase-dependent cloning, only a few published reports using gap-repair cloning in E. coli exist [12–17]. Some studies elaborate this cloning principle in hyper-recombinogenic E. coli strains in which the RecE and RecT homologous recombination pathway is over-expressed (the ET cloning method) [12]. A few other published reports propose specific applications for gap-repair cloning in E. coli. For example, homologous recombination was successfully used for high-throughput cloning of 1302 ORFs amplified from the Campylobacter jejuni genome [13]. The strategy was to supply PCR fragments with 21 bp flanks identical to the ends of a cloning vector prepared with restriction enzymes. Similarly, Klock et al. described the gap-repair cloning of 448 full-length and 2143 truncated ORFs from 30 different bacterial sources [14]. However, in this case, both the vector and the ORFs were amplified by PCR. The authors proposed that a different mechanism than in vivo homologous recombination explained the obtained results. It was suggested that in normal PCR reactions, a large fraction of amplified fragments have ssDNA stretches on both termini due to incomplete primer extension during the PCR elongation step (therefore the method was termed PIPE, Polymerase Incomplete Primer Extension) [14]. Upon mixture, vector and insert complementary ssDNA regions anneal and a single recombinant DNA molecule is sealed in vivo following transformation in E. coli. The efficacy of gap-repair cloning through the PIPE protocol has been recently confirmed in a study in which this procedure was compared with the SLIC and OEC techniques [15]. The FastCloning method is another recombination-mediated approach that is very similar to the PIPE protocol [16]. It describes a quick and efficient way of cloning PCR fragments containing 16 bp flanking regions overlapping the ends of a linear cloning vector. Both fragments and vectors are first obtained by PCR. A mixture of vector and insert is subsequently treated with DpnI to eliminate the template plasmids background and then readily transformed in E. coli to obtain the desired clones. In line with the PIPE description, the authors of the FastCloning protocol speculate that the cloning mechanism relies on vector and inserts having complementary ssDNA overhangs that are fortuitously generated by the DNA polymerase 3′ to 5′ exonuclease activity [16].

The examples described above illustrate that recombinational cloning in E. coli is a highly efficient technique that presents remarkable advantages over conventional cloning with restriction endonucleases and DNA ligase. However, it is puzzling that this molecular cloning principle has so far been ignored by a wider community of molecular biologists. Not only is the amount of publications reporting the use of gap-repair in E. coli very little, but the interpretation of the cloning mechanism itself has been contradictory between them [10,11,14,16]. In an attempt to unify those different reports and to popularize recombinational cloning in E. coli as a practical method, we present here a study of the different parameters that modulate the efficiency of this technique. We also provide evidence dismissing the assumption that ssDNA complementary tracts, supposedly generated on the vector and inserts by DNA polymerase incomplete primer extension [14] or exonuclease activity [16], are critically needed for gap-repair cloning to occur in E. coli. Finally, to support the notion that recombination cloning in E. coli might be the method of choice for assembling single and double PCR fragments into a plasmid, we optimized the cloning parameters to obtain 94% efficiency during the construction of five different recombinant DNAs.


What Exactly Are Restriction Enzymes?

Restriction enzymes are also known as restriction endonucleases. The first of these enzymes was found by observing that a phage was able to be grown in one strain of bacteria, but not another (1). After investigation, it was discovered that the reason the phage growth was restricted (hence the name) was certain bacteria lines had the ability to chop up the phage DNA, preventing infection. Upon further study, an enzyme was found that was ultimately responsible for cutting the phage DNA not present in phage-susceptible bacteria lines.

Today, researchers have found thousands of different enzymes that can be used for cutting DNA, hundreds of which can be purchased from suppliers with easy-to-use buffer systems. Additionally, there are plenty of tools online, such as NEBcutter that can be used to find restriction enzyme cut sites in a given DNA sequence.


DISCUSSION

With the rapid development of the next-generation sequencing technology, more plant genomes will be sequenced in the near future. How to rapidly determine the function of the identified genes on a large scale is a daunting challenge. The ability to efficiently make constructs to transiently and stably express specific genes in cells, tissues, or whole plants is a fundamental aspect and bottle neck of plant functional genomics research. Traditionally, the cloning vectors for plant research carry a multicloning site (MCS) within their target gene expression cassettes. The restriction sites in the MCS are rather limited, making cloning of most target genes difficult. Although TA cloning vectors have been widely used for cloning of PCR-amplified fragments, the system has not yet been incorporated in the cloning vectors for transient and stable expression of target genes because of the technical challenge of generating low-background TA cloning vectors. The Gateway system has been a popular choice for generating various constructs because it allows the gene of interest to be easily cloned into specifically designed plasmids without DNA restriction digestions. The two-step cloning and expensive reagents, however, make the Gateway system impractical for large-scale cloning in most individual laboratories when the entry clone collections are not available. The ZeBaTA system described here overcomes the limitations of both the TA and Gateway cloning systems. After two XcmI recognition sites have been introduced into the MCS, any PCR fragments with a T-overhang can be easily cloned into a ZeBaTA vector. With the introduction of the negative selection marker gene ccdB between the two XcmI sites, any self-ligation transformants are eliminated. Using this technology, we constructed a set of 12 transient and 12 stable transformation vectors for plant gene expression studies and tested the vectors in rice or Arabidopsis in our laboratories. These vectors can be used in a wide range of functional genomics projects in plants and will be distributed to the research community upon request.

Under certain conditions, cloning with T-vectors generated by digestion with AhdI or XcmI gave low efficiency and the T residue of the insert-vector junction in the recombinant clones is often missing ( Mead et al., 1991 Chen et al., 2006a). Chen et al. (2006a) speculated that this may be due to the presence of unknown factors that, during digestion and preparation of the T-vectors, influence the stability of 3′-T overhangs. In this study, we found that the main factor affecting successful cloning is the use of an appropriate T4 DNA ligase. We tested the Promega T4 DNA ligase, which is included in the pGEM-T easy vector system, and the T4 DNA ligase from USB Corporation. The ligations using Promega T4 DNA ligase consistently gave a very high cloning efficiency most of the ligations using USB Corporation T4 DNA ligases yielded low efficiency. Many of the recombinant plasmids from the latter ligations missed a T residue in the insert-vector junction, consistent with observations by Mead et al. (1991) and Chen et al. (2006a). The T residue is missing mainly because regular commercial T4 DNA ligases contain exonuclease activities that can remove the 3′-T tails from the vector, as reported in the technical manual of the pGEM-T and pGEM-T easy vector systems (http://www.promega.com/tbs/tm042/tm042.pdf) removal of the 3′-T tails from the vector results in very low cloning efficiency. When the Promega T4 DNA ligase was used for ligation, we never observed the absence of the T residue in the recombinant plasmids (data not shown), suggesting that the 3′-T overhangs of the T-vectors generated by XcmI digestion were stable. Our cloning tests also indicated that the size of T-vectors, vector overdigestion, and the insert-to-vector molar ratios had a minimal impact on cloning efficiency.

In recent years, Gateway cloning technology has become increasingly popular for functional genomics in plants ( Karimi et al., 2002, 2005, 2007a, 2007b Curtis and Grossniklaus, 2003 Earley et al., 2006 Hilson, 2006 Himmelbach et al., 2007 Lei et al., 2007). In comparison to the Gateway system, ZeBaTA offers several significant advantages. First, it allows simple and highly efficient one-step cloning for PCR-amplified genes/fragments of interest into a specialized ZeBaTA expression vector. Second, the ZeBaTA constructs have a short vector-insert junction with only 8 bp (CCANNNNN), whereas the constructs generated by Gateway recombination contain a longer att linker sequence with 25 bp or even up to 125 bp ( Lei et al., 2007). For example, protein-tagging constructs generated by Gateway cloning technology usually result in a long linker with at least eight or sometimes up to 20 amino acids ( Dubin et al., 2008) between the tag and the protein that could interfere with protein function or expression. In contrast, ZeBaTA-based constructs for protein tagging result in a short linker with only three amino acids ( Supplemental Fig. S1 ), thereby reducing the potential for interference with the tag-protein fusion. Last, the XcmI recognition sequences CCANNNNN/NNNNTGG, where N can be any pyrimidine or purine, is more flexible for designing special vectors for high-throughput functional analysis.

In this article, we demonstrate the flexibility of this system by developing a ZeBaTA-amiRNA vector for rice functional genomics ( Fig. 5C Supplemental Fig. S3 ). With the introduction of a single base mutation of GCA→CCA and TGC→TGG on the 5′ and 3′ stemloop backbones of an endogenous rice miRNA precursor Osa-MIR528 ( Warthmann et al., 2008), respectively, the modified vector allows for generating an expression T-vector with preassembled 5′ and 3′ Osa-MIR528 stemloop backbone sequences. Thus, amiRNA expression constructs can be simply generated by cloning an amiRNA-amiRNA* fragment generated from a single-step PCR. Prediction of the original and modified Osa-MIR528 stemloop gave exactly the same secondary structures ( Supplemental Fig. S4 ), suggesting that the modification of the stemloop backbone did not affect the structural features of the miRNA precursor. Further stable transformation using vectors based on the original or modified stemloop backbones gave a similar high efficiency for targeted OsPDS silencing, indicating that without changing structural features, the modified precursor stemloop backbone can functionally express a miRNA to trigger targeted gene silencing. Although we did not apply the approach to other miRNA precursors, our results suggest that this approach should be amenable to different amiRNA systems in other plants.


PCR Cloning Method

PCR cloning differs from traditional cloning in that the DNA fragment of interest, and even the vector, can be amplified by the Polymerase Chain Reaction (PCR) and ligated together, without the use of restriction enzymes. PCR cloning is a rapid method for cloning genes, and is often used for projects that require higher throughput than traditional cloning methods can accommodate. It allows for the cloning of DNA fragments that are not available in large amounts.

Typically, a PCR reaction is performed to amplify the sequence of interest, and then it is joined to the vector via a blunt or single-base overhang ligation prior to transformation. Early PCR cloning often used Taq DNA Polymerase to amplify the gene. This results in a PCR product with a single template-independent base addition of an adenine (A) residue to the 3' end of the PCR product, through the normal action of the polymerase. These "A-tailed" products are then ligated to a complementary T-tailed vector using T4 DNA ligase, followed by transformation.

High-fidelity DNA polymerases are also now routinely used to amplify sequences with the PCR product containing no 3' extensions. The blunt-end fragments are joined to a plasmid vector through a typical ligation reaction or by the action of an "activated" vector that contains a covalently attached enzyme, typically Topoisomerse I, which facilitates the vector:insert joining. Some PCR cloning systems contain engineered "suicide" vectors that include a toxic gene into which the PCR product must be successfully ligated to allow propagation of the strain that takes up the recombinant molecule during transformation.

A typical drawback common to many PCR cloning methods is a dedicated vector that must be used. These vectors are typically sold by suppliers, like NEB, in a ready-to-use linearized format and can add significant expense to the total cost of cloning. Also, the use of specific vectors restricts the researcher's choice of antibiotic resistance, promoter identity, fusion partners, and other regulatory elements.

  • High efficiency, with dedicated vectors
  • Amenable to high throughput
  • Limited vector choices
  • Higher cost
  • Lack of sequence control at junction
  • Multi-fragment cloning is not straight forward
  • Directional cloning is difficult

PCR Cloning

Note that times are based on estimates for moving a gene from one plasmid to another. If the source for gene transfer is gDNA, add 2 hours to calculation for the traditional cloning method. Total time does not include transformation, isolation or analysis.

Abstract

Long fragment cloning is a challenge for its difficulty in accurate amplifying and tendency to get unwanted mutation. Here we discuss Restriction-based Multiple-fragment Assembly Strategy's advantages and limitations. In this strategy, rather than PCR amplifying the entire coding sequence (CDS) at one time, we amplified and sequenced smaller fragments which are shorter than 1.5kb spanning the CDS. After that, the sequence-proved fragments were assembled by digestion-ligation cloning to the target vector. We test its universality in our script programmed in Python. Our data shows that, among the entire human and mouse CDS, at least 70% of long CDS cloning will benefit from this strategy.

Keywords: PCR, ligation, cloning strategy, CDS, endonuclease.


Vector preparation

    using (preferably) two restriction enzymes.
  • Dephosphorylation of the ends using calf intestine or shrimp alkaline phosphatase. This reduces the background of non-recombinants due to self-ligation of the vector (especially when a single site was used for cloning).
  • Purification of the digested vector by agarose electrophoresis to remove residual nicked and supercoiled vector DNA and the small piece of DNA that was cut out by the digestions. This usually reduces strongly the background of non-recombinants due to the very efficient transformation of undigested vector.

Methods

Strains, plasmids and reagents

Strains and plasmids used in this study are listed in Table S1. Recombineering-proficient E. coli strain GB05-dir which is chromosomally integrated recE and recT genes was used for plasmids construction 41 . E. coli GB2005 was used as cloning host. All E. coli strains were cultured in Luria-Bertani (LB) broth at 37 °C. P. luminescens subsp. laumondii TT01 and A. tumefaciens C58 (hereafter C58) were grown with aeration in LB broth at 30 °C.

The plasmid pGB-hyg-Ptet-gbaA which consists of tetracycline inducible promoter (Ptet), broad-host-range RK2 replicon (oriV and trfA gene), ColE1 origin, hygromycin and ampicillin resistance genes was used to construct Cre expression plasmid by replacing redα-redβ-redγ-recA operon with cre gene. The resultant plasmid was designated as pGB-hyg-Ptet-cre.

The widely used BAC vector, pBeloBAC11 was used to carry and stably maintain large DNA fragments containing the gene clusters of interest. The chloramphenicol resistance gene of pBeloBAC11 was replaced by kanamycin resistance gene. One loxP site was incorporated into the forward oligo for amplifying ampicillin resistance gene and introduced into pBeloBAC11 simultaneously. There exist EcoRІ, SacІ, BamHІ and HindIII unique restriction sites between kanamycin resistance gene and loxP site, which can be used to linearize the modified pBeloBAC11. DNA fragment containing homology arm (HA1) corresponding to the 5′ end of the gene clusters of interest was recombined into the modified pBeloBAC11 to generate pBeloBAC-HA1.

Another loxP site was incorporated into the reverse oligo for amplifying apramycin resistance gene and introduced into pUC19 plasmid simultaneously to generate pUC-Apr. Two DNA fragments containing homology arms (HA2 and HA3) corresponding to the 3′ end of the gene clusters of interest were recombined to pUC-Apr sequentially to obtain pUC-Apr-HA2-HA3. The HA2-Apr-loxP-HA3 fragment could be released from pUC-Apr-HA2-HA3 by restriction digestion.

Restriction enzymes, Phusion polymerase and DNA marker were purchased from New England Biolabs. Anhydrotetracycline and antibiotics were purchased from Sigma-Aldrich. Appropriate antibiotics were added at the following concentrations: for E. coli strains, ampicillin at 50 μg ml −1 (liquid) or 100 μg ml −1 (solid), hygromycin at 25 μg ml −1 (liquid) or 30 μg ml −1 (solid), apramycin at 10 μg ml −1 (liquid) or 20 μg ml −1 (solid), kanamycin at 15 μg ml −1 (both liquid and solid) for TT01, ampicillin at 100 μg ml −1 (both liquid and solid), kanamycin at 15 μg ml −1 (both liquid and solid), apramycin at 25 μg ml −1 (both liquid and solid) for C58, hygromycin at 50 μg ml −1 (both liquid and solid), apramycin at 50 μg ml −1 (both liquid and solid), kanamycin at 50 μg ml −1 (both liquid and solid).

Electroporation procedures

For E. coli, electroporation was performed as previously described 41 . For TT01 and C58, 60 μl of overnight culture was transferred into 1.4 ml fresh LB medium supplemented with appropriate antibiotics if necessary and shaken at 30 °C for 3 h (for TT01) or 4 h (for C58) until the OD600nm was around 0.6

0.8. Cells were harvested by centrifugation at 4 °C, 10000 rpm for 30 s. The supernatant was discarded and the cell pellet was resuspended in 1 ml ice-cold GH buffer (10% glycerol, 2-mM HEPES, pH 6.5) and centrifuged. The ice-cold GH buffer washing procedure repeated once. Cells were finally resuspended in 30 μl ice-cold GH buffer and DNA was added. The mixture of electro-competent cells and DNA was transferred into an ice-cold 1 mm cuvette and shocked once using Eppendorf electroporator 2510 (Eppendorf, Germany) set at 1350 v. After electroporation, 1 ml LB medium was added into the cuvette and the cells were recovered at 30 °C for 2 hours with shaking and then plated on LB plates appropriate antibiotics. This was followed by incubation at 30 °C for 24 h (for TT01) or 48 h (for C58). Colony PCR was used to check recombinants and purified PCR products were subjected to sequencing to ensure the validity of every integration event.

Recombineering in E. coli GB05-dir

Recombineering was performed according to the method established by our group 41 . Briefly, 40 μl of overnight culture of GB05-dir was transferred into 1.4 ml fresh LB medium and grown at 30 °C for 2 h. Then, 20 μl of 10% L-arabinose was added into the culture to induce the expression of Red recombinases and cells were shifted to 37 °C and shaken for another 40 min. Cells were harvested by centrifugation, washed twice with ice-cold sterile water and finally resuspended in 50 μl ice-cold sterile water. Recombineering proficient competent cells mixed with DNA were transferred to an ice-cold 1 mm cuvette and shocked once using Eppendorf electroporator 2510 (Eppendorf, Germany) set at 1250 v. After electroporation, 1 ml LB medium was added into the cuvette and cells were recovered at 37 °C for 1 h with shaking and plated on LB plates supplemented with appropriate antibiotics, followed by an incubation at 37 °C for 24 h.

Induction of Cre recombinase expression, DNA isolation and electroporation

TT01 or C58 recombinant strains which contain two loxP sites flanking gene cluster of interest and harbor Cre expression plasmid pGB-hyg-Ptet-cre were inoculated into 2 ml LB medium supplemented with appropriate antibiotics and shaken overnight at 30 °C. Different amounts of anhydrotetracycline solution (final concentrations of 1, 2, 4, 6, 8 μg ml −1 respectively) were added into the saturated overnight cultures and grow cells at 30 °C for another 1 h, 2 h, 4 h or 6 h to induce the expression of Cre recombinase to mediate excision between two parallel loxP sites.

After induction, an aliquot (50 μl) was spread onto LB agar plate and incubated at 30 °C. Single colonies appeared on LB plate were picked and screened for ampicillin and apramycin resistance. The excision efficiency was calculated by dividing the number of total colonies by the number of ampicillin and apramycin sensitive colonies. The rest culture was subjected to DNA isolation by using the alkaline lysis method. Isolated DNAs were immediately electroporated into GB2005. Transformants appeared on LB plates supplemented with ampicillin and apramycin were counted and subjected to DNA isolation and restriction analysis.

Quantitative growth curve

The growth curve of E. coli GB2005 recombinant strain GB2005(pBeloBAC-AgS) containing siderophore gene cluster from C58 was measured and compared to that of GB2005 under both normal and Fe-depleted conditions. An overnight culture of GB2005 or GB2005(pBeloBAC-AgS) was diluted 1:50 into fresh LB medium supplemented with or without 0.15 mM 2, 2′-dipyridyl (DIP) and shaken at 37 °C for 24 h. A 1 ml of aliquot was taken every 2 h and OD600nm was measured.

Plasmid stability test

GB2005(pBeloBAC-pluT3SS) or GB2005(pBeloBAC-AgS) was grown overnight at 37 °C in LB supplemented with ampicillin plus apramycin. The overnight culture was 100 times diluted with fresh LB and incubated at 37 °C with shaking and then subcultured 20 times (approximately 240 generations) at the same conditions. At subculturings, diluted cells were plated on LB agar plates without antibiotic and single colonies were picked and subjected to restriction analysis.


Results and discussion

Construction of a broad-host-range metagenome vector system

The broad-host-range fosmid and BAC vector (pRS44, Fig. 1) was constructed using the commercially available pCC1FOS vector as a starting point. pCC1FOS has two origins of replication, ori2 from the F plasmid and oriV from RK2. ori2 functions in E. coli and is active during construction of libraries in this host, while it is not active in most other hosts. In pCC1FOS oriV is included to be able to produce large quantities of vector DNA in a particular E. coli host. This host carries a chromosomally integrated mutant version (copy-up) of the gene encoding the replication initiation and copy number control protein TrfA that can be activated by expression from the inducible pBAD promoter. Instead here we use oriV as a tool for replication of metagenome libraries in non-E. coli hosts. In addition we have inserted the origin of conjugative transfer (oriT) to allow efficient transfer of the libraries from E. coli to such alternative hosts. This is a very important feature because it is well known that the efficiency of conjugational transfer is usually much better than transformation of naked DNA. We predicted that vector stability could become an important feature and we therefore also inserted the stabilization element parDE from RK2, because it is known that parDE stabilizes RK2 vectors across species barriers ( Sia et al., 1995 Blatny et al., 1997a). Finally, the kanamycin resistance gene was inserted to provide an alternative selection marker, potentially useful in some hosts. The BamHI and HindIII (pTA44, see legend to Fig. 1) sites are useful for sticky-end BAC cloning, while the Eco72I site is used for blunt-end fosmid cloning. The lac system for blue–white screening was kept from pCC1FOS.

Plasmid vectors for use in genomic library constructions (pRS44/pTA44) and for support of vector replication in hosts other than Escherichia coli (pRS48). In pTA44 the HindIII site in the Km r gene is removed, making it possible for cloning of HindIII-digested DNA into the remaining HindIII site. pRS44/pTA44 can replicate as a single copy replicon via ori2 and repE, while oriV contributes to a medium copy number if its replication initiation protein TrfA is expressed in the same cell. pRS44/pTA44 DNA can easily be prepared in large quantities in the E. coli strain EPI300 by expressing a mutant trfA gene from an arabinose-induced promoter, as described by Wild et al. (2002). cosN is the site used for packaging of the environmental DNA library in bacteriophage λ particles, BamHI and Eco72I sites are used for BAC and fosmid cloning respectively, and NotI is suitable for size determination of the inserts. The trfA-gene is inserted into the chromosome of hosts of interest by the transposon present in the narrow-host-range plasmid pRS48. The inside and the outside ends of the transposon (designated TnRS48) are marked I and O, respectively. tpn, gene encoding the transposase, which is not a part of the transposon XylS, gene encoding activator of PmG5 transcription in the presence of benzoic acid-type inducers, like m-toluate OriT, origin of conjugative transfer. For further details see Table 1 and the text.

Plasmid vectors for use in genomic library constructions (pRS44/pTA44) and for support of vector replication in hosts other than Escherichia coli (pRS48). In pTA44 the HindIII site in the Km r gene is removed, making it possible for cloning of HindIII-digested DNA into the remaining HindIII site. pRS44/pTA44 can replicate as a single copy replicon via ori2 and repE, while oriV contributes to a medium copy number if its replication initiation protein TrfA is expressed in the same cell. pRS44/pTA44 DNA can easily be prepared in large quantities in the E. coli strain EPI300 by expressing a mutant trfA gene from an arabinose-induced promoter, as described by Wild et al. (2002). cosN is the site used for packaging of the environmental DNA library in bacteriophage λ particles, BamHI and Eco72I sites are used for BAC and fosmid cloning respectively, and NotI is suitable for size determination of the inserts. The trfA-gene is inserted into the chromosome of hosts of interest by the transposon present in the narrow-host-range plasmid pRS48. The inside and the outside ends of the transposon (designated TnRS48) are marked I and O, respectively. tpn, gene encoding the transposase, which is not a part of the transposon XylS, gene encoding activator of PmG5 transcription in the presence of benzoic acid-type inducers, like m-toluate OriT, origin of conjugative transfer. For further details see Table 1 and the text.

To allow replication of pRS44 in new hosts, expression of trfA (we used wild type) is needed. However, we decided not to include trfA as part of pRS44, as such a vector would contain two active replication systems in E. coli, possibly leading to plasmid instability. Instead we constructed a suicide vector (pRS48, Fig. 1), which can be used to insert a derivative of transposon Tn5 expressing the TrfA protein from the inducible PmG5 promoter. This promoter is a mutant derivative of wild-type Pm, which is known to be active in many hosts ( Mermod et al., 1986 Ramos et al., 1988 Keil & Keil, 1992). The inducibility in addition allows for modification of the amount of TrfA produced. pRS48 replicates in the E. coli strain S17.1(λpir), which expresses the Pir protein needed for replication initiation of the plasmid R6K origin, oriR6K ( De Lorenzo et al., 1993).

Plasmid stability in the absence of selection

Plasmid stability may potentially become critical for the functioning of the metagenome cloning vector described here, and to quantify this we measured the rate by which it became lost in the absence of antibiotic selection in E. coli EPI300 ( Fig. 2). As controls in this experiment we used the native RK2 plasmid and pCC1FOS. By the repeated transferring, growth was monitored over about 230 generations, a number which enormously exceeds the number of generations taking place in laboratory-scale batch cultures. The experiments showed that plasmid loss could easily be detected for pCC1FOS, while both pRS44 and a derivative of it containing a 36-kb control DNA insert (pRS49) were remarkably stable, like whole RK2. This experiment therefore clearly confirmed the relevance of introducing parDE into the vector.

Plasmid stability of pRS44 and pRS49 in the absence of antibiotic selection. Exponentially growing cells in shake flasks (in the presence of selection) were diluted 10 5 times in medium lacking antibiotics. The cultures were then grown overnight and the dilution procedures were repeated until about 230 generations had elapsed. After each growth step cells were plated on LB-agar lacking antibiotics. From each step, 184 colonies were picked and duplicated into 96-well plates containing media with and without chloramphenicol. ▪, pRS44 □, pRS49 Δ, RK2 and ○, pCC1FOS.

Plasmid stability of pRS44 and pRS49 in the absence of antibiotic selection. Exponentially growing cells in shake flasks (in the presence of selection) were diluted 10 5 times in medium lacking antibiotics. The cultures were then grown overnight and the dilution procedures were repeated until about 230 generations had elapsed. After each growth step cells were plated on LB-agar lacking antibiotics. From each step, 184 colonies were picked and duplicated into 96-well plates containing media with and without chloramphenicol. ▪, pRS44 □, pRS49 Δ, RK2 and ○, pCC1FOS.

Construction of a metagenomic library in pRS44

Initial experiments showed that the standard 36 kb insert used as a control for the commercially available vectors was packaged and established in E. coli at similar frequencies for pRS44 as for pCC1FOS. However, we wanted to test the vectors with DNA from the environment, because it is well known that it may be difficult or inefficient to clone DNA from such samples ( Robe et al., 2003), and because we wanted to test the behaviour of the environmental DNA inserts in new hosts.

A fosmid library consisting of about 20 000 clones was constructed using DNA isolated from marine sediments as cloning material. End sequencing and restriction digest analysis of a selection of the clones confirmed that the library consists of plasmids with different environmental inserts of the expected size of around 35 kb. To explore the microbial diversity represented by this library, regions of 16S rRNA genes were amplified and analysed by DGGE. More than 10 different bands could be clearly distinguished, indicating that the DNA in the library originates from many different genotypes, as expected. In addition, sequencing of 24 amplified 16S rRNA gene fragments resulted in identification of 10 different genotypes, none of which were identical to existing sequences in the databases (data not shown). It could therefore be concluded that the library contains environmental DNA of diverse origins, and that pRS44 has the features required for metagenomic studies.

Verification of heterologous expression of environmental proteins in P. fluorescens by 2-DE and MS

In order to study if cloned genes in the metagenomic library are expressed in a non-E. coli host, crude protein extracts from five P. fluorescens library clones and cells harbouring the vector only were subjected to 2-DE in duplicate. Protein patterns of all six crude protein extracts were analysed with the delta- 2 d software in order to identify protein spots present in both duplicates of only one environmental DNA-containing strain but not in any other clone or the strain containing the vector only. Even though gels were generally remarkably similar, such individual spots could easily be identified, in some cases also in parts of the gel that were not heavily crowded by host proteins. As an example, Fig. 3 shows the 2-DE images of the control containing the vector only (P. fluorescens pRS44) and the P. fluorescens metagenome library clone P-13. As can be seen there is a particular very clear spot that is present in both technical replicates from the strain with the metagenomic insert, while missing in the vector control strain. The protein in this spot was subjected to LC-MS/MS measurements and the respective spectra were searched against a P. fluorescens specific database or the entire SwissProt database, respectively. Whereas clear cut matches to the P. fluorescens database were observed for all host proteins tested, no significant match against the P. fluorescens or the SwissProt database was obtained for the presumable insert DNA-encoded protein depicted in Fig. 3, even if the search was performed with information-rich spectra. This observation proves the concept by clearly indicating expression of an additional, non-P. fluorescens protein encoded by the environmental DNA insert.

Comparative analysis of protein patterns of Pseudomonas fluorescence-containing the vector pRS44 or the vector with environmental DNA inserts. Crude protein extracts of the control P. fluorescence pRS44 and a strain (P-13) containing pRS44 with an environmental DNA insert were separated by 2-DE and protein patterns were compared with the software package delta- 2 d after staining with silver nitrate. In the lower part detailed zoomed regions of the two technical replicates of each strain containing insert-encoded proteins are shown. An insert-encoded protein is located in the centre of the enlarged section.

Comparative analysis of protein patterns of Pseudomonas fluorescence-containing the vector pRS44 or the vector with environmental DNA inserts. Crude protein extracts of the control P. fluorescence pRS44 and a strain (P-13) containing pRS44 with an environmental DNA insert were separated by 2-DE and protein patterns were compared with the software package delta- 2 d after staining with silver nitrate. In the lower part detailed zoomed regions of the two technical replicates of each strain containing insert-encoded proteins are shown. An insert-encoded protein is located in the centre of the enlarged section.

Construction of BAC clones with inserts up to around 200 kb

To test the capability of the vector (as a BAC) to hold even larger inserts than those described above high-molecular-weight DNA from plant nuclei was cloned into pTA44. This resulted in BAC clones with inserts up to 200 kb. PFGE was used to determine the size of the inserts, and Fig. 4 shows the result for 22 of the obtained BAC clones. The ligation and transformation efficiencies were similar to what was observed in parallel experiments with the commercially available BAC cloning vector pIndigoBAC5. These experiments confirmed that pRS44/pTA44 has retained the capacity of the parent pCC1FOS to carry and maintain very large inserts in E. coli, and the performance of the new vector in alternative hosts could therefore be tested.

Pulsed field gel electrophoretic analysis of 22 NotI-digested BAC clones (lanes 1–22). S, molecular weight standard (kb) (New England BioLabs).

Pulsed field gel electrophoretic analysis of 22 NotI-digested BAC clones (lanes 1–22). S, molecular weight standard (kb) (New England BioLabs).

Transfer of fosmid library and BAC clones from E. coli to P. fluorescens and X. campestris

The E. coli strain EPI300 does not contain the tra genes required for mobilized oriT-mediated conjugation to new hosts, and for this reason the entire fosmid library and selected BACs were first transformed into strain S17.1, which has the RK2 tra genes integrated into the chromosome. Direct library construction in S17.1 can also be carried out but is more complicated due to lower frequencies of transformation. However, it was easy to obtain the required number of transformants from the pre-existing library in EPI300 thus, this additional step will apparently not significantly reduce the representativity of the clones in the original libraries.

Before the fosmid and BAC plasmids were transferred to two selected hosts, P. fluorescens (fosmid library and selected BAC clones) and X. campestris (fosmid clones), the transposon in pRS48 (carrying the trfA gene) was inserted into their chromosomes by electroporation. The entire fosmid library and the selected BAC-clones were then conjugatively transferred to the new hosts. In all cases kanamycin-(P. fluorescens) and chloramphenicol-(X. campestris) resistant clones were obtained at high frequencies, demonstrating that complete fosmid libraries and BAC clones with large inserts could efficiently be transferred.

Plasmids were isolated from independent transconjugants, retransformed into E. coli EPI300, digested with HindIII and analysed by agarose gel electrophoresis. Fosmids (30–35-kb inserts) and BACs with inserts up to 130 kb could be recovered in an intact state from the respective hosts ( Fig. 5). Plasmids with the largest inserts (around 190 kb) were difficult to retransform into E. coli, presumably due to low levels of plasmid recovery and also inefficient transformation. However, Southern hybridization analyses showed that also these plasmids were present in the plasmid state in P. fluorescens (data not shown).

Agarose gel electrophoretic analysis of two HindIII-digested BAC clones before and after passage through Pseudomonas fluorescens::TnRS48. Lanes 1 and 2, plasmid B9 before and after transfer, respectively lanes 3 and 4, plasmid B19 before and after transfer, respectively S, molecular weight standard (kb) (Fermentas).

Agarose gel electrophoretic analysis of two HindIII-digested BAC clones before and after passage through Pseudomonas fluorescens::TnRS48. Lanes 1 and 2, plasmid B9 before and after transfer, respectively lanes 3 and 4, plasmid B19 before and after transfer, respectively S, molecular weight standard (kb) (Fermentas).

Based on all these experiments it could be concluded that pRS44/pTA44 has retained the capacity of the parent plasmid pCC1FOS to hold and maintain very large inserts, that it is more stably maintained than its parent and that it has the capacity to be efficiently transferred to and stably maintained both as a fosmid and a BAC in presumably a large number of non-E. coli hosts. To our knowledge, this is the only available vector system that combines all these features, and pRS44 should therefore represent a very useful tool for functional screening across species barriers.