Helpful Hints: The Transposome Strategy for Creating Knockouts in Bacteria
When using EZ-TN5 Transposomes to create gene knockouts or to insert open reading frames, control
elements, regulatory elements and antibiotic resistances into bacteria, there are number of considerations
that must be made in order to plan for a successful transposition event in vivo. The three main
considerations given below should be examined prior to performing in vivo transposition reactions using
the EZ-TN5 transposomes.
The organism must have a good transformation efficiency using electroporation.
First and foremost, the bacterium or other cell type you plan to use must be easily transformed by
electroporation. Transposomes will not work when using chemical transformation (e.g., calcium
chloride/Hanahan method). We suggest that the “basic” transformation efficiency for your bacterium be at
6
least 10 cfu/µg when transforming a simple plasmid into the cells (as an example, E. coli and pUC19,
which gives a base transformation efficiency of 108 to 1010 cfu/µg using electroporation). Transformation
efficiencies lower than this will result in either no or very few transposition clones obtained.
Electroporation conditions will vary from organism to organism. For a standard E. coli strain, we
recommend using 50 µl of electrocompetent cells, 0.2 cm (2 mm) electroporation cuvette, 2500 volts, and
5 msec time constant. These have been shown to work very well in a number of bacterial types, and the
reason for using a 2mm cuvette is to reduce as much as possible the potential for arc-ing (and thus
failure of the electroporation). Even though the transposomes are delivered to the cells in water, a narrow
cuvette gap, such as 1 mm (0.1 cm) increases the likelihood of arcing during the electroporation step.
Gram-positives are difficult to electroporate in general but a recent article (Pajunene, et al.: “Generation of
transposon insertion mutant libraries in Gram positive bacteria by electroporation of phage Mu DNA
transposition complexes”, Microbiology 151: 1209-1218 [2005]) gives good details on making gram
positive organisms electrocompetent. Note that this article presents infromation on Mu complexes but the
same information can be used with EZ-Tn5 Transposome complexes.
The antibiotic resistance marker must function in the organism of interest.
The resistance marker must be able to express in the cell. In Gram-negative cell types, the Kanamycin
resistance gene as set up in our KAN-2 transposon will almost always be the selectable marker of choice.
The KAN marker is also active in certain gram positive bacteria, and not active in others. This can be due
to the nature of the promoter (it may or may not be recognized by a particular cell type, or will have much
weaker activity that is seen in E. coli). In some gram positives, some researchers have had success in
making their own transposons using “alternative” resistance markers, such as spectinomycin, hygromycin,
and erythromycin.
If you are getting ready to create an in vivo knockout library and plan to use transposons with the
Kanamycin resistance marker, we urge you to consider the R6Kγ/KAN-2 transposome, which is very
useful in performing gene rescue studies. While it may seem that creating a knockout library is the only
goal of the research, the ability to rescue a gene once it is knocked out is more and more becoming the
goal in many genomics projects, so it is very useful to consider downstream research when creating the
original transposition library.
Beware of endogenous restriction endonucleases!
In many cells (especially those isolated from “wild” locations), there exists a sort of immune system that
protects the organism from unwanted DNA brought into the cells through transfection, mating, or other
gene transfer event. These systems, called restriction and modification systems, permit the host
organism to degrade unwanted, unprotected DNA once it enters the cell. There are two types of
restriction/modification systems: Type I and Type II.
Type I restriction systems (cited from Murray, N.M., Microbiol Mol Biol Rev. 2000 June; 64 [2]: 412–434):
“Restriction enzymes are well known as reagents widely used by molecular biologists for genetic
manipulation and analysis, but these reagents represent only one class (type II) of a wider range of
enzymes that recognize specific nucleotide sequences in DNA molecules and detect the provenance of
the DNA on the basis of specific modifications to their target sequence. Type I restriction and modification
(R-M) systems are complex; a single multifunctional enzyme can respond to the modification state of its
target sequence with the alternative activities of modification or restriction. In the absence of DNA
modification, a type I R-M enzyme behaves like a molecular motor, translocating vast stretches of DNA
towards itself before eventually breaking the DNA molecule”.
Type II restriction systems (From Pingold and Jeltsch, Nucleic Acids Research 2001; 29 (18): 3705–3727)
“More than 3000 type II restriction endonucleases have been discovered. They recognize short, usually
palindromic, sequences of 4–8 bp and, in the presence of Mg2+, cleave the DNA within or in close
proximity to the recognition sequence. The orthodox type II enzymes are homodimers which recognize
palindromic sites. Depending on particular features subtypes are classified. All structures of restriction
enzymes show a common structural core comprising four β-strands and one α-helix. Furthermore, two
families of enzymes can be distinguished which are structurally very similar (EcoRI-like enzymes and
EcoRV-like enzymes). Like other DNA binding proteins, restriction enzymes are capable of non-specific
DNA binding, which is the prerequisite for efficient target site location by facilitated diffusion. Non-specific
binding usually does not involve interactions with the bases but only with the DNA backbone. In contrast,
specific binding is characterized by an intimate interplay between direct (interaction with the bases) and
indirect (interaction with the backbone) readout....The precise mechanism of cleavage has not yet been
established for any enzyme, the main uncertainty concerns the number of Mg2+ ions directly involved in
cleavage. Cleavage in the two strands usually occurs in a concerted fashion and leads to inversion of
configuration at the phosphorus. The products of the reaction are DNA fragments with a 3′-OH and a 5′phosphate.”
There are two ways to defeat the effect restriction endonucleases have on transposon DNA
electroporated into the organism: using a protein that in effect “plugs” the active site of type I restriction
enzymes and employing a process we call “laundering”, which results in the protection of DNA from Type
I AND Type II restriction digestion. Note also that restriction enzyme digestion can be a major problem,
restriction enzyme recognition sites are very sequence-specific. If the restriction endonuclease does not
find a recognition site in the transposon DNA, transposition into host cell DNA will not be inhibited.
However, in many “wild” organisms, the restriction/modification systems have undefined recognition sites
and even extensive characterization of the organism may not be able to predict the recognition sites
attacked by the restriction enzyme.
There are procedures available for testing novel organisms to be transposed for the presence of
restriction enzymes. The protocol for testing is quite simple:
1.
Grow bacterial strain in desired growth medium to late logarithmic growth stage.
2.
Pellet 1.5 ml cells in microfuge tube (7.5X for 5 min in Beckman Microfuge 11). Decant
supernatant.
3.
Wash cells in 10 mM Tris (pH 8), 1mM EDTA. Pellet cells. Decant supernatant.
4.
Add 1 ml 1X low salt restriction enzyme buffer.
5.
Place tube in ice for 5 min.
6.
Sonicate sample with microprobe:
30 sec. 50% pulsed #2 setting
30 sec. 50% pulsed #3 setting
30 sec. 50% pulsed #4 setting
7.
Pellet sonicated lysate (13.5X for 2 min.)
8.
Transfer 500 ul of supernatant from top to new tube.
9.
Assay:
10 ul Lambda DNA (50 ng/ul in 1X low salt restriction enzyme buffer)
10 ul sonicated lysate
10.
Incubate 2 hr at 37°C. Heat inactivate with 5 min at 68 C. Place on ice.
11.
Run sample on gel.
Lambda DNA stock: 55 ul DNA (0.03 ug/ul sol'n), 40 ul 10X low salt, 305 µl water which will yield a DNA
solution of about 0.004 µg/µl.
In addition to the Lambda DNA, one can include the transposon/pMOD constructs in the testing to
determine the effect any restriction enzymes might have on the transposon DNA.
A. TypeOne Restriction Inhibitor (Catalog number TY0261H: 100 µg)
EPICENTRE offers TypeOne™, a phage protein from the “ocr” gene that acts as a molecular “decoy” that
blocks the binding of the transposon (or any unprotected DNA for that matter) to any Type I restriction
endonuclease, essentially buying time for the transposon to randomly insert into the host cell DNA.
Transformation/transposition frequencies can be improved by as much as three orders of magnitude in E.
coli MG1655 (see http://www.epibio.com/item.asp?ID=392), See the EPICENTRE Forum, Issue 9:2, page
8: http://www.epibio.com/pdfforum/9_2typeone.pdf
B. Laundering transposon DNA: a method to protect transposons once they enter the hostile environment
inside a r+m+ cell
The laundering process is a method that is used to generate methylated transposon DNA that will resist
digestion by endogenous restriction endonucleases. The process involves several steps including
transformation of a transposon-containing plasmid construct into the desired target cell selection with an
antibiotic and purification of the transposon DNA from the donor backbone (plasmid vectors). The
technique is most easily used with the pMOD-2 and pMOD-3 transposon construction vectors in Gram
negative bacteria, but can be used in Gram positive bacteria provided that a functional gram-positive
origin of replication is provided in the vector.
The transposons that EPICENTRE sells are PCR products and as such are quite susceptible to restriction
digestion.
Here is a sample protocol for laundering transposon DNA, using a fictitious transposon construct in our
pMOD-2 vector with a chloramphenicol resistance marker into a Gram-negative cell that is r+m+:
1. Using electroporation, transform the chloramphenicol resistant plasmid into the cell using standard
techniques (see above).
2. Select the cells on LB medium containing chloramphenicol. Note that in the step, the apparent
transformation efficiency will be very low (perhaps as few as one or two colonies). Do not be alarmed by
this apparent lack of success.
3. Recover one of the chloramphenicol-resistant clones and re-streak onto chloramphenicol-containing
medium to confirm resistance to the antibiotic.
4. Pick a portion of one Cm-resistant colony and inoculate into 1 ml of broth medium containing the
antibiotic and incubate overnight.
5. Isolate the plasmid DNA (containing the transposon) using miniprep methods and confirm present of
the transposon by PvuII digestion and agarose gel electrophoresis.
6. Once the intact transposon has been confirmed, grow a larger culture (50-100 ml) of the plasmidcontaining cells and isolate the plasmid DNA (midiprep/maxiprep).
7. Take a small sample of the plasmid DNA isolated in step 6 and re-transform the restriction
endonuclease-containing bacterium and plate on the appropriate antibiotic-containing medium. The
apparent transformation efficiency should now be MUCH higher (closer to what the basic transformation
efficiency should be). If this is indeed the case, the DNA is now “protected” (methylated).
8. Take the remaining transposon-containing pMOD construct, digest with either PvuII or PshAI, to purify
the transposon DNA using agarose gel electrophoresis (being careful not to expose the transposon DNA
to UV light and Ethidium bromide),
9. Mix with the EZ-TN5 Transposase to make the Transposome.
10. Electroporate the Transposome into your organism, select for the appropriate transposon insertion
mutants.
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Additional Tips and Tricks
The EZ-Tn5 TET-1 “Transposome”
While EPICENTRE does not sell a bona fide TET Transposome, one can easily be constructed using
techniques outlined above. The main reason we do not sell TET-1 Transposomes is that the TET marker
works far better in high-copy applications than in single-copy applications (such as Fosmid mutagenesis
and sequencing applications). However, recent applications research by EPICENTRE scientists has led
to a useful and simple protocol modification that will increase the number of TET-resistant colonies
generated when a cell is mutagenized using s TET-1 Transposome.
This simple protocol modification involves the addition of 5mM of either MgSO4 or MgCl2 to the plating
medium, along with the standard amount (usually 10 µg/ml) tetracycline. Note that while the product
literature for the EZ-Tn5 TET-1 Insertion Kit cautions against the use of magnesium in the medium;
testing has shown that an increase in the number of TET-resistant clones can be achieved. The result will
vary based on electroporation conditions and cell type being used. Recently, a paper was puiblsihed that
documents the use of TET Transposomes in the bacterium Pseudomonas aeruginosa, strain PAO1, that
has been reproduced by a number of our customers ( Filiatrault, MJ, et al.: “ Identification of
Pseudomonas aeruginosa Genes Involved in Virulence and Anaerobic Growth”. Infection and Immunity V.
74, No. 7, 4237-4245 (2006).
The EZ-Tn5 TET-1 Insertion Kit (Catalog number EZI921T) provides all of the needed reagents (except
for glycerol) required for synthesis of the TET-1 Transposome.
The basic protocol for making Transposomes is quite simple, and is outlined on Page 3 of the Product
literature for the EZ-Tn5 Transposase: http://www.epibio.com/item.asp?ID=292
Production of EZ-Tn5 Transposomes
Making Transposomes using the EZ-Tn5 Transposase and transposon DNA is very straightforward. The
protocol, reproduced below, was taken from the EZ-Tn5 Transposase product literature:
http://www.epibio.com/item.asp?ID=292.
Production of stable EZ-Tn5 Transposomes can only be accomplished in the absence of Mg+2.
Do not use the EZ-Tn5 10X Reaction Buffer provided with the EZ-Tn5 Transposase to prepare
EZ-Tn5 Transposomes.
1. Prepare the transposome reaction mixture by adding in the following order:
2 µl EZ-Tn5 Transposon DNA (100 mg/ml in TE Buffer [10 mM Tris-HCl (pH 7.5), 1 mM EDTA])
4 µl EZ-Tn5 Transposase
2 µl 100% glycerol
8 µl Total reaction volume
2. Mix by vortexing. Incubate for 30 minutes at room temperature.
3. Store the solution at -20°C.
The solution will not freeze stored at -20°C and is stable for at least one year.
4. Use 1 ml of the EZ-Tn5 Transposome for electroporation into a competent bacterial strain and
plate on selective media as dictated by the transposon insert.
The EZ-Tn5 Transposome production protocol can be scaled up or scaled down as needed.
One thing we have seen on occasion is that incubating the Transposome constructed for a longer period
of time (over the weekend, for example) can increase the efficiency of the transposition reaction. At this
time we do not understand why this occurs and it’s not consistent with all Transposomes made.
Making your own custom EZ-Tn5 Transposon
1. Using the EZ-Tn5 pMOD-series transposon construction vectors
EPICENTRE makes it easy for anyone to make their own transposons for use in in vitro OR in vivo
mutagenesis/knockout experiments. In addition to providing a range of cloning vectors, we also provide
technical assistance for making the transposons by PCR using a number of different procedures.
The pMOD cloning vectors are pUC19-based plasmids that use the standard pUC backbone and have
the following features:
A. The pUC19 MCS multiple cloning site
B. The required Mosaic Ends that are essential for transposition (synaptic complex formation
between the transposon and the Transposase enzymes)
C. The R6Kγ origin of replication – the key tool for doing “rescue cloning”
Almost ANY DNA can be cloned into the pMOD vector and be used to make a transposon. The longest
transposons that we are aware of are approximately 12-13 kb. One should also understand that the
longer the transposon the lower the efficiency will be. The longest of our standard transposons is just over
2.0 kb; efficiency remains very good under standard conditions with transposons as long as 5 kb
(unpublished data).
The following transposon construction vectors are available from EPICENTRE:
EZ-Tn5 pMOD2 and pMOD-4 Transposon Construction vectors
http://www.epibio.com/item.asp?id=291
These vectors contain the two transposon “Mosaic end” sequences for transposon binding and an MCS.
The pMOD-4 version has been developed with a single R6Kγ origin of replication, which requires growth
in a pir-containing E. coli strain (unlike the pMOD-2 vector, which has a standard ColEI origin that will
grow in many different E. coli strains).
The reason for the development of the pMOD-4 vector is to reduce the amount of “background clones”
seen in a number of in vitro or in vivo transposition reactions. These background clones have been found
to contain whole pMOD-2 constructs that were not digested or incompletely digested with the restriction
enzymes PvuII or PshAI and were not sufficiently purified away from the transposon itself…leading to
“false positive” transposition clones. As one will see below, purification of the transposon away from the
vector is a key step in assuring excellent results from your transposon experiments.
The second class of transposon construction vectors (containing the R6Kγ origin of replication) are
intended for future rescue cloning operations. Once the transposon has been inserted into the desired
target DNA (either a plasmid/episome or into chromosomal DNA), the transposon and the surrounding
DNA can be sheared or digested mechanically or by using judiciously chosen restriction enzymes,
followed by religation of the DNA to generate plasmid containing the transposon and the surrounding
DNA for “rescue cloning” experiments.
Link to the EZ-Tn5 pMOD-3 and pMOD-5 Transposon Construction vector on-line information:
http://www.epibio.com/item.asp?id=291
Methods for purifying transposon DNA
The key to making a good transposon is to make sure that the transposon is “clean” and free of
contaminating vector and extraneous DNA.
A functional EZ-Tn5 Transposon can be isolated either by restriction enzyme digestion or PCR
amplification. If the transposon is prepared by restriction enzyme digestion we recommend using pMOD-4
<MCS> which contains an R6Kγ origin of replication rather than a colE1 origin of replication. Replication
from the R6Kγ origin is dependent on the pir gene product produced by TransforMax EC100D
pir+ and pir-116 E. coli cells (sold separately). Since most bacterial strains do not contain a pir gene,
the uncut plasmid DNA that contaminates these transposon preparations can’t replicate and this type
of background problem is eliminated.
Purification of transposon DNA by Gel electrophoresis
This protocol below is a slight modification of the DNA gel purification method we use for our Cosmid,
Fosmid, and CopyControl Fosmid Kits. It has been adapted for use with transposon DNA that has been
cut out of a pMOD-2 or pMOD-3 vector using PvuII or PshAI. By eliminating the ethidium and UV
exposure, you will note an excellent increase in transformation and transposition efficiency when the
transposon DNA is used in either in vitro insertion or in vivo Transposome mutagenesis.
After digestion of the pMOD-based chimera with PvuII or PshAI, analyze the digested DNA on a 1%
agarose gel. If doing standard agarose gel electrophoresis, be certain to cast a long gel (at least 14 cm,
preferably 20 cm... but NOT a minigel). Perform electrophoresis overnight, at 1.5 Volt/cm, using the length
of the gel box to measure the “cm”.... that is, not the length of the gel, but the length of the electrophoresis
box). This will give optimal separation of your DNA
Fractionate the DNA on a low melting temperature agarose gel. It is important to perform this
electrophoresis in the absence of Ethidium Bromide (do not add Ethidium Bromide to the gel). The
DNA that will be cloned should not be exposed to UV light under any circumstances. This can decrease
the cloning efficiency by 100-fold or more. A diagram on our method is shown in Figure 1.
NOTE 1: Even 30 seconds exposure to 302 nm UV light will cause a 100-fold to 200-fold drop in ligation
and cloning efficiency.
Note 2: The protocol below is designed for use with our GELase agarose digestion preparation, and thus
requires low-melting-point agarose. Standard high-melt agarose can also be used and the DNA extracted
from the gel slices with other methods (Qiagen, Gene-clean, etc.).
1. Prepare a 1% LMP agarose gel in 1X TAE or 1X TBE buffer.
Note: Do not include ethidium bromide in the gel solution.
2. Load the DNA Size Marker into each of the outside lanes of the gel and fill the rest of the wells with
PvuII or PshAI-digested pMOD transposon construct.
3. Resolve the samples by gel electrophoresis (for example at room temperature overnight) at a
constant voltage of 30-35 V. Visualizing and excising the end-repaired DNA can be done by
one of two methods described below.
4. Following electrophoresis, cut off the outer lanes of the gel containing the DNA Size Marker, and a
small portion of the next lanes that contain your digested DNA. (see Figure 1).
Stain the cut-off sides of the gel with ethidium bromide and visualize the with UV light. Mark the position
of the desired size DNA in the gel using a Pasteur pipet. Note: Do not expose the sample DNA to UV
irradiation! Even short UV exposure can decrease cloning efficiencies by 100-1000 fold.
Reassemble the gel and excise a 2-4 mm wide gel slice containing sample DNA that migrated with and
just slightly above (higher MW) the appropriate position of the DNA markers.
Transfer the gel slice to a tared, sterile, 15 ml screw-cap tube for extraction, either by using the GELase
method, or other desired method for isolating DNA from agarose gels (see figure 1 for graphic).
Figure 1.
Purification of transposon DNA (PCR Product or restricted transposon clone) by gel
electrophoresis.
Stain this
side
Do Not Stain
center
section!!
Stain this
side
DNA Size
standards
Mark
Gel
here
Mark
Gel
here
Cut here
Keep this part of the gel,
but do NOT stain it!
Cut here
PCR Amplification of Transposon DNA
Note: Make sure that the PCR primers, no matter which of the three methods you choose, have a
5’-phosphate on both forward and reverse primers.
There are three standard methods for making transposons using PCR. They are:
1. Using a long-standing protocol using the pMOD PCR primers provided with the EZ-Tn5 pMODseries Transposon Construction vectors.
Linearize the pMOD transposon construct using an enzyme that will cut only in the vector backbone. An
excellent choice, when possible, would be an enzyme that cuts in the origin of replication. Linear DNA is a
better template for PCR than circular DNA. A very good enzyme to use (is AlwNI since it cuts in the origin
of replication on the pMOD vector. The enzyme we would suggest for this is AlwNI (cuts in the ColEI
replicon of the vectors pMOD-2 and pMOD-3). This linearization should not be required when using the
transposon construction vectors pMOD4 or pMOD-5.
Amplify the transposon region using the pMOD<MCS> Forward and Reverse PCR Primers provided
with the vector. A suggested cycling profile is outlined below.
a. Initially, denature the template at 94°C for 2 minutes.
b. Perform 30 cycles of:
Denature at 94°C for 30 seconds.
Anneal at 60°°C for 45 seconds.
Extend at 72°C for 1 minute for every kb of expected product.
We recommend PEG precipitation to remove small molecules (e.g. primers, nucleotides) that may
interfere with transposition. Alternatively, a standard ethanol precipitation can be used.
a. Dilute the PCR reaction to 500 μl with TE.
b. Add 250 μl of 5 M NaCl and 250 μl of 30% PEG 8000/1.5 M NaCl.
c. Mix well and incubate at 4oC for at least 30 minutes.
d. Centrifuge at 4oC for 10 minutes at 10,000 x g. Discard the supernatant, centrifuge again
for a few seconds, and discard any remaining supernatant.
e. Dissolve the DNA in a suitable amount of TE.
For those researchers who wish to gel-purify their transposons after synthesis by PCR, we strongly
recommend following the procedure shown in Figure 1 above.
2. PCR using the “ME-Plus” primers with transposon constructs created in the pMOD-series vectors.
This is a straightforward method that serves as an alternative to PvuII/PshAI digestion, especially if the
transposon construct has PvuII and PshAI sites within the transposon construct. It also serves as an
alternative to using the pMOD PCR primers, which require “pruning” of the ends of the PCR product with
PvuII or PshAI for maximum transposition efficiency. The pMOD transposon construct, after building using
normal recombinant DNA procedures and grown up in E. coli, is purified using any alkaline lysis-based
plasmid preparation kit. We recommend EPICENTRE’s PlasmidMAX Plasmid DNA Purification Kit
(which uses a method that virtually eliminates contaminating chromosomal DNA from the plasmid
preparation).
1) Linearize the pMOD transposon construct using an enzyme that will cut only in the vector backbone.
An excellent choice, when possible, would be an enzyme that cuts in the origin of replication. Linear DNA
is a better template for PCR than circular DNA. A very good enzyme to use (is AlwNI since it cuts in the
origin of replication on the pMOD vector. The enzyme we would suggest for this is AlwNI (cuts in the
ColEI replicon of the vectors pMOD-2 and pMOD-3).
2) Perform PCR using primers which incorporate the reverse complement of the Mosaic Element. We use
the reverse complement of the ME plus 9 bases.
Sequences of the two primers we use:
ME Plus9 – 3’ primer
5'-CTGTCTCTTATACACATCTCAACCATCA-3'
ME Plus9 – 5’ primer
5'-CTGTCTCTTATACACATCTCAACCCTGA-3'
Cycling protocol is
94°C for 1 min, followed by:
25-30 cycles of:
94°C for 30 sec.
55°C for 1 min, and
72°C for X min*** (One minute per kb PCR product).
(***You will want to modify the elongation time for the size construct you have made.)
As far as what buffer to use, I imagine each template will be different. You will want to use a high fidelity
enzyme. Our FailSafe enzyme would be an excellent choice as it will give you 3X the fidelity of Taqw but
does not have the processivity issues of many pure proofreaders. We strongly recommend to use the
FailSafe PCR Selection Kit, use all twelve of the FailSafe 2X PCR PreMixes, and find out the best one to
use for each template.
Here is the link to the product information in case you have not heard of FailSafe before:
http://www.epibio.com/item.asp?id=294
One more suggestion, after finding the best 2X PCR PreMix for the transposon amplification, we would
recommend performing PCR with 10-50 tubes of the same PCR to get enough Transposon DNA.
Then pool all PCR products together, PEG precipitate the DNA, and resuspend in final concentration of
~100 ng/ul**, which is what we sell.
**Adjust concentration appropriately depending on the size of your custom transposon.
3. PCR using primers constructed to cover the desired genes for transposition that are specific for
the desired DNA and tailed with the 19-base mosaic ends.
This method, using PCR primers that are “ME-tailed”, is described in a recent EPICENTRE Forum article:
Prepare Custom EZ::TN™ Transposons by PCR Using Primers with Transposase-Specific Mosaic
End (ME) Sequences
(as shown in EPICENTRE Forum Volume 8, issue 3, page 4)
http://www.epibio.com/pdfforum/8_3me.pdf
Again, for maximum efficiency, a 5’-phosphate is required on these primers!