levin moran, 2011

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Through her pioneering work in maize, Barbara McClintock was the first to realize that eukaryotic genomes are not static entities but contain transposable elements (TEs) that have the ability to move from one chromosomal location to another 1 . It now is clear that  virtually al l orga nisms ha rbour TEs that hav e amp lified in copy number over evolutionary time via DNA or RNA intermediates. On occasions, TEs have been sporadically co-opted by the host to perform crucial cellular func- tions (for example, see REFS 2–5). However, most TEs are likely to be finely tuned genomic parasites that mobilize to ensure their own survival 6–9 .The genomic revolution, coupled with new DNA sequencing technologies, now provides an unprecedented wealth of data documenting TE content and mobility in a broad array of organisms. In multicellular eukaryotes, TEs must mobilize within gametes or during early development to be transmitted to future generations. In humans, there are at least 65 documented cases of diseases resulting from de novo TE insertions, and t hese events account for approxima tely 1 in 1,000 spontaneous cases of disease 5,10 . Indeed, new genomic technologies combined with cell-culture-based experiments have demonstrated that active TEs are more prevalent in the human population than was previously appreciated 11–18 . A growing body of evidence further suggests that mammalian TE integration occurs during early development 19–21 . In addition, studies of neurogen- esis and some forms of cancer have raised the int riguing possibility that TE activity may have an impact on the biology of particular somatic cells 12,22–24 . It is likely that we have observed only the tip of the iceberg and that we are still underestimating the contribution of TE-mediated events to the inter- and intra-individual structural variation in mammalian genomes. TE mobility poses a serious challenge to host fitness. Paradoxically, TE insertions that are harmful to the host  jeopardize TE sur vival. Thus, many TE s have evolved highly specific targeting mechanisms that direct their integration to genomic ‘safe havens’ , thereby minimizing their damage to the host (for example, see REFS 25–29). Nevertheless, host genomes have evolved potent restric- tion mechanisms, such as the met hylation of TE DNA sequences and the expression of small RNAs or cytidine deaminases, to restrict TE activity in the germ line and perhaps in somatic cells (for example, see REFS 30–33). Interestingly, a growing number of examples suggest that TEs may become activated under certain environ- mental conditions, such as stress. Stress has been shown to induce TE transcription, integration or to redirect TE integration to alternative target sites 34–38 . These findings are consistent with Barbara McClintock’ s hypothesis t hat environmental challenges may induce transposition and that transposition, in turn, may create genetic diversity to overcome threats to host survival 39 . We begin this review with a brief description of the types of TEs and their modes of mobility . We then describe the latest understanding of TE integration mech- anisms and how the host defends against these attacks. Finally, we discuss exciting new research that suggests that TE mobility may affect the biology of somatic cells. From the growing understanding of target-site selec- tion to the discovery of new, active TE copies in human populatio ns, it is clear that the f ield of transposon biology continues to yield new insights about genome biology. *Section on Eukaryotic Transposable Elements, Laboratory of Gene Regulation and Development, Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20892, USA. Departments of Human Genetics and Internal Medicine and Howard Hughes Medical Institute, University of Michigan Medical School, Ann Arbor, Michigan 48109–6518, USA. e-mails: [email protected] ; [email protected] doi:10.1038/nrg3030 Dynamic interactions between transposable elements and their hosts Henry L. Levin* and John V. Moran Abstract | Transposable elements (TEs) have a unique ability to mobilize to new genomic locations, and the major advance of second-generation DNA sequencing has provided insights into the dynamic relationship between TEs and their hosts. It now is clear that TEs have adopted diverse strategies — such as specific integration sites or patterns of activity — to thrive in host environments that are replete with mechanisms, such as small RNAs or epigenetic marks, that combat TE amplification. Emerging evidence suggests that TE mobilization might sometimes benefit host genomes by enhancing genetic diversity, although TEs are also implicated in diseases such as cancer. Here, we discuss recent findings about how, where and when TEs insert in diverse organisms. REVIEWS NATURE REVIEWS | GENETICS VOLUME 12 | SEPTEMBER 2011 | 615 © 2011 Macmillan Publishers Limited. All rights reserved

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Page 1: Levin Moran, 2011

7/17/2019 Levin Moran, 2011

http://slidepdf.com/reader/full/levin-moran-2011 1/13

Through her pioneering work in maize, BarbaraMcClintock was the first to realize that eukaryoticgenomes are not static entities but contain transposableelements (TEs) that have the ability to move from onechromosomal location to another1. It now is clear that

 virtually all organisms harbour TEs that have amplifiedin copy number over evolutionary time via DNA or RNAintermediates. On occasions, TEs have been sporadicallyco-opted by the host to perform crucial cellular func-tions (for example, see REFS 2–5). However, most TEs arelikely to be finely tuned genomic parasites that mobilizeto ensure their own survival6–9.The genomic revolution,coupled with new DNA sequencing technologies, nowprovides an unprecedented wealth of data documentingTE content and mobility in a broad array of organisms.

In multicellular eukaryotes, TEs must mobilize withingametes or during early development to be transmittedto future generations. In humans, there are at least 65documented cases of diseases resulting from de novo TE

insertions, and these events account for approximately1 in 1,000 spontaneous cases of disease5,10. Indeed, newgenomic technologies combined with cell-culture-basedexperiments have demonstrated that active TEs are moreprevalent in the human population than was previouslyappreciated11–18. A growing body of evidence furthersuggests that mammalian TE integration occurs duringearly development19–21. In addition, studies of neurogen-esis and some forms of cancer have raised the intriguingpossibility that TE activity may have an impact on thebiology of particular somatic cells12,22–24. It is likely thatwe have observed only the tip of the iceberg andthat we are still underestimating the contribution of

TE-mediated events to the inter- and intra-individualstructural variation in mammalian genomes.

TE mobility poses a serious challenge to host fitness.Paradoxically, TE insertions that are harmful to the host

 jeopardize TE survival. Thus, many TEs have evolvedhighly specific targeting mechanisms that direct theirintegration to genomic ‘safe havens’, thereby minimizingtheir damage to the host (for example, see REFS 25–29).Nevertheless, host genomes have evolved potent restric-tion mechanisms, such as the methylation of TE DNAsequences and the expression of small RNAs or cytidinedeaminases, to restrict TE activity in the germ line andperhaps in somatic cells (for example, see REFS 30–33).

Interestingly, a growing number of examples suggestthat TEs may become activated under certain environ-mental conditions, such as stress. Stress has been shownto induce TE transcription, integration or to redirect TEintegration to alternative target sites34–38. These findingsare consistent with Barbara McClintock’s hypothesis that

environmental challenges may induce transposition andthat transposition, in turn, may create genetic diversityto overcome threats to host survival39.

We begin this review with a brief description ofthe types of TEs and their modes of mobility. We thendescribe the latest understanding of TE integration mech-anisms and how the host defends against these attacks.Finally, we discuss exciting new research that suggeststhat TE mobility may affect the biology of somatic cells.From the growing understanding of target-site selec-tion to the discovery of new, active TE copies in humanpopulations, it is clear that the field of transposon biologycontinues to yield new insights about genome biology.

*Section on Eukaryotic

Transposable Elements,

Laboratory of Gene

Regulation and Development,

Eunice Kennedy ShriverNational Institute of Child

Health and Human

Development, National

Institutes of Health, Bethesda,

Maryland 20892, USA.‡Departments of Human

Genetics and Internal

Medicine and Howard

Hughes Medical Institute,

University of Michigan

Medical School, Ann Arbor,

Michigan 48109–6518, USA.

e-mails: [email protected] ;

[email protected]

doi:10.1038/nrg3030

Dynamic interactions betweentransposable elements and their hostsHenry L. Levin* and John V. Moran‡

Abstract | Transposable elements (TEs) have a unique ability to mobilize to new genomic

locations, and the major advance of second-generation DNA sequencing has provided

insights into the dynamic relationship between TEs and their hosts. It now is clear that TEs

have adopted diverse strategies — such as specific integration sites or patterns of activity

— to thrive in host environments that are replete with mechanisms, such as small RNAs orepigenetic marks, that combat TE amplification. Emerging evidence suggests that TE

mobilization might sometimes benefit host genomes by enhancing genetic diversity,

although TEs are also implicated in diseases such as cancer. Here, we discuss recent

findings about how, where and when TEs insert in diverse organisms.

R E V I E W S

NATURE REVIEWS | GENETICS  VOLUME 12 | SEPTEMBER 2011 | 615

© 2011 Macmillan Publishers Limited. All rights reserved

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Long terminal repeat

(LTR). A terminal repeated

sequence present at the

ends of LTR retrotransposons.

The LTR contains cis-acting

sequences that allow

the transcription and

polyadenylation of

retrotransposon mRNA.

LTRs also have crucial roles in

the reverse transcription of

LTR retrotransposon mRNA.

Short interspersed elements

(SINEs). A family of

non-autonomous

retrotransposons that require

functional proteins encoded

by long interspersed elements

(LINEs) to mediate their

retrotransposition.

hAT elements

A family of transposons named

after the hobo, Activator  

and Tam3 elements.

LINE-1

(L1). An abundant family of

autonomous, long interspersed

element (LINE) non-LTR

retrotransposons in

mammalian genomes. In

humans, L1 elements comprise

~17% of genomic DNA.

The vast majority of L1s

are inactive; however, it is

estimated that an average

human genome contains

~80–100 active elements.

Long interspersed elements

(LINEs). A family of

autonomous non-long

terminal repeat (non-LTR)

retrotransposons that mobilizeby retrotransposition.

 Alu

An abundant class of short

interspersed elements (SINEs)

that comprise ~10% of human

genomic DNA. Alu elements

require the endonuclease and

reverse transcriptase activities

contained within the long

interspersed element 1 (L1)

ORF2-encoded protein to

mediate their mobility. Some

 Alu elements remain active in

the human genome.

The diversity and abundance of transposons

 Mobilization mechanisms. TEs mobilize by remarkablydiverse replication strategies40 (FIG. 1; TABLE 1). ManyDNA transposons mobilize by a non-replicative ‘cutand paste’ mechanism, whereby an element-encodedenzyme, the transposase, recognizes sequences at ornear TE inverted terminal repeats to cut the TE from itsexisting genomic location and then to paste the excisedDNA into a new genomic location41 (FIG. 1a).

Retrotransposons mobilize by the reverse transcrip-tion of an RNA intermediate; however, different typesof retrotransposons carry out this process by distinctmechanisms. Long terminal repeat (LTR) retrotranspo-sons42–44 (FIG. 1b) and non-LTR retrotransposons45 (FIG. 1c) use element-encoded enzymes to mediate their mobility.In addition, the endonuclease and reverse transcriptaseactivities of non-LTR retrotransposons also have a cen-tral role in mobilizing non-autonomous short interspersed

elements  (SINEs)46–48, certain classes of non-codingRNAs49–52 and messenger RNAs, the latter of whichresults in the formation of processed pseudogenes53,54.

Transposon activity across species. Examples of DNATEs include Tn5 and Tn7  of Escherichia coli 55,56, P  ele-ments of Drosophila melanogaster 57, and Tc1 elements ofCaenorhabditis elegans58. Although they thrive in bacteriaand simpler eukaryotes, DNA TE activity appears to beextinct in most mammals, which has fuelled specula-tion that DNA TEs have a limited role in the ongoingevolution of mammalian genomes59. However, recentstudies have suggested that DNA TEs, including hAT

elements and helitrons, are active in certain bat species60–62.Thus, these studies highlight how new DNA sequenc-ing technologies can facilitate fundamental discoveriesabout the impact of different TE families on genomeevolution and serve as a cautionary note against derivinggeneral conclusions regarding TE activity from relativelyfew ‘reference’ sequences.

LTR retrotransposons are particularly abundantin eukaryotes. For example, D. melanogaster  containsapproximately 20 distinct families of LTR retrotranspo-sons that comprise ~1% of the genome63, whereas maizecontains ~400 families of LTR retrotransposons thatcomprise ~75% of the genome64,65. In addition, the mousegenome contains multiple active LTR-retrotransposonfamilies. Indeed, the ongoing retrotransposition ofboth autonomous LTR retrotransposons and their non-autonomous derivatives is estimated to account for

approximately 10–12% of sporadic mutations in themouse66. By comparison, there seems to be little LTRretrotransposon activity in human genomes59; however,a small number of human endogenous retroviruses arepolymorphic with respect to their presence or absenceat a given genomic location, suggesting that they haveretrotransposed (integrated into new locations) relativelyrecently in human evolution67.

Non-LTR retrotransposons are widespread amongeukaryotes, but have been especially prolific in mam-malian genomes. For example, LINE-1 (long interspersed

element 1; abbreviated here to L1) and the non-autonomousSINEs that they mobilize (for example ,  Alu  and

SINE-R–VNTR– Alu elements (SVA elements))47,48 compriseapproximately 30% of human genomic DNA sequence59.Recent research has used a combination of transposondisplay 68,69, second-generation DNA sequencing12,15–17 andanalyses of genomic DNA sequences from the HumanStructural Variation project13,70–72, the 1000 GenomesProject18,73,74 and clinical cohorts14. This research hasrevealed that L1 presence/absence dimorphisms, aswell as non-allelic recombination between L1 and Alu elements, account for an appreciable proportion of theinter-individual structural variation that is observedamong humans and continue to have a profound effect onthe human genome (see REF. 5 for a detailed review).

The diverse patterns of integration sites

Transposons exhibit a remarkable diversity of integra-tion behaviours. Some TEs preferentially integrate intogene-dense regions of the genome, whereas others targetregions such as heterochromatin, telomeres or ribosomalDNA arrays, and some seem to insert throughout thegenome. Below, we describe several examples of TE inte-

gration and what is known about how TEs target specificsites in genomic DNA.

Integration into gene-rich regions. Many TEs integrateinto gene-rich regions, although they use mechanismsthat prevent the disruption of ORFs. An extreme exam-ple is the E. coli Tn7  DNA TE. Tn7  encodes the sequence-specific DNA-binding protein TnsD, which mediatesTn7  integration into a specific position in the host chro-mosome, termed attnTn7 , and thereby avoids damagingthe host genome75,76. A second targeting protein, TnsE,can alter Tn7  target preference by directing integrationto plasmids that are transferred between E. coli by conju-gation77 or to dsDNA breaks and DNA structures formedduring DNA replication78. By comparison, the D. mela-nogaster  P  element avoids disrupting ORFs by integrat-ing within 500 bp upstream of transcription start sites ofgenes79. However, the mechanism by which P  elementstarget these sites requires elucidation.

Certain non-LTR retrotransposons encode endonu-cleases that target specific sites in genomic DNA. Forexample, the R1 and R2 elements of insects encodesequence-specific endonucleases that cleave at specificpositions within the 28S rDNA locus to initiate target-

site-primed reverse transcription (TPRT)28,45. However,these endonucleases operate by distinct mechanisms.R1 encodes an endonuclease that shares sequence simi-

larity with an apurinic or apyrimidinic site DNA repairendonuclease (APE)80,81, whereas R2 encodes a type IISrestriction endonuclease82. Thus, these elements appar-ently have evolved convergent mechanisms to integrateinto ribosomal DNA arrays.

LTR retrotransposons also have evolved strategies tointegrate into gene-rich regions while ensuring minimaldamage to their hosts. For example, the Ty1 and Ty3 ret-rotransposons of Saccharomyces cerevisiae specificallytarget gene-free windows that are located immediatelyupstream of RNA polymerase III-transcribed genes,such as tRNAs25,27,83,84; Ty3 is directed to integration sites2–3 bp upstream of such genes by the transcription factor

R E V I E W S

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′ 

′ 

        

    

      

      

 

Figure 1 | The diverse mechanisms of transposon mobilization. a  | DNA transposons. Many DNA transposons are

flanked by terminal inverted repeats (TIRs; black arrows), encode a transposase (purple circles), and mobilize by a

‘cut and paste’ mechanism (represented by the scissors). The transposase binds at or near the TIRs, excises the

transposon from its existing genomic location (light grey bar) and pastes it into a new genomic location (dark grey

bar). The cleavages of the two strands at the target site are staggered, resulting in a target-site duplication (TSD)

typically of 4–8 bp (short horizontal black lines flanking the transposable element (TE)) as specified by the

transposase. Retrotransposons (b and c) mobilize by replicative mechanisms that require the reverse transcription

of an RNA intermediate. b | LTR retrotransposons contain two long terminal repeats (LTRs; black arrows) and

encode Gag, protease, reverse transcriptase and integrase activities, all of which are crucial for retrotransposition.

The 5′ LTR contains a promoter that is recognized by the host RNA polymerase II and produces the mRNA of the

TE (the start-site of transcription is indicated by the right-angled arrow). In the first step of the reaction,

Gag proteins (small pink circles) assemble into virus-like particles that contain TE mRNA (light blue), reversetranscriptase (orange shape) and integrase. The reverse transcriptase copies the TE mRNA into a full-length dsDNA.

In the second step, integrase (purple circles) inserts the cDNA (shown by the wide, dark blue arc) into the new

target site. Similarly to the transposases of DNA transposons, retrotransposon integrases create staggered cuts at

the target sites, resulting in TSDs. c | Non-LTR retrotransposons lack LTRs and encode either one or two ORFs. As for

LTR retrotransposons, the transcription of non-LTR retrotransposons generates a ful l-length mRNA (wavy, light blue

line). However, these elements mobilize by target-site-primed reverse transcription (TPRT). In this mechanism, an

element-encoded endonuclease generates a single-stranded ‘nick’ in the genomic DNA, liberating a 3 ′-OH that is

used to prime reverse transcription of the RNA. The proteins that are encoded by autonomous non-LTR

retrotransposons can also mobilize non-autonomous retrotransposon RNAs, as well as other cellular RNAs (see the

main text). The TPRT mechanism of a long interspersed element 1 (L1) is depicted in the figure; the new element

(dark blue rectangle) is 5′ truncated and is retrotransposition-defective. Some non-LTR retrotransposons lack

poly(A) tails at their 3′ ends. The integration of non-LTR retrotransposons can lead to TSDs or small deletions

at the target site in genomic DNA. For example, L1s are generally flanked by 7–20 bp TSDs.

SINE-R–VNTR– Alu 

elements

(SVA elements). Composite,

non-autonomous

retrotransposons that also

require long interspersed

element 1 (L1)-encoded

proteins to mediate their

mobility. SVA elements are less

abundant than Alu elements,

and certain families of SVA

elements remain active

in the human genome.

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Table 1 | Classes of transposable elements and their mobility mechanisms

Class of TE Structural features Replication mechanism Variant forms Active examples

DNA transposons   •TIRs•Transposase

Transposase-mediatedexcision of donor dsDNAfollowed by insertion intothe target site

•Some DNA transposons also mobilize viareplicative mechanisms

• ssDNA transposons lack TIRs: donor ssDNAis inserted into target-site ssDNA, such as forIS608 ofHelicobacter pylori 

•Tn7 inEscherichia coli•P elements in Drosophila

melanogaster •Tc1 elements in

Caenorhabditis elegans

LTRretrotransposons

•LTRs•Gag, protease,

reversetranscriptase andintegrase

Within virus-like particles,reverse transcriptasecopies the mRNA of the TEinto a full-length cDNA;integrase inserts thecDNA into target sites

•Solo LTRs are commonly found in genomesand are a result of LTR–LTR recombination

•Ty1, Ty3 and Ty5 inSaccharomyces cerevisiae

•Tf1 and Tf2 inSchizosaccharomycespombe

•Tnt1 in tobacco

Non-LTRretrotransposons

•One or two ORFs•5′ truncations

and inversion/deletion (formammalianL1 elements)

•Some end in poly(A)tails (for example,L1s); others do not(for example, R2)

An element-encodedendonuclease mediatesTPRT. The endonucleasenicks the DNA at thetarget site and uses the 3′ nicked end for the primeras it reverse transcribes TEmRNA

•Non-autonomous, non-LTR retrotransposons(for example, Alu and SVA elements, aswell as other eukaryotic SINEs) rely on theendonuclease and reverse transcriptase ofan autonomous non-LTR retrotransposon tomediate retrotransposition

•The L1 retrotransposition machinery canalso mobilize mRNAs (to generate processedpseudogenes) and certain non-coding RNAs(for example, the U6 snRNA)

•L1 in human, mouse, andother mammals

• I factor in D.melanogaster 

•Zorro3 inCandida albicans

•R1 and R2 in insects

L1, long interspersed element 1; LTR, long terminal repeat; SINE, short interspersed element; snRNA, small nuclear RNA; SVA, SINE-R–VNTR– Alu; TE, transposableelement; TIR, terminal inverted repeat; TPRT, target-site-primed reverse transcription.

Target-site-primed reversetranscription

(TPRT). The mechanism of

mobility that is generally used

by long interspersed elements

(LINEs) and short interspersed

elements (SINEs). An

endonuclease, encoded by

the LINE, nicks genomic

DNA to expose a 3′-OH at

the target site that can be used

as a primer to initiate the

reverse transcription of the

retrotransposon RNA by a

LINE-encoded reverse

transcriptase.

complexes TFIIIB and TFIIIC85,86 (FIG. 2a). However, inthe case of the snR6  non-coding RNA gene, which doesnot depend on TFIIIC for its expression, the TFIIIBfactors Brf1 and TATA-box binding protein (TBP) aresufficient to direct Ty3 integration87,88.

Ty1  integrates into a ~700 bp window upstreamof tRNA genes with a periodicity of 80 bp 27,89 (FIG. 2b).Although the factors that direct Ty1 to tRNA genes remainunknown, the unusual periodicity of integration dependson the amino-terminal domain of Bdp1, which is anotherTFIIIB factor90. The ability to integrate upstream of RNApolymerase III-transcribed genes can also regulate hostand TE gene expression. For example, Ty1 and Ty3 inser-tions can stimulate the transcription of downstream RNApolymerase III-transcribed genes and transcription ofthe RNA polymerase III target genes can reciprocate byrepressing Ty1 transcription91,92. Clearly, determining howTy1 targets integration sites and exploring how integrationalters gene regulation remain areas for future study.

The ability to target RNA polymerase III-transcribedgenes is not unique to LTR retrotransposons. For exam-ple, the Dictyostelium discoideum non-LTR retrotranspo-son DRE (also known as TRE5A) preferentially inserts

~48 bp upstream of tRNA genes, whereas the retrotrans-poson Tdd3 (also known as TRE3A) inserts downstreamof tRNA genes29,93. Indeed, experimental evidence sug-gests that the TRE5A ORF1-encoded protein directlyinteracts with subunits of TFIIIB to direct its integrationto tRNA genes94.

The Schizosaccharomyces pombe retrotransposonTf1 preferentially integrates into the promoters of RNApolymerase II-transcribed genes and provides anotherexample of how TEs target gene-rich regions95–97 (FIG. 2c). Tf1 integration has been studied by examiningits integration into promoters contained within extra-chromosomal replicating plasmids26. For example, the

fructose-1,6-bisphosphatase ( fbp1) promoter is inducedwhen the activating transcription factor Atf1 binds toan 8 bp upstream activating sequence (UAS1)98. Tf1 integration generally occurs 30 bp or 40 bp downstreamof UAS1; however, mutating six nucleotides of UAS1 ordeleting the atf1 gene disrupts Tf1 integration specificity,causing integration to occur throughout the plasmid26.Although cells lacking Atf1 show little reduction in theoverall Tf1 retrotransposition frequency 99, the abovedata, as well as the finding that Atf1 forms a complexwith Tf1 integrase, indicate that specific transcriptionfactors such as Atf1 can play a crucial part in directingTf1 integration to a specific target site. Notably, experi-ments conducted with a synthetic promoter revealedthat RNA polymerase II transcription is not sufficientto target Tf1 integration99.

The recent development of second-generationsequencing technology has allowed the in vivo examina-tion of Tf1 integration sites en masse. Characterizationof 73,125 Tf1 integration events from four independ-ent experiments revealed a highly reproducible pattern:approximately 95% of integration events are clusteredupstream of ORFs96. Interestingly, the most frequently

targeted promoters are associated with genes that areinduced by environmental stresses. The targeting ofgenes that respond to stress, coupled with the abilityof Tf1 to induce the expression of adjacent genes26, sug-gests that Tf1 integration has the potential to improvethe survival of specific cells that are exposed to environ-mental stress. Likewise, the transcription of Tf2, anotherLTR retrotransposon in S. pombe, is induced by oxidativeand osmotic stress or by growth in low oxygen concen-trations34,100. Clearly, understanding the consequences ofstress-induced retrotransposition will yield insights abouthow TE mobility can lead to genetic diversity, which mayaffect the ability of an organism to cope with stress.

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Finally, certain retroviruses, which are descendedfrom LTR retrotransposons101, also exhibit preferentialintegration in gene-rich regions. For example, HIV-1preferentially integrates into RNA polymerase II-transcribed genes, whereas murine leukaemia virusshows a strong integration preference near transcrip-tional start sites102–104. Structural and biochemicaldata demonstrate that HIV-1 integrase interacts withthe cellular lens epithelium-derived growth factor(LEDGF; also known as p75) host factor, and there isevidence that this interaction has an important role inproviral DNA integration105,106.

Integration into heterochromatin. Some TEs targetheterochromatic sequences that contain relatively fewgenes. For example, chromoviruses, which are related toTy3–Gypsy  LTR retrotransposons, reside in the hetero-chromatin of eukaryotes f rom fungi to vertebrates107.The integrase enzymes of these viruses contain achromodomain near the carboxyl terminus that isrelated to the heterochromatin protein HP1, which

binds to histone H3 methylated at lysine 9 (REF. 107).Furthermore, chromovirus integrase chromodomains

fused to GFP colocalize with heterochromatin108, whichsuggests that the chromodomain has a principal role indirecting integration. Indeed, fusion of one such chro-modomain to the C terminus of Tf1 integrase directsintegration of this TE to heterochromatin108.

The Ty5 LTR retrotransposon also targets gene-poor regions in S. cerevisiae. Approximately 90% of Ty5 integration events occur within the silent mating typeloci or near silent heterochromatin at telomeres109–111 

(FIG. 2d). Genetic and biochemical experiments indi-cate that a nine-amino-acid targeting domain in theTy5 integrase C terminus directly binds to a structuralcomponent of heterochromatin, Sir4, to target integra-tion35,112,113. Moreover, fusing Sir4 to the DNA bind-ing domain of the LexA repressor protein causes Ty5 integration to be redirected to LexA binding sites114.Thus, the Ty5 integrase targeting domain, by inter-acting with Sir4, directs integration to heterochroma-tin. Interestingly, genetic and biochemical evidenceindicate that the Ty5 targeting domain evolved itsinteraction with Sir4 by mimicking residues in a host

factor, Esc1p, that binds to the same amino acids ofSir4 (REF. 115).

Figure 2 | Mechanisms that position integration.

a | The Ty3 targeting mechanism. Integration of Ty3 

occurs one or two base pairs upstream of tRNA genes.

This pattern requires Brf1 and TATA-box binding protein

(TBP), which are components of the transcription factor

complex TFIIIB that recruits integrase (grey oval) to

the target site. b | The Ty1 targeting mechanism. The

transcription factor Bdp1 is a component of TFIIIB and is

required to recruit the chromatin remodelling complex

Isw2 (light blue semicircle). Orange cylinders indicatenucleosomes and black lines represent DNA. Integration

of Ty1 occurs with an 80 bp periodicity in a 700 bp

window upstream of tRNA genes. The periodicity

requires both Bdp1 and Isw2. c | The mechanism of Tf1 

targeting. Tf1 integrates into promoters that are

transcribed by RNA polymerase II. Transcription factors,

such as Atf1, bind to the promoter and recruit integrase

to the insertion sites. d | The mechanism of Ty5 

integration. In the absence of a stress condition, the

targeting domain of integrase is phosphorylated (P).

This directs the integration to heterochromatin by

binding Sir4 (green ovals), a structural component

of the heterochromatin. When cells are exposed to

environmental stress the integrase is dephosphorylated

and integration occurs in gene-rich regions.

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Desmoid tumours

Soft tissue tumours that can

arise in the abdomen as well

as in other parts of the body.

They are typically benign

and grow slowly.

Although the ability of Ty5 integrase to target het-erochromatin suggests that the TE dictates target-siteintegration, there are also indications that TE–hostinteractions can alter Ty5 target-site preference. Massspectroscopy revealed that phosphorylation of the inte-grase targeting domain at Ser1095 is crucial for bindingSir4 (REF. 35) and mutating Ser1095 redirected integra-tions to expressed regions of the genome. Although thehost-encoded kinase has not been identified, studiesusing a phospho-specific antibody indicate that stresses,such as nitrogen deprivation, can downregulate Ser1095phosphorylation35. Thus, stress conditions may alterthe phosphorylation state of Ty5 integrase, therebyredirecting the Ty5 integration specificity. This elegantexample provides a plausible mechanism for how stresscan alter transposon mobilization in a manner thatmight provide an advantage for the host. It remains to bedetermined whether retargeting Ty5 to gene-rich regionsbenefits Ty5 by allowing newly retrotransposed copiesto reside in permissive expression contexts, or benefitsthe host by generating genetic diversity, thus offering the

potential for the host to adapt to stress.

Integration into telomeres. Some TEs exclusively integrateat or near telomeric chromosome ends. For example, theHeTA, TART and TAHRE non-LTR retrotransposonscomprise the ends of D. melanogaster   chromosomesand probably substitute for the function of telomerasein maintaining chromosome end integrity 2,116,117. TheSART1 and TRAS1 non-LTR retrotransposons may havea similar role in Bombyx mori118. The proteins encodedby TART, TAHRE, SART1 and TRAS1 have an APE-likedomain118,119, and it is likely that the SART1 and TRAS1endonuclease proteins direct their integration into telom-eric repeats118; however, the functional role of the putativeendonuclease domains encoded by TART and TAHREremains unknown.

Excitingly, recent studies have revealed that certainretrotransposons can target telomeric sequences for inte-gration. For example, by an alternative endonuclease-independent retrotransposition mechanism, human L1retrotransposons containing missense mutations in theL1 endonuclease active site can integrate at endogenousDNA lesions and dysfunctional telomeres in Chinesehamster ovary cell lines that are deficient for p53 func-tion and for factors that are important in the non-homologous end-joining pathway of DNA repair120,121.Similarly, members of the Penelope clade of retrotrans-

posons, which encode a reverse transcriptase that lacksan obvious endonuclease domain, reside at telomeres inorganisms from four eukaryotic kingdoms122. The RNAsencoded by these terminal Penelope elements also con-tain sequences that are complementary to telomeric DNAsequences, suggesting that base pairing between the TERNA and telomeric ssDNA is crucial for integration.Interestingly, both of the above cases can be consideredas a type of RNA-mediated DNA repair that seems curi-ously similar to the mechanism used by telomerase120,122.Future studies could elucidate whether host factors arecrucial for the localization of these retrotransposonsto DNA lesions and/or chromosomal termini.

Dispersed patterns of integration. In contrast to someTEs that use elegant mechanisms for targeted integrationinto specific regions of the genome, other TEs appear tolack target-site specificity. For example, L1s and the non-autonomous elements that they mobilize are interspersedthroughout the genome59. Indeed, ~30% of engineeredhuman L1 retrotransposition events in cultured cells, anda similar proportion of recently discovered full-length,dimorphic human-specific L1s, are near or within theintrons of genes13,50,123,124. Because protein-coding genesconstitute ~40% of the human genome125,126, these find-ings suggest a lack of robust mechanisms that are used byL1s or the host to prevent L1 retrotransposition into genes.

The interspersed nature of L1 and  Alu sequencesprobably reflects the fact that the L1 endonuclease hasrelatively weak target-site specificity, preferentiallycleaving the sequence 5′-TTTT/A-3′ (and variants ofthat sequence) to initiate TPRT80,121,127,128. Interestingly,although ‘young’  Alu and L1 insertions have similarinterspersed integration patterns, cytogenetic studies andexamination of the human genome reference sequence

have revealed that evolutionarily ‘older’ L1s and Alus havedistinct genomic distributions59,129. Older L1s preferen-tially reside in gene-poor, AT-rich sequences, whereasolder Alus preferentially reside in gene-rich, GC-richregions of the genome59.

The distinct distributions of older L1s and  Alus arelikely to result from post-integration selective processesthat have operated on the genome for millions of years59.However, how these skewed distributions arose remainsa mystery. Some researchers have suggested that Alusmay possibly have an advantageous, albeit undefined,role in gene-rich regions of the genome59. Others havesuggested that L1 retrotransposition events into genicregions may exert a greater fitness cost to the host thanfor Alu insertions130. If so, negative selection would leadto the removal of detrimental L1 alleles from the popula-tion. Consistent with this hypothesis, data suggest thatevolutionarily recent human full-length L1 insertionsare detrimental to the host131,132, whereas in vitro studieshave revealed that L1s contain cis-acting sequences thatcan reduce gene expression133,134. Clearly, further studiesare needed to explain how the distributions of L1 and Alu elements have diverged over evolutionary time.

Despite their interspersed distribution, a small bodyof evidence suggests that there may be preferred, albeitrare, L1 integration sites. For example, independentL1-mediated retrotransposon insertions (that is, an

SVA element and an Alu element) at the same nucleotideposition in the Bruton agammaglobulinaemia tyrosinekinase (BTK ) gene have resulted in two sporadic cases ofX-linked agammaglobulinaemia135. Similarly, independ-ent L1 and Alu insertions associated with colorectal anddesmoid tumours, respectively, have occurred at the samenucleotide position in the adenomatous polyposis coli( APC ) gene22,135,136. Additionally, two independent Alu insertions at the same nucleotide position in the F9 genehave caused haemophilia B135,137. Thus, it would not besurprising to find that chromatin structure and acces-sibility have an impact on L1-mediated retrotransposontarget preference138.

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RNA-directed DNA

methylation

(RdDM). A pathway in which

24 nucleotide small RNAs

interact with a de novo 

methyltransferase to mediate

the methylation and

transcriptional silencing of

homologous genomic loci

in plants.

Small interfering RNAs

(siRNAs). Small (~21–24

nucleotide) RNAs that are

generated from dsRNA

‘triggers’ by Dicer-dependent

and Dicer-independent

mechanisms. They bind to

Argonaute proteins and guide

the resultant complex tocomplementary mRNAs

to mediate silencing.

PIWI-interacting RNAs

(piRNAs). A family of small

(~24–35 nucleotide) RNAs

that are processed from

piRNA precursor mRNAs. The

mature piRNAs interact with

specialized Argonaute proteins

(from the PIWI clade), to

mediate RNA silencing.

Dicer

A family of RNase III proteins

that possess an endonuclease

activity that can process

dsRNA ‘triggers’ into small

interfering RNAs (siRNAs) or

microRNAs (miRNAs).

 Argonaute proteins

Proteins that bind to small

RNAs and are the defining

component of the

RNA-induced silencing

complex (RISC); they have an

ssRNA binding domain (PAZ)

and a ribonuclease domain

(PIWI). The small RNAs guide

Argonaute proteins to target

mRNAs in order to mediate

post-transcriptionaldegradation and/or

translational silencing.

PIWI clade of proteins

A specialized class of

Argonaute proteins that

interact with PIWI-interacting

RNAs (piRNAs) to mediate

transposable element

silencing. Members include:

PIWI, Aubergine and Argonaute

3 in D. melanogaster ; MIWI1,

MIWI2 and MILI in mice, and

HIWI1, HIWI2, HIWI3 and HILI

in humans.

How the host defends against transposons

Although many TEs have evolved mechanisms to limitgenome damage, TE integration still poses a potentialthreat to the host. Thus, it is not surprising that hostorganisms have evolved many diverse mechanismsto combat TE activity. However, the host must beable to discriminate TE sequences from host genes toaccomplish this feat. Below we discuss the mechanisticstrategies used by the host to restrict TE mobilization.

DNA methylation. Cytosine methylation (to 5-methyl-cytosine) is an important DNA modification in eukary-otes with genomes >500 Mb, which include vertebrates,flowering plants and some fungi. Most cytosine meth-ylation in plants and mammals, and almost all cytosinemethylation in the fungus Neurospora crassa, occurswithin repetitive elements and is correlated with thetranscriptional repression of retrotransposons in somaticand germline cells139,140.

Experiments in mammals and plants have demon-strated that the global demethylation of genomic DNA

strongly reactivates TE transcription141–144. For exam-ple, deletion of the DNA cytosine-5-methyltransferase3-like (Dnmt3l ) gene in mice leads to the loss of de novo cytosine methylation of both LTR and non-LTR retro-transposons, reactivation of TE expression in sper-matocytes and spermatogonia and meiotic catastrophein male germ cells145. Determining whether TE mobi-lization is directly responsible for the meiotic defectsrequires further study. Moreover, recent data demon-strate that the inactivation of cytosine methylation in Arabidopsis thaliana causes a burst of retrotransposonand DNA TE activity, resulting in substantial increasesin TE copy number144. Thus, epigenetic mechanisms actto control the expression, and perhaps the mobility, of

 various TEs.Multiple lines of evidence indicate that DNA meth-

ylation inhibits TE transcription. Patterns of DNAmethylation are established during gametogenesisand are mediated by DNMT3A and the non-catalyticparalogue DNMT3L in mammals, but how TEs arerecognized as methylation substrates requires furtherstudy 146. By comparison, during plant development,small, 24 nucleotide (nt) RNAs target paralogous DNAsequences that share high levels of homology (such asTEs) for cytosine methylation. The mechanism of RNA-

directed DNA methylation (RdDM) is not fully understoodbut it appears to require the canonical RNA interference

(RNAi) machinery (see below and FIG. 3), the DNAmethyltransferase DRM2 and two plant-specific RNApolymerases, Pol IV and Pol V146.

Small RNAs inhibit TEs. Small-RNA-based mechanisms— including those involving endogenous small interfering

RNAs (endo-siRNAs) and PIWI-interacting RNAs (piRNAs))— also act to defend eukaryotic cells against TEs. Themechanisms by which these small RNAs are gener-ated and how they inhibit TEs remain an active area ofinvestigation in various model organisms. Mechanisticdetails regarding these processes can be found inmany outstanding reviews on this topic (for example, 

REFS 30,147–152); here we briefly summarize commonthemes that have emerged from the above studies.

Endo-siRNAs have the potential to inhibit TE mobil-ity through the post-transcriptional disruption of trans-poson mRNA. For example, ‘trigger’ dsRNAs can bederived from: the complementary inverted terminalrepeats in DNA transposons, structured mRNA tran-scripts or overlapping regions contained within conver-gent transcription units30,147,148,153. The resultant dsRNAscan then be processed into ~21–24 nt endo-siRNAs bymembers of the Dicer family of proteins30,154 (FIG. 3a).These endo-siRNAs are loaded onto an Argonaute protein,and the ‘passenger’ RNA strand (typically the sensestrand of a TE) is degraded. The remaining complexof an ssRNA and Argonaute is called the RNA-inducedsilencing complex (RISC); the RNA directs RISC to com-plementary sequences in target mRNAs, leading to theirpost-transcriptional degradation. Importantly, the RNAimachinery has the capability to inhibit any TE that gen-erates a dsRNA trigger that can serve as a substrate forthe RNAi machinery.

By a different mechanism, piRNAs can be generatedfrom genomic loci that encode long precursor RNAscontaining the remnants of different families of TEs151.In general, processing of these precursor RNAs leads tothe production of a mature ~24–35 nt piRNAs (FIG. 3b).A subfamily of Argonaute proteins, known as the PIWI

clade of proteins, predominantly binds to mature antisensepiRNAs and directs them to complementary sequencesin TE mRNA. An endonuclease activity that is associatedwith the PIWI protein cleaves the TE mRNA to release asense-strand piRNA, which can interact with other PIWIclade proteins. Binding of this complex to the originalpiRNA precursor RNA then re-iterates this amplificationcycle through a ‘ping-pong’ mechanism151,155. In additionto this type of mechanism that restricts TE mobility in thegerm line, recent studies suggest that specialized piRNApathways that do not operate via a ping-pong mechanismmight restrict somatic TE activity 32,155–157.

Examples of TEs that are controlled by small-RNA-based mechanisms include Tc1 transposons in C. elegans and P  elements in D. melanogaster 153,158. Also, 21 nt siR-NAs, which are derived from the Athila family of LTRretrotransposons in the vegetative nucleus of pollengrains in A. thaliana, are delivered to the sperm cells toinhibit the expression of transposons that, in principle,could mobilize in the germ line159.

Small-RNA-based mechanisms may also be crucial

for silencing mammalian L1 elements. For example,an antisense promoter located within the human L1 5′ UTR allows the production of an antisense RNA that, inprinciple, could base pair with sense-strand L1 mRNAto establish a dsRNA substrate for Dicer160. Furthermore,mouse mutants lacking the murine PIWI family proteinsMILI or MIWI2 exhibit a loss of methylation of L1 andintracisternal A particle (IAP) LTR retrotransposonDNA; this loss correlates with their transcriptional acti-

 vation in male germ cells161. Similarly, mice lacking aMILI-interacting protein, Tudor-containing protein 1,exhibit a similar loss of methylation of L1 DNA and areactivation of L1 expression162. Finally, mouse mutants

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′′

′   ′

′′

′′   ′′

′   ′

′′

     

Figure 3 | The degradation of transposon mRNA by RNAi. a | The small interfering RNA (siRNA) pathway.

dsRNA ‘triggers’ (represented by the hairpin), which are derived from the inverted terminal repeats of a DNA

transposon in the case illustrated, are processed and then cleaved into 21–24 nucleotide (nt) siRNAs by the Dicer

family of proteins (light green shape). A single-strand siRNA (short red line), complementary to the transposon mRNA,

is selectively incorporated into the Argonaute (dark green shape)-containing RNA-induced silencing complex (RISC).

The siRNA directs RISC to complementary sequences in the transposon mRNA (long red line), leading to its

post-transcriptional degradation. The figure is drawn based on concepts presented in REFS 147,148,152. b | The

PIWI-interacting RNA (piRNA) pathway. A primary piRNA transcript (wavy, blue line) generated from a piRNA cluster 

(blue rectangles) that contains sequences derived from transposable elements (TEs; dark blue rectangle) is processed

into mature 24–35 nt piRNAs (small blue line). Binding of the mature piRNA by the PIWI or Aubergine (AUB) proteins

allows it to be directed to complementary sequences in TE mRNA (red line). Endonucleolytic cleavage of the mRNA

(represented by the scissors), 10 nt from the 5′ end of the small RNA, and 3′ cleavage/processing liberates a secondary

sense-strand transposon piRNA (small red line), which associates with the Argonaute 3 (AGO3) protein. The binding of

this complex to complementary sequences in the original precursor piRNA, followed by endonucleolytic cleavage,

regenerates an antisense piRNA that can be directed to TE mRNA. This iterative cycle (known as  a ‘ping-pong’ cycle)

can lead to the destruction of transposon mRNA in the germ line. The piRNA model was redrawn based on concepts

and models presented in REFS 30,151,157. The example illustrated is for Drosophila melanogaster ; however, a similar

pathway probably operates in mammalian cells (see the main text for details).

piRNA cluster

A genomic DNA locus that

encodes PIWI-interacting RNA

(piRNA) precursor RNAs. Many

piRNA clusters contain sense

and antisense sequences that

are derived from mobile

genetic elements.

lacking the non-canonical Maelstrom protein, a compo-nent of the nuage complex (a germline-specific perinu-clear structure) that may be important for small RNAbiogenesis, exhibit derepression of L1 transcription,an increase in the levels of L1 ribonucleoprotein par-ticle intermediates in spermatids and a chromosomalsynapsis defect during male meiosis163. Together, the

above examples provide compelling evidence that small-RNA-based pathways probably control the expressionof certain TEs in the mammalian germ line.

Finally, it is noteworthy that other antisense-RNA-based mechanisms may be involved in TE silencing. Forexample, antisense transcripts from S. cerevisiae Ty1 elements reduce Ty1 integrase and reverse transcriptase

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 Aicardi–Goutieressyndrome

A rare, autosomal recessive

genetic disorder that leads to

brain dysfunction as well as

other symptoms. The early

onset form of the disease can

be caused by mutations in the

TREX1 gene and is usually fatal.

Hybrid dysgenesis

In Drosophila melanogaster ,

a sterility-inducing syndrome

that is induced by the

mobilization of P  elements

in crosses between females

lacking P  elements andmales carrying P  elements.

 Virus-like particle

(VLP). A cytoplasmic particle

that comprises long terminal

repeat (LTR) retrotransposon

mRNA, the LTR retrotranspo-

son-encoded proteins and

host factors that are required

for reverse transcription

of LTR retrotransposon mRNA.

LTR retrotransposon mRNA is

reverse transcribed into a

double-stranded cDNA

within VLPs.

protein levels by a post-translational mechanism; thisleads to the inhibition of Ty1 mobility and thus controlsTy1 copy number164. Because S. cerevisiae lacks RNAimachinery, these results suggest that genomes haveevolved other RNA-dependent strategies to 'tame' TEs.

Cytosine deaminases and DNA repair factors restrict

TEs. Proteins that are involved in nucleic-acid metabo-lism and/or DNA repair can also restrict TE mobility.For example, members of the apolipoprotein B mRNA-editing enzyme 3 (APOBEC3) family of cytidine deami-nases can restrict the retrotransposition of various retro-

 viruses and LTR and non-LTR retrotransposons33. Forretroviruses and LTR retrotransposons, APOBEC3 pro-teins generally deaminate cytidines during the first strandcDNA synthesis, which leads to either cDNA degradationor the integration of a mutated provirus. The mechanismsby which particular APOBEC3 proteins restrict non-LTRretrotransposons still require elucidation. Similarly, over-

expression of the 3′-repair exonuclease 1 (Trex1) gene,mutations in which cause Aicardi–Goutieres syndrome,can inhibit L1 and IAP retrotransposition in culturedcell assays165,166 but the mechanism of TREX1-mediatedTE repression requires elucidation.

Other mechanisms are also likely to restrict themobility of non-LTR retrotransposons. For example,the overwhelming majority of L1 elements in mamma-lian genomes are 5′ truncated and are thus essentially‘dead on arrival’ because they cannot synthesize proteinsthat are crucial for retrotransposition167. It has been pro-posed that 5′ truncation may be due to the low proces-sivity of the L1-encoded reverse transcriptase. However,

recent work on the reverse transcriptase encoded by theR2 non-LTR retrotransposon of B. mori demonstratedthat this enzyme is more processive than the reversetranscriptases encoded by retroviruses168. Alternatively,L1 5′ truncation might result if host factors cause the dis-sociation of the L1 reverse transcriptase from the nascentcDNA and/or degrade the L1 mRNA during integration.In these scenarios, to generate a full-length insertion theL1 reverse transcriptase would need to complete inte-gration before the TPRT intermediate is recognized asDNA damage by the host50,169. Indeed, proteins that areinvolved in the non-homologous end-joining pathway ofDNA repair seem to restrict the retrotranspositionof a zebrafish LINE-2 element in DT40 chicken cells170,whereas members of DNA excision repair pathway(such as the ERCC1–XPF complex) might restrict L1retrotransposition in cultured human cells171.

Finally, in addition to recognizing the L1 integrationintermediate as a form of DNA damage, recent data sug-gest that retrotransposition indicator cassettes, deliveredby engineered L1s in human embryonic carcinoma cell

lines, can be epigenetically silenced during or imme-diately after their integration into genomic DNA172.Because L1 is an ancient ‘stowaway’ in mammaliangenomes, it is likely that the host has evolved multiplemechanisms to combat L1 mobility at discrete steps inthe retrotransposition pathway and that some of thesemechanisms operate in a context-dependent manner.Continued studies should reveal new and more diversehost mechanisms to restrict TE mobility.

Developmental triggers of transposition

Despite mechanisms to combat TE mobility, TEs con-tinue to thrive in many host genomes. Thus, TEs musthave evolved ways to either overwhelm or counteractthese host defences. TEs must mobilize in germ cells orduring early development to ensure their survival (FIG. 4).However, some TEs can mobilize in somatic cells, provid-ing a potential mechanism to generate intra-individualgenetic variation.

Transposition in the germ line or during early devel-

opment. D. melanogaster P  elements provide one of thebest-studied cases of cell-type-specific transposition173.P  element transposition occurs when females lacking P  elements mate with males carrying P  elements; P  ele-ment mobilization can cause hybrid dysgenesis in theoffspring. In the reciprocal cross, eggs from females

containing P  elements produce a repressor protein andpiRNAs that inhibit P  elements transposition57,158. Therepressor is an alternatively spliced, truncated form ofthe transposase. Importantly, the repressor not onlycontrols which crosses produce germline integrationbut also inhibits transposition in the soma.

The D. melanogaster  Gypsy  element is another exam-ple of a TE that exhibits tissue-specific control174,175.Gypsy  transcription is induced in somatic follicle cellsthat surround the oocyte. The TE mRNA assemblesinto virus-like particles that are thought to traffic to theoocyte to carry out transposition. It remains unclearwhether the transfer of Gypsy  virus-like particles to the

Figure 4 | Timing of transposition. Germline transposable element (TE) integration

events can result from TE mobility in cells that give rise to gametes or from TE mobility

post-fertilization during early development. Embryonic TE mobility in cells that do not

contribute to the germ line or mobility at later developmental stages can, in principle,

lead to somatic TE integration events. The overlapping brackets signify that some TEinsertions in early development can contribute to the germ line, whereas others may

not. Endogenous long interspersed element 1 (L1) retrotransposition events can occur

in certain tumours, and experiments using engineered human L1s suggest that L1

retrotransposition may also occur during mammalian neurogenesis. Examples of the

developmental timing of TE integration events are described in the main text.

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X-linked choroideraemia

A recessive degenerative

retinal disease.

oocyte occurs via an enveloped particle (similarly toretroviruses) or by a form of endocytosis. However, theFlamenco locus encodes piRNAs that silence Gypsy  ele-ments in follicle cells, thereby preventing the spread ofthese TEs to the surrounding germ cells155.

Relatively little is known about the developmentaltiming of L1 retrotransposition in mammals. The sheernumbers of L1 and Alu retrotransposons that populatemammalian genomes provide immediate evidence thatthey mobilize in the germ line. Various studies, usingendogenous and engineered L1s, provide strong experi-mental evidence to support this assertion (also reviewedin REF. 5). For example, full-length mouse L1 RNA andthe mouse L1 ORF1-encoded protein are co-expressedin leptotene and zygotene spermatocytes during meioticprophase176. In addition, the mouse ORF1-encoded pro-tein is expressed in the cytoplasm during specific stagesof development in oocytes177. Similarly, human oocytesexpress L1 RNA and support the retrotransposition ofan engineered human L1 element178. Finally, transgenicmouse experiments demonstrated that an engineered

human L1 retrotransposon, which was driven from thepromoter of the mouse RNA polymerase II large subunitgene, could retrotranspose in male germ cells179.

Unexpectedly, a growing body of experimental evi-dence suggests that L1 retrotransposition might occurfrequently during early development (FIG. 4)  (alsoreviewed in REF. 5). For example, human embryonic stemcells can express L1 RNA and ORF1-encoded protein,and can accommodate the retrotransposition of engi-neered L1s, albeit at lower levels than in other types oftransformed human cells19,180. In addition, studies of amale patient with X-linked choroideraemia revealed thathis mother had mosaicism for the mutagenic L1 inser-tion in both germline and somatic tissues20. Thus, theinitial retrotransposition event must have occurred dur-ing early embryogenesis in the mother. Finally, recenttransgenic experiments conducted in rats and mice ledto the conclusion that most L1 retrotransposition occursduring early embryogenesis and that most of the resultantevents are not heritable21. Intriguingly, these data suggestthat L1 ribonucleoprotein particles can be deposited intozygotes by either the sperm or the egg to undergo retro-transposition during early development, thereby provid-ing a possible mechanism to generate somatic mosaicismand intra-individual genetic variation (see below).

Somatic transposition. Classical experiments in maize

revealed that DNA TE activity in somatic tissues couldlead to variegated corn colour phenotypes1,181. Sincethen, somatic TE events have been reported in otherorganisms. For example, it is well established that Tc1 transposition in the Bergerac strain of C. elegans pref-erentially occurs in somatic cells182. Similarly, a recentstudy has revealed that somatic transposition of theHatvine1-rrm DNA TE — into the promoter region ofthe VvTFL1A gene of the grapevine cultivar Carnigan— affects the grapevine branching pattern and the sizeof fruit clusters183. Also, a mutagenic L1 insertion wasidentified in the APC  gene in tumour tissue, but not inthe surrounding tissue, of a patient with colon cancer,

suggesting a role for the insertion in cancer develop-ment22. Together with the transgenic L1 experiments(discussed above), these findings establish that somaticTE mobility can lead to phenotypic changes in the host.

Intriguingly, several lines of evidence suggest thatsomatic L1 retrotransposition may also occur in themammalian nervous system (reviewed in REF. 5). First,an engineered human L1 can retrotranspose in neuro-genic zones of the brain in transgenic mice24 when itsexpression is driven by a promoter contained withinits native 5′ UTR 184. Second, engineered human L1s canretrotranspose in cultured rat neuronal progenitor cells(NPCs), in human embryonic stem cell-derived NPCsand at low levels in human fetal-derived NPCs23,24. Third,sensitive multiplex quantitative PCR experiments havesuggested a modest increase in L1 copy number inpost-mortem brain tissue when compared to heart andliver tissue that was derived from the same individual23.Finally, the retrotransposition of an engineered humanL1 is elevated in a mouse model of Rett syndrome (aneurodevelopmental disorder), and induced pluripotent

stem cells derived from Rett syndrome patients exhibitan increase in L1 DNA copy number when comparedto normal controls, suggesting a potential increase inendogenous L1 retrotransposition185.

The above studies strongly suggest that certain neu-ronal cells may be permissive for L1 retrotransposition.However, additional research is needed to truly under-stand the effect of L1 retrotransposition in the brain. Forexample, recent advances in DNA sequencing technol-ogy should provide a means to directly test whether theL1 DNA copy number changes that were detected inquantitative PCR experiments represent actual de novo endogenous retrotransposition events or instead resultfrom other forms of genomic instability that have beenreported in neurons186,187. Similarly, it remains unclearwhether endogenous L1 retrotransposition events rep-resent a type of ‘genomic noise’ or whether they have anyfunctional impact on neuronal development. Finally, itremains a mystery why neuronal cells may accommo-date L1 retrotransposition at apparently higher levelsthan other somatic cells. Nonetheless, these studies haveunveiled a new area of investigation that is likely to bethe subject of future work.

Deregulated L1 retrotransposition in cancer cells. Agrowing body of evidence suggests that L1 retrotrans-position may become deregulated in certain cancers. For

example, early studies revealed that hypomethylation ofthe L1 promoter is correlated with increased L1 expres-sion and/or the production of the L1 ORF1-encodedprotein in certain tumours188–190. Moreover, engineeredhuman L1s readily retrotranspose in a variety of trans-formed human and mouse cell lines but generallyshow lower levels of retrotransposition activity in nor-mal human cells such as fibroblasts (for example, seeREFS 11,191,192). Consistent with this, recent findingsusing second-generation DNA sequencing revealeda total of nine de novo L1 retrotransposition events in6 out of 20 examined non-small-cell lung tumours12.Intriguingly, the tumours that contained the new L1

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insertions also exhibited a specific genome-wide hypo-methylation signature, which is consistent with thenotion that altering the epigenome can create a permis-sive environment for L1 expression and/or retrotrans-position and perhaps for the retrotransposition of otherclasses of non-LTR retrotransposons. Clearly, furtherinnovations in DNA sequencing of heterogeneous cellpopulations will be crucial to reveal patterns of TE activ-ity in diverse tumours. Then the challenge will be todetermine whether all these TE insertions are ‘passenger’mutations that are a consequence of the altered cellularmilieu of cancer cells or whether some act as ‘drivers’ topromote tumorigenesis.

Closing remarks

It is undeniable that TEs have played an important partin structuring genomes and generating genetic diversity.By understanding how, when and where TEs integrate,

and how the host responds to this ever-present threat,we might unveil the dynamic forces that shape ourgenomes. Indeed, we are now able to critically evaluatethe McClintock doctrine, and future experiments couldprovide valuable insight into whether the increases inTE transcription that are caused by environmental stresslead to higher levels of TE integration and whether theseinsertions have an impact on host phenotypes and/orsurvival.

It remains a curiosity why sequences without anyapparent purpose continue to thrive in genomes. What isclear is that an understanding of TE biology is necessaryto understand genome biology. It is intriguing to specu-late that some phenotypic differences among organismsor between individuals are due to the effects of TEs.These speculations require rigorous experimental tests.However, the coming years should be an exciting timefor TE biology.

1. McClintock, B. The origin and behavior of mutable loci

in maize. Proc. Natl Acad. Sci. USA 36, 344–355

(1950).

2. Levis, R. W., Ganesan, R., Houtchens, K., Tolar, L. A. &

Sheen, F. M. Transposons in place of telomeric repeats

at a Drosophila telomere. Cell  75, 1083–1093 (1993).

3.  Agrawal, A., Eastman, Q. M. & Schatz, D. G.

Transposition mediated by RAG1 and RAG2 and its

implications for the evolution of the immune system.

Nature 394, 744–751 (1998).

4. Feschotte, C. Transposable elements and the evolution

of regulatory networks. Nature Rev. Genet. 9,

397–405 (2008).

5. Beck, C. R., Garcia-Perez, J. L., Badge, R. M. &

Moran, J. V. LINE-1 elements in structural variation

and disease. Annu. Rev. Genomics Hum. Genet. 

18 Jul 2011 (doi:10.1146/annurev-genom-

082509-141802).

6. Orgel, L. E., Crick, F. H. & Sapienza, C. Selfish DNA.

Nature 288, 645–646 (1980).

7. Doolittle, W. F. & Sapienza, C. Selfish genes, the

phenotype paradigm and genome evolution. Nature 

284, 601–603 (1980).8. Bestor, T. H. Sex brings transposons and genomes into

conflict. Genetica 107, 289–295 (1999).

9. Hickey, D. A. Selfish DNA: a sexually-transmitted

nuclear parasite. Genetics 101, 519–531 (1982).

10. Goodier, J. L. & Kazazian, H. H. Jr. Retrotransposons

revisited: the restraint and rehabilitation of parasites.

Cell  135, 23–35 (2008).

11. Moran, J. V. et al. High frequency retrotransposition

in cultured mammalian cells. Cell  87, 917–927

(1996).

12. Iskow, R. C. et al. Natural mutagenesis of human

genomes by endogenous retrotransposons. Cell  141,

1253–1261 (2010).

13. Beck, C. R. et al. LINE-1 retrotransposition activity in

human genomes. Cell  141, 1159–1170 (2010).

14. Huang, C. R. et al. Mobile interspersed repeats are

major structural variants in the human genome.

Cell  141, 1171–1182 (2010).15. Witherspoon, D. J. et al. Mobile element scanning

(ME-Scan) by targeted high-throughput sequencing.BMC Genomics 11, 410 (2010).

16. Hormozdiari, F. et al.  Alu repeat discovery and

characterization within human genomes. Genome Res. 

21, 840–849 (2011).

17. Ewing, A. D. & Kazazian, H. H. Jr. High-throughput

sequencing reveals extensive variation in human-

specific L1 content in individual human genomes.

Genome Res. 20, 1262–1270 (2010).

18. Ewing, A. D. & Kazazian, H. H. Jr. Whole-genome

resequencing allows detection of many rare LINE-1

insertion alleles in humans. Genome Res. 21,

985–990 (2011).19. Garcia-Perez, J. L. et al. LINE-1 retrotransposition in

human embryonic stem cells. Hum. Mol. Genet. 16,

1569–1577 (2007).

20. van den Hurk, J. A. et al. L1 retrotransposition can

occur early in human embryonic development.

Hum. Mol. Genet. 16, 1587–1592 (2007).

21. Kano, H. et al. L1 retrotransposition occurs mainly in

embryogenesis and creates somatic mosaicism.

Genes Dev. 23, 1303–1312 (2009).

References 19–21 provide evidence that L1s can

retrotranspose during early embryonic development.

22. Miki, Y. et al. Disruption of the APC  gene by a

retrotransposal insertion of L1 sequence in a colon

cancer. Cancer Res. 52, 643–645 (1992).

23. Coufal, N. G. et al. L1 retrotransposition in human

neural progenitor cells. Nature 460, 1127–1131

(2009).

24. Muotri, A. R. et al. Somatic mosaicism in neuronal

precursor cells mediated by L1 retrotransposition.

Nature 435, 903–910 (2005).

25. Sandmeyer, S. Integration by design. Proc. Natl Acad.

Sci. USA 100, 5586–5588 (2003).

26. Leem, Y. E. et al. Retrotransposon Tf1 is targeted to

pol II promoters by transcription activators. Mol. Cell  

30, 98–107 (2008).

This paper demonstrates a mechanism by which

the S. pombe Tf1 retrotransposon can target RNA

polymerase II-transcribed genes.

27. Devine, S. E. & Boeke, J. D. Integration of the yeastretrotransposon Ty1 is targeted to regions upstream

of genes transcribed by RNA polymerase III.

Genes Dev. 10, 620–633 (1996).

28. Eickbush, T. H. in Mobile DNA II  (eds Craig, N. L.,

Craigie, R., Gellert, M. & Lambowitz, A. M.) 813–835

(ASM Press, Washington DC, 2002).

29. Winckler, T., Dingermann, T. & Glockner, G.

Dictyostelium mobile elements: strategies to amplify

in a compact genome. Cell. Mol. Life Sci. 59,

2097–2111 (2002).

30. Malone, C. D. & Hannon, G. J. Small RNAs as

guardians of the genome. Cell  136, 656–668 (2009).

31. Bestor, T. H. & Bourc’his, D. Transposon silencing and

imprint establishment in mammalian germ cells.

Cold Spring Harb. Symp. Quant. Biol. 69, 381–387

(2004).

32. Golden, D. E., Gerbasi, V. R. & Sontheimer, E. J.

 An inside job for siRNAs. Mol. Cell  31, 309–312

(2008).

33. Chiu, Y. L. & Greene, W. C. The APOBEC3 cytidinedeaminases: an innate defensive network opposing

exogenous retroviruses and endogenous retroelements.

 Annu. Rev. Immunol. 26, 317–353 (2008).

34. Sehgal, A., Lee, C. Y. & Espenshade, P. J. SREBP controls

oxygen-dependent mobilization of retrotransposons in

fission yeast. PLoS Genet. 3, e131 (2007).

35. Dai, J., Xie, W., Brady, T. L., Gao, J. & Voytas, D. F.

Phosphorylation regulates integration of the yeast Ty5 

retrotransposon into heterochromatin. Mol. Cell  27,

289–299 (2007).

This paper demonstrates that TEs can respond to

stress by changing their target sites. When cells

lack access to nitrogen, Ty5 no longer integrates

into heterochromatin but instead targets coding

sequences in the genome.

36. Grandbastien, M. A. et al. Stress activation and genomic

impact of Tnt1 retrotransposons in Solanaceae.

Cytogenet. Genome Res. 110, 229–241 (2005).

37. Hirochika, H. Activation of tobacco retrotransposons

during tissue culture. EMBO J. 12, 2521–2528 (1993).

38. Courtial, B. et al. Tnt1 transposition events are

induced by in vitro transformation of Arabidopsis

thaliana, and transposed copies integrate into genes.

Mol. Genet. Genomics 265, 32–42 (2001).

39. McClintock, B. The significance of responses of the

genome to challenge. Science 226, 792–801 (1984).

40. Craig, N. L., Craigie, R., Gellert, M. & Lambowitz, A. M.

(eds) Mobile DNA II  (ASM Press, Washington DC, 2002).

41. Kleckner, N. Regulation of transposition in bacteria.

 Annu. Rev. Cell Biol. 6, 297–327 (1990).

42. Garfinkel, D. J., Boeke, J. D. & Fink, G. R. Ty element

transposition: reverse transcriptase and virus-like

particles. Cell  42, 507–517 (1985).

43. Boeke, J. D., Garfinkel, D. J., Styles, C. A. & Fink, G. R.

Ty elements transpose through an RNA intermediate.

Cell  40, 491–500 (1985).

44. Eichinger, D. J. & Boeke, J. D. The DNA intermediate

in yeast Ty1 element transposition copurifies with

virus-like particles: cell-free Ty1 transposition. Cell  54,

955–966 (1988).

45. Luan, D. D., Korman, M. H., Jakubczak, J. L. &Eickbush, T. H. Reverse transcription of R2Bm RNA is

primed by a nick at the chromosomal target site: a

mechanism for non-LTR retrotransposition. Cell  72,

595–605 (1993).

46. Kajikawa, M. & Okada, N. LINEs mobilize SINEs in the

eel through a shared 3′ sequence. Cell  111, 433–444

(2002).

47. Dewannieux, M., Esnault, C. & Heidmann, T.

LINE-mediated retrotransposition of marked Alu 

sequences. Nature Genet. 35, 41–48 (2003).

48. Hancks, D. C., Goodier, J. L., Mandal, P. K.,

Cheung, L. E. & Kazazian, H. H. Jr. Retrotransposition

of marked SVA elements by human L1s in cultured

cells. Hum. Mol. Genet. 2 Jun 2011 (doi:10.1093/

hmg/ddr245).

49. Garcia-Perez, J. L., Doucet, A. J., Bucheton, A.,

Moran, J. V. & Gilbert, N. Distinct mechanisms for

trans-mediated mobilization of cellular RNAs by

the LINE-1 reverse transcriptase. Genome Res. 17,

602–611 (2007).50. Gilbert, N., Lutz, S., Morrish, T. A. & Moran, J. V.

Multiple fates of L1 retrotransposition intermediates

in cultured human cells. Mol. Cell. Biol. 25,

7780–7795 (2005).

51. Buzdin, A. et al. A new family of chimeric retrotranscripts

formed by a full copy of U6 small nuclear RNA fused to

the 3′ terminus of L1. Genomics 80, 402–406 (2002).

52. Weber, M. J. Mammalian small nucleolar RNAs are

mobile genetic elements. PLoS Genet. 2, e205 (2006).

53. Wei, W. et al. Human L1 retrotransposition: cis 

preference versus trans complementation. Mol. Cell.

Biol. 21, 1429–1439 (2001).54. Esnault, C., Maestre, J. & Heidmann, T. Human LINE

retrotransposons generate processed pseudogenes.

Nature Genet. 24, 363–367 (2000).

55. Reznikoff, W. S. Tn5 transposition: a molecular tool

for studying protein structure-function. Biochem. Soc.

Trans. 34, 320–323 (2006).

R E V I E W S

NATURE REVIEWS | GENETICS  VOLUME 12 | SEPTEMBER 2011 | 625

© 2011 Macmillan Publishers Limited. All rights reserved

Page 12: Levin Moran, 2011

7/17/2019 Levin Moran, 2011

http://slidepdf.com/reader/full/levin-moran-2011 12/13

56. Peters, J. E. & Craig, N. L. Tn7 : smarter than we

thought. Nature Rev. Mol. Cell Biol. 2, 806–814

(2001).

57. Rio, D. C. in Mobile DNA II  (eds Craig, N. L., Craigie, R.,

Gellert, M. & Lambowitz, A. M.) 484–518 (ASM Press,

Washington DC, 2002).

58. Plasterk, R. H. The Tc1/mariner  transposon family.

Curr. Top. Microbiol. Immunol. 204, 125–143 (1996).

59. Lander, E. S. et al. Initial sequencing and analysis of

the human genome. Nature 409, 860–921 (2001).

60. Ray, D. A., Pagan, H. J., Thompson, M. L. &

Stevens, R. D. Bats with hATs: evidence for recent DNAtransposon activity in genus Myotis. Mol. Biol. Evol. 

24, 632–639 (2007).

61. Pritham, E. J. & Feschotte, C. Massive amplification of

rolling-circle transposons in the lineage of the bat

Myotis lucifugus. Proc. Natl Acad. Sci. USA 104,

1895–1900 (2007).

62. Ray, D. A. et al. Multiple waves of recent DNA

transposon activity in the bat, Myotis lucifugus.

Genome Res. 18, 717–728 (2008).

References 60–62 reveal that certain DNA

transposons are active in the Myotis genus of bats.63. Bowen, N. J. & McDonald, J. F. Drosophila 

euchromatic LTR retrotransposons are much younger

than the host species in which they reside. Genome

Res. 11, 1527–1540 (2001).

64. Schnable, P. S. et al. The B73 maize genome:

complexity, diversity, and dynamics. Science 326,

1112–1115 (2009).

65. SanMiguel, P. et al. Nested retrotransposons in the

intergenic regions of the maize genome. Science 274,

765–768 (1996).66. Maksakova, I. A. et al. Retroviral elements and their

hosts: insertional mutagenesis in the mouse germ line.

PLoS Genet. 2, e2 (2006).

67. Moyes, D., Griffiths, D. J. & Venables, P. J. Insertional

polymorphisms: a new lease of life for endogenous

retroviruses in human disease. Trends Genet. 23,

326–333 (2007).68. Badge, R. M., Alisch, R. S. & Moran, J. V. ATLAS: a

system to selectively identify human-specific L1

insertions. Am. J. Hum. Genet. 72, 823–838 (2003).

69. Sheen, F. M. et al. Reading between the LINEs:

human genomic variation induced by LINE-1

retrotransposition. Genome Res. 10, 1496–1508

(2000).70. Kidd, J. M. et al. Mapping and sequencing of

structural variation from eight human genomes.

Nature 453, 56–64 (2008).

71. Kidd, J. M. et al. A human genome structural variation

sequencing resource reveals insights into mutational

mechanisms. Cell  143, 837–847 (2010).72. Korbel, J. O. et al. Paired-end mapping reveals

extensive structural variation in the human genome.

Science 318, 420–426 (2007).

73. Durbin, R. M. et al. A map of human genome variation

from population-scale sequencing. Nature 467,

1061–1073 (2010).

74. Mills, R. E. et al. Mapping copy number variation by

population-scale genome sequencing. Nature 470,

59–65 (2011).

References 12–18 and 70–74 used

second-generation DNA sequencing and/or modern

genomic approaches to demonstrate that TEs

continue to have an ongoing impact on human

genome structural variation.

75. Kuduvalli, P. N., Rao, J. E. & Craig, N. L. Target DNA

structure plays a critical role in Tn7  transposition.

EMBO J. 20, 924–932 (2001).

76. Craig, N. L. in Mobile DNA II  (eds Craig, N. L.,

Craigie, R., Gellert, M. & Lambowitz, A. M.) 423–456

(ASM Press, Washington DC, 2002).77. Wolkow, C. A., DeBoy, R. T. & Craig, N. L. Conjugating

plasmids are preferred targets for Tn7 . Genes Dev. 10,

2145–2157 (1996).

78. Peters, J. E. & Craig, N. L. Tn7  transposes proximal to

DNA double-strand breaks and into regions where

chromosomal DNA replication terminates. Mol. Cell  6,

573–582 (2000).

79. Bellen, H. J. et al. The Drosophila gene disruption

project: progress using transposons with distinctive

site-specificities. Genetics 188,731–743 (2011).

80. Feng, Q., Moran, J., Kazazian, H. & Boeke, J. D.

Human L1 retrotransposon encodes a conserved

endonuclease required for retrotransposition. Cell  87,

905–916 (1996).

81. Feng, Q., Schumann, G. & Boeke, J. D.

Retrotransposon R1Bm endonuclease cleaves the

target sequence. Proc. Natl Acad. Sci. USA 95,

2083–2088 (1998).

82.  Yang, J., Malik, H. S. & Eickbush, T. H. Identification of

the endonuclease domain encoded by R2 and other

site-specific, non-long terminal repeat

retrotransposable elements. Proc. Natl Acad. Sci. USA 

96, 7847–7852 (1999).

83. Bushman, F. D. Targeting survival: integration site

selection by retroviruses and LTR-retrotransposons.

Cell  115, 135–138 (2003).

84. Lesage, P. & Todeschini, A. L. Happy together:

the life and times of Ty retrotransposons and their

hosts. Cytogenet. Genome Res. 110, 70–90 (2005).

85. Kirchner, J., Connolly, C. M. & Sandmeyer, S. B.Requirement of RNA polymerase III transcription

factors for in vitro position-specific integration of a

retroviruslike element. Science 267, 1488–1491

(1995).86. Chalker, D. L. & Sandmeyer, S. B. Ty3 integrates

within the region of RNA polymerase III

transcription initiation. Genes Dev. 6, 117–128

(1992).

87.  Yieh, L., Hatzis, H., Kassavetis, G. & Sandmeyer, S. B.

Mutational analysis of the transcription factor IIIB-DNA

target of Ty3 retroelement integration. J. Biol. Chem. 

277, 25920–25928 (2002).

88.  Yieh, L., Kassavetis, G., Geiduschek, E. P. &

Sandmeyer, S. B. The Brf and TATA-binding protein

subunits of the RNA polymerase III transcription

factor IIIB mediate position-specific integration of

the gypsy-like element, Ty3. J. Biol. Chem. 275,

29800–29807 (2000).

89. Bachman, N., Eby, Y. & Boeke, J. D. Local definition of

Ty1 target preference by long terminal repeats and

clustered tRNA genes. Genome Res. 14, 1232–1247

(2004).

90. Bachman, N., Gelbart, M. E. , Tsukiyama, T. &

Boeke, J. D. TFIIIB subunit Bdp1p is required for

periodic integration of the Ty1 retrotransposon and

targeting of Isw2p to S. cerevisiae tDNAs. Genes Dev. 

19, 955–964 (2005).

91. Bolton, E. C. & Boeke, J. D. Transcriptional

interactions between yeast tRNA genes, flanking

genes and Ty elements: a genomic point of view.

Genome Res. 13, 254–263 (2003).

92. Kinsey, P. T. & Sandmeyer, S. B. Adjacent pol II and pol

III promoters: transcription of the yeast

retrotransposon Ty3 and a target tRNA gene. Nucleic

 Acids Res. 19, 1317–1324 (1991).

93. Hofmann, J. et al. Transfer RNA genes from

Dictyostelium discoideum are frequently associated

with repetitive elements and contain consensus boxes

in their 5′ and 3′-flanking regions. J. Mol. Biol. 222,

537–552 (1991).

94. Chung, T., Siol, O., Dingermann, T. & Winckler, T.Protein interactions involved in tRNA gene-specific

integration of Dictyostelium discoideum non-long

terminal repeat retrotransposon TRE5-A. Mol. Cell.

Biol. 27, 8492–8501 (2007).95. Behrens, R., Hayles, J. & Nurse, P. Fission yeast

retrotransposon Tf1 integration is targeted to 5′ ends

of open reading frames. Nucleic Acids Res. 28,

4709–4716 (2000).

96. Guo, Y. & Levin, H. L. High-throughput sequencing

of retrotransposon integration provides a

saturated profile of target activity in

Schizosaccharomyces pombe. Genome Res. 20,

239–248 (2010).

This study describes the sequencing of a saturated

set of insertion sites that are actively targeted

by a TE.

97. Singleton, T. L. & Levin, H. L. A long terminal repeat

retrotransposon of fission yeast has strong

preferences for specific sites of insertion. Eukaryotic

Cell  1, 44–55 (2002).98. Neely, L. A. & Hoffman, C. S. Protein kinase A and

mitogen-activated protein kinase pathways

antagonistically regulate fission yeast fbp1

transcription by employing different modes of action

at two upstream activation sites. Mol. Cell. Biol. 20,

6426–6434 (2000).

99. Majumdar, A., Chatterjee, A. G., Ripmaster, T. L. &

Levin, H. L. The determinants that specify the

integration pattern of retrotransposon Tf1 in the fbp1

promoter of Schizosaccharomyces pombe. J. Virol. 

85, 519–529 (2010).

100. Chen, D. et al. Global transcriptional responses of

fission yeast to environmental stress. Mol. Biol. Cell  

14, 214–229 (2003).

101. Malik, H. S., Henikoff, S. & Eickbush, T. H. Poised for

contagion: evolutionary origins of the infectious

abilities of invertebrate retroviruses. Genome Res. 10,

1307–1318 (2000).

102. Mitchell, R. S. et al. Retroviral DNA integration:

 ASLV, HIV, and MLV show distinct target site

preferences. PLoS Biol. 2, e234 (2004).

103. Ciuffi, A. Mechanisms governing lentivirus integration

site selection. Curr. Gene Ther. 8, 419–429 (2008).

104. Ciuffi, A. & Bushman, F. D. Retroviral DNA integration:

HIV and the role of LEDGF/p75. Trends Genet. 22,

388–395 (2006).

105. Engelman, A. & Cherepanov, P. The lentiviral integrase

binding protein LEDGF/p75 and HIV-1 replication.

PLoS Pathog. 4, e1000046 (2008).

106. Poeschla, E. M. Integrase, LEDGF/p75 and HIVreplication. Cell. Mol. Life Sci. 65, 1403–1424 (2008).

107. Malik, H. S. & Eickbush, T. H. Modular evolution of the

integrase domain in the Ty3/Gypsy class of LTR

retrotransposons. J. Virol. 73, 5186–5190 (1999).108. Gao, X., Hou, Y., Ebina, H., Levin, H. L. & Voytas, D. F.

Chromodomains direct integration of retrotransposons

to heterochromatin. Genome Res. 18, 359–369

(2008).

109. Zou, S. & Voytas, D. F. Silent chromatin determines

target preference of the Saccharomyces 

retrotransposon Ty5. Proc. Natl Acad. Sci. USA 94,

7412–7416 (1997).

110. Zou, S., Wright, D. A. & Voytas, D. F.

The Saccharomyces Ty5 retrotransposon family is

associated with origins of DNA replication at the

telomeres and the silent mating locus HMR. Proc. Natl

 Acad. Sci. USA 92, 920–924 (1995).

111. Zou, S., Ke, N., Kim, J. M. & Voytas, D. F.

The Saccharomyces retrotransposon Ty5 integrates

preferentially into regions of silent chromatin at the

telomeres and mating loci. Genes Dev. 10, 634–645

(1996).

112. Gai, X. & Voytas, D. F. A single amino acid change in

the yeast retrotransposon Ty5 abolishes targeting to

silent chromatin. Mol. Cell  1, 1051–1055 (1998).

113. Xie, W. et al. Targeting of the yeast Ty5 

retrotransposon to silent chromatin is mediated by

interactions between integrase and Sir4p. Mol. Cell.

Biol. 21, 6606–6614 (2001).

114. Zhu, Y., Dai, J., Fuerst, P. G. & Voytas, D. F.

Controlling integration specificity of a yeast

retrotransposon. Proc. Natl Acad. Sci. USA 100,

5891–5895 (2003).

115. Brady, T. L., Fuerst, P. G., Dick, R. A., Schmidt, C. &

 Voytas, D. F. Retrotransposon target site selection by

imitation of a cellular protein. Mol. Cell. Biol. 28,

1230–1239 (2008).

116. George, J. A., Traverse, K. L., DeBaryshe, P. G.,

Kelley, K. J. & Pardue, M. L. Evolution of diverse

mechanisms for protecting chromosome ends by

Drosophila TART telomere retrotransposons.Proc. Natl Acad. Sci. USA 107, 21052–21057 (2010).

117. Biessmann, H. et al. HeT-A, a transposable element

specifically involved in “healing” broken chromosome

ends in Drosophila melanogaster . Mol. Cell. Biol. 12,

3910–3918 (1992).

118. Fujiwara, H., Osanai, M., Matsumoto, T. & Kojima, K. K.

Telomere-specific non-LTR retrotransposons and

telomere maintenance in the silkworm, Bombyx mori .

Chromosome Res. 13, 455–467 (2005).

119. Gladyshev, E. A. & Arkhipova, I. R. A subtelomeric

non-LTR retrotransposon Hebe in the bdelloid rotifer

 Adineta vaga is subject to inactivation by deletions

but not 5′ truncations. Mob. DNA 1, 12 (2010).

120. Morrish, T. A. et al. Endonuclease-independent LINE-1

retrotransposition at mammalian telomeres. Nature 

446, 208–212 (2007).

121. Morrish, T. A. et al. DNA repair mediated by

endonuclease-independent LINE-1 retrotransposition.

Nature Genet. 31, 159–165 (2002).

122. Gladyshev, E. A. & Arkhipova, I. R. Telomere-associated endonuclease-deficient Penelope-like

retroelements in diverse eukaryotes. Proc. Natl Acad.

Sci. USA 104, 9352–9357 (2007).

References 120 and 122 reveal pathways by which

retrotransposons can use telomeric sequences as

integration substrates.

123. Gilbert, N., Lutz-Prigge, S. & Moran, J. V.

Genomic deletions created upon LINE-1

retrotransposition. Cell  110, 315–325 (2002).

124. Symer, D. E. et al. Human L1 retrotransposition is

associated with genetic instability in vivo. Cell  110,

327–338 (2002).

125. Harrow, J. et al. GENCODE: producing a reference

annotation for ENCODE. Genome Biol. 7 (Suppl. 1), 4

(2006).126. Coffey, A. J. et al. The GENCODE exome: sequencing

the complete human exome.Eur. J. Hum. Genet. 19,

827–831 (2011).

R E V I E W S

626 | SEPTEMBER 2011 | VOLUME 12 www.nature.com/reviews/genetics

© 2011 Macmillan Publishers Limited. All rights reserved

Page 13: Levin Moran, 2011

7/17/2019 Levin Moran, 2011

http://slidepdf.com/reader/full/levin-moran-2011 13/13

127. Jurka, J. Sequence patterns indicate an enzymatic

involvement in integration of mammalian retroposons.

Proc. Natl Acad. Sci. USA 94, 1872–1877 (1997).

128. Cost, G. J. & Boeke, J. D. Targeting of human

retrotransposon integration is directed by the specificity

of the L1 endonuclease for regions of unusual DNA

structure.Biochemistry 37, 18081–18093 (1998).

129. Korenberg, J. R. & Rykowski, M. C. Human genome

organization: Alu, LINES, and the molecular structure

of metaphase chromosome bands. Cell  53, 391–400

(1988).

130. Brookfield, J. F. Selection on Alu sequences?Curr. Biol. 11, R900–R901 (2001).

131. Boissinot, S., Entezam, A. & Furano, A. V. Selection

against deleterious LINE-1-containing loci in the

human lineage. Mol. Biol. Evol. 18, 926–935 (2001).132. Boissinot, S., Davis, J., Entezam, A., Petrov, D. &

Furano, A. V. Fitness cost of LINE-1 (L1) activity in

humans. Proc. Natl Acad. Sci. USA 103, 9590–9594

(2006).

133. Han, J. S., Szak, S. T. & Boeke, J. D. Transcriptional

disruption by the L1 retrotransposon and implications

for mammalian transcriptomes. Nature 429,

268–274 (2004).

134. Perepelitsa-Belancio, V. & Deininger, P.

RNA truncation by premature polyadenylation

attenuates human mobile element activity. Nature

Genet. 35, 363–366 (2003).

135. Conley, M. E., Partain, J. D., Norland, S. M.,

Shurtleff, S. A. & Kazazian, H. H. Jr. Two independent

retrotransposon insertions at the same site within the

coding region of BTK. Hum. Mutat. 25, 324–325

(2005).136. Halling, K. C. et al. Hereditary desmoid disease in a

family with a germline Alu I  repeat mutation of the

 APC gene. Hum. Hered. 49, 97–102 (1999).

137. Vidaud, D. et al. Haemophilia B due to a de novo 

insertion of a human-specific Alu subfamily member

within the coding region of the factor IX gene.

Eur. J. Hum. Genet. 1, 30–36 (1993).138. Cost, G. J., Golding, A., Schlissel, M. S. & Boeke, J. D.

Target DNA chromatinization modulates nicking by L1

endonuclease. Nucleic Acids Res. 29, 573–577 (2001).

139. Selker, E. U. et al. The methylated component of the

Neurospora crassa genome. Nature 422, 893–897

(2003).

140. Yoder, J. A., Walsh, C. P. & Bestor, T. H.

Cytosine methylation and the ecology of intragenomic

parasites. Trends Genet. 13, 335–340 (1997).

141. Goll, M. G. & Bestor, T. H. Eukaryotic cytosine

methyltransferases. Annu. Rev. Biochem. 74,

481–514 (2005).

142. Schaefer, C. B., Ooi, S. K., Bestor, T. H. & Bourc’his, D.Epigenetic decisions in mammalian germ cells. Science 

316, 398–399 (2007).

143. Maksakova, I. A., Mager, D. L. & Reiss, D.

Keeping active endogenous retroviral-like elements in

check: the epigenetic perspective. Cell. Mol. Life Sci. 

65, 3329–3347 (2008).

144. Tsukahara, S. et al. Bursts of retrotransposition

reproduced in Arabidopsis. Nature 461, 423–426

(2009).

This paper used modern genomic approaches to

reveal that certain classes of LTR retrotransposon

are mobilized in a DDM1 (decrease in DNA

methylation 1) mutant background. It provides

an example of how epigenetic changes can lead

to TE mobility.145. Bourc’his, D. & Bestor, T. H. Meiotic catastrophe and

retrotransposon reactivation in male germ cells

lacking Dnmt3L. Nature 431, 96–99 (2004).146. Law, J. A. & Jacobsen, S. E. Establishing, maintaining

and modifying DNA methylation patterns in plantsand animals. Nature Rev. Genet. 11, 204–220 (2010).

147. van Rij, R. P. & Berezikov, E. Small RNAs and the

control of transposons and viruses in Drosophila.

Trends Microbiol. 17, 163–171 (2009).

148. Ghildiyal, M. & Zamore, P. D. Small silencing RNAs:

an expanding universe. Nature Rev. Genet. 10,

94–108 (2009).

149. Slotkin, R. K. & Martienssen, R. Transposable

elements and the epigenetic regulation of the genome.

Nature Rev. Genet. 8, 272–285 (2007).

150. Czech, B. & Hannon, G. J. Small RNA sorting: matchmaking

for Argonautes. Nature Rev. Genet. 12, 19–31 (2011).151. Aravin, A. A., Hannon, G. J. & Brennecke, J. The Piwi-

piRNA pathway provides an adaptive defense in the

transposon arms race. Science 318, 761–764 (2007).

152. Fischer, S. E. Small RNA-mediated gene silencing

pathways in C. elegans. Int. J. Biochem. Cell Biol. 42,

1306–1315 (2010).

153. Sijen, T. & Plasterk, R. H. Transposon silencing in the

Caenorhabditis elegans germ line by natural RNAi.

Nature 426, 310–314 (2003).

154. Tabara, H. et al. The rde-1 gene, RNA interference,

and transposon silencing in C. elegans. Cell  99,

123–132 (1999).

155. Malone, C. D. et al. Specialized piRNA pathways act in

germline and somatic tissues of the Drosophila ovary.

Cell  137, 522–535 (2009).156. Olivieri, D., Sykora, M. M., Sachidanandam, R.,

Mechtler, K. & Brennecke, J. An in vivo RNAi assay

identifies major genetic and cellular requirements forprimary piRNA biogenesis in Drosophila. EMBO J. 29,

3301–3317 (2010).

157. Zamore, P. D. Somatic piRNA biogenesis. EMBO J. 29,

3219–3221 (2010).158. Brennecke, J. et al. An epigenetic role for maternally

inherited piRNAs in transposon silencing. Science 

322, 1387–1392 (2008).

159. Slotkin, R. K. et al. Epigenetic reprogramming and

small RNA silencing of transposable elements in

pollen. Cell  136, 461–472 (2009).

References 155, 156, 158 and 159 highlight some

pathways by which small RNAs can inhibit TE

mobility in either the germ line or soma.

160. Yang, N. & Kazazian, H. H. Jr. L1 retrotransposition

is suppressed by endogenously encoded small

interfering RNAs in human cultured cells. Nature

Struct. Mol. Biol. 13, 763–771 (2006).

161. Kuramochi-Miyagawa, S. et al. DNA methylation of

retrotransposon genes is regulated by Piwi family

members MILI and MIWI2 in murine fetal testes.

Genes Dev. 22, 908–917 (2008).

162. Reuter, M. et al. Loss of the Mili-interacting Tudor

domain-containing protein-1 activates transposons

and alters the Mili-associated small RNA profile.

Nature Struct. Mol. Biol. 16, 639–646 (2009).

163. Soper, S. F. et al. Mouse maelstrom, a component of

nuage, is essential for spermatogenesis and transposon

repression in meiosis. Dev. Cell  15, 285–297 (2008).

References 161–163 highlight how the small RNA

machinery influences the expression, and perhaps

mobility, of certain TEs in mammalian cells.

164. Matsuda, E. & Garfinkel, D. J. Posttranslational

interference of Ty1 retrotransposition by antisense

RNAs. Proc. Natl Acad. Sci. USA 106, 15657–15662

(2009).

This paper reveals a novel RNA-based mechanism

that inhibits Ty1 retrotransposition in S. cerevisiae.

165. Crow, Y. J. et al. Mutations in the gene encoding the

3′-5′ DNA exonuclease TREX1 cause Aicardi-Goutieres

syndrome at the AGS1 locus. Nature Genet. 38,

917–920 (2006).166. Stetson, D. B., Ko, J. S., Heidmann, T. & Medzhitov, R.

TREX1 prevents cell-intrinsic initiation of

autoimmunity. Cell  134, 587–598 (2008).

167. Grimaldi, G. & Singer, M. F. Members of the KpnI

family of long interspersed repeated sequences join

and interrupt alpha-satellite in the monkey genome.

Nucleic Acids Res. 11, 321–338 (1983).

168. Eickbush, T. H. & Jamburuthugoda, V. K. The diversity

of retrotransposons and the properties of their reverse

transcriptases. Virus Res. 134, 221–234 (2008).

169. Babushok, D. V. & Kazazian, H. H. Jr. Progress in

understanding the biology of the human mutagen

LINE-1. Hum. Mutat. 28, 527–539 (2007).

170. Suzuki, J. et al. Genetic evidence that the non-

homologous end-joining repair pathway is involved in

LINE retrotransposition. PLoS Genet. 5, e1000461

(2009).

171. Gasior, S. L., Roy-Engel, A. M. & Deininger, P. L.

ERCC1/XPF limits L1 retrotransposition. DNA Repair

(Amst.) 7, 983–989 (2008).172. Garcia-Perez, J. L. et al. Epigenetic silencing of

engineered L1 retrotransposition events in human

embryonic carcinoma cells. Nature 466, 769–773

(2010).

This paper describes a host mechanism that is able

to epigenetically silence reporter genes delivered

into genomic DNA by L1 retrotransposition.

173. Rio, D. C. Regulation of Drosophila P  element

transposition. Trends Genet. 7, 282–287 (1991).

174. Pelisson, A. et al. Gypsy transposition correlates

with the production of a retroviral envelope-like

protein under the tissue-specific control of the

Drosophila flamenco gene. EMBO J. 13, 4401–4411

(1994).

175. Prud’homme, N., Gans, M., Masson, M., Terzian, C. &

Bucheton, A. Flamenco, a gene controlling the gypsy 

retrovirus of Drosophila melanogaster. Genetics 139,

697–711 (1995).

176. Branciforte, D. & Martin, S. L. Developmental and cell

type specificity of LINE-1 expression in mouse testis:

implications for transposition. Mol. Cell. Biol. 14,

2584–2592 (1994).

177. Trelogan, S. A. & Martin, S. L. Tightly regulated,

developmentally specific expression of the first

open reading frame from LINE-1 during mouse

embryogenesis. Proc. Natl Acad. Sci. USA 92,

1520–1524 (1995).178. Georgiou, I. et al. Retrotransposon RNA expression

and evidence for retrotransposition events in human

oocytes. Hum. Mol. Genet. 18, 1221–1228 (2009).179. Ostertag, E. M. et al. A mouse model of human L1

retrotransposition.Nature Genet. 32, 655–660

(2002).

180. Macia, A. et al. Epigenetic control of retrotransposon

expression in human embryonic stem cells. Mol. Cell.

Biol. 31, 300–316 (2011).

181. Federoff, N. in Mobile DNA II  (eds. Craig, N. L.,

Craigie, R., Gellert, M. & Lambowitz, A. M.)

997–1007 (ASM Press, Washington DC, 2002).

182. Emmons, S. W. & Yesner, L. High-frequency excision

of transposable element Tc1 in the nematode

Caenorhabditis elegans is limited to somatic cells.

Cell  36, 599–605 (1984).

183. Fernandez, L., Torregrosa, L., Segura, V., Bouquet, A.

& Martinez-Zapater, J. M. Transposon-induced gene

activation as a mechanism generating cluster shape

somatic variation in grapevine. Plant J. 61, 545–557

(2010).

184. Swergold, G. D. Identification, characterization, and

cell specificity of a human LINE-1 promoter. Mol. Cell.

Biol. 10, 6718–6729 (1990).185. Muotri, A. R. et al. L1 retrotransposition in neurons is

modulated by MeCP2. Nature 468, 443–446 (2010).

References 23, 24 and 185 suggest that

engineered human L1s, and perhaps endogenous

L1s, can retrotranspose in somatic cells of the

mammalian nervous system.

186. Rehen, S. K. et al. Chromosomal variation in neurons

of the developing and adult mammalian nervous

system. Proc. Natl Acad. Sci. USA 98, 13361–13366

(2001).

187. Westra, J. W. et al. Neuronal DNA content variation

(DCV) with regional and individual differences in the

human brain. J. Comp. Neurol. 518, 3981–4000

(2010).188. Belancio, V. P., Roy-Engel, A. M. & Deininger, P. L.

 All y’all need to know ‘bout retroelements in cancer.

Semin. Cancer Biol. 20, 200–210 (2010).

189. Alves, G., Tatro, A. & Fanning, T. Differential

methylation of human LINE-1 retrotransposons in

malignant cells. Gene 176, 39–44 (1996).190. Asch, H. L. et al. Comparative expression of the

LINE-1 p40 protein in human breast carcinomas

and normal breast tissues. Oncol. Res. 8, 239–247

(1996).

191. Kubo, S. et al. L1 retrotransposition in nondividing

and primary human somatic cells. Proc. Natl Acad.

Sci. USA 103, 8036–8041 (2006).

192. Shi, X., Seluanov, A. & Gorbunova, V. Cell divisions are

required for L1 retrotransposition. Mol. Cell. Biol. 27,

1264–1270 (2007).

 AcknowledgementsWe thank J. Kim, J. Garcia-Perez and members of the Moran

laboratory for critical reading of the manuscript. H.L.L. was

supported in part by the Intramural Research Program of the

US National Institutes of Health (NIH) from the Eunice

Kennedy Shriver National Institute of Child Health and

Human Development. He received additional support from

the Intramural AIDS Targeted Antiviral Program. J.V.M. was

supported in part by grants from the NIH (GM060518 andGM082970) and is an Investigator of the Howard Hughes

Medical Institute.

Competing interests statementThe authors declare competing financial interests: see Web

version for details.

FURTHER INFORMATIONHenry L. Levin’s homepage: https://science.nichd.nih.gov/

confluence/display/pcrm/Henry+Levin

 John V. Moran’s homepage at the Howard Hughes Medical

Institute:  http://www.hhmi.org/research/investigators/

moran_bio.html

 John V. Moran’s homepage at the University of Michigan: 

http://www.hg.med.umich.edu/faculty/john-v-moran-phd

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