Patent Application
20130315886
This application contains a paper copy and an electronic form of a sequence listing. The contents of sequence listing are incorporated herein by reference.
1. Field of the Invention
The invention relates to retrotransposons in neural cells.
2. Related Art
L1s are abundant retrotransposons that comprise approximately 20% of mammalian genomes (1-3). Recently-evolved L1s are polymorphic, resulting in individual variations in retrotransposition capacity (4,5). Although most L1s are retrotransposition-defective (6,7), active L1 retrotransposons can impact the genome in a variety of ways, creating insertions, deletions, new splice sites or gene expression fine-tuning (8-10). Previous data showed that, during neuronal differentiation, an EGFP-tagged L1 element could insert near or within neuron-associated genes, affecting gene expression and cell function. An analysis of the sequence data from several L1 insertions in neuronal precursor cells (NPCs), derived from neural stem cells (NSCs), indicated that the integration process might be regulated (11). Thus, neuronal networks may be affected by de novo L1 insertions during brain development (12,13).
In one aspect, a method of treating non-LTR retrotransposition that occurs in neural cells is provided. The method includes exposing a neural cell to a retrotransposition inhibitor in an amount sufficient to decrease non-LTR retrotransposition occurring in the neural cell or a progeny of the neural cell. The neural cell can be a neural stem cell or a neural precursor cell. In particular embodiments, the neural cell is a mammalian cell such as a human cell, and can be a fetal or embryonic cell. The neural cell can be identified with a nervous system condition that results from non-LTR retrotransposition in neural cells. In some embodiments, the neural cell is in a patient, which can be a newborn, a child or an adult. In some embodiments, the neural cell is in an embryo or fetus in a pregnant patient. The non-LTR retrotransposition can involve at least one L1 retrotransposon. A decrease in non-LTR retrotransposition can be determined by comparison to a control cell, for example, by comparison to a comparable neural cell not exposed to a retrotransposition inhibitor.
Nervous system conditions resulting from non-LTR retrotransposition in neural cells include, but are not limited to, autism or autism spectrum disorders, schizophrenia, Rett syndrome, Tourette syndrome, ataxia telangiectasia and other ataxias, xeroderma pigmentosum, Cockyne syndrome, fragile x, aspergers syndrome, childhood disintegrative disorder, tuberous sclerosis complex, or psychiatric disorders such as neurogiromatosis, Prader-Willi, Angelman, Joubert, Down, Williams or Cowdern syndrome or other psychiatric disorders, or any combination of conditions thereof.
Transposition inhibitors include, but are not limited to, anti-retroviral drugs, inhibitors of RNA stability, inhibitors of reverse transcription, inhibitors of L1 endonuclease activity, stimulators of DNA repair machinery, zinc-fingers that target the L1 promoter region, enzymes that inhibit L1, repressors that inhibit L1, or any combination thereof.
In another aspect, a method of assaying retrotransposition in neural cells is provided. The method includes sorting synchronized neural cells of the same genetic background into single neural cells, and subjecting one or more of the sorted single neural cells to quantitative polymerase chain reaction (“qPCR”) amplification of at least one retrotransposon. The synchronized neural cells can be neural stem cells or neural precursor cells. In some embodiments, the content of the at least one retrotransposon determined by the qPCR amplification is compared to the content of the at least one retrotransposon in one or more control cells. The control cells can be neural or non-neural cells depending on the type of comparison, and are of the same, or comparable, genetic background as the synchronized neural cells. In various embodiments, the retrotransposon is a non-LTR retrotransposon, and can be an L1 retrotransposon.
In a further aspect, a method of identifying an inhibitor of retrotransposition is provided. The method includes exposing one or more neural precursor cells to a candidate inhibitor, determining the content of at least one retrotransposon in the one or more neural precursor cells or in progeny of the one or more neural precursor cells, or both, and comparing the content of the at least one retrotransposon in the one or more neural precursor cells, or their progeny, or both, to the content of the at least one retrotransposon in one or more control cells not exposed to the candidate inhibitor. Depending on the type of comparison, the control cells can be neural precursor cells, or their progeny, or both. In this method, a decrease in the content of the retrotransposon in the one or more neural precursor cells, or their progeny, or both, compared to the one or more control cells is indicative of inhibition of retrotransposition. In various embodiments, the retrotransposon is a non-LTR retrotransposon, and can be an L1 retrotransposon.
In another aspect, a method of identifying a neural condition associated with non-LTR retrotransposition is provided. The method includes determining the content of at least one non-LTR retrotransposon in a neural cell in comparison to the content of the at least one non-LTR retrotransposon in one or more control cells. In this method, the neural cell is of a genotype associated with a nervous system condition. An increase in non-LTR retrotransposition content in neural cells versus control cells is an indication that the nervous system condition is associated with non-LTR retrotransposition. The nervous system condition can be autism or autism spectrum disorders, schizophrenia, Rett syndrome, Tourette syndrome, ataxia telangiectasia and other ataxias, xeroderma pigmentosum, Cockyne syndrome, fragile x, aspergers syndrome, childhood disintegrative disorder, tuberous sclerosis complex, or neurogiromatosis, Prader-Willi, Angelman, Joubert, Down, Williams and Cowdern syndrome or other psychiatric disorders, or any combination of conditions thereof. In some embodiments, the neural cell is from a knockout animal or an individual having the nervous system condition. In various embodiments, the retrotransposon is a non-LTR retrotransposon, and can be an L1 retrotransposon.
In a further aspect, highly efficient methods to measure Line-1 retrotransposition in tissue samples and single cells are provided. The methods and procedures provided herein may be used to measure Line-1 retrotransposition in a single cell as well as in multi-cellular samples. The assay may be used to monitor Line-1 retrotransposition in individual cells derived from fresh or frozen tissue samples, biopsies, fertilized eggs, induced pluripotent stem cells as well as tumor cells and therefore provides a valuable tool to monitor genomic mosaicism and genomic rearrangement. The present invention provides a novel diagnostic tool to monitor genomic rearrangement in cells. Examples for areas of application are cancer diagnosis, genomic screening in the context of in vitro fertilization, preventative screening, diagnosis of neurologic disorders as well as measuring neural plasticity.
In another aspect, a method of measuring Line-1 retrotransposition activity in single cells is provided. The method includes separating a tissue into single cells and isolating genomic DNA from the single cells, thereby forming single cell DNA samples. The single cell DNA samples are incubated with Line-1 primers and control primers. A Line-1 DNA is amplified with the Line-1 primers, thereby forming an amplified Line-1 DNA and a control DNA is amplified with the control primers, thereby forming an amplified control DNA. An amount of the amplified Line-1 DNA is compared with an amount of the amplified control DNA, thereby measuring Line-1 retrotransposition activity in the single cells.
In another aspect, a kit for measuring Line-1 retrotransposition activity in a single cell is provided. The kit includes Line-1 specific primers, control primers and a single cell.
In a further aspect, a kit for measuring Line-1 retrotransposition activity in a tissue is provided. The kit includes Line-1 specific primers, control primers and a tissue.
For a more complete understanding of the present invention, reference is now made to the following descriptions taken in conjunction with the accompanying drawings.
Various embodiments of methods are provided involving the transposition of retrotransposons, and in particular, non-LTR retrotransposons, in neural cells. As used herein, a “neural cell” is a neuroepithelial cell, a neural stem cell, a neural precursor cell, a neuron, a nerve cell, or a neurocyte. Retrotransposons include both long terminal repeat (“LTR”) retrotransposons and non-LTR retrotransposons. Non-LTR retrotransposons include LINE1 (long interspersed nucleotide elements, or L1) retrotransposons, SINE (short interspersed nucleotide elements) retrotransposons, and SVA (SINE-R, VNTR, Alu) retrotransposons. As is known, L1 retrotransposons are autonomous transposons containing many of the activities necessary for their mobility, while SINE and SVA retrotransposons are non-autonomous elements mobilized by L1 retrotransposons.
In some embodiments, the content of one or more retrotransposons in neural cells or their progeny is compared to the content of one or more retrotransposons in a control neural or non-neural cell. By “content” is meant the amount of DNA encoding a retrotransposon per cell, or the copy number of the retrotransposon per cell.
Some embodiments involve treating non-LTR retrotransposition in a cell. By “treating” is meant to decrease the level of non-LTR retrotransposition occurring in the cell. Although non-LTR retrotransposition may be a normal process of the developing nervous system, in some cases non-LTR retrotransposition can be associated with certain nervous system conditions. In those cases, decreasing non-LTR retrotransposition would be beneficial. In particular cases, the nervous system condition may be affected by, or be the result of, increased non-LTR retrotransposition above that which normally occurs during nervous system development, and in such cases, decreasing non-LTR retrotransposition would also be beneficial. In some embodiments, the treated cell is in a patient, and in such embodiments, “treating” can also mean to lessen the symptoms of a condition, or a total avoidance of a condition, in the patient.
Examples of nervous systems conditions for treatment include, but are not limited to, autism or autism spectrum disorders, schizophrenia, Rett syndrome, Tourette syndrome, ataxia telangiectasia and other ataxias, xeroderma pigmentosum, Cockyne syndrome, fragile x, aspergers syndrome, childhood disintegrative disorder, tuberous sclerosis complex, or neurogiromatosis, Prader-Willi, Angelman, Joubert, Down, Williams and Cowdern syndrome or other psychiatric disorders, or any combination of conditions thereof. A neural cell “identified” with a nervous system condition means the neural cell has a genotype, genetic background, and/or phenotype that causes or predisposes an individual to a nervous system condition.
In embodiments involving treatment, a neural cell is exposed to a retrotransposition inhibitor. The term “expose” means bringing the exterior and/or interior of a cell in contact with the inhibitor. A neural cell in culture can be exposed to a retrotransposition inhibitor by adding the inhibitor to the culture medium. A neural cell in a patient can be exposed to an inhibitor by administering the inhibitor to the patient. The routes of administration will vary, naturally, with the particular patient, condition and retrotransposition inhibitor, and can include, e.g., intradermal, transdermal, parenteral, intravenous, intramuscular, intranasal, subcutaneous, percutaneous, intratracheal, intraperitoneal, intratumoral, perfusion, lavage, direct injection, and oral administration and formulation. For exposure of neural cells in an embryo or fetus in a pregnant patient, routes of administration can include in utero or perinatal administration, injections into the maternal vasculature, or through or into maternal organ including the uterus, cervix and vagina, and into embryo, fetus, neonate and allied tissues and spaces such as the amniotic sac, the umbilical cord, the umbilical artery or veins and the placenta. Both bolus and continuous administration of an inhibitor are contemplated. The dose or quantity to be administered, the particular route and formulation, and the administration regimen are within the skill of those in the clinical arts.
In addition, prior to treatment of a patient, the genotype and/or phenotype of the patient can be determined with respect to the nervous system condition being treated. For example, with respect to Rett syndrome, the MeCP2 genotype and the Rett syndrome phenotype of the patient can be determined. Similarly, prior to exposure of neural cells in a patient to a retrotransposition inhibitor, the genotype and/or phenotype of the neural cells in the patient can be determined with respect to the nervous system condition being treated. Also, the genotype and/or phenotype of siblings of the patient, or the genotype and/or phenotype of progeny or children of the patient, can be determined with regard to the nervous system condition being treated. Separately or in combination, these determinations can indicate which patient or neural cells in a patient are to be treated or exposed.
Examples of retrotransposition inhibitors include, but are not limited to, an anti-retroviral drug (such as AZT, tenofovir, or nevirapine); an inhibitor of RNA stability; an inhibitor of reverse transcription (such as ddI or ddC); an inhibitor of L1 endonuclease activity; an inhibitor of DNA repair machinery (such as ATM inhibitor CP466722); a zinc-finger that targets the L1 promoter region; an enzyme that inhibits L1 (such as the protein APOBEC3G); and a repressor that inhibit L1 (such as MePC2 and/or Sox2); and any combination thereof.
The retrotransposition inhibitors can be formulated in neutral or salt forms, and with one or more carriers. Pharmaceutically-acceptable salts include the acid addition salts (formed with the free amino groups of the protein) and those which are formed with inorganic acids such as, for example, hydrochloric or phosphoric acids, or such organic acids as acetic, oxalic, tartaric, mandelic, and the like. Salts formed with the free carboxyl groups can also be derived from inorganic bases such as, for example, sodium, potassium, ammonium, calcium, or ferric hydroxides, and such organic bases as isopropylamine, trimethylamine, histidine, procaine and the like. Upon formulation, solutions can be administered in a manner compatible with the dosage formulation and in such amount as is therapeutically effective to reduce non-LTR retrotransposition. The formulations can be administered in a variety of dosage forms such as injectable solutions, drug release capsules and the like.
The term “carrier” includes any and all solvents, dispersion media, vehicles, coatings, diluents, antibacterial and antifungal agents, isotonic and absorption delaying agents, buffers, carrier solutions, suspensions, colloids, and the like. The use of such media and agents for pharmaceutical active substances is well known in the art. Except insofar as any conventional media or agent is incompatible with the active ingredient, its use in the therapeutic compositions is contemplated. Supplementary active ingredients can also be incorporated into the compositions.
In embodiments involving the assaying of retrotransposition in neural cells, the content of a retrotransposon in a neural cell is compared to the content of the retrotransposon in a control cell. The nature of the control cell depends on the type of comparison. For example, the assayed neural cell can be a mutant neural cell, such as an MeCP2 knockout (“KO”) cell, while the control cell is an MeCP2 wild type neural cell. Alternatively, the control cell can be a non-neural cell, such as a fibroblast, a heart cell, a hepatocyte, a muscle cell, or another non-neural cell.
Embodiments that involve identifying an inhibitor of retrotransposition also compare the content of a retrotransposon in a neural cell to a control cell. In these embodiments, the control cell is typically a similar type of neural cell that has not been exposed to the inhibitor, and can be of the same or comparable genetic background as the neural cell.
In embodiments involving the identification of a neural condition associated with non-LTR retrotransposition by comparison to control cells, the nature of the control cell depends on the type of comparison. For example, the assayed neural cell can be a mutant neural cell, such as an MeCP2 knockout (“KO”) cell, while the control cell is an MeCP2 wild type neural cell. Alternatively, the control cell can be a non-neural cell, such as a fibroblast, a heart cell, a hepatocyte, a muscle cell, or another non-neural cell. In addition, the content of the neural cell can be determined by specific methods such as copy number determination by polymerase chain reaction, or by measuring the hybridization signal of a probe. The identification of a neural condition associated with non-LTR retrotransposition can be part of a subject's diagnostic or treatment regimen, where the diagnosed neural condition is then treated by exposing the subject's neural cells to a transposition inhibitor in an amount sufficient to decrease non-LTR retrotransposition in the neural cells.
The neural cells in various embodiments can be in culture, in an organ, or in an individual.
It is contemplated that any method or composition described herein can be implemented with respect to any other method or composition described herein. Also, the use of the word “a” or “an” when used in conjunction with the term “comprising” in the claims and/or the specification may mean “one,” but is also consistent with the meaning of “one or more,” “at least one,” and “one or more than one.”
Rat NSCs were isolated, characterized and cultured as described (31,32). For neuronal differentiation, cells were cultured in N2 medium (Invitrogen) containing retinoic acid (RA, 1 μM, Sigma) and forskolin (5 μM, Sigma) for 4 days (11). Freshly isolated neuroepithelial cells from time-pregnant midgestation (E11.5) telencephalons from WT and MeCP2 KO sibling mouse embryos from the same background (C57BL/6J) were briefly cultured for 2-3 passages in FGF-2 as described elsewhere (19). Primary skin fibroblasts were isolated from tail biopsy and cultured in DMEM (Invitrogen) with 10% FBS (Invitrogen). Samples of all isolated cells were used for genotyping using previously described primers (33). Plasmid transfections were done by electroporation following the manufacturer's instructions (Amaxa Biosystem).
In Vitro Methylation, Luciferase Assay and siRNA Sequences
The L1 5′UTR-Luc plasmid was methylated by Hpa II (NEB) according to the manufacturer's protocol. Complete methylation was checked by digestion with Hpa II restriction enzyme. Luciferase activity was measured with the Dual-Luciferase reporter assay system (Promega) according to the manufacturer's protocol. Luciferase activity was usually measured 48 h after transfection. A plasmid containing the Renilla luciferase gene was used as an internal control. All the experiments were done at least 3 times independently and transfection efficiency was about 30% for all samples. The siRNAs used in this study were purchased from Dharmacon and used according to the manufacturer's protocol.
ChIP assay was done essentially following the manufacturer's protocol using a ChIP assay kit (Upstate). Antibodies used were anti-MeCP2 (Upstate), Sox2 (Chemicon), and IgG. Purified DNA was amplified by PCR using primers for the rat L1 5′UTR promoter region (L1.3, accession # X03095).
Genomic DNA from NSCs was isolated using standard phenol-chloroform extraction techniques. Subsequently, DNA was digested with the restriction enzyme EcoRI and the bisulfite conversion reaction was performed using the Epitect kit (Quiagen), following manufacturer's instructions. Primers were designed based on the rat L1 sequence Mlvi2, using Methyl Primer Express; primers for L1 converted 5′UTR region: forward 5-AACAAAGTAACACTAGAGATAA-3′ (SEQ ID NO:1) and reverse 5′-TTTGGTGGGAGAATTGGGCT-3′ (SEQ ID NO:2). PCR products were cloned into TOPO TA 2.1 plasmids (Invitrogen) and 40 bacterial colonies were analyzed by sequencing.
Immunofluorescence for EGFP was performed as previously described (11). A non-transgenic animal was used to measure the background fluorescence in the brain and to establish a threshold for detection. Western blotting was carried out using standard protocols with the following antibodies: mouse anti-Actin (1/500, Ambion), rabbit anti-MeCP2 (1/1000, Imgenex or Upstate), and rabbit anti-Sox2 (1/100, Chemicon). All secondary antibodies were purchased from Jackson ImmunoResearch. For co-immunoprecipitation, the Nuclear Complex Co-IP kit (Active Motif) was used, following the manufacturer's protocol with the highest stringency buffers.
Single-Cell Genomic Quantitative PCR (qPCR)
Cell cycle-arrested cells were subjected to a Fluorescent Activated Cell (FACs) sorting in which matched passage number single cells were sorted into an Optical 96-well reaction plate (MicroAmp™-Applied Biosystems, CA) suitable for use in Real Time PCR. The plates containing 1 cell/well were then snap frozen at −70° C. until the day of the qPCR. The qPCR was performed using the protocol available on the manufacturer's website. Briefly, a solution containing forward/reverse primers and SYBR® Green PCR Master Mix was added to the previously sorted cells and the detection of DNA products was carried out in a ABI PRISM® 7700 Sequence Detection System. The SYBR green dye fluoresces only upon binding to the minor groove of double stranded DNA. Thus, the mass of DNA generated by PCR could be quantified and verified by a dissociation (melting) curve analysis that was performed using the Dissociation Curves application software. The primers used for qPCR were designed using specifications on the Primer Express software. Computational estimates using BLAT (May 2006 mouse genome assembly, see Methods) indicated that at least 1,290 endogenous active L1 elements could be detected using ORF2 primers (ORF2-F: 5′-ctggcgaggatgtggagaa-3′ (SEQ ID NO:3), ORF2-R: 5′-cctgcaatcccaccaacaat-3′ (SEQ ID NO:4)). Primers were designed to amplify a product of 52-57 bp. Amplicons of the predicted size were detected in most single cells analyzed (
Multiplex qPCR in Human Tissues
Oligonucleotide PCR primers and TaqMan-MGB probes were designed using Primer Express software (Applied Biosystems). Primers were purchased from Allele Biotech, and probes were purchased from Applied Biosystems. L1 primers were verified using the L1 database (http://llbase.molgen.mpg.de/) and matched at least 140 of 145 identified full length retrotransposition competent L1s. Human tissues were obtained from the NICDH Brain and Tissue Bank for Developmental Disorders at the University of Maryland, Baltimore, Md. Patients were between 17 and 22 years of age. Human genomic DNA was extracted and purified from human tissues using a Blood & Tissue kit (Quiagen), according to the manufacturer's instructions. PCR reactions were carried out using 80 pg of DNA and were verified empirically as amplifying with a CT between 20 and 25 (n=16). Quantitative PCR experiments were performed using an ABI Prism 7000 sequence detection system and Taqman Gene Expression Mastermix from Applied Biosystems. Data analysis was performed using the SDS 2.3 software (Applied Biosystems). The multiplexing reaction was optimized by limiting reaction components until both reactions amplified as completely as each individual reaction. Primer efficiency was verified using a PCR standard curve of plasmid DNA to have a slope of near −3.32. Standard curves of genomic DNA ranging from 2 ηg to 3.2 pg were performed to verify that the 80 pg dilution used was within the linear range.
ORF2-1 primers match 5,543 L1's in the genome, and align with 4,560 L1 sequences in the genome. These primers match 144 of the full length L1's in an L1 database (on the Internet at llbase.molgen.mpg.de/). ORF2-1 probe: ctgtaaactagttcaaccatt (SEQ ID NO:11), ORF2-1F: 5′-tgcggagaaataggaacactttt-3′ (SEQ ID NO:12), ORF2-1R: 5′-tgaggaatcgccacactgact-3′ (SEQ ID NO:13). ORF2-2 primers match 3,447 L1's in the genome and align with 2,918 L1 sequences in the genome. ORF2-2 probe: 5′-aggtgggaattgaac-3′ (SEQ ID NO:14), ORF2-2F: 5′-caaacaccgcatattctcactca-3′ (SEQ ID NO:15), ORF2-2R: 5′-cttcctgtgtccatgtgatctca-3′ (SEQ ID NO:16). L15′UTR probes: L15′UTR-1 primers match 1,299 L1's in the genome, and align with 965 μl sequences in the genome. L15′UTR-1 probe: 5′-aaggcttcagacgatc-3′ (SEQ ID NO:17), L15′UTR-1F: 5′-gaatgattttgacgagctgagagaa-3′ (SEQ ID NO:18), L1 5′UTR-1R: 5′-gtcctcccgtagctcagagtaatt-3′ (SEQ ID NO:19). L15′UTR-2 primers match 1,442 L1's in the genome, and align with 876 μl sequences therein. L15′UTR-2 probe: 5′-tcccagcacgcagc-3′ (SEQ ID NO:20), L15′UTR-2F: 5′-acagctttgaagagagcagtggtt-3′ (SEQ ID NO:21), L15′UTR-2R: agtctgcccgttctcagatct-3′ (SEQ ID NO:22).
Satellite alpha (SATA) primers match millions of tandem copies in the genome, with little sequence variability. SATA-probe: 5′-tcttcgtttcaaaactag-3′ (SEQ ID NO:23), SATA-F: 5′-ggtcaatggcagaaaaggaaat-3′ (SEQ ID NO:24), SATA-R: 5′-cgcagtttgtgggaatgattc-3′ (SEQ ID NO:25). There are 47 copies of the 5S ribosomal RNA gene found in the genome, with 35 probe matches. 5S RNA-probe 5′-agggtcgggcctgg-3′(SEQ ID NO:26), 5S-F: 5′-ctcgtctgatctcggaagctaag-3′ (SEQ ID NO:27), 5S-R: 5′-gcggtctcccatccaagtac-3′ (SEQ ID NO:28). There are 316 primer matches to human endogenous retrovirus H (HERVH), a repetitive non-mobile element in the genome, with 99 probe matches. HERVH-probe: 5′-cccttcgctgactctc-3′ (SEQ ID NO:29), HERVH-F: 5′-aatggccccacccctatct-3′ (SEQ ID NO:30), HERH-R: 5′-gcgggctgagtccgaaa-3′ (SEQ ID NO:31).
MeCP2 KO mice (33) were obtained. The generation of the L1-EGFP animals has been previously described (11). The L1-EGFP transgene was incorporated in the MeCP2 KO background by crossing L1-EGFP males to MeCP2+/− females. Six gender-matched mice, from the same C57BL/6J background, were used per group. Tissues were prepared from adult animals (8 weeks old) as previously described (11). Quantification of EGFP-positive cells in whole brain slices was done by individuals blinded to mice genotypes. EGFP-positive cells were counted in a one-in-six series of sections (approximately 240 μm apart). Images were taken by a z-step of 1 μm using a Biorad radiance 2100 confocal microscope. All experimental procedures and protocols were approved by the Animal Care and Use Committees of The Salk Institute, La Jolla, Calif.
Whole slides containing multiple 40-μm sections of brain from a representative individual from each genetic background were scanned on an iCyte™ fluoro-chromatic imaging cytometer (CompuCyte Corperation, Cambridge, Mass.). The iCyte is a research imaging cytometer based on an inverted platform utilizing lasers, photomultiplier tubes and a scatter detector. The stained slides were scanned at 20× using an argon laser. Sequential images were captured across the entire slide in the X, Y and Z planes. Image processing was completed using iBrowser™ (CompuCyte Corperation) software to stitch the images together into a high-resolution single image of the entire slide. Image Pro Plus™ software (MediaCybernetics, Silver Springs, Md.) was used to extract the images of individual serial sections of brain. The stained cells were manually tagged using Image Pro to identify the X and Y coordinates prior to performing three-dimensional reconstruction. Three-dimensional brain reconstruction was possible using MATLAB software expanded with the MATLAB Image Processing and Virtual Reality Toolboxes. A combination of small, custom programs designed to automate the process of reconstruction from two-dimensional slice images first performed an image contrast calibration on the image stack to increase the illumination of each slice with respect to the slide background. Then, imaged slices were ordered to correspond to a two-dimensional coronal representation of the Allen Reference Atlas. A MATLAB algorithm performed an approximate overlay of the positive cells onto the reference atlas slices, and then the complete data set was converted into Virtual Reality Modeling Language (VRML) format for three-dimensional display. This method was favored over a three-dimensional reconstruction of the actual imaged slices because tears and imperfect alignment of the original slices produced a reconstruction that was more difficult to interpret.
In rat undifferentiated NSCs, the repressor complex in the L1 5′UTR includes the transcriptional factor Sox2 and the histone deacetylase 1 (HDAC1) protein (11), a well-characterized partner of MeCP2 (14,15). MeCP2 was shown to bind to methylated CpG islands in the L1 promoter and reduce retrotransposition in an artificial, non-neural in vitro system (16). Therefore, the role of MeCP2 in the promoter activity of L1 elements in rat NSCs cultured in the presence of FGF-2 was investigated. For that purpose, the human L1 5′UTR promoter region was cloned upstream to the luciferase gene, generating the L1 5′UTR-Luc plasmid (11). Reduction of MeCP2 protein levels by approximately 65%, using specific siRNA against MeCP2 transcripts, led to a 3-fold increase in luciferase activity from the in vitro methylated L1 5′UTR-Luc plasmid (
Immunoprecipitation of MeCP2 in protein extracts isolated from rat NSCs revealed Sox2 enrichment in Western blot analysis. The reverse was also true: immunoprecipitating Sox2 enriched the detection of MeCP2 in NSCs (
Using chromatin immunoprecipitation (ChIP), high levels of MeCP2 were detected in association with the L1 promoter region (i.e., 5′UTR) in rat NSCs compared to differentiated neurons (
To study L1 regulation by MeCP2 in vivo, the MeCP2 KO mouse model was used to compare the brains of the L1-EGFP transgenic mice in WT and MeCP2 KO genetic backgrounds. The L1-EGFP transgenic mice have an L1 indicator cassette that will only activate the expression of the EGFP reporter after retrotransposition (11) (
To better visualize the distribution of EGFP-positive cells in the brain, high-resolution, three-dimensional maps of both MeCP2 KO and WT brains were generated. Analysis of six brains and a representative three-dimensional brain reconstruction indicated that, despite MeCP2 KO brain sections had an average of 3.5-fold more EGFP-positive cells than WT, certain brain structures are more prone to L1 retrotransposition. Specifically, the cerebellum, striatum and olfactory bulb contained 2.2±0.5, 6.7±1.2 and 4.6±1.7-fold more EGFP-positive neurons, respectively, in the MeCP2 KO genetic background compared to WT (
Next asked was whether endogenous L1 retrotransposition is indeed increased in the MeCP2 KO brain. Although the L1-EGFP transgenic animals provide an accessible way to visualize L1 retrotransposition in vivo, they only represent the activity of a single L1 element. In contrast, the mouse genome is estimated to contain at least 3,000 active L1s located on different chromosomes and subject to distinct chromatin context regulation (17,18). To confirm that the L1-EGFP behaves similar to the endogenous L1 elements, a technique was developed based on single-cell genomic quantitative PCR (qPCR) that measures the frequency of mouse L1 sequences (
A control experiment was performed using individual fibroblast cells isolated from the two genetic backgrounds (
Mutations on the MeCP2 gene cause Rett Syndrome (RTT), a severe X-linked neurological disease characterized by impaired motor function; half of RTT patients develop seizures and autistic behavior at different levels of intensity (21,22). To analyze the amount of L1 retrotransposition in tissue samples with clinical diagnostics of RTT and controls, brain and other somatic tissues were obtained from the same individuals. After DNA extraction, a Taqman multiplex qPCR approach was used to compare the number of L1 ORF2 sequences normalized by non-mobile repetitive sequences in the human genome (for instance, the 5S ribosomal RNA repeats), giving a normal distribution of ORF2/5S inverted ratios (
Increased L1 Retrotransposition in NPCs Derived from iPSCs-RTT
Mutations on the MeCP2 gene cause Rett Syndrome (RTT), a severe X-linked neurological disease characterized by impaired motor function; half of RTT patients develop seizures and autistic behavior at different levels of intensity (21, 22) UDATED MS. To determine if L1 retrotransposition can occur in NCPs derived from RTT patients, induced pluripotent stem cells (iPSCs) were generated from a RTT patient's fibroblasts carrying a frameshift MeCP2 mutation and from a control, non-affected healthy individual. The resultant iPSCs were isogenic to the donor cells, providing a valuable opportunity to study early stages of human development in the context of complex genetic diseases (23, 24). Clones of iPSCs derived from the RTT patient and a healthy normal individual (WT) control were pluripotent and able to generate mature, electrophysiologically active neurons in culture. Moreover, RTT-derived neurons showed reduced numbers of glutamatergic synapses and spine density as well as altered intracellular Ca2+ influx, indicating that the human iPSCs system can recapitulate some of the disease onset in culture. Moreover, similar to neuroepithelial cells from mice (
NPCs were then electroporated with an active L1-element tagged with the EGFP reporter construct (L1RE3-EGFP) (25, 26). EGFP expression was detected after 5-7 days in both WT and RTT-derived NPCs (
These data show that MeCP2, together with Sox2, is a potent suppressor of L1 expression in NSCs. Depending on the cellular context, Sox2 protein can function as an activator or repressor (23). The fact that MeCP2 is associated with Sox2 proteins confirms its repressor nature in the maintenance of NSC proliferation, adding a new factor to Sox proteins' molecular versatility. Using two different strategies, it has also been shown that L1 retrotransposition can be modulated by MeCP2 in vivo, characterizing L1 retroelements as genuine MeCP2 targets. First, it was demonstrated that L1 retrotransposition from a transgenic animal carrying an L1-EGFP indicator element was significantly higher in the brains of a MeCP2 KO genetic background compared to a WT sibling animal. Such an approach allowed visualization of de novo L1 retrotransposition in neurons. However, such approach probably underestimates the actual capacity of neuronal retrotransposition, since the engineered L1-EGFP used here represents only one of at least 3,000 active L1 elements in the mouse genome (17,18). Moreover, the L1 EGFP-indicator system does not take into account insertions that truncate or silence the reporter cassette, in trans retrotransposition of Alus or other RNAs (24-26). Second, a new technique was developed, based on single-cell genomic qPCR, to measure relative endogenous L1 sequences. The method is sensitive enough to detect differences between WT and MeCP2 KO genetic backgrounds. MeCP2 KO neuroepithelial cells, but not fibroblasts, have more L1 sequences in the genome when compared to WT cells. These results indicate that endogenous mouse L1 elements can retrotranspose during development but may be restricted to the nervous system. The qPCR data reflect a snapshot of a specific moment during brain development. If a portion of the cells that support retrotransposition survives and the rates of retrotransposition are similar during the entire development, the impact of L1 insertions may be significant, especially in the MeCP2 KO genetic background.
To study L1 retrotransposition during early stages of human development, iPSCs from a RTT patient with a MeCP2 frameshift mutation and from a normal control were derived. NPCs derived from both WT and RTT iPSCs could support L1-EGFP de novo insertions. However, RTT-NPCs showed a higher frequency of L1 retrotransposition compared to WT control cells, confirming that MeCP2 is a repressor of active human L1 retrotransposons.
A similar qPCR experiment extended these observations to human brain samples from heterozygote females RTT patients compared to normal controls and other somatic tissues. However, because mature human brain tissues, sampling from a non-homogeneous population of cells, including neurons and astrocytes, could mask more dramatic differences between neurons and non-neuronal cells. Moreover, the effects of X-chromosome inactivation status, allowing the expression of the WT MeCP2 allele only in some neurons, are likely to contribute to a subtle difference between L1 ORF2 sequences in RTT patients and controls. Finally, in the human brain, cells that survived brain development were analyzed, whereas the embryonic neuroepithelial cells isolated from the mouse will face a strong selection process wherein many cells undergo programmed cell death.
It has been hypothesized that DNA methylation and methyl-binding proteins protect the genome against retrotransposition in germ cells (27). However, the discovery that Piwi/piRNAs can suppress transposition in germ cells suggested that it may not be the only mechanism (28). The data here provides strong evidence for a role of DNA methylation-dependent MeCP2 activity in controlling transposable elements activity. Interestingly, such activity may be specific to NSC, de-repressing retroelements during neuronal differentiation, raising the question why neurons support L1 retrotransposition. Recently, re-activation of MeCP2 expression in both embryonic and adult KO mice led to prolonged life span and delayed onset or reversal of certain neurological symptoms (29,30). Since L1 insertions are genetically stable, the new insertions may have a small contribution to the reversible RTT syndrome phenotype in mouse. The high rates of neuronal retrotransposition in the MeCP2 KO mice and RTT brains may be a consequence, rather than a cause, of the disease process. However, a more effective L1 silencing may have an important impact as a modulator of neighboring gene expression. L1 sequences may function as master regulators of chromatin structure through heterochromatin silencing of discrete chromosomal regions close to neuronal genes. In that context, new somatic insertions in the MeCP2 KO mice brain and in RTT brains may contribute to the epigenetic status of neurons, affecting neuronal networks and behavior.
The data presented here are the first to demonstrate intrinsic tissue-specific somatic genetic variation outside the immune system in humans. These findings add a new layer of complexity to the understanding of genomic plasticity, revealing that neuronal genomes can accommodate somatic mutations caused by L1 retrotransposition. These observations have direct implications for genetic, non-heritable neurological diseases and individual responses to drug treatment or environmental cues.
Long Interspersed Element-1 (LINE-1 or L1) retrotransposons have dramatically impacted the human genome. Retrotransposons constitute approximately 40% of the mammalian genome and play an important role in genome evolution. Their prevalence in genomes reflects a delicate balance between their further expansion and the restraint imposed by the host. L1s must retrotranspose in the germ-line or during early development to ensure their evolutionary success. Yet the extent to which this process impacts somatic cells is poorly understood. It has been previously demonstrated that engineered human L1s can retrotranspose in adult rat hippocampus progenitor cells (NPCs) in vitro and in the mouse brain in vivo (34). Here it is demonstrated that NPCs isolated from human fetal brain and NPCs derived from human embryonic stem cells (hESCs) support the retrotransposition of engineered human L1s in vitro. Furthermore, a quantitative multiplex polymerase chain reaction is described that detects an increase in the copy number of endogenous L1s in the hippocampus and in several regions of adult human brains when compared to the copy number of endogenous L1s in heart or liver genomic DNAs from the same donor. The data indicate that de novo L1 retrotransposition events may occur in the human brain and, in principle, have the potential to contribute to individual somatic mosaicism.
Fetal hCNS-SCns lines (36) and hESCs (57,59) were cultured as previously described. Neural progenitors were derived from hESCs as previously described (47,60). NPCs were transfected by nucleofection (Amaxa Biosystems), and either maintained as progenitors in the presence of FGF-2 or differentiated as previously described (47). Cells were transfected with L1s containing an EGFP retrotransposition cassette in pCEP4 (Invitrogen) that lacks the CMV promoter and contains a puromycin resistance gene (40). The hCNS-SCns lines FBR BR1, BR4 and BR3 were cultured as previously described and were a kind gift from Stem Cells Inc. (Palo Alto, Calif.) (36). hCNS-SCns, also known as huCNS-SC (human CNS stem cells grown as neurospheres), were derived from fetal brain by FACS using the following cell surface markers: (CD133)+, (5E12)+, (CD34)−, (CD45)−, and CD24−/lo. This combination of markers enriches for progenitor neurosphere-initiating cells capable of differentiating into cells of both the neuronal and glial lineages (36). The hCNS-SCns were cultured in X-Vivo 15 media (Lonza Bioscience) supplemented with 20 ng/mL FGF-2, 20 ng/mL epidermal growth factor (EGF), 10 ng/mL leukemia inhibitor factor (LIF), N2 supplement, 0.2 mg/mL heparin, and 60 mg/mL N-acetylcysteine. For differentiation experiments, hCNS-SCns were dissociated using Liberase Blendzymes (Roche) and plated on laminin/polyornithine-coated plates. Mitogens were withdrawn and cells were differentiated by retrovirus-mediated transduction with Neurogenin 1 (NGN1), a pan-neuronal helix-loop-helix transcription factor. NGN1 was a kind gift from Dr. David Turner and was cloned into a murine Moloney leukemia retrovirus-based plasmid and expressed under the control of the ubiquitously expressed CAG promoter as previously described (63). Virus was made in human embryonic kidney 293T cells and collected by ultracentrifugation (63). hCNS-SCns were infected 48 hrs before differentiation at an approximate efficiency of 70% and allowed to differentiate for 3-4 weeks. Karyotype analysis of hCNS-SCns lines indicated grossly normal karyotype (
Primary human neo-natal dermal fibroblasts (cat# CC-2509) and primary adult astrocytes (cat# CC-2565) were commercially obtained and cultured using instructions provided by the manufacturer (Lonza Bioscience). Karyotype and fluorescence in situ hybridization analyses were performed at Cell Line Genetics (Madison, Wis.). The hESC lines HUES6 and H9 were cultured as previously described (on world wide web at mcb.harvard.edu/melton/HUES/) by the Gage group (59). Under these experimental conditions, hESCs exhibited a grossly normal karyotype (
The NIH-approved hESC lines (WA07 (i.e., H7), WA09 (i.e., 119), WA13B (i.e., H13B), and BG01) were cultured as previously described by the Moran group (57). Briefly, hESCs were grown on irradiated MEFs and then were passaged by manual dissection using the StemPro EZPassage passaging tool (Invitrogen). A protocol based on Zhang et al. was used to derive NPCs (60). hESCs first were seeded in a suspension culture dish (Corning) in hESC media lacking FGF2 to generate embryoid bodies. After 4-6 days, the resulting embryoid bodies were seeded in a Petri dish coated with gelatin and cultured in NeuroSphere (NS) media for 14-16 days. NS culture medium contains DMEM F12 (Invitrogen) supplemented with 20 ng/ml FGF2, N2 supplement, and 2 μg/mL Heparin (Sigma). After 14-16 days, the resulting rosettes were picked manually, trypsinized, and then plated to form neurospheres. Neurospheres were passaged by single cell dissociation using a pulled Pasteur pipette once a week. To induce differentiation, a single cell suspension of NPCs was plated on polyornithine-coated plates in DMEM/F12 with N2 and 1% FBS and allowed to differentiate for 6 days. NPCs derived using either protocol expressed neural stem cell markers (
Cells were transfected with L1s containing an EGFP retrotransposition cassette in a modified version of pCEP4 (Invitrogen) that lacks the CMV promoter and contains a puromycin resistance gene instead of a hygromycin selection gene (40). Prior to transfection, DNAs were checked for superhelicity by electrophoresis on 0.7% agarose-ethidium bromide gels. Only highly supercoiled preparations of DNA (>90%) were used in transfection experiments. L1RP is a full-length retrotransposition-competent L1 whose expression is driven by the native L1 5′ UTR (64,40). LRE3 is a previously described full-length retrotransposition-competent L1 (65). JM111/L1RP is a derivative of L1RP containing two missense mutations (RR261-262AA) in the RNA binding domain of the ORF1-encoded protein that reduce L1 retrotransposition by greater than three orders of magnitude (38,40). In UB-LRE3 and UB-JM111, the expression of the L1 is driven by the ubiquitin C promoter (a 1.2-kb fragment of the human UBC gene nucleotides 123964272-123965484 from chromosome 12). All constructs contained the CMV-EGFP retrotransposition cassette (40). The LRE3-neo and LRE3-blasticidin constructs contained the mneoI or blasticidin retrotransposition cassettes, respectively (38,44).
hCNS-SCns and HUES6- and H9-derived NPCs (one passage after neural rosette selection) were transfected by Nucleofection using the Amaxa rat NSC nucleofector solution and program A-31. The transfection efficiency was determined using an EGFP-expressing plasmid control 2 days post transfection by FACS analysis. The transfection efficiency ranged from 50-70% for hCNS-SCns and from 50-80% for hESC-derived NPCs. Cells were cultured as progenitors in the presence of mitogens. For differentiation studies, cells were dissociated and plated for differentiation 18 days after the initial transfection. H7-, H13B-, H9-, and BG01-derived NPCs were transfected using the Amaxa mouse NSC nucleofector solution and program A-33 and cultured as progenitors. In some experiments puromycin (0.2 μg/mL) was added 2 days post transfection for 5-7 days prior to scoring for retrotransposition. Primary human fibroblasts and astrocytes were transfected using Fugene6 (Roche) per manufacturer's instructions. Cells were monitored for EGFP expression by fluorescence microscopy. For FACS analysis, cells were dissociated and analyzed on a Becton-Dickinson LSR I in the presence of 1 μg/mL propidium iodide for live/dead cell gating. All assays were performed in triplicate. JM111/L1RP transfected cells were used as a negative control for gating purposes. The criterion to determine L1 insertional silencing was a 10-fold increase in EGFP expression after the addition of 500 nM trichostatin-A for 16 hours on day 7 post-transfection with the L1 construct. NPCs transfected with L1s containing the mneoI or blasticidin retrotransposition indicator cassettes were subjected to either G418 or blasticidin selection beginning 4-7 days post-transfection. Cells were selected with 50 μg/ml of geneticin (G418, Invitrogen) for 1 week and with 100 μg/ml of G418 the following week, or with 2 μg/mL of blasticidin (InvivoGen) for 2 weeks.
HUES6-derived NPCs were electroporated with the LRE3-EGFP pCEP4 plasmid, allowed to proliferate for 7 additional days, and subsequently differentiated. Whole-cell perforated patch recordings were performed on EGFP-expressing cells after 10 weeks of differentiation. The recording micropipettes (tip resistance 3-6 MΩ) were tip-filled with internal solution composed of 115 mM K-gluconate, 4 mM NaCl, 1.5 mM MgCl2, 20 mM HEPES, and 0.5 mM EGTA (pH 7.4) and then back-filled with the same internal solution containing 200 μg/ml amphotericin B. Recordings were made using Axopatch 200B amplifier (Axon Instruments). Signals were sampled and filtered at 10 kHz and 2 kHz, respectively. The whole-cell capacitance was fully compensated, whereas the series resistance was uncompensated but monitored during the experiment by the amplitude of the capacitive current in response to a 5-mV pulse. The bath was constantly perfused with fresh HEPES-buffered saline composed of 115 mM NaCl, 2 mM KCl, 10 mM HEPES, 3 mM CaCl2, 10 mM glucose and 1.5 mM MgCl2 (pH 7.4). For current-clamp recordings, cells were clamped at ˜−60 to −80 mV. For voltage-clamp recordings, cells were clamped at −70 mV. All recordings were performed at room temperature. Amphotericin B was purchased from Calbiochem. All other chemicals were from Sigma.
Cells were fixed in 4% paraformaldehyde, and immunocytochemistry was performed as previously described (57,66). Antibodies and dilutions were as follows: βIII tubulin, mouse monoclonal, 1:400 or rabbit polyclonal, 1:500 (both Babco/Covance); Map (2a+2b), mouse monoclonal, 1:500 (Sigma); GFAP rabbit polycolonal, 1:300 (DAKO); GFAP, guinea pig polyclonal, 1:1000 (Advanced Immunochemical); Nestin, mouse monoclonal, 1:800 (Chemicon); Musashi-1, rabbit polyclonal, 1:200 (Chemicon); Sox1, rabbit polyclonal, 1:200 (Chemicon); Sox1, goat polyclonal, 1:200 (R&D); TH, rabbit polyclonal, 1:500 (Pel-Freez); Ki-67, rabbit monoclonal, 1:500 (VectorLabs); Sox2, rabbit polyclonal, 1:500 (Sigma), Sox3, rabbit polyclonal, 1:500 (a generous gift from Dr. M. W. Klymkowsky, Denver, Colo.). Secondary antibodies were purchased from Jackson ImmunoResearch or Invitrogen and all were used at 1:250. Cells were imaged using a CARVII spinning disk confocal imaging system (BD).
Luciferase activity was measured with the Dual-Luciferase reporter assay system according to instructions provided by the manufacturer (Promega). In all assays, a plasmid expressing the Renilla luciferase gene was used as an internal control. The assays were replicated independently at least three times. The L1 5′ UTR luciferase construct has been previously described (34,67). The Synapsin-1 promoter region was a kind gift from G. Thiel. All promoters were subcloned into the pGL3-basic vector (Promega).
Southern blotting was performed following standard protocols (68) on hCNS-SCns line FBR4 collected 3 months post-transfection with the L1RP pCEP4 plasmid. Briefly, 20 μg of genomic DNA was digested with CM, a restriction enzyme that digests the tagged-L1 both at 5980 bp (20 bp 5′ to start of the retrotransposition cassette) and at 8517 bp (in the 3′ UTR). The L1.3 plasmid containing the indicator cassette yields a 2547 bp band, whereas a retrotransposed L1 integrated into a genomic sequence that lacks the intron in the EGFP expression cassette yields a 1645 bp band. This methodology collapses all the tagged L1 insertions into a single imaged band. The probe was a full-length EGFP DNA fragment that was radioactively labeled with γ-32P-dCTP using the Random Prime Labeling Kit according to instructions provided by the manufacturer (Roche).
hESC or NPCs were harvested and lysed as previously described with 1 ml of 1.5 mM KCl, 2.5 mM MgCl2, 5 mM Tris-Hcl pH 7.4, 1% deoxycolic acid, 1% Triton X-100, and 1× Complete Mini EDTA-free Protease Inhibitor cocktail (Roche) (41). Cell debris was removed by centrifugation at 3,000×g at 4° C. for 5 minutes, and 10% of the supernatant fraction was saved (i.e., Whole Cell Lysate or WCL fraction). A sucrose cushion then was prepared with 8.5% and 17% w/v sucrose in 80 mM NaCl, 5 mM MgCl2, 20 mM Tris-Hcl pH 7.5, and 1 mM DTT, which was supplemented with 1× Complete Mini EDTA-free Protease Inhibitor cocktail (Roche). WCLs were centrifuged at 39,000 rpm for 2 hours at 4° C. using a Sorvall SW-41 rotor. After centrifugation, the pelleted material (i.e., the ribonucleoprotein particle (RNP) sample) was resuspended in 50 μL of purified water supplemented with 1× Complete Mini EDTA-free Protease Inhibitor cocktail (Roche). Total protein concentration was determined by Bradford assay according to instructions provided by the manufacturer (BioRad). WCL and/or RNP samples (8 μg of each sample) were loaded on 10% SDS-PAGE gels (BioRad). Antibodies and dilutions were as follows: Anti-ORF1, rabbit polyclonal antibody, 1:10,000 dilution (a generous gift from Dr. Thomas Fanning); anti-S6 ribosomal protein, rabbit polyclonal antibody, 1:1,000 dilution (Cell Signaling); anti-Sox3 antibody, rabbit polyclonal, 1:1,000 dilution (a generous gift from Dr. M. Klymkowsky); anti-Sox1, goat polyclonal, 1:500 dilution (R&D). All HRP conjugated secondary antibodies were used at a 1:20,000 dilution (abeam).
Ribonucleoprotein particles were isolated and analyzed as previously described (41). Luciferase assays were performed as previously described (34). Chromatin immunoprecipitation was performed utilizing primers towards the L1 5′UTR and a ChIP assay kit (Upstate/Millipore) as per manufacturer's protocol.
RNA was isolated from various cell and tissue types with RNABee (Tel-test Inc., Friendswood Tex.) following the manufacturer's directions. RNA quality was verified by gel electrophoresis, and cDNA was synthesized using the cells-to-cDNA H kit (Ambion/Applied Biosystems) per manufacturer's instructions. Quantitative RT-PCR was performed with the same ORF2 #1 Taqman primer/probe combination utilized for genomic DNA analysis. Standardization was performed using the beta-actin Taqman Detection Kit (Applied Biosystems). RT-PCR analysis was performed using the following primers towards ORF1:
and PCR products from RT-PCR and QPCR reactions were cloned into the PCR TOPO II vector (Invitrogen) and sequenced.
Fetal tissues were obtained from the Birth Defects Research Lab at the Univ. of Washington. Bisulfite conversions were performed by manufacturer's instructions utilizing the Epitect kit (Quiagen). BLASTN (available on the Internet at blast.ncbi.nlm.nih.gov/Blast.egi) was used to align sequences to a database of full-length L1s. Fetal tissues were obtained from donations resulting from voluntary pregnancy terminations and were collected by the Birth Defects Research Lab at the University of Washington, Seattle, Wash. (NIH HD 000836). Genomic DNAs from 80-day-old female and 82-day-old male fetuses were isolated from brain and skin tissue using standard phenol-chloroform extraction techniques. The resulting DNA was digested with the restriction enzyme DraI and the bisulfite conversion reaction was performed using the Epitect kit according to instructions provided by the manufacturer (Qiagen). The bisulfite conversion was performed two times, consecutively, to achieve a CpG conversion rate of >90% in the LINE-1 repeat regions. The L1 5′ UTR contains a CpG island that has a G+C content greater than 60% and a CpG frequency ratio of greater than 0.6 (observed/expected CpGs) (16). The sequence of all full-length Ta-subfamily L1s was used to design oligonucleotide primers that allowed us to amplify a 363 bp region from a constellation of L1 s, which included both young Ta-1 and older subfamilies of the L1Hs/L1PA1 family such as Ta-0 due to the high degree of L1 sequence conservation (42,43). Thus, the following primers were designed against sequences in the L1 5′ UTR using Methyl Primer Express:
and the resulting PCR products were cloned into the TOPO TA 2.1 plasmid (Invitrogen) and 100 bacterial colonies were sequenced from each tissue sample.
After bisulfite treatment, BLASTN was used to align the L1 5′ UTR sequences to a database of full-length L1s with two intact open reading frames that was extracted from the May 2004 assembly of the human genome (hg17). The BLASTN alignment used a mismatch penalty of −1 and a match reward of +1. The best match for each brain or skin sequence to the genomic L1 database was determined (the database consisted of known RC-L1s). The alignment excluded cytosine nucleotides in the L1 database to prevent bias due to the bisulfite conversion. The fraction of CpG sites that were unmethylated was calculated by computationally comparing CpG dinucleotides in the L1 database to the corresponding sequences from the brain and skin samples. The fraction of CpG sites converted by the bisulfite analyses was measured as the proportion of TG dinucleotides in brain and skin sequences at CpG sites in the genomic L1 database to total number of CpG sites in the region. To determine differences in methylation between brain and skin L1 s, a cumulative distribution (CDF) plot was generated for all the sequences that aligned above an alignment cutoff. The alignment cutoff was one standard deviation below the mean of the alignment identity score for all sequences aligned. Conversion efficiency was assessed by analyzing the conversion rate at genomic cytosine nucleotides that were not upstream of a guanine nucleotide. The same analysis was carried out for all possible dinucleotides and possible conversions of the first nucleotide. A two-sample Kolmogorov-Smirnov test indicated a statistically significant difference between skin and brain. Comparison of each dinucleotide pair within each sequence revealed a statistically significant difference in the CpG bisulfite conversion efficiency (i.e., CpG to TpG nucleotide changes) between the brain and skin samples but not in any of the other dinucleotide pairs in the L1 5′ UTR (
Chromatin immunoprecipitation (ChIP) was performed following the manufacturer's protocol and a ChIP assay kit (Millipore/Upstate). The protocol was modified such that antibody hybridization was performed twice to decrease background. Antibodies used were anti-Sox2 (Chemicon), anti-MeCP2 (Chemicon), and IgG. Resultant purified DNA was amplified with the following primers designed to the active LRE3 element (64,65), amplifying the SOX2 binding sites:
and towards the CpG island region:
the Sox2 primers were used with ChIP utilizing antibodies towards SOX2, and primers designed towards the CpG island were utilized with MeCP2 immunoprecipitated DNA.
Adult human tissues were obtained from the NICDH Brain and Tissue Bank for Developmental Disorders (University of Maryland, Baltimore, Md.). Taqman probes and primers were designed using L1 Base (on Internet at llbase.molgen.mpg.de/) and copy number estimates were based on the UCSC genome browser (on Internet at genome.ucsc.edu). Experiments were performed on an ABI Prism 7000 sequence detection system (Applied Biosystems). For each tissue, three separate tissue samples were extracted and considered as repeated measures. Whole genome size was estimated based on the equation, cell genomic DNA content=3*109(#bps)*2(diploid)*660 (MW 1 bp)*1.67*1012 (weight 1 dalton), resulting in the approximation that one cell contains 6.6 pg genomic DNA (61). Therefore, the 80 pg of genomic DNA utilized per reaction is derived from approximately 12 cells. Inverse PCR was performed as previously described (34,57). Genomic DNA from transfected NPCs and hCNS-SCns was isolated using the DNeasy Blood & Tissue kit according to instructions provided by the manufacturer (Qiagen). Genomic DNA was collected 8 days post-transfection from NPCs and 2 months post-transfection from hCNS-SCns. To assay for removal of the intron from the retrotransposition indicator cassette, 200 ng of genomic DNA was used in a 25 μL PCR reaction with the following primers: EGFP968s and EGFP1013 as (in experiments conducted with EGFP-tagged L1s), NEO437s and NEO1808 as (in experiments conducted with mneoI-tagged L1s), or Blast-Fw (5′-GCTGTCCATCACTGTCCTTCA (SEQ ID NO:40)) and Rv (5′-CCATCTCTGAAGACTACAGCG (SEQ ID NO:41)) primers (in experiments conducted with blasticidin-tagged L1s). PCR cycling conditions were described previously, and the blasticidin cycling conditions were identical to those utilized for the mneoI PCR (38,40). The marker ladder utilized in all gel pictures is a 1 kB plus ladder (Invitrogen, catalog#10787-081). Sequence analysis of all L1 PCR products was performed with the USCS genome browser (on world wide web at genome.ucsc.edu) and Repeatmasker (on world wide web at repeatmasker.org).
Initially fluorescent activated cell sorting (FACS) was used to isolate EGFP-positive and EGFP-negative NPCs 18 days post-transfection. However, no clones grew to confluence in a 96-well plate, and after whole genome amplification and inverse PCR (34,44,57) only a single retrotransposition event was characterized (Table 3). Single EGFP-positive cells were sorted into 96-well plates and allowed to proliferate for 6-8 weeks. Cells then were trypsinized using 10 mL of Tryple reagent (Invitrogen). Since the DNA yield for the single colonies was very low, whole genome amplification was performed using the Genomiphi kit according to instructions provided by the manufacturer (GE Life Sciences).
Cells harboring retrotransposition events derived from native, full-length, LRE3-tagged element insertions were grown in NPC medium for 18-20 days. The resulting cells were dissociated with trypsin and sorted on a Becton-Dickenson FACscan. A total of 40,000 EGFP-positive cells and EGFP-negative cells was sorted and expanded in culture for three passages. This experiment was replicated independently using a second sample of independently derived NPCs. As expected, genomic DNA from the EGFP-positive cells yielded a PCR product corresponding to the retrotransposed EGFP gene (
Inverse PCR (IPCR) was performed as previously described (34,57). Briefly, 5-10 μg of genomic DNA was digested overnight with either SspI or XbaI. The digested DNA then was ligated under dilute conditions in a final volume of 1 mL with 3,200 U of T4 DNA ligase (NEB) overnight at 4° C. The circular ligated DNA was concentrated to 50 μL using a Microcon 100 column (Millipore), and then was subjected to IPCR using previously described conditions (34,57). PCR products were gel-isolated, cloned into the TOPO TA 2.1 plasmid (Invitrogen) and sequenced. Identification of the L1 pre-integration sites and other DNA sequence analyses were performed using the UCSC genome browser (March 2006 assembly) (69).
Oligonucleotide PCR primers were purchased from Allele Biotech and TaqMan-MGB probes from Applied Biosystems and were designed using Primer Express software (Applied Biosystems). L1 primers were verified using the L1 database L1 Base and matched a minimum of 140 of 145 full-length L1s with two intact open reading frames in the database. Human tissues were obtained from the NICDH Brain and Tissue Bank for Developmental Disorders (University of Maryland, Baltimore, Md.). Donors were between 17 and 45 years old. Dissection of the subventricular zone (SVZ), dentate gyrus (DG), CA1 and CA3 regions was performed from human brain sections. Human genomic DNAs were extracted and purified from tissues using the DNeasy Blood & Tissue kit according to instructions provided by the manufacturer (Qiagen). PCR reactions were carried out using 80 pg of genomic DNA and were verified empirically as amplifying with a cycle threshold (CT) value between 20 and 25 (n=16). Whole genome size was estimated based on the equation, cell genomic DNA content=3*109(#bps)*2(diploid)*660 (MW 1 bp)*1.67*1012 (weight 1 dalton), resulting in the approximation that one cell contains roughly 6.6 pg genomic DNA (61).
Quantitative PCR experiments were performed using an ABI Prism 7000 sequence detection system and Taqman Gene Expression Mastermix (Applied Biosystems). Data analysis was performed with SDS 2.3 software (Applied Biosystems). The multiplexing reaction was optimized by limiting reaction components until both reactions amplified as well as each individual reaction. Standard curves of genomic DNA ranging from 2 ng to 16 pg were performed to verify the 80 pg dilution used is within the linear range of the reaction. Primer efficiency and multiplexing effectiveness was verified by linear regression to the standard curve and indicated a slope near −3.32, representing acceptable amplification of both PCR products and matched primer efficiencies. ORF2 probes were conjugated to the fluorophore label VIC and all other probes were conjugated with 6FAM. For the control assay depicted in
For each tissue type—hippocampus, cerebellum, heart, and liver—three individual tissue samples were taken and considered as repeated measures from each of the three individuals. The four tissues were compared utilizing a repeated measures one-way ANOVA with a Bonferroni correction. Statistically significant findings (p<0.05) are indicated in
The protocol of Frisen (Spalding, et al. (2005) Cell 122(1): 133) has been optimized for nuclei isolation from very small amounts of tissue (<0.1 g) from fresh-frozen tissue samples stored at −80 C. Nuclei isolation is performed quickly as tissue is beginning to thaw. All solutions' are stored at 4° C. and the procedure is performed on ice. Frozen tissue (<0.5 g) is placed in 0.5 mL Lysis Buffer [0.32 M sucrose, 5 mM CaCl2, 3 mM Mg acetate, 0.1 M EDTA, 10 mM Tris pH 8.0, 0.1% triton, 1 mM DTT], triturated slightly, and transferred to a small dounce homogenizer. Homogenization is accomplished in 10-12 strokes, after which the homogenate is transferred to 2 mL of Sucrose Buffer [1.8 M sucrose, 3 mM Mg acetate, Tris pH 8.0, 1 mM DTT]. The homogenizer is then rinsed with an additional 0.5 mL of Lysis buffer which is also added to the Sucrose Buffer containing homogenate. The mixture is then combined well by several inversions. Nuclei are separated from other tissue debris by centrifugation on a sucrose cushion. The 3 mL homogenate mixture from above is layered onto a cushion of 6 mL sucrose solution in a conical ultracentrifuge tube (Beckman part #358126). It is critical that layering be performed slowly and carefully, furthermore care is taken when loading these tubes into an ultracentrifuge rotor (Beckman SW28) so that a sharp interface between the cushion and the homogenate is maintained. Centrifugation at 12:9 K rpm, 4° C., for 2 hrs leads to a nuclei pellet in the centrifuge tube and cellular debris at the sucrose interface. After removing the sucrose cushion and cellular debris, nuclei are resuspended on 0.5 mL Nuclei Storage Buffer [15% sucrose, 2 mM MgCl2, 70 mM KCl, 10 mM Tris pH 8.0, 1 mM DTT, 1× protease inhibitor cocktail with EDTA (Roche)]. Nuclei in this buffer are stored at 4 C until sorting, typically <4 days.
Staining for nuclear antigens (e.g. NeuN) can be performed at this point following standard protocols. Otherwise, or afterward, stored nuclei are resuspended and diluted 1:5 in PBS containing 10 mM propidium iodide, then filtered through 20 um nylon mesh. Sorting is performed using a FACS Vantage SE DiVa (Becton-Dickenson). Gates are adjusted to obtain G1 nuclei with a diploid DNA content. Further dilution of the nuclei prep using PBS is performed if the solution is to concentrated. Sorting of highly concentrated preps leads to occasional sorting of debris rather than nuclei into wells. Nuclei are sorted into 96 well plates that are suitable for future analysis using quantitative PCR. Aside from actual sorting, care is taken to keep these plates clean by performing subsequent steps in a laminar flow hood. Before sorting, 5 μL of TE buffer (10 mM Tris pH 8.0, 1 mM EDTA) is placed in each well using a high throughput reagent dispenser (Multidrop 384, Thermo Scientific). The last column (8 wells) of the plate does not typically contain sorted nuclei and these wells are analyzed as DNA-free controls. Twelve plates are typically sorted for each tissue.
Quantitative PCR (qPCR)
Multiplex qPCR is performed using Taqman methodology (Applied Biosystems). Primer probe combinations are listed in an additional table. A master mix is prepared so that when 10 μL of mastermix is added to 5 μL containing a single nuclei in TE the final concentration of reagents is: 1× gene expression mastermix (Applied Biosystems), 1 mM control primers (e.g. 5S RNA), 0.1 mM experimental primers (e.g. L1 Orf2), and 0.2 mM taqman probes. The fluorophore 6FAM is typically used for the less abundant template (e.g. 5S RNA) and the fluorophore VIC is typically used for the more abundant template (e.g. Orf2). Mastermix is prepared and dispensed using the high throughput reagent dispenser in a laminar flow hood. The high throughput reagent dispenser requires an additional “dead” volume of 5 mL mastermix, this dead volume seems stable for several days and is often combined with “fresh” mastermix on subsequent days to reduce expenses.
The following primer pairs were used for quantitative PCR assays as well as the primer pairs previously listed in the methods section:
qPCR results are obtained as a “Ct” value. This is the cycle number at which amplification of each template crosses a defined threshold. The threshold is defined as a point during which exponential amplification is observed, typically at 0.1 arbitrary fluorescence units. A dCt value is obtained for each well by subtracting the Ct of the more abundant template from the Ct of the less abundant template (e.g. dCt=Ct5S−CtORF2). In order to calculate the number of de novo RT events one must calculate a fold-change in dCt in an experimental (e.g. brain, iPS-derived neurons) versus a control (e.g. heart, small intestine, pre-iPS fibroblasts). The first component of calculating fold-change is a “ddCt,” obtained by subtracting the dCt from one control nucleus from one experimental nucleus. These may be paired individually either randomly or by rank; or an average of control nuclei can be used for each experimental nucleus. Fold-change is calculated as 2̂−ddCt; 2 assume perfect efficiency in the reactions, if the reactions are found to be less efficient the value 2 should be changed accordingly (e.g. 95% efficiency=1.9̂−ddCt). The number of initial L1 sequences in the reference genome is obtained using BLAT (UC Santa Cruz), and de novo events are calculated based on a fold-change from this reference value.
The human nervous system is complex, containing approximately 1015 synapses with a vast diversity of neuronal cell types and connections that are influenced by complex and incompletely understood environmental and genetic factors (35). Neural progenitor cells (NPCs) give rise to the three main lineages of the nervous system: neurons, astrocytes, and oligodendrocytes. To determine if human NPCs can support L1 retrotransposition, human fetal brain stem cells (hCNS-SCns) (
A low level of L1RP retrotransposition, averaging 8-12 events per 100,000 cells, was observed in three different hCNS-SCns lines (BR1, BR3 and BR4;
Next two different protocols were used to derive NPCs from five human embryonic stem cell lines (hESCs;
Characterization of EGFP-positive neurons revealed that some expressed subtype-specific markers (tyrosine hydroxylase (
Several studies have reported an inverse correlation between L1 expression and the methylation status of the CpG island in their 5′ UTRs (48,49). Thus, bisulfite conversion analyses on genomic DNAs derived from matched brain and skin tissue samples from two 80- to 82-day-old fetuses were performed (
Previous data suggested that Sox2 and MeCP2 could associate with the L1 promoter and repress L1 transcription under some experimental conditions (34,50). Two putative SRY/Sox2 binding sites are located in the L1 5′ UTR immediately 3′ to the CpG island (
Although NPCs are useful to monitor L1 activity, they only allow monitoring a single L1 expressed from a privileged context. By comparison, the average human genome contains ˜80-100 active L1s whose expression may be affected by chromatin structure (37). Therefore, a quantitative multiplexing PCR strategy was developed to investigate endogenous L1 activity in the human brain, hypothesizing that active retrotransposition would result in increased L1 content in the brain as compared to other tissues (
To independently corroborate the observed increase in L1 copy number in the hippocampus and cerebellum samples, 80 pg of liver and heart genomic DNA were spiked (approximately 12 genomes) from individual 1846 with a calculated quantity of L1 plasmid, then the multiplexing approach was repeated to assay ORF2 quantity relative to 5S rDNA internal control (
Experimental results are summarized in the tables referred to herein.
Table 1 provides the variation of L1 ORF2 sequences in the human brain and heart tissue from normal and RTT patients; numbers correspond to inverse CT values and were obtained from the multiplex qPCR strategy.
Table 2 provides the results of L1 retrotransposition assays in hESC-derived NPCs. From left to right, column 1 indicates the hESCs cell line from which NPCs were derived, column 2 indicates the lab where the experiments were performed, column 3 indicates if selection (puromycin 0.2 μg/mL) was used in the assay, and column 4 indicates the percentage of EGFP-expressing cells with s.d. The variation likely depends on the individual NPC preparation, the differentiation protocol, whether the NPCs were subjected to puromycin selection prior to assaying for retrotransposition, and if the resultant retrotransposition event was subjected to silencing (indicated by the (* in column 1)). It was observed that L1 retrotransposition events could be efficiently silenced in some hESC-derived NPCs (column 1, marked H13B*). This silencing could be overcome by treating the cells with histone deacetylase inhibitors and it may reflect idiosyncrasies that arise during the differentiation protocol. In the table: a—Gage (G) or Moran (M) groups. HUES6 is a private cell line all others are federally approved; b—puromycin, 0.2 ugfmL; (*) these NPC derivations exhibited silencing of the Li insertions, whereas other NPCs derivations did not; (*) either a JM1 il/L1RP construct or untransfected samples (in triplicate) were used to determine baseline background fluorescence; UB=where the ubiquitin C promoter drives the expression of the LI. In all other experiments, the LRE3 is driven from its native 5′ UTR.
Table 3 provides an analysis of L1 insertions in hESC-derived NPCs. From left to right: column 1: if the insertion was characterized from a clone or from FACS-sorted cells (derivations 1 and 2 are from separate NPC derivations, and separate transfections of L1); column 2: if the insertion characterization was full or partial; column 3: the truncation site of the retrotransposed tagged L1; column 4: the estimated length of the poly (A) tail; column 5: the sequence of the actual or inferred LINE-1 endonuclease bottom strand cleavage site; column 6: the chromosomal locus of the insertion; column 7: the insertion target site of the tagged L1. Note that eight insertions were characterized completely; however, only the 3′ end was characterized of the remaining insertions because the restriction enzyme utilized in the ligation step of the inverse PCR protocol was also present in the retrotransposed L1 sequence. In the table: 8—clone, or from either the first or second independent NPC derivation; b—full insertions: both 5 and Y documented; partial insertions: only 3′ genomic location isolated; c—the nucleotide position in RC-L1mEGFPI where the retrotransposition event is truncated; NA=not analyzed.
Tables 4A and 4B provide a sequencing analysis of QPCR genomic DNA products. PCR products from both ORF2#1 and ORF2#2 primer sets were cloned and sequenced from PCR reactions run with both hippocampus and liver genomic DNA. Percentage sequence identity to an RC-L1 consensus sequence was determined. Sequence analysis using the UCSC genome browser and Repeatmasker indicates that the majority of amplified sequences belong to the L1Hs subfamily of elements. Notably, due to their short length, some amplicons could not be definitively assigned to a single L1 subfamily.
Tables 5A and 5B provide a sequencing analysis of QPCR products from L1 RT-PCR. Quantitative RT-PCR products from ORF2 #1 primer sets were cloned from three sample types: fetal brain, hCNS-SCns, and HUES6-derived NPCs. Percentage sequence identity to an RC-L1 consensus was determined, and sequence analysis using UCSC genome browser and Repeatmasker indicated that most sequences belonged to the L1Hs subfamily of elements. Complete sequence of the QPCR product is indicated in Table 5A.
Tables 6A and 6B provide a sequence analysis of actively transcribed ORF1 fragments from RT-PCR. RT-PCR fragments (see
The following documents are incorporated by reference herein.
Although the present invention has been described in connection with the preferred embodiments, it is to be understood that modifications and variations may be utilized without departing from the principles and scope of the invention, as those skilled in the art will readily understand. Accordingly, such modifications may be practiced within the scope of the following claims.
This application claims the benefit of Provisional Patent Application No. 61/231,663, filed on Aug. 5, 2009, and Provisional Patent Application No. 61/273,599, filed on Aug. 5, 2009, which are both incorporated by reference herein.
This invention was made with Government support under Grant No. R56 MH082070 from the National Institute of Health. The Government has certain rights in this invention.
| Filing Document | Filing Date | Country | Kind | 371c Date |
|---|---|---|---|---|
| PCT/US2010/044358 | 8/4/2010 | WO | 00 | 8/8/2013 |
| Number | Date | Country | |
|---|---|---|---|
| 61273599 | Aug 2009 | US | |
| 61231663 | Aug 2009 | US |
