Temporal self-regulation of transposition through host-independent transposase rodlet formation
Nucleic Acids Research
Temporal self-regulation of transposition through host-independent transposase rodlet formation
Lauren E. Woodard 0 1 2
Laura M. Downes 1 2
Yi-Chien Lee 0 1
Aparna Kaja 0 1
Eyuel S. Terefe 0 1
Matthew H. Wilson 0 1 2
0 Department of Veterans Affairs , Houston, TX 77030 , USA and Department of Medicine, Baylor College of Medicine , Houston, TX 77030 , USA
1 Published by Oxford University Press on behalf of Nucleic Acids Research 2016. This work is written by (a) US Government employee(s) and is in the public domain in the US
2 Department of Veterans Affairs , Nashville, TN 37212 , USA and Department of Medicine, Vanderbilt University , Nashville, TN 37232 , USA
Transposons are highly abundant in eukaryotic genomes, but their mobilization must be finely tuned to maintain host organism fitness and allow for transposon propagation. Forty percent of the human genome is comprised of transposable element sequences, and the most abundant cut-andpaste transposons are from the hAT superfamily. We found that the hAT transposase TcBuster from Tribolium castaneum formed filamentous structures, or rodlets, in human tissue culture cells, after gene transfer to adult mice, and ex vivo in cell-free conditions, indicating that host co-factors or cellular structures were not required for rodlet formation. Timelapsed imaging of GFP-laced rodlets in human cells revealed that they formed quickly in a dynamic process involving fusion and fission. We delayed the availability of the transposon DNA and found that transposition declined after transposase concentrations became high enough for visible transposase rodlets to appear. In combination with earlier findings for maize Ac elements, these results give insight into transposase overproduction inhibition by demonstrating that the appearance of transposase protein structures and the end of active transposition are simultaneous, an effect with implications for genetic engineering and horizontal gene transfer.
Transposition is an ancient and influential force on the
genome of nearly every organism, reshaping chromosomes
by an inherently mutagenic process (1). Transposons
commonly make up the majority of eukaryotic genomes, with
maize having the most transposon-rich genome (>85%) (2).
The transposase protein coding genes are the most
abundant and ubiquitous genes in nature (3). Using these
diverse transposases as genetic starting material, hosts have
tamed them into “domesticated transposases” with diverse
and crucial functions (4).
Class II DNA transposons cut-and-paste DNA
sequences in the genome and are useful for gene transfer
applications. Transposons with a linear dsDNA intermediate
(5) propagate mainly by replication of the host genome, in
the process also replicating the transposon. In this study,
we report data on two DD(E/D)-transposons of insect
origin: Tribolium castaneum Buster (TcBuster; (6) and
piggyBac (7). TcBuster is a newly discovered member of the hAT
(hobo, Ac, Tam3) transposon superfamily that includes the
transposon Hermes which has a known crystal structure
(8,9). PiggyBac is the namesake for its own family (10).
We found that TcBuster has a high level of activity in
human tissue culture cells but displayed an overproduction
inhibition/negative dosage effect––meaning that high doses
of transposase resulted in suboptimal transposition (11).
In fact, in a careful review of the literature the majority of
DD(E/D) transposons tested displayed overproduction
inhibition (12), indicating that the cause of overproduction
inhibition must be: (i) independent of the details of
transposase structure or exact mechanism of DNA cleavage; (ii)
present in all transposons and (iii) independent of the host.
The overproduction inhibition/negative dosage effect
was first described by McClintock (13). This effect has
presented the need for optimization for the successful creation
of transposon vectors for genome modification, most
notably for the Sleeping Beauty transposon (12,14,15).
Despite the difficulty this presents in genome engineering (16),
in nature transposon self-regulation is expected so that
suicidal autointegration events are limited (17).
Additionally, since the health of the host is crucial for
transposon replication, transposons cannot act as purely ’selfish’
genes: this adjective implies unmitigated harm to the host
on which the transposon relies (18). DNA transposons keep
their activity in check through an assortment of
mechanisms, including complementary regulatory RNA (19),
repressors (20,21), and concentration-dependent negative
feedback mechanisms (22). The relationship between
transposase concentration and the rate of transposition could be
thought of as taking three possible courses: a linear or
exponential increase in activity, overproduction inhibition in
which activity decreases above a certain transposase
concentration, and saturated transposition activity by
transposase overconcentration where the increasing amount of
transposase beyond a certain point does not affect the rate
of transposition. In 33 out of 50 manuscripts reviewed by
Bire et al., classic overproduction inhibition was reported;
therefore, the most common outcome of a surplus of
transposase protein across a wide variety of DNA cut-and-paste
transposons from different families is a decrease in
transposition rate (12). Our data expand upon previous studies
that linked overproduction inhibition or negative dosage
effect to high transposase protein concentration by linking
this inhibition to the creation of host-independent
structures. To further investigate these rodlet structures, we made
several tagged versions of TcBuster transposase and
discovered they were created in a dynamic process in human tissue
culture cells, in cell-free solutions ex vivo, and in mice. We
also demonstrated that transposition occurred prior to the
detection of transposase rodlets and that the presence of
aggregates for both TcBuster and piggyBac signaled the end of
a window of transposition.
MATERIALS AND METHODS
Plasmids were prepared by endotoxin-free maxiprep
(Qiagen, Valencia, CA, USA) or midiprep (Zymo Research
Corp., Irvine, CA, USA). The filler DNA plasmid pUC19
(Invitrogen, Carlsbad, CA, USA) is commercially
available. The construction of the piggyBac plasmids
pCMVpiggyBac (23), pCMV-HA-piggyBac (24), pTpB (23), and
pTpBCAGLuc (referred to as pT-CAGLuc in (25) and the
TcBuster plasmids pCMV-TcBuster and pTcBNeo (11) have
been described elsewhere.
pCMV-HA-TcBuster was created concurrently with
pCMV-TcBuster (11), the only difference being the forward
primer SpeITCB2 (GCTGACTAGTATGATGCTGAAT
TGGCTGAAAAGC) used to clone the insert into the SpeI
site of pCMV-HA-piggyBac to allow retention of the HA
tag at the N-terminus to create pCMV-HA-TcBuster.
pCMV-Flag-TcBuster was created by PCR of
pCMVTcBuster with the primers TcB-Flag-F (CCCCGCGG
TcB-Flag-R (CTATGCGTCGGCTGATAGTG) to create
a Flag-tagged insert fragment that was digested and cloned
into the SacII and BstBI sites of pCMV-TcBuster.
To create pCMV-TcBuster-GFP, a similar PCR of
pCMV-TcBuster with the primers TcB-c3GFP-for
and TCB-c3GFP-rev (GGTGAGATTTCTGGGCCT
GCTTG) synthesized an insert containing the
TcBuster reading frame that was TOPO cloned via the
pCDNA3.1/CT-GFP-TOPO kit (Invitrogen), which
was also transformed alone to create the
pCMVGFP empty vector control. To create
pCMV-GFPTcBuster, a PCR of pCMV-TcBuster was performed
with the primers TcB-c3GFP-for and RevKpnITcB2
and TOPO cloned into pCDNA3.1/NT-GFP-TOPO
(Invitrogen). The same linker (GGSGGSGGSGGSGTS)
we have previously employed (15,26) was introduced via
annealed oligos into the AgeI site in pCMV-GFP-TcBuster
and the XbaI site in pCMV-TcBuster-GFP.
The luciferase-expressing TcBuster transposon
pTcBCAGLuc was created by digest of the pTpBCAGLuc
plasmid with BsaI, SpeI, and BamHI to liberate the
CAGLuc cassette. This insert was then cloned into the
TcBuster transposon pXL-TcB-D-GFP/Bsd (27) digested
with BglII and SpeI.
HEK-293 and HeLa cells (ATCC, Manassas, VA) were
tured in cellgro Minimum Essential Medium, Alpha 1×
with Earle’s Salts, ribonucleosides, deoxyribonucleosides
and L-glutamine (Mediatech, Inc., Manassas, VA) to which
10% fetal bovine serum (Thermo Scientific, Logan, UT) and
1% penicillin/streptomycin (Invitrogen) were added.
Immunofluorescent staining and live cell imaging of tissue
For staining of the HA tag, HEK-293 cells were plated
onto poly-D-lysine-coated 12 mm glass coverslips (BD
Biosciences, San Jose, CA) or HeLa cells were plated onto
uncoated 18 mm glass coverslips. Cells were transfected
via FuGene 6 (Roche, Indianapolis, IN) and fixed in 4%
paraformaldehyde in phosphate-buffered saline (PBS; USB
Corporation, Cleveland, OH) for 20 min. Fixation was
followed by five washes of PBS, permeabilization in PBST
(PBS with 0.1% Triton-X 100) for 10 min, blocking in
PBSBT (PBS with 3% bovine serum albumin, 0.1% Triton-X
100 and 0.02% sodium azide) for 30 min, incubation in
primary rat -HA antibody (3F10, Roche) or rat IgG (Santa
Cruz Biotechnology, Dallas, TX) diluted 1:100 in PBSBT
for 45 min, three washes of PBSBT, and incubation in
secondary goat -rat Alexa Fluor 594 antibody (Invitrogen)
diluted 1:1000 in PBSBT for 30 min. After three washes of
PBSBT, the coverslips were mounted in ProLong Gold
Antifade Reagent plus DAPI (Invitrogen). The following day,
the coverslips were sealed with nail polish.
For co-staining of the Flag and HA tags, the above
procedure was carried out with the addition of a Flag staining
step after blocking and before incubation with -HA.
Either mouse IgG (Santa Cruz Biotechnologies) or mouse M2
monoclonal -Flag (Sigma, St. Louis, MO) were diluted
1:500 in PBSBT, clarified, and incubated on the cells for 45
min followed by three washes in PBSBT. The secondary
antibody was goat -mouse Alexa Fluor 488 (Invitrogen) for
30 minutes followed by 3 washes in PBSBT.
For imaging of GFP and DAPI together, cells grown on
coverslips were mounted directly in ProLong Gold Antifade
Reagent with DAPI. For live cell imaging of GFP-tagged
rodlets, HEK-293 cells were grown and transfected in 35
mm dishes with glass windows (MatTek Corporation,
Ashland, MA) in media without phenol red. They were
maintained in a heated incubator affixed to an inverted confocal
microscope and imaged every 10 min overnight.
Images were acquired on an Olympus BX51, Nikon
Eclipse 80i, LSM510 Meta Inverted, or a DeltaVision
Image Restoration Microscope. Images were merged in
Photoshop (Adobe, San Jose, CA). For quantification of rodlet
number, five pictures per coverslip were obtained and
picture names were coded. We then counted number of DAPI+
nuclei, the number of nuclei that have rodlets, and the
total number of rodlets per picture in a blinded fashion. Data
were analyzed and lines of best fit determined in Excel
(Microsoft, Seattle, WA).
HEK-293 cells seeded onto six-well plates were
transfected at 50–80% confluence in triplicate with FuGene6
(Roche) at a 3:1 ratio of FuGene6 to DNA according to
the manufacturer’s instructions. Two days later, the cells
were trypsinized, reconstituted in 1 ml of media, and 13.33
l was transferred onto 10 cm plates containing 10 ml
of media containing 1 mg/ml geneticin (Invitrogen) for a
1:750 dilution (unless noted differently in the figure legend).
Two weeks later, the cells were fixed and stained with 50%
methanol (VWR, Radnor, PA) and 1% methylene blue
(Invitrogen). After 1 h, the plates were washed twice with PBS
and colonies were counted by hand following drying. Error
bars represent the standard error of the mean.
Protein purification and cell-free chamber system
We transfected eight 10 cm plates of HEK-293 cells with
8 g of pCMV-Flag-TcBuster plasmid each 48 h prior to
protein lysis in 125 l of CellLytic M buffer (Sigma) with
mini cOmplete ULTRA protease inhibitor cocktail tablet
(Roche) per plate. The Flag M Purification Kit was used to
absorb the Flag-TcBuster protein onto ANTI-FLAG M2
affinity gel followed by elution by competition with the
supplied 3xFlag peptide (Sigma). A clean flow chamber was
created as previously described (28). A solution
containing 32% purified protein in modified PEM buffer (90 mM
PIPES, 1.1 mM ethylene glycol tetraacetic acid (EGTA)
and 0.1 mM MgCl2, pH 6.9) with or without 0.4 ng/ l
pTcBNeo was flowed into the chamber. After evaporation
overnight at 37◦C, 10 mM PIPES buffer was introduced into
the chamber which was sealed with wax and the rodlets were
imaged by confocal differential interference contrast (DIC)
microscopy on a LSM 510 Meta Inverted microscope.
For Figure 1, our procedure for obtaining cytoplasmic and
nuclear protein fractions has been described previously (29).
To obtain the insoluble protein fraction, the pellet that was
not soluble in either the hypertonic or nuclear lysis buffers
was resuspended in the same volume of cell lysis buffer as
used to obtain soluble protein fractions (550 l). The 10%
Bis–Tris NuPage Gel (Invitrogen) was loaded with 20 g
of soluble protein and 40 l of insoluble suspension,
transferred via iBLOT (Invitrogen) to a nitrocellulose membrane,
immunoblotted for HA and -actin, and imaged as
previously described (29) except that rat -HA (3F10, Roche)
diluted 1:1000 and goat anti-rat 800 diluted 1:15 000 were
used to detect HA. For Figure 7, cells were harvested by
scraping in PBS, washed once in PBS, and frozen. RIPA
(Sigma) supplemented with PhosSTOP (Roche) and
cOmplete ULTRA Tablets (Roche) inhibitors was added to each
thawed cell pellet and cells were further lysed by freeze-thaw.
Protein lysates were analyzed by Pierce BCA Protein Assay
Kit (Thermo Scientific) and 25 g protein was combined
with NuPAGE sample buffer and reducing agent (Life
Technologies), loaded onto 4–12% Bis–Tris gels (Life
Technologies), and processed further as described above. Image
Studio software (LiCOR) was used for quantification and
further analysis was performed in Excel and GraphPad Prism.
For detection of Flag, the buffers and washes were Tris
and milk-based as described elsewhere (30). The amount
of unpurified protein sample loaded was 30 l while the
amount of purified protein sample loaded was 20 l. The
primary rabbit -Flag antibody was diluted 1:1000 (Cell
Signalling, Danvers, MA) and the secondary antibody was
goat anti-rabbit HRP diluted 1:10 000 (Bio-Rad, Hercules,
CA). The membrane was developed with SuperSignal West
Pico Chemiluminescent Substrate (Thermo Pierce) and
imaged on a ChemiDoc XRS+ (Bio-Rad). The membrane
was stained with Ponceau S Staining Solution (Tocris
Biosciences, Bristol, UK).
Gene delivery in vivo, live animal imaging, and liver
Plasmid DNA was prepared by endo-free maxiprep
(Qiagen). For long-term luciferase imaging, female FVB
8week-old mice obtained from Baylor College of Medicine
(Houston, TX) received hydrodynamic tail vein injections
of TransIT-QR hydrodynamic delivery solution (Mirus Bio,
Madison, WI) containing 25 g of transposon DNA with
or without transposase DNA (2.5 g for pCMV-TcBuster
or 5 g for pCMV-piggyBac). Mice were imaged
periodically under isoflurane on an IVIS machine (Perkin Elmer,
Waltham, MA) as previously described (25). Mice that were
sacrificed at 24 h were 4-month-old male FVB mice
obtained from Charles River. Hydrodynamic tail vein
injections contained 10 g of pTcBCAGLuc and either 2.5 g
or 25 g of pCMV-HA-TcBuster (n = 3 per group) in 2
ml of TransIT-QR solution. Luciferase positive mice (n =
2 per group) were sacrificed 24 h post-injection and their
livers fixed in 10% formalin at 4◦C overnight. Samples were
stained for the HA antigen as previously described (29). The
Institutional Care and Use Committees of Baylor College
of Medicine and Vanderbilt University approved all animal
experiments. Error bars represent the standard error of the
TcBuster transposase localization in human cells
We initially sought to evaluate the subcellular localization
of TcBuster transposase in human cells to determine if the
transposase was able to localize to the nucleus. We
constructed a plasmid to add the hemagglutinin (HA) tag to
the N-terminus of the transposase, pCMV-HA-TcBuster, to
permit detection of the protein with antibodies to the tag.
We tested the relative activity of HA-TcBuster transposase
by drug-resistant colony assay to measure the integration
efficiency of a transposon carrying the neomycin-resistance
gene (pTcBNeo) and found it to be indistinguishable from
untagged TcBuster transposase (Figure 1A). By
immunofluorescent staining of transfected HEK-293 cells, we
observed rod-shaped structures (rodlets) that colocalized with
the 4 ,6-diamidino-2-phenylindole (DAPI)-stained genomic
DNA (Figure 1B). The vast majority of expressing cells have
several rodlets between 1–5 m in length with their
localization limited to the nuclei. Rarely, cells could be found
that were overexpressing the protein with bright inclusions
throughout the cytoplasm (Figure 1C). To better visualize
the nuclear rodlet structures, we used deconvolution
microscopy to remove the light scatter emanating from the
fluorescent particles. The rodlets were shown by this method
to be thin and filamentous (Figure 1D and E). To confirm
the localization observed by immunofluorescent imaging,
we also obtained cytoplasmic and nuclear protein fractions
along with the insoluble pellet from cells transfected with
pCMV-HA-TcBuster and subjected these fractions to
immunoblotting. We found that the TcBuster transposase was
not detectable in the cytoplasm but was present in the
nuclear fraction with a substantial portion remaining in the
insoluble pellet (Figure 1F). We concluded that TcBuster
transposase formed nuclear rodlet protein structures in the
The number of rodlets increases with transposase plasmid
We previously found a non-linear relationship between
transposase dose and colony count when the transposon
amount was held constant (11). With increasing
transposase dose at low concentrations, the number of colonies
increased steadily. When 100 ng of transposase was
transfected, the maximal number of colonies was reached and
thereafter the number of colonies decreased with a 500 ng
dose of transposase producing approximately half the
number of colonies as the 100 ng dose (11). To determine the
number of rodlets present at various doses, we repeated the
transfections done for the colony assay (11) with
pCMVHA-TcBuster on HEK-293 cells seeded onto coverslips. The
total number of nuclei, total number of rodlets and the
number of nuclei with rodlets were counted blindly (Figure 2).
Representative images of cells given 0 ng (Figure 2A), 50 ng
(Figure 2B), 100 ng (Figure 2C) and 500 ng (Figure 2D)
of transposase plasmid along with 500 ng of transposon
plasmid are shown. The number of rodlets inside of
positive nuclei increased modestly from 1–2 to 3–4 with
increasing dosage (Figure 2E). The percentage of nuclei that had
rodlets (Figure 2F) and the total number of rodlets per
nuclei (Figure 2G) increased linearly. For instance, at the 100
ng transposase dose, peak transpositional activity declined
after transposase concentrations became high enough for
visible transposase rodlets to appear in >5% of cells (Figure
2F) (11). Therefore, we found that the presence of rodlets
coincided with the inhibition of activity at high transposase
doses. These data suggest that the negative dosage effect was
related to the presence of the rodlets.
The dynamics of rodlet formation
We created a N-terminally Flag-tagged version of TcBuster
transposase (pCMV-Flag-TcBuster) and found that it
retained activity when compared to untagged TcBuster in a
colony assay (Figure 3A). If the rodlets were formed slowly
over time, we would expect a ’seeding’ to take place
during which the rodlet would form from a static starting
location, with polymerization continuing out from this initial
seed either from one or both ends. To test this hypothesis,
we transfected HEK-293 cells with either first the HA, then
the Flag-tagged transposase, or vice versa, 6 h apart and
stained the following day (Figure 3B and C). We found cells
with rodlets that stained for only HA (red rodlets, Figure
3B and C) or Flag (green rodlets, Figure 3B and C),
presumably because they were transfected in only one of the
two transfections. Many cells expressed both HA and Flag
at varying ratios (yellow rodlets, Figure 3B and C), but all
of the yellow rodlets examined appeared at this resolution
to co-stain throughout the rodlet for both HA and Flag
(inset, Figure 3B and C), suggesting that the rodlets may not be
slowly formed from a starting seed but rather are a dynamic
structure that is flexible in composition.
In order to examine how rodlets form in real time, we
created a series of GFP fusions to TcBuster. All of the GFP
fusions changed the rodlet phenotype from a rodlet to diffuse
speckled inclusions, as shown for pCMV-TcBuster-GFP in
Figure 4A (GFP images for each fusion are shown in the
supplementary data, Figure S1A). By introducing untagged
or HA-tagged TcBuster transposase along with the
GFPtagged transposase, we were able to ’lace’ the rodlets with
GFP to permit live cell imaging. GFP-positive cells
exhibited a spectrum of phenotypes ranging from cells having
speckles without rodlets to cells with rodlets
indistinguishable from those seen when HA or Flag tags were used to
detect them. This spectrum likely represents the many
random ratios of plasmids that could occur during a lipid
transfection of both the GFP-tagged and untagged transposase.
The average cell exhibited an intermediate phenotype where
the rodlets were shortened and increased in number (Figure
4B and C).
Although still slightly above the background
pCMVGFP, GFP fusion to either the N- or C-terminus severely
blunted the activity of TcBuster transposase as measured
by colony assay (Figure 4D). It should be noted that since
the pCMV-GFP helper plasmid vector contains a neomycin
resistance cassette and some will randomly integrate, more
colonies were found when using pCMV-GFP alone even
without transposase present. To increase the activity of
the protein, we introduced a linker region that we have
previously used to retain transposase activity when fused
to DNA-binding domains (15,26). We found that the
Nterminal pCMV-GFP-LK-TcBuster was similarly active to
untagged TcBuster (Figure 4D), so we used this construct
to study GFP-labeled rodlets by live imaging (Figure S1B).
The GFP-laced rodlets were generally confined to the
nucleus, colocalizing with H2B-mCherry fusion protein (31)
to define the nucleus (Figure S1C).
We proceeded to carry out time-lapse live cell DIC and
fluorescent confocal imaging at a 1:1 ratio of
pCMVTcBuster to pCMV-GFP-LK-TcBuster in HEK-293 cells.
Snapshots beginning at 32 h post-transfection are shown in
Figure 4E–I in 30-min increments. In Figure 4E, a HEK-293
cell is shown containing one rodlet in the upper right
portion of the cell and one round inclusion at the bottom left. In
Figure 4F, the upper rodlet changed position by ∼90◦ and
the lower round inclusion split off into three short rodlets. In
Figure 4G, the middle rodlet elongated whilst the lower two
remained in close proximity. In Figure 4H, the two lowest
inclusions reformed a larger, brighter round inclusion and
the middle rodlet faded, and was hardly visible in Figure
4I. Therefore, the transposase rodlets were created in a fast,
dynamic process involving fission and fusion.
Ex vivo formation of TcBuster rodlets
When we first found them, we wondered if the rodlets were
labeling a pre-existing nuclear structure. Slowly we began to
acquire mounting evidence that rodlets were generated by
the transposase alone. In support of this hypothesis are the
following points: (i) the TcBuster rodlets are unique
structures that only look similar to those filamentous structures
others have observed for Ac transposase (32); (ii) we have
found the TcBuster rodlets in animal cells while Ac rodlets
were observed in plant cells (32), which are different
kingdoms of life; (iii) the rodlet length was decreased
dramatically through fusion to GFP (Figure 4A); (iv) the rodlet
length could be manipulated in a linear fashion depending
on the ratio of GFP fusion protein to untagged protein
provided to the cells (Figure 4B) and (v) transposon DNA was
not required for rodlets to form.
To further test the hypothesis that the rodlet structures
are formed from TcBuster protein rather than the
transposase protein being drawn to a pre-existing cellular
structure, we purified Flag-TcBuster from HEK-293 cells that
were overexpressing the protein at 48 h post-transfection.
The Ponceau Red staining for nonspecific protein revealed
many more bands in the Input lane and none in the
Purified protein lane as expected (Figure 5A). A band at the
expected size (75.8 kDa) was weakly present in the Input
lane after a long exposure and gamma manipulation of the
image; this band was enriched in the purified protein lane
(Figure 5A). A smaller 50 kDa band present in the purified
protein lane was not present in the input or in other samples
analyzed (Figures 1 and 7), therefore, we conclude the band
is either an endogenous protein that cross-reacted with the
Flag epitope or a cleavage product generated during protein
To observe rodlet formation ex vivo we followed
microtubule polymerization protocols (28). Chamber slides were
prepared from cleaned coverslips and a solution of ∼1/3
protein in a PEM-like buffer was flowed into the chamber.
After incubation at 37◦C, rodlets formed both in the
absence (Figure 5B) and presence (Figure 5C) of transposon
DNA. The rodlets were thin (average diameter 0.86 m;
95% confidence interval of 0.81–0.99 m) and measured
several microns in length, indistinguishable from those that
appear in cells. The rodlets in the presence of DNA
(Figure 5C) were somewhat thicker than those without (Figure
5B). Additionally there were small ring-like structures
visible only in preparations that included DNA (Figure 5C). In
the absence of DNA, the ends of the rodlets appear
somewhat frayed (Figure 5B), an effect that we also saw in cells
more frequently for Flag-TcBuster rodlets than the
HATcBuster rodlets (Figure 3).
TcBuster activity and localization in mouse liver in vivo
To test the utility of TcBuster for liver-directed gene
transfer, we compared TcBuster to piggyBac for long-term gene
expression after hydrodynamic injection of mice. First,
we created pTcBCAGLuc, a plasmid carrying TcBuster
IRs and the same luciferase expression cassette as
pTpBCAGLuc (25). The luciferase expression cassette consists
of the CAGGS chicken -actin enhancer with a minimal
CMV promoter driving expression of the firefly luciferase
protein. The transposon alone or transposon plus
transposase for piggyBac or TcBuster were introduced into the
mouse liver by hydrodynamic tail vein injection, a
commonly used method to transfect mouse hepatocytes in vivo
(33,34). Luciferase expression was measured in reflective
light units by live animal imaging over a 168-day
timecourse (Figure 6A). The difference in the groups was
statistically significant by one-way analysis of variance. To better
compare the two systems, the last three time-points (Day
112, Day 140, and Day 168) representing stable long-term
gene expression were normalized to Day 1 to account for
differences in transfection efficiency between groups and
the promoter activity within the 5 inverted repeat of
piggyBac (35)(Figure 6B). For both systems, the mice given
transposase had higher levels of gene expression at the later
timepoints than mice receiving transposon alone (piggyBac,
P < 0.01; TcBuster, P < 0.0005). However, no significant
difference was found between the TcBuster and piggyBac
groups. The mice were monitored for gross evidence of
cancer or other illness and remained healthy until sacrifice. We
used PCR to confirm that TcBuster excision occurred in the
mouse liver in vivo. Three mice were given hydrodynamic
injection and plasmid DNA was purified from their livers
the following day and used as a template for excision PCR.
Excision PCR products of the correct size were sequenced
and confirmed TcBuster activity in vivo (Figure S2).
TcBuster transposition in mouse liver successfully produced
long-term gene expression in vivo.
To assess if rodlet formation is also found upon
transposase introduction to a living organism, we performed
hydrodynamic tail vein injection of the pCMV-HA-TcBuster
plasmid in low (2.5 g) and high (25 g) doses along with
the 10 g of the pTcBCAGLuc plasmid to monitor injection
efficacy by live animal luciferase imaging.
Immunofluorescent staining for the HA-TcBuster protein was performed
on 24-h samples of liver obtained from mice that received
a successful injection. No positive cells were present on the
IgG control (Figure 6C). Only a few positive cells could be
found in sections from the mice receiving 2.5 g of
pCMVHA-TcBuster (Figure 6D). Other mice received 10 times
this dose, 25 g, which is a standard hydrodynamic dose for
the transgene to be integrated but high for the transposase
DNA. Sections from these mice were abundant in positive
cells, many of which contained rodlets. However, some cells
had cytoplasmic inclusions only, nuclear and cytoplasmic
inclusions, or more diffuse staining patterns without
obvious inclusions (Figure 6E). Confocal microscopy of
hepatocyte nuclei (Figure 6F) revealed a similar rodlet pattern to
that observed in transfected tissue culture cells (Figure 1D).
These results demonstrate that the formation of TcBuster
transposase into rodlets occurred in somatic tissues and that
overall the properties of TcBuster transposase rodlets were
consistent regardless of the setting (cell-free, tissue culture
cells or live mice).
Transposition is complete when transposase rodlets appear
To examine protein expression over time, HEK-293 cells
were transfected with a low amount of either transposase
and cells were harvested at 0, 4, 8, 12, 16, 24 or 48
h post-transfection. Immunoblotting to detect the
HAtagged transposases revealed a greater overall RIPA-soluble
protein amount for HA-piggyBac than for HA-TcBuster
(Figure 7A). Protein began to be detectable by western blot
at 12 h for HA-piggyBac and 16 h for HA-TcBuster, with the
transposase protein abundance increasing greatly between
24 and 48 h post-transfection for both (Figure 7B, right
We have previously compared the activity of the TcBuster
and piggyBac systems and found that at low doses of
transposase plasmid the systems appear comparable in activity
whereas at high transposase doses piggyBac is able to yield
a higher number of drug-resistant colonies (11). We
transfected HEK-293 cells with 100 ng of the pCMV-piggyBac
or pCMV-TcBuster transposase plasmids. After 0, 4, 8, 12,
16, 24 or 48 h, cells were transfected a second time with 1 g
of the matching transposon carrying a neomycin-resistance
cassette to produce drug-resistant colonies. We found that
the cells are transposition-competent for a much shorter
time for TcBuster than for piggyBac (Figure 7B, left y-axis).
For TcBuster, transposition was greatly attenuated between
8 and 12 h following transfection of the pCMV-TcBuster
plasmid, with transposition completely ceased at 24 h
(Figure 7B). For piggyBac, transposition dropped sharply
between 12 and 16 h following transfection with
pCMVpiggyBac plasmid, with transposition over by 48 h after
transfection (Figure 7B). Therefore, one reason that
piggyBac is more active than TcBuster simply because it retains
the ability to transpose for a longer time.
To examine the localization of the transposases at the
same timepoints and doses as the colony assay in Figure
7A, HEK-293 cells were transfected with 100 ng of
either pCMV-HA-piggyBac or pCMV-HA-TcBuster.
Interestingly, the number of positive cells at each timepoint was
approximately equal for either transposase. No positive cells
were found at 0, 4, 8 or 12 h (data not shown; Figure 7C),
and very few positive cells were seen at 16 or 24 h
(Figure 7C). At 48 h, the number of positive cells was higher
with many cells having bright positive staining, consistent
with the immunoblots (Figure 7A and B). While the
number of positive cells was approximately the same for the
different transposases, localization differed considerably. At
24 h, the HA-piggyBac samples had diffuse nuclear
staining while at 48 h there were a variety of localization
phenotypes present including cytoplasmic, cytoplasmic and
nuclear, or inclusions (white arrows) in the nucleus.
Additionally, at 48 h most cells had the characteristic
HATcBuster rodlets (white arrows), but some had cytoplasmic
non-nuclear staining. In both cases there was a diversity of
localization phenotypes present at 48 h that we did not
observe at 24 h, when all HA-piggyBac-expressing cells had
diffuse nuclear staining and all HA-TcBuster cells had
nuclear rodlets. Therefore, we conclude that the end of
active transposition is signaled by the presence of structures
formed by the transposase.
We found that TcBuster localized to long nuclear rodlet
structures. Rodlet formation was not affected by staining
parameters such as the fixation method (formaldehyde
versus methanol, data not shown), cell type (HEK-293 (Figure
1B) versus HeLa (Figure 1D), or the presence of
transposon plasmid DNA (absent, Figure 1B versus present, Figure
2D). Similarities between Hermes and TcBuster transposase
domains and amino acid sequences (9) might suggest that
TcBuster forms similar octamers to Hermes (8), but at this
point we have no information as to whether the active
octameric structures are present within the rodlet structures
as subunits or if the protein forms another conformation in
the rodlets. If anything, the inhibitory aspect of the rodlets
suggests the later. The published octameric ring structure of
Hermes transposase is only 20 nm in diameter (8), while the
TcBuster rodlets are much larger structures (∼860 nm
diameter by ∼5000 nm in length) that, for reference, are similar
in size to the primary cilia. Herein, we could not detect signs
of a seed and growth mechanism (Figures 3 and 4). Rodlet
formation was disrupted by addition of the GFP sequence,
and the rodlet phenotype was partially rescued when
untagged TcBuster transposase was introduced to create
GFPlaced rodlets of approximately half the length. This suggests
that rodlets are formed by a polymeric mechanism of some
sort that depends on the strength of protein-protein
interactions between transposase molecules: in the case of
fusion proteins, such as to GFP, these interactions are
weakened, resulting in shorter rodlets. Rodlets branched off from
a large inclusion (Figure 4E and F) and also combined to
become one large inclusion (Figure 4G and H). Therefore,
the TcBuster transposase rodlets exhibit a propensity for
fission and fusion. Rodlets form without host cofactors, a
conclusion supported by the cross-species nature of rodlet
formation (Figures 6 and 7). By the time transposase rodlets
were detectable by immunofluorescence in some of the cells,
transposition was essentially terminated (Figure 7).
We first observed similarities between TcBuster and the
classic hAT element Ac in the negative dose response that
they exhibit (11,36). Now, further parallels between
TcBuster and Ac transposase can be drawn with regard to the
unique filamentous structures to which the transposase
protein is localized. Similar rodlets were found in Ac
localization experiments from synthetic constructs that were carried
out in a number of diverse species: insect (37), petunia (32),
maize (32,38), tobacco (22) and zebrafish (39). The
recombinant transposases can be difficult to purify due to protein
insolubility (40–42). It is possible that smaller, precursor
aggregated forms are inhibitory rather than the larger
aggregates we visualized; however, such hypothetical structures
are beyond detection with the techniques we utilized. A
detailed review of the association between transposase
concentration and activity for different transposons has been
published recently in which there are several transposases
(Mos1, POKEY, mPB, SB10 and HSB16) that when
overexpressed as GFP fusion proteins (12) exhibited punctate
staining similar to our TcBuster GFP fusions (Figure 4,
Figure S1A). It is possible that these punctate structures may,
like TcBuster, be modulated in shape when untagged
transposases are introduced to ’lace’ the rodlets with GFP rather
than expressing only the GFP fusions.
Yeast two-hybrid studies on truncated and mutated forms
of Ac transposase identified the hAT ’C-terminal
dimerization domain (38)’. This highly conserved region of the hAT
transposases is now known from the Hermes crystal
structure to be critical for maintaining the overall structure of
the transposase but, surprisingly, was not directly involved
in the dimerization interfaces of the transpososome in
octameric form (8,9,38). One dominant negative mutant in
a highly conserved residue within this region of Ac
transposase, R733A, formed aggregates that resembled
hexagonal crystals rather than filamentous rodlets (38).
Therefore, it is possible that the ’dimerization domain’ could play
a role in the oligomerization that may occur in the
context of hAT transposase rodlet formation, rather than in
the direct transposase–transposase interfaces of the active
complex. For Ac transposase, filamentous aggregation is
thought to limit transposition and has been proposed as a
protective mechanism (32). Scofield et al. put forward the
hypothesis that Ac transposase accumulates during a phase
in which transposition is encouraged by monomers that
bind to the transposon to form the DNA-protein
transpososome, but eventually protein levels exceed some threshold,
at which time the transposition reaction becomes inhibited
by protein-protein interactions (22). However, this
hypothesis does not address the complete cessation of transposition
that we observed (Figure 7).
The termination of transposition after rodlets appear
suggests a window or timer effect. This timer begins when
transposase is made in undetectable amounts in the cells
and ends when an “off switch” is flipped by transposase
aggregation. It is striking how complete the mechanism is
(Figure 7) and that the length of the timer correlated
perfectly with well-established overall levels of activity since
piggyBac is more active than TcBuster (11,27) and on a
longer timer (Figure 7). We found an inverse relationship
between colonies formed and the transposase concentration
in the cell at any given timepoint for both transposons
(Figure 7). For TcBuster, transposition was essentially over by
the time the protein was detectable by western blot while
for piggyBac, detectable protein expression overlapped with
colony formation at the 12, 16 and 24 h timepoints. We
previously analyzed piggyBac protein expression over the week
following transfection and found protein expression slowly
dropped after the 48-h peak (43). Different transposases are
expected to reach this ’point of no return’ earlier or later due
to their solubility properties, providing an important factor
by which we may be able to increase transposase activity
in our favor for gene transfer or mutagenesis. Higher
activity transposase mutants or fusions could be identified with
an extended active window that is achieved through greater
solubility rather than changing the transposase-DNA
As with other filamentous proteins, purification of
TcBuster monomers together with advanced imaging
techniques may allow us to further elucidate the mechanism
of rodlet formation. TcBuster transposase expression in the
red flour beetle should be studied to understand if rodlets
form and attenuate transposition in the native setting.
Fusion of other domains to the N-terminus of TcBuster
transposase with a linker may allow the enzyme to maintain
activity while directing it with a DNA-binding domain to
direct transposition to particular sites in the genome. Finally,
we hope to find TcBuster mutants in which rodlet
formation is disrupted such that the negative dosage effect can be
mitigated to increase transposition for gene transfer
Supplementary Data are available at NAR Online.
We thank Nancy Craig, Peter Atkinson, Reinhard Kunze,
Joseph Gall, Allison Hickman and Fred Dyda for helpful
discussions regarding the TcBuster rodlets. We thank the
Baylor College of Medicine Integrated Microscopy Core
and the Vanderbilt Cell Imaging Shared Resource for use
of their microscopes. H2B-mCherry was a gift from Robert
Benezra (Addgene plasmid #20972). We thank Ruth Ann
Veach and Felisha M. Williams for their technical
National Institutes of Health [5T32DK062706 and
2T32DK060445-11]; a fellowship from Dr and Mrs Harold
Seltzman; Career Development Award from the
Department of Veterans Affairs [BX002797 to L.E.W.]; Vanderbilt
Student Research Training Program in Diabetes and
Obesity, Kidney Disease and Digestive Disease funded by
the Vanderbilt Short Term Research Training Program
for Medical Students from the National Institutes of
Health [DK007383]; Vanderbilt Diabetes Research and
Training Center [DK20593 to L.M.D.]; Baylor College of
Medicine Integrative Molecular and Biomedical Sciences
(to Y.C.L.); Baylor College of Medicine SMART PREP
program funded by the National Institutes of Health
[GM069234-2003-2014 to E.S.T.]; National Institutes
of Health [DK093660]; Department of Veterans Affairs
[BX002190]; Vanderbilt Center for Kidney Disease (to
M.H.W.). Funding for open access charge: VA and NIH
Conflict of interest statement. None declared.
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