Identification of Ty1 integrase binding partners.
To identify yeast proteins associated with Ty1 IN that could regulate Ty1 replication, we adapted the tandem chromatin affinity purification procedure after in vivo cross-link (TChAP), which we developed previously (41) (Fig. S1A). IN protein was expressed with a C-terminal histidine-biotin-histidine (HBH) tag (42), allowing ectopic IN-HBH purification under denaturing conditions. Proteins that co-purified with IN-HBH were compared to those detected in the control yeast strain expressing untagged IN, and IN-HBH specific partners were sorted according to the settings of TChAP selection criteria (41). Ultimately, we identified a set of 265 IN binding proteins (Table S1), which expands the number of candidates previously described (37). Many of these proteins were nuclear. Their recovery was consistent with IN function in Ty1 integration into the nuclear genome and with the TChAP chromatin purification step. In addition, several peptides mapped to Gag, PR, IN and RT protein sequences from Ty1 and Ty2 retrotransposons, which are closely related, suggesting that ectopic IN-HBH interacts with endogenous Ty1 or Ty2 proteins (Table S2).
To get insights into the cellular activity associated with IN binding partners, the proteins identified by TChAP were classified using Gene ontology (GO) enrichment analysis and the GOrilla visualization tool (Fig. 1B, Tables S3A-D) and revealed a limited number of highly enriched biological processes (p-value < 6.53 E-8), including transposition, RNA Pol I and Pol III transcription (Fig. S1B).
GO cellular components analysis revealed a specific enrichment of RNA Pol I and Pol III polymerases, with the identification of many of their subunits. These data are consistent with previous studies showing a physical interaction between IN and both RNA polymerases (30, 37) (Fig. 1B). In particular, their common subunit AC40 was overrepresented and had the largest protein coverage in the TChAP experiment (Table S1), a finding consistent with the previously described IN-AC40 interaction that targets Ty1 integration upstream of Pol III-transcribed genes (17, 30). Other enriched GO biological processes were related to chromatin dynamics and histone modifications, suggesting that IN may interact with factors modulating chromatin state and content (Fig. S1B and Tables S3A-B). IN was also associated with other evolutionary conserved transcription complexes, including PAF1 (Polymerase-Associated Factor 1) and FACT (FAcilitates Chromatin Transcription) (Tables S3C-D and Fig. 1B). Several subunits of PAF1 were previously shown to restrict Ty1 retrotransposition, especially at coding genes (35), while FACT interacts with HIV-1 IN and stimulates HIV-1 infectivity and integration. Proteasome components were also found in our GO term analyses suggesting that IN may be regulated by proteasome-dependent degradation or that the proteasome degrades overexpressed IN-HBH. Finally, the four subunits of the serine/threonine/tyrosine protein kinase CK2 were identified (Table S1). The Ckb2 subunit was previously identified in a proteomic screen for Ty1 IN cofactors (37) and in a genetic screen for Ty1 repressors (35) but its role in Ty1 replication was not explored.
In conclusion, we identified by TChAP a large number of proteins that are good candidates to interact with IN in vivo and regulate Ty1 replication. We decided to further explore the role of CK2 in Ty1 retrotransposition because the kinase i) is present at tRNA genes, which are primary targets for Ty1 integration (44, 45), ii) phosphorylates several components of the Pol III machinery and iii) positively regulates Pol III transcription (46–48), suggesting that CK2 could interact with IN and control Ty1 integration at tRNA genes.
The CK2 holoenzyme interacts with Ty1 integrase in vivo.
CK2 mainly exists in S. cerevisiae as a tetramer composed of two regulatory subunits, Ckb1 and Ckb2, and two catalytic subunits, Cka1 and Cka2 either as homo or hetero-dimer (49) but free Cka2 has also been detected (50). No single CK2 subunit is essential. Only the combined absence of the catalytic subunits Cka1 and Cka2 is lethal to the cell (51). To validate the association between IN and CK2, we performed co-immunoprecipitation (co-IP) analyses. IN-HBH was expressed in a WT strain or in different strains, each expressing a TAP-tag version of Cka1, Cka2, Ckb1 or Ckb2 subunits or Tup1 as a negative control. IN-HBH co-purified with each of the TAP-tagged CK2 subunits immunoprecipitated from exponentially growing cells, but not with either Tup1-TAP or a control protein extract prepared from the WT strain lacking TAP-tag proteins (Fig. 1C). These data strongly suggest that Ty1 IN is associated with CK2 holoenzymes in vivo.
To determine which CK2 subunit interacts with IN, a two-hybrid assay using Ty1 IN as bait and the four CK2 subunits as preys was conducted and revealed an interaction between Ty1 IN and the Ckb2 regulatory subunit (Fig. 1D). This interaction could only be detected when the full-length Ty1 IN carried a C-tag at its C-terminus, suggesting that this tag promotes a structural conformation of Ty1 IN that allows its interaction with Ckb2 (Fig. 1E). We further delineated the Ckb2 binding sequence in Ty1 IN by showing that the interaction with Ckb2 occurred in the absence of the bNLS and the targeting domain (Fig. 1E, GAD-IN1 − 578) and that Ty1 IN amino acids 386 to 511 were sufficient for the interaction (Fig. 1E, GAD-IN386 − 511). In contrast, no physical association could be detected between IN and any of the catalytic subunits. We could not conclude for Ckb1 as GBD-Ckb1 self-activated the expression of the reporter gene. Together, these results point to a physical interaction between IN and CK2 holoenzymes that likely depends on the binding of Ckb2 to a sequence located in the C-terminal domain (CTD) of IN.
To investigate whether CK2 also interacts with IN expressed from a functional Ty1 element, we immunoprecipitated CK2 in yeast cells expressing TAP-tagged Cka2 or Ckb1 proteins and a Ty1 element under the control of a Tet-off promoter to increase cellular IN protein levels (Fig. S2). IN was associated with the two TAP-tagged CK2 subunits but not with TAP-YKR011C as a negative control and was not detected in the absence of TAP-proteins. However, the co-IP was difficult to detect presumably because of the low level of processed IN produced by Ty1 compared with ectopically expressed IN-HBH (Fig. 1C).
Altogether, these data indicate that in yeast cells, the protein kinase CK2 interacts with Ty1 IN expressed from ectopic or endogenous Ty1 copies.
CK2 phosphorylates Ty1 integrase in vitro.
The interaction between Ty1 IN and CK2 observed in vivo suggested that CK2 could directly phosphorylate the integrase. Ty1 IN (635 amino acids) harbours 78 serines and 52 threonines that could be phosphorylated. A minimal consensus sequence for CK2 phosphorylation (S/T–X–X–D/E) has been described but deviations from the consensus have already been observed (39). To determine whether Ty1 IN is a substrate of CK2, we performed in vitro kinase assays with recombinant Ty1 IN produced in E. coli in the presence of either yeast CK2 holoenzymes or commercial recombinant human rCK2. With both kinases, a radiolabelled signal was detected in the presence of γ32P-ATP for Maf1, a known substrate of CK2 (47), as well as for the full-length IN. This confirms that Ty1 IN is a substrate of CK2 in vitro. In agreement with our two-hybrid assay, a C-terminal fragment of IN (amino acids 578–635), which harbours potential phosphorylation sites, encompasses the bipartite NLS and the Ty1 targeting domain but does not interact with Ckb2, did not show radioactive incorporation, indicating that this region of Ty1 IN is not a substrate for CK2 (Fig. 2A, Fig. S3 and Fig. 1E).
To identify the amino acids phosphorylated by CK2, we performed a mass spectrometry analysis of in vitro phosphorylated Ty1 IN and identified 12 amino acids, including 9 serines, 2 threonines and one tyrosine (Fig. 2B and Table S4B). To confirm the role of these residues in the phosphorylation by CK2, we performed in vitro kinase assays using similar amounts of purified IN variants, where the candidate serine or threonine residues were changed into alanine and the tyrosine (Y472) residue into phenylalanine to prevent their phosphorylation (Fig. 2C, left and right panels). No radioactive signal could be detected when all 12 amino acids were mutated simultaneously (Fig. 2C, mutant M6). The same loss of signal was observed when we mutated together the 8 amino acids present in a CK2 consensus (Netphos analysis, Table S4A), indicating that these residues are the only ones efficiently phosphorylated by CK2 in vitro (Fig. 2C, mutant M5). In comparison, the residual radioactive signal observed in M3 and M4 mutants revealed that S499 and S43 were phosphorylated by CK2 in vitro. Finally, the reduction in the autoradiographic signal of the M1 and M2 mutants relative to IN indicated that both serine clusters (S469, S471 and S473 or S347, S354 and S360, respectively) were targeted by CK2 in vitro with no clear definition of the substrate serines.
Together, these results indicate that S43, S499 and one or more serines in the S347-S354-S360 and S469-S471-S473 clusters account for the majority, if not all, of CK2-mediated IN phosphorylation in vitro.
IN is phosphorylated during Ty1 replication.
To determine whether IN was also phosphorylated in a cellular context, we purified ectopic IN-HBH expressed in yeast cells and analyzed its phosphorylation profile by mass spectrometry. All in vivo phosphorylated amino acids were located within or near IN residues 386 to 511 that interact with Ckb2 in two-hybrid experiments (Fig. 1E and Table S4B). The data also indicated that all but one amino acid identified as CK2 targets in vitro were also phosphorylated in vivo. The only exception concerned residue S43, located in a region that was absent from the peptide sequences detected by mass spectrometry, hampering any conclusion about its modification in vivo. Ten additional phosphopeptides were detected only in the cellular context (Table S4B). All the amino acids phosphorylated in ectopically expressed IN, both in vivo and in vitro, were also present in the PhosphoGRID database, which compiles experimentally verified in vivo protein phosphorylation sites in S. cerevisiae (Table S4B and http://www.phosphogrid.org/ (52)), indicating that endogenous and ectopic INs harbour the same modifications. Finally, we compared the phosphorylation sites identified by the different approaches (in vitro and in vivo with IN expressed endogenously or ectopically) with the consensus sequence for phosphorylation by CK2 and obtained a final list of 12 amino acids potentially phosphorylated by CK2 in the endogenous IN.
To assess the contribution of CK2 to IN phosphorylation in vivo, we expressed IN from a pTet-off inducible vector in ste12∆ and cka2∆ ckb2∆ ste12∆ mutant cells. Of note, STE12 deletion prevents Ty1 endogenous expression (53) and the cka2∆ ckb2∆ double mutation affects CK2 holoenzyme formation (49). On a conventional SDS-PAGE gel, ectopically expressed IN was detected as a single band with similar mobility in ste12∆ and cka2Δ ckb2Δ ste12Δ protein extracts (Fig. 2D, upper panel, compare lanes 3 and 4). By contrast, on Phos-tag™ SDS-PAGE, which allows a better separation of phosphorylated and non-phosphorylated proteins, IN migrated as diffuse protein bands in ste12∆ cells, and mainly as a single discrete band in cka2∆ ckb2∆ ste12∆ cell extracts (Fig. 2D, lower panel, compare lanes 3 and 4). This discrete band is probably the dephosphorylated IN, while the diffuse bands could correspond to different phosphorylated forms of IN. Therefore, the difference in the migration profiles between the two strains indicates that ectopically expressed IN is phosphorylated in vivo, and that this phosphorylation is dependent on CK2. It also suggests additional phosphorylation of other residues in vivo, which is in agreement with the mass spectrometry data (Table S4B).
We also compared the electrophoretic mobility of IN and INM6 mutant (Fig. 2D, compare lanes 2 and 3). In INM6, the 12 putative phosphorylated residues present in endogenous IN are changed in alanine or phenylalanine (Y472) to block their phosphorylation (Table S4B and Fig. 2C). Surprisingly, there was no significant difference in the migration profile of the two proteins on Phos-tag™ SDS-PAGE gels, revealing that ectopically expressed IN is not highly phosphorylated in the cell.
Finally, since CK2 also interacted with IN expressed from a Ty1 element (Fig. S2), we asked whether CK2 could phosphorylate IN produced during Ty1 replication cycle. In yeast cells, IN is produced by successive cleavages of the precursor polyprotein Gag-Pol in Ty1 VLPs (23). The Gag-Pol polyprotein has three distinct cleavage sites at the Gag/PR, PR/IN and IN/RT junctions. The Gag/PR site is cleaved first, but no further processing order was identified (23) and thus, in addition to the mature IN, various Gag-Pol intermediates including IN-RT and PR-IN could be detected by an antibody raised against IN (Fig. 2E, upper panel and (23). To examine IN phosphorylation during Ty1 replication, Ty1 or Ty1M6 harbouring the same mutations in the IN sequence as described in Fig. 2C and 2D, were expressed from a pTet-off promoter in WT and cka2∆ ckb2∆ cells. The endogenous Gag-Pol and Pol precursors, but not the processed IN, could be observed when the expression of Ty1 was repressed in the presence of doxycycline (Fig. 2E, lane 1 in both panels). In the absence of doxycycline, processed IN and additional IN precursors or intermediates were detected (Fig. 2E, lanes 2–4). On conventional SDS-PAGE, the mature INs expressed from Ty1 and Ty1M6 migrated as a single band, as observed when the two proteins were ectopically expressed (Fig. 2E, upper panel, lanes 2 and 3), and the migration profile of IN was similar in WT and cka2∆ ckb2∆ cell extracts (Fig. 2E, upper panel, lanes 3 and 4). The sizes of the upper bands were consistent with Gag-Pol (200 kDa), Pol (160 kDa) and IN-RT (134 kDa), while the two bands above IN may correspond to Pol intermediates including PR-IN (92 kDa) (Fig. 2E, upper panel). The pattern observed on Phos-tag™ SDS-PAGE was different with the presence of multiple bands and smears in both WT and cka2∆ ckb2∆ extracts expressing Ty1 or Ty1M6, which migrated slower than ectopic IN from cka2∆ ckb2∆ ste12∆ cell extracts (Fig. 2E, lower panel, compare lanes 2–4 with lane 5). This complex pattern reflects the presence of various phosphorylated mature and unprocessed INs. First, the slight difference in profiles between Ty1 or Ty1M6 in WT cell extracts suggests that some of the phospho-ablative mutations in Ty1M6 partially prevent the phosphorylation of IN expressed from Ty1 (Fig. 2E, lower panel, compare lanes 2 and 3). Second, the faster mobility of smear bands in cka2∆ ckb2∆ cell extracts compared to WT are consistent with a role for CK2 in IN phosphorylation during Ty1 replication (Fig. 2E, lower panel, compare lanes 3 and 4). Third, the reduced mobility of IN from Ty1M6 mutant in WT cell extracts compared to that of IN from Ty1 in cka2∆ ckb2∆ cell extracts, suggests that other residues of IN are phosphorylated by kinases. The activity of these kinases may or may not depend on CK2 (54).
In conclusion, IN expressed from a Ty1 element is a substrate for CK2 in vivo. Noteworthy, the detection of several distinct bands by Western blot, mainly when IN is expressed from a Ty1 element, suggests that mature IN produced in vivo during Ty1 replication is a better substrate for kinases than that expressed ectopically (Fig. 2E and 2D, compare lanes 2 and 3).
The CK2 protein kinase inhibits Ty1 retrotransposition.
The interaction between IN and CK2 suggested that the kinase could regulate Ty1 retrotransposition. Accordingly, Ckb2 was previously found to repress Ty1 retrotransposition in a genetic screen (35). To investigate further the role of CK2 in the control of Ty1 retrotransposition, we performed a quantitative retrotransposition assay, based on the presence of a Ty1-his3AI reporter, which is naturally integrated in the genome and confers histidine prototrophy to cells that have undergone a new integration event (55). We measured retrotransposition frequencies in strains harbouring single or double deletions of CK2 subunits, except the cka1∆ cka2∆ combination that is lethal (51). The absence of either of the catalytic subunits Cka1 or Cka2 caused little or no increase in retrotransposition, respectively (Fig. 3A). In contrast, an 8 to 15-fold increase was observed in the absence of the regulatory subunits Ckb1 or Ckb2. No additive effect could be detected when both subunits were deleted. Both Ckb1 and Ckb2 are required for the formation of CK2 holoenzymes, while the catalytic subunits Cka1 and Cka2 are partially redundant within the complex (51). These data strongly argue that it is primarily the CK2 holoenzymes that control Ty1 retrotransposition. The deletion of CKA2 in the ckb1∆ and ckb2∆ mutants further increased Ty1 retrotransposition up to 80 to nearly 200-fold, respectively (Fig. 3A). This strong derepression suggests that Cka2 might also participate to the restriction of Ty1 mobility in the absence of holoenzymes in good agreement with the existence of free form of Cka2 in vivo (49).
To determine whether IN phosphorylation by CK2 could repress Ty1 retrotransposition, we introduced the phospho-ablative mutations that prevent IN phosphorylation by CK2 in vitro (Fig. 2C, M1, M2 and M6 mutants) into a Ty1-his3AI reporter element. To avoid possible trans-complementation by endogenous Ty1 elements (56), we expressed the different mutants from the heterologous PSP2 promoter in a ste12∆ mutant. Unlike Ty1 promoter, the PSP2 promoter was not stimulated by the Ste12 transcription factor. Instead, we observed a slight increase in PSP2 promoter activity in the absence of STE12 (Fig. S4A and S4B). Moreover, this promoter has a relatively weak activity (57) such that in WT cells the retrotransposition frequency of Ty1-his3AI expressed from the PSP2 promoter was of the same order of magnitude as that obtained with a Ty1-his3AI chromosomal element expressed from the Ty1 promoter (Compare WT in Fig. 3A and Fig. 3B), thus approaching physiological retrotransposition conditions. With this construct, we observed a ~ 14-fold increase in Ty1 retrotransposition in the ste12∆ cka2∆ ckb2∆ mutant compared to the ste12∆ mutant, indicating that CK2 represses pPSP2-Ty1-his3AI retrotransposition (Fig. 3B). This increase in retrotransposition was lower than with the chromosomal Ty1-his3AI reporter expressed from Ty1 promoter (~ 200-fold, Fig. 3A), suggesting that some of the repression by CK2 could occur at the transcriptional level. However, there was no difference in His+ frequency between the WT and the three M1, M3 and M6 phospho-ablative mutants, in both ste12∆ and ste12∆ cka2∆ ckb2∆ cells. These results show that the repression of Ty1 retrotransposition by CK2 is independent of the phosphorylation of the IN residues that are targeted by the kinase in vitro.
Ty1 integrates primarily in a 1-kb window upstream of Pol III-transcribed genes but subtelomeres are secondary target sites when the interaction between Ty1 IN and RNA Pol III is abolished (17, 30). To investigate whether CK2 could play a role in Ty1 integration site selection, we performed a dedicated PCR assay to compare the integration profile of endogenous Ty1 elements in WT and different CK2 mutants upstream of SUF16, a hotspot of Ty1 integration (58). In this assay, proficient integration at SUF16 is characterized by a ~ 70–base pair periodic banding pattern that corresponds to specific insertion events in nucleosomal DNA (17, 44, 45). This profile was difficult to detect in WT and cka2∆ cells, probably because endogenous Ty1 retrotransposition was too low. On the contrary, it was observed in ckb2∆, cka2∆ ckb1∆ and cka2∆ ckb2∆ mutants (Fig. 3C), which have the highest Ty1 integration frequency (Fig. 3A). These data indicate that SUF16 remains a hotspot of Ty1 integration in CK2 mutants. Furthermore, when we analyzed Ty1 integration at four representative subtelomeric homologous genes (HXT13, HXT15, HXT16 and HXT17), we also detected more de novo insertions at the HXT loci in ckb2∆, cka2∆ ckb1∆ and cka2∆ ckb2∆ mutants than in WT or cka2∆ cells (Fig. 3C). Therefore, the increase in Ty1 integration frequency observed in CK2 mutants is associated with an increase in de novo insertion events at both primary and secondary Ty1 integration sites.
The above results indicate that CK2 does not participate in the recognition of Ty1 integration preferred sites but they do not exclude that CK2 could control Ty1 integration at other locations in the genome. Such a scenario has been reported for the retrotransposon Ty5 whose phosphorylation of the integrase targeting domain is essential for its interaction with the Sir4 protein and the integration of Ty5 into heterochromatin. In the absence of phosphorylation, Ty5 integrates throughout the genome and can cause mutations (59). To investigate the impact of CK2 on Ty1 genome-wide integration profile, libraries of His+ selected de novo Ty1 insertion events were generated in WT and cka2∆ ckb2∆ cells expressing Ty1-his3AI elements from the GAL1 promoter, and Ty1-HIS3 de novo insertion events were characterized by high-throughput sequencing, as done previously (17, 30). Of note, the pGAL1-Ty1-his3AI element retrotransposes at a much higher frequency than the Ty1-his3AI chromosomal element and with similar frequencies in WT and cka2∆ ckb2∆ cells (Fig. S4C). The percentage of Ty1-HIS3 insertions, calculated for representative features of the yeast genome, indicated that de novo Ty1-HIS3 insertions occurred mainly upstream of Pol III-transcribed genes and in retrotransposon sequences associated with these sites in both WT and cka2∆ ckb2∆ cells (Fig. 3D). In addition, both strains displayed the same characteristic periodic pattern with two insertion sites per nucleosome (Fig. S4D), in the three nucleosomes located upstream of Pol III-transcribed genes (44, 45). Together, these data confirm that CK2 mutants do not alter Ty1 integration targeting at Pol III-transcribed genes. We did not identify any enrichment of de novo Ty1-HIS3 insertions in different chromosome features (ORFs, subtelomeres, telomeres or ARS) in the cka2∆ ckb2∆ cells compared to WT cells (Fig. 3D). Therefore, the increase in Ty1 insertions observed by PCR at the HXT subtelomeric loci is most likely a mere consequence of increased retrotransposition frequency and not of a change in Ty1 integration preferences.
Collectively, these results indicate that CK2 represses Ty1 retrotransposition without altering Ty1 integration profile; this repression is independent of IN amino acids phosphorylated by CK2. Therefore, the kinase controls Ty1 activity at other stage(s) of its replication cycle.
The repression of Ty1 retrotransposition by CK2 occurs mainly at the level of transcription.
Several studies describe a role for CK2 in the regulation of transcription by altering chromatin dynamics (60, 61). Interestingly, our data indicate a smaller increase in retrotransposition in cka2∆ ckb2∆ mutants when Ty1-his3AI was expressed from the heterologous PSP2 promoter instead of Ty1's own promoter. This discrepancy supports the hypothesis that the kinase controls Ty1 transcription. To address this point, we measured Ty1 and Ty1-his3AI RNA levels expressed from Ty1 promoter by RT-qPCR in the different CK2 deletion mutants that impact chromosomal Ty1-his3AI retrotransposition frequency to varying degrees. Our assay revealed an approximately 3-fold increase in the level of both RNA species in the cka2∆ ckb1∆ and cka2∆ ckb2∆ mutants that showed the greatest increase in Ty1 retrotransposition (Fig. 4A-B, compare with Fig. 3A). Conversely, Ty1 or Ty1-his3AI RNA levels did not increase in the cka2∆ mutant and slightly in the ckb2∆ mutant, whose retrotransposition frequencies remain unchanged or moderately enhanced, respectively. These results demonstrate that CK2 represses Ty1 transcription.
Since Ty1 RNA plays a central role in the retrotransposition by serving as a template for both Ty1 protein translation and reverse transcription into cDNA, increased Ty1 mRNA levels could explain the strong derepression of Ty1 retrotransposition in CK2 mutants. Thus, we decided to evaluate the impact of Ty1 mRNA derepression on the production of Ty1 proteins and cDNA. Anti-VLP polyclonal antibodies were used to reveal Gag proteins by Western blot, while an anti-IN monoclonal antibody was used for Gag-Pol and IN detection (Fig. 4C). The levels of Gag and Gag-Pol remained unchanged in cka2∆ or ckb2∆ single mutants compared with WT, but slightly increased in cka2Δ ckb1Δ and cka2Δ ckb2Δ mutants. On the other hand, IN, which was barely apparent in WT cells and in the single mutants (Fig. 4C), was readily detected in cka2Δ ckb1Δ and cka2Δ ckb2Δ mutants. This analysis indicates that the activation of Ty1 transcription in CK2 mutants leads to higher levels of Ty1 proteins.
We also assessed the levels of unintegrated Ty1 cDNA in a subset of CK2 deletion mutants. Using a qPCR methodology we developed recently (62), we found that the amounts of cDNA in the various mutants correlated with the transposition efficiency in these strains (Fig. 4D). Specifically, the cka2Δ ckb1Δ and cka2Δ ckb2Δ mutants that displayed the highest derepression of Ty1-his3AI retrotransposition (Fig. 3A) also had the highest levels of unintegrated cDNA.
Importantly, Ty1-his3AI mRNA levels and His+ retrotransposition frequencies were strongly correlated in the different CK2 mutants (Pearson correlation coefficient r = 0.824) (Fig. 4E). A similar correlation was observed between Ty1 cDNA levels and Ty1 mRNA levels (Pearson correlation coefficient r = 0.944) with Ty1 mRNA accumulation leading to an exponential increase in unintegrated cDNA (Fig. 4F). Collectively, these results establish that the increase in Ty1 reverse transcription and retrotransposition observed in CK2 mutants is mainly due to Ty1 RNA accumulation in these mutants.